Abstract
Macropinocytosis is a conserved endocytic process used by Dictyostelium amoebae for feeding on liquid medium. To further Dictyostelium as a model for macropinocytosis, we developed a high-throughput flow cytometry assay for macropinocytosis, and used it to identify inhibitors and investigate the physiological regulation of macropinocytosis. Dictyostelium has two feeding states: phagocytic and macropinocytic. When cells are switched from phagocytic growth on bacteria to liquid media, the rate of macropinocytosis slowly increases, due to increased size and frequency of macropinosomes. Upregulation is triggered by a minimal medium of 3 amino acids plus glucose and likely depends on macropinocytosis itself. Bacteria suppress macropinocytosis while their product, folate, partially suppresses upregulation of macropinocytosis. Starvation, which initiates development, does not of itself suppress macropinocytosis: this can continue in isolated cells, but is shut down by a conditioned-medium factor or activation of PKA signalling. Thus macropinocytosis is a facultative ability of Dictyostelium cells, regulated by environmental conditions that are identified here.
Summary A high-throughput flow cytometry assay shows that macropinocytosis in D. discoideum is upregulated in the presence of nutrients and absence of bacteria. Development and bacteria induce cells to downregulate macropinocytosis.
Introduction
Macropinocytosis, first described in the 1930s (Lewis, 1931), is a process of large-scale, non-specific fluid uptake carried out by a wide variety of cells. Actin-driven protrusions from the plasma membrane form cup-shaped circular ruffles that can be several microns in diameter. When a ruffle closes it engulfs and delivers extracellular material to the cell interior in macropinosomes. Macropinosomes proceed through the endocytic system where their contents can be broken down by digestive enzymes and useful metabolites extracted (Buckley and King, 2017).
In the immune system, dendritic cells and macrophages use macropinocytosis to sample environmental antigens for presentation to B and T cells (Sallusto et al., 1995, Norbury et al., 1995). Certain bacteria and viruses can utilise macropinocytosis to invade host cells (Marechal et al., 2001, Nanbo et al., 2010, Hardt et al., 1998), while other bacteria stimulate macropinocytosis to promote toxin internalisation (Lukyanenko et al., 2011). Prions and neurodegenerative protein deposits also invade new host cells through macropinocytosis (Magzoub et al., 2006, Fevrier et al., 2004, Munch et al., 2011, Falcon et al., 2015). Tumour cells can maintain a high rate of macropinocytosis (Lewis, 1937), with Ras-activated cancer cells obtaining a substantial part of their nutrition in this way (Commisso et al., 2013).
Considering its widespread importance, the basic biology of macropinocytosis is poorly understood. It has been studied most intensively in tissue culture cells, particularly macrophages, although genetic screens have been performed in C. elegans (Fares and Greenwald, 2001) and Dictyostelium discoideum (Bacon et al.,1994). Dictyostelium in particular has great potential as a model because of the high constitutive rate of macropinocytosis maintained by cells in the right circumstances and because the evolutionary distance from mammalian cells should allow conserved, core features to be discerned.
The high rate of macropinocytosis by standard axenic strains of Dictyostelium used in the laboratory is due to deletion of the RasGAP, NF1 (Bloomfield et al., 2015). This mutation allows cells to grow in nutrient media without a bacterial food source (hence axenic). Wild isolates also perform macropinocytosis, although the rate of fluid uptake is too low to allow growth in the standard media for laboratory-adapted axenic strains. These strains can, however, grow in medium supplemented with additional nutrients (Maeda, 1983, Bloomfield et al., 2015).
Axenic strains form frequent, large macropinosomes. The macropinocytic cups are organized around intense patches of active Ras, Rac and plasmanylinositol (3,4,5)-trisphosphate (PIP3) (Hoeller et al., 2013, Parent et al., 1998, Veltman et al., 2016) (In Dictyostelium PIP3 is a plasmanylinositide, rather than a phosphatidylinositide (Clark et al., 2014)), with SCAR/WAVE and WASP localised to their periphery (Veltman et al., 2016). SCAR/WAVE and WASP activate the Arp2/3 complex to polymerise actin and form a macropinocytic cup, which is also known as a crown, or circular ruffle. The cup may be supported by actin polymerization around the base driven by a Ras-activated formin (Junemann et al., 2016).
The rate of fluid uptake through macropinocytosis by axenic cells is regulated by environmental factors, principally whether the cells’ nutrient source is growth media or bacteria (Kayman and Clarke, 1983, Aguado-Velasco and Bretscher, 1999) and their developmental state (Maeda, 1983, Katoh et al., 2007). Macropinocytosis is additionally affected by the stage of the cell cycle and the concentration of bacterial peptone in the medium (Maeda, 1988), as well as the incubation temperature and the pH (Maeda and Kawamoto, 1986). For certain mutants, fluid uptake is dependent upon whether cells are attached to a surface or in shaking suspension (Novak et al., 1995).
Fluid uptake by standard axenic strains of Dictyostelium, such as Ax2, is almost entirely due to macropinocytosis and can be accurately measured by following the uptake of fluorescent dextran as a fluid phase marker (Kayman and Clarke,1983, Thilo and Vogel, 1980, Hacker et al., 1997). We have developed a high-throughput assay to measure macropinocytosis in Dictyostelium, identified useful inhibitors and sought to better understand how macropinocytosis is physiologically regulated during the switch between macropinocytic and phagocytic feeding and the growth-to-development transition.
Results
Measurement of uptake by high-throughput flow cytometry
Macropinocytosis accounts for more than 90% of fluid uptake by axenic strains of Dictyostelium, and can therefore be followed by measuring fluid uptake (Hacker et al., 1997). However, existing methods based on processing individual cell pellets after uptake of fluorescent dextran are of relatively low throughput. We therefore developed a high-throughput assay using flow cytometry to measure TRITC-dextran uptake. The assay is performed in 96-well plates and, after loading with TRITC-dextran, the cells are washed in-situ by ‘dunk-banging’ and detached using sodium azide (Glynn and Clarke, 1984) (Figure 1A), which also prevents exocytosis of internalized dextran (Figure 1B). Plates are analysed by flow cytometry using a High-Throughput Sampling attachment to load the flow cytometer, and subsequent analysis is performed using Flowjo, which easily distinguishes Dictyostelium cells from beads and bacteria, but not yeast (Figure 1C). A principle advantage of flow cytometry is that the fluorescence of internalized TRITC-dextran can be determined for single cells (Figure 1D). The accumulation of TRITC-dextran proceeds in a uniform fashion across the population over time, with an extended lagging edge of cells with lower uptake. Median fluid internalisation over time by Ax2 cells is quantified in figure 1E.
Controls show the efficiency of the wash step (Figure S1A); that Ax2 cells take up similar volumes of liquid whether in suspension or attached to a surface (as in the assay-Figure S1B), although this is not true for all strains (Novak et al.,1995); and the range of cell numbers that can be accommodated per well (Figure S1C). The assay is calibrated in terms of volume taken up per cell by reference to measurements of uptake by the same cell population using a fluorimeter (Figure S1D) and standardised over time using Flow-Set fluorosphere calibration beads (Beckman Coulter). Phagocytosis of beads (Figure S1E) and bacteria (Figure S1F) as well as membrane uptake (Figure S1G) can be readily measured.
Effect of inhibitors on macropinocytosis
Inhibitors are powerful tools for acutely interfering with biological processes, but relatively few are currently known for macropinocytosis. We therefore tested a number of inhibitors affecting both the cytoskeleton and cellular signalling. These were added to Ax2 cells growing in HL5 medium at the start of the uptake assay, with the TRITC-dextran. The internalised fluorescence was measured 1 hour later (Table 1). A number of inhibitors were without effect, although whether this is due to lack of inhibitor uptake, target interaction or the target not functioning in macropinocytosis is unknown.
As macropinocytosis is an actin-dependent process, we tested a number of inhibitors of actin dynamics. Latrunculin B efficiently inhibited macropinocytosis at standard concentrations (Figure S2A), as expected from its profound effects on the actin cytoskeleton (Pramanik et al., 2009). Cytochalasin A, as previously reported (Hacker et al., 1997), inhibited macropinocytosis (Figure S2B). Inhibitors of the Arp2/3 complex (CK666, Figure S2C), WASP (Wiskostatin, Figure S2D) and formins (SMIFH2, Figure S2E) were all potent inhibitors of macropinocytosis, consistent with the localization of the target proteins to macropinosomes, the macropinocytosis defects in WASP (Veltman et al., 2016) and ForG (Junemann et al., 2016) mutants and the axenic growth defect of an ArpB mutant (Langridge and Kay, 2007).
The microtubule inhibitors nocodazole (Figure S2F) and thiabendazole (Figure S2G) both partially inhibited fluid uptake, indicating a role for microtubules. The myosin II inhibitor blebbistatin had no effect on macropinocytosis, in contrast to previously published data (Shu et al., 2005). This may be because, in our hands, blebbistatin readily precipitated at concentrations above those used.
Macropinosomes are organised around active Rac/Ras/PIP3 patches (Hoeller et al., 2013, Veltman et al., 2016) and, accordingly, the PI3-kinase (PI3K) inhibitor LY294002 (Figure S2H) inhibits fluid uptake. TGX221, which targets the mammalian p110β PI3K isoform, inhibited macropinocytosis (Figure S2I) whereas inhibitors targeting α and γ isoforms did not. We found the Racinhibitor EHT1864 (Shutes et al., 2007) is a potent inhibitor of fluid uptake (Figure S2J).
In mammalian cells macropinocytosis of free amino acids, notably leucine, induces mTORC1 activation (Yoshida et al., 2015), allowing cell proliferation. Rapamycin, a TORC1 specific inhibitor, did not affect fluid uptake when applied acutely, as found previously, although it does prevent proliferation (Rosel et al., 2012). It has been suggested that there are functions of mTORC1 that are not inhibited by rapamycin, but are by more potent, less specific, mTor inhibitors (Thoreen and Sabatini, 2009). We therefore tried alternative Tor inhibitors and observed an inhibition of macropinocytosis in cells treated with torin 1 (Figure S2K), but not palomid 529 or PP242. Whether this is due to greater inhibition of TORC1, inhibition of TORC2, or both is not clear: TORC2 has previously been described as having no function in Dictyostelium macropinocytosis (Rosel et al., 2012), however we see a reduction in macropinocytosis when TORC2 components are knocked out in the Ax2 strain used here (not shown).
The nearest to diagnostic inhibitors for macropinocytosis in mammalian cells are amiloride and EIPA, which block the plasma membrane Na+/H+ exchanger, thus affecting sub-membranous pH (Koivusalo et al., 2010). Although Dictyostelium possesses two Na+/H+ exchangers (Patel and Barber, 2005, Fey et al., 2013), it is not known whether they are sensitive to these drugs and we find that the drugs do not affect macropinocytosis. The removal of extracellular calcium by EGTA inhibits constitutive macropinocytosis in immune cells (Canton et al., 2016), but had no effect on macropinocytosis by Dictyostelium incubated in a calcium-free medium (50 mM lysine, 55 mM glucose in 50 mM MES, pH 6.5) indicating that extracellular calcium is not required. Indeed, high extracellular calcium concentrations are reported to inhibit Dictyostelium macropinocytosis (Maeda and Kawamoto, 1986).
These results support previous genetic studies showing that macropinocytosis depends on PI3K, Rac and actin dynamics controlled through SCAR/WAVE, WASP and formins. On the other hand, regulation through extracellular calcium is not part of a core, conserved mechanism of macropinocytosis across species, while roles for the Na+/H+ exchanger and Tor have not been confirmed.
Slow switching between feeding strategies
Ax2 cells grown on bacteria have a low rate of macropinocytosis, which increases greatly when they are switched to HL5 growth medium (a complex medium containing peptone, yeast extract and glucose). A similar, although much reduced, increase is seen in wild-type NC4 cells (which have an intact NF1 gene) in media enriched with protein (Maeda, 1983).
We confirmed the upregulation of macropinocytosis in Ax2 cells switched from growth on bacteria to HL5 (Figure 2A). It is slow, taking about 10 hours (Figure 2B), similar to Ax3 (Kayman and Clarke, 1983), and involves both an increased rate of macropinosome formation (Figure 2C) and size (Figure 2D). A 50% increase in diameter, as seen here, would lead to a 3-4 fold increase in macropinosome volume.
Wild-type DdB cells (the parent of the standard Ax2, Ax3 and Ax4 strains) with an intact NF1 gene do not noticeably upregulate macropinocytosis in HL5 (Figure 2A). However, if DdB cells are switched to HL5 supplemented with 10% FCS (Gibco, providing ~4 mg ml-1 additional protein), in which they can proliferate (Bloomfield et al., 2015), they substantially upregulate macropinocytosis, although not as much as Ax2. The increased fluid uptake by DdB cells in this case appears to be due only to an increased rate of macropinosome formation (Figure 2E), with no detectable increase in size (Figure 2F). Thus the macropinocytic rate of wild-type cells is also controlled by the availability of environmental nutrients, as in Ax2 cells.
Ax2 and other axenic cells can consume yeast and other large particles, but this ability depends on the loss of the NF1 gene and is not shared by wild-type cells (Bloomfield et al., 2015). Thus the abilities to phagocytose large particles and to take in large volumes of fluid by macropinocytosis are linked as both depend on the loss of NF1. We asked whether this linkage is also seen at a physiological level. It is apparent from Figure S3A that it is: Ax2 cells adapted to HL5 medium and with a high macropinocytic rate can take up yeast or large beads (2 micron diameter) much better than the same cells grown on bacteria, which have a low macropinocytic rate. Uptake of either particle by DdB cells, which have an intact NF1 gene, was unaltered by the nutritional environment they came from (Figure S3B), consistent with the unaltered macropinosome size observed in figure 2F.
The macropinocytic and phagocytic states are not mutually exclusive, as we found that Ax2 cells fully adapted to HL5 maintain a relatively high rate of phagocytosis of bacteria (Figure S3C). We therefore asked what happens when Ax2 cells are presented with both bacteria and liquid medium for food. In this case, irrespective of whether the cells had been grown on bacteria or HL5, they adopted a low rate of macropinocytosis (Figure S3D).
These results show that Dictyostelium has two basic feeding modes: the preferred mode is phagocytosis, which is seen with cells growing on bacteria. Alternatively, cells in liquid media without bacteria adopt a second mode, in which macropinocytosis is upregulated, although the potential for phagocytosis of bacteria is retained.
A minimal set of soluble nutrients can stimulate the upregulation of macropinocytosis
Ax2 cells do not sustainably increase their rate of macropinocytosis when they are switched from bacteria to buffer (Figure 2B), but require HL5 medium, or some components of it, to do so. To identify such stimulatory components, we first showed that HL5 could be replaced by the SIH defined medium (Figure 3A) and then dissected this defined medium to find the active components. Leaving out blocks of components showed that vitamins and micro-minerals are not necessary for macropinocytic upregulation and that the effect is accounted for by amino acids and glucose alone (Figure 3B). Testing amino acids individually showed that only arginine, lysine and glutamate induce macropinocytosis upregulation at the tested concentrations (Table S1). Consistent with this, removal of these amino acids from SIH severely impairs the ability of cells to upregulate macropinocytosis, which is restored when the amino acids are returned to the medium (Figure 3C). Testing different sugars showed that only glucose and other metabolizable sugars that can support cell growth permit macropinocytosis upregulation (Table S2) (Watts and Ashworth, 1970,Ashworth and Watts, 1970).
Based on these results, a simplified medium for macropinocytosis upregulation, SUM (Simple Upregulation Media) was devised, consisting of KK2MC plus 55 mM glucose, 4 mM arginine, 3.7 mM glutamate and 8.5 mM lysine (the same concentrations as SIH) at pH 6.5. SUM induces nearly the same level of macropinocytosis as complete SIH, with faster upregulation kinetics (Figure 3D). Although cells remain healthy in SUM for several days, it does not support longterm growth. SUM has very low background fluorescence, and we have found it very useful for microscopy, particularly for cells with weakly expressed markers, such as knock-ins. Cells can be grown rapidly on bacteria before transfer to SUM a few hours prior to microscopy, during which time macropinocytosis is greatly upregulated.
These results show that macropinocytosis upregulation can be induced by only a handful of the components present in defined medium, while the requirement for the sugar to be metabolizable hints that sugars may be sensed through their effects on metabolism, rather than by dedicated receptors.
Macropinocytosis is required for efficient upregulation of macropinocytosis
We envisioned that nutrients that cause macropinpocytosis upregulation might either be sensed by dedicated receptors, such as for glutamate, or indirectly through their effect on metabolism, or a combination of both. Since nutrients obtained by macropinocytosis can only be utilized after internalization and digestion, this second route implies that macropinocytic upregulation would depend on macropinocytosis itself. To test this idea we turned to the inhibitors identified earlier in this work to inhibit macropinocytosis during upregulation.As this experiment requires prolonged inhibitor treatment, we first tested how well cells recover from the inhibitors. Ax2 cells growing in HL5 recover quite well from prolonged treatment with LY294002, TGX221 (both PI3K), CK666 (Arp2/3 complex), EHT1864 (Rac) and torin 1 (Tor) (Figure S4A-E). Prolonged incubation with other inhibitors was too deleterious to make them useful.
We next used the inhibitors to determine to what extent the upregulation of macropinocytosis is affected by inhibiting macropinocytosis (Figure 4 ‘raw’ curves), also making a correction for the relatively small deleterious effects of long-term exposure of cells to the inhibitors (Figure 4, ‘corrected’ curves). Although these inhibitors affect macropinocytosis through different targets, they all inhibit upregulation (measured after 10 hours incubation in HL5) in a dose-dependent manner (Figure 4A-E). The effect remains even after correcting for the long-term effects of the inhibitors. Upregulation is not completely abolished by the inhibitors, reflecting their incomplete inhibition of macropinocytosis.Thus these results suggest that the upregulation of macropinocytosis by nutrient media depends on delivery of nutrients into the cell through macropinocytosis.
We considered the possibility that the ingested nutrients delivered by macropinocytosis might be detected through the TORC1 complex, similar to other organisms. Although rapamycin does not inhibit macropinocytosis acutely, it does somewhat inhibit upregulation (Figure 4F), with extremely mild effects on control cells (Figure S4F). Torin1, has a stronger effect on upregulation (Figure 4E), but as it is less specific, some of this might be due to inhibition of the TORC2 complex. In summary, these results suggest that nutrients causing cells to increase their rate of macropinocytosis are detected in the macropinocytic pathway, possibly by TORC1.
Sensing of bacteria
Bacteria have two distinct effects on the regulation of macropinocytosis. They inhibit upregulation of macropinocytosis by cells transferred to HL5 (Figure 5A) and promote downregulation of macropinocytosis by cells transferred from HL5 to buffer, where it otherwise remains high (Figure 5B).
Bacteria can be sensed through their release of folate, which is a chemoattractant for Dictyostelium and acts through the G-protein coupled receptor fAR1 (Pan et al., 2016). We found that folate inhibits the upregulation of macropinocytosis when cells are transferred from bacteria to HL5 (Figure 5C), but has no effect when cells are transferred from HL5 to buffer (not shown). fAR1-cells were essentially blind to this inhibitory effect of folate (Figure 5D), as were mutants of the Gβ and Gα4 (Hadwiger and Firtel, 1992) subunits of the cognate hetero-trimeric G-protein (Figure 5E) (Hadwiger and Srinivasan, 1999) and the downstream MAP kinase, ErkB (Figure 5F) (Nichols et al., in preparation). Thus bacteria can exert some, but clearly not all, of their effects on feeding behaviour through canonical folate signalling.
Developmental regulation of macropinocytosis
Development in Dictyostelium is triggered by starvation and over the first 8-10 hours the cells aggregate by chemotaxis to cyclic-AMP. Macropinocytosis is downregulated during this period (Maeda, 1983, Katoh et al., 2007), and it was therefore surprising that macropinocytosis continues at a high rate for at least 24 hours in cells starved under buffer (Figure 5B). However, compared to standard developmental conditions, these cells were starved at low density, likely causing attenuation of developmental signalling. This suggests that the downregulation of macropinocytosis during development requires a developmental signal in addition to starvation.
Figure 6A confirms that macropinocytosis is strongly downregulated by starving Ax2 cells (previously grown in HL5) at high density in shaking suspension and pulsed with cyclic-AMP to mimic developmental signalling. By 5 hours of development, fluid uptake is negligible. Similar results were obtained when developing cells on non-nutrient agar (not shown). Similarly if the cell density in 96-well plates is increased from 5000 to 50,000 cells per well, the cells can be seen to aggregate (not shown) and downregulate macropinocytosis (Figure 6B).
We tested the effects of known developmental signals on macropinocytosis using cells starving at low cell density. As shown in table S3, the developmental signals cyclic-AMP, ATP (Ludlow et al., 2008, Traynor and Kay, 2017), adenosine and the polyketides DIF-1, DIF-2 and MPBD (Morris et al., 1987, Morris et al., 1988, Saito et al., 2006) were without effect, as was the high cell density signal, polyphosphate (Suess and Gomer, 2016). However, conditioned medium (CM) prepared by shaking starving cells at high density for 8 hours was effective, with the active component(s) being heat-labile and retained by a 30 kDa cut-off membrane and therefore likely to be protein(s) (Figure 6C). Most likely this signal is one of the known proteins controlling early developmental events in Dictyostelium, but unfortunately these were unavailable for testing.
To gain insight into how developmental signals suppress macropinocytosis, we examined possible signal transduction routes, focussing on cyclic-AMP dependent protein kinase (PKA), which is a crucial mediator of both early and late events in development (Mann and Firtel, 1991, Harwood et al., 1992, Kay, 1989). PKA can be directly activated using the membrane-permeable analogue of cyclic-AMP, 8-bromo-cyclic-AMP (8-Br-cAMP) and we found that this, unlike cyclic-AMP, causes up to a 50% downregulation of macropinocytosis in starving, low-density cells (Figure 6D). High concentrations are required, but these are comparable to those used previously (Kay, 1989).
The involvement of PKA is strongly supported by mutants with elevated intracellular cyclic-AMP levels, due to defective breakdown. The hybrid cyclic-AMP phosphodiesterase RegA is activated by a His/Asp phospho-relay in which RdeA is the essential phosphate carrier protein (Shaulsky et al., 1998, Thomason et al., 1999, Thomason et al., 1998, Chang et al., 1998). Elimination of either protein results in strong downregulation of macropinocytosis in starving cells at low density (Figure 6E).
Conversely eliminating PKA activity by mutation of the catalytic subunit (pkaC-cells; (Primpke et al., 2000)) results in cells where macropinocytosis remains high for at least 24 hours after starvation, even if they are at high cell density or treated with CM (Figure 6F). Combined, these results strongly argue that macropinocytosis is downregulated in starving cells by PKA activation. These results are also relevant to the interpretation of recent work (Scavello et al., 2017) showing that pkaC-cells have a strong defect in chemotaxis to cyclic-AMP (see Discussion).
Discussion
To facilitate work on macropinocytosis in Dictyostelium, we have developed a high-throughput assay to measure fluid, membrane or particle uptake with single cell resolution. While this is by no means the first use of flow cytometry for these purposes (Bacon et al., 1994, Pan et al., 2016, King et al., 2013), the high-throughput nature of the assay provides a distinct advantage over previous techniques. A screen of inhibitors provides new tools for acute inhibition of macropinocytosis and further supports the involvement of PI3K, Rac, WASP, formins and the Arp2/3 complex expected from genetic and subcellular localization studies (Hoeller et al., 2013, Langridge and Kay, 2007, Dumontier et al., 2000, Veltman et al., 2016, Junemann et al., 2016).
Macropinocytosis in Dictyostelium occurs at a high rate in conditions where the cells are able to proliferate in liquid medium. However it is under physiological control with cells able to slowly transition between high and low macropinocytic states according to whether bacteria or soluble nutrients are available. In these transitions the frequency of macropinosome formation is altered: in axenic cells, where the active Ras patches are unconstrained by NF1, macropinosome size is additionally increased. Wild-type cells with an intact NF1 gene also transition between low and high macropinocytic states according to the nutrients available, showing that this regulation is not just a feature of axenic strains (this work, Maeda, 1983). The presence of a high macropinocytic state in wild-type cells suggests there are ecological circumstances where macropinocytosis is used for feeding, though these are yet to be defined.
Ax2 cells in the low macropinocytic state can sense bacteria through their secretion of folic acid, inhibiting macropinocytic upregulation accordingly. However, due to the relatively modest effects of folate, and the fact that it does not induce downregulation of macropinocytosis, it seems certain that other sensory pathways also play a prominent role. It has recently been reported that certain bacteria secrete cyclic-AMP, which functions as a chemoattractant for vegetative Dictyostelium (Meena and Kimmel, 2017), however cyclic-AMP did not affect macropinocytosis up- or downregulation.
We find that four nutrients from defined medium are largely responsible for inducing macropinocytosis upregulation in Ax2 cells: arginine, glutamate, lysine and a metabolisable sugar. None of the other amino acids appears effective individually, and even in combination they only have a modest effect. Arginine and lysine are essential amino acids, but glutamate is not (Marin, 1976, Franke and Kessin, 1977). Dictyostelium has several receptors similar to metabotropic glutamate receptors (Taniura et al., 2006, Fey et al., 2013), but it seems likely that the major route for nutrient sensing is intracellular, with nutrients delivered by macropinocytosis.
In mammalian cells, free amino acids obtained by macropinocytosis are sensed through mTORC1 (Yoshida et al., 2015), which is recruited to the lysosome by Rag proteins and activated (Sancak et al., 2010). Activation of mTORC1 inhibits autophagy. Dictyostelium autophagy can be induced within minutes by removal of arginine and lysine (King et al., 2011), the same amino acids that induce macropinocytosis upregulation. This argues that TORC1 is activated by arginine and lysine in Dictyostelium to both upregulate macropinocytosis and inhibit autophagy. Though the TORC1-specific inhibitor rapamycin does not induce Dictyostelium autophagy (Dominguez-Martin et al., 2017), other mTor inhibitors are more effective (Cardenal-Munoz et al., 2017), suggesting that rapamycin is a relatively poor inhibitor for some Dictyostelium TORC1 processes, as has been suggested in mammalian cells (Thoreen and Sabatini, 2009). The more potent, but less specific, inhibitor torin 1 inhibits both macropinocytosis and macropinocytosis upregulation, although this is likely to be at least partially due to TORC2 inhibition.
As only metabolizable sugars induce upregulation of macropinocytosis, it is probable that the sensing of these is through a general metabolic readout, such as the presence of ATP, which is produced during glycolysis. Increased levels of AMP and ADP, as in nutrient poor conditions (such as without sugar), activate AMP-kinase. Overexpression of a constitutively active AMP-kinase α subunit in Dictyostelium inhibits growth but does not affect macropinocytosis (Bokko et al., 2007), similar to what we observe in low-density starvation conditions. It may therefore be the case that in the absence of a sugar source AMP-kinase is activated, which could prevent full macropinocytosis upregulation.
Our results show that the cessation of macropinocytosis during early development requires a developmental signal that most likely acts through PKA. Macropinocytosis does not cease immediately when cells are starved, but decreases over several hours and so may occur at reduced levels in cells that are used for studying chemotaxis to cyclic-AMP, particularly in mutants with a defect in early development. This can be a confounding influence on studies of chemotaxis, since macropinocytosis uses the same actin machinery as pseudopods and thus impairs chemotaxis (Veltman, 2015). In particular, we found that macropinocytosis continues at a high rate in mutants of the PKA catalytic subunit, possibly accounting for the strong chemotactic defect of these strains (Scavello et al., 2017). This could also be a confounding issue for some other strains with early developmental defects (Khosla et al., 2005, Wu et al., 1995, Rodriguez et al., 2008, Lee et al., 2005).
Many of the molecular components required for macropinocytosis in Dictyostelium are the same as those in mammalian cells: actin, Arp2/3, PI3K, SCAR/WAVE, WASP, Rac and Ras proteins. This conservation of the core components suggests that macropinocytosis may have first arisen in simple protists as a way of feeding in the absence of bacterial prey. In mammalian cells there are additional levels of regulation, some of which are cell-type specific (such as the calcium requirement in immune cells) and others that are more generic (such as growth factor stimulated macropinocytosis). We believe that Dictyostelium is an excellent model organism for establishing the core, conserved elements of macropinocytosis and their function.
Materials and Methods
Cell culture and materials
Cells were cultivated at 22°C. HL5, SIH, variants of SIH and SM media were from Formedium. Unless otherwise specified, cells were grown on Klebsiella aerogenes (Ka) lawns on SM plates and harvested for experiments from the feeding front, washing three times with KK2 (16.6 mM KH2PO4, 3.8 mM K2HPO4, pH 6.1) by centrifugation (280 g, 3 min) to remove the bacteria. Cells were also grown in tissue culture plates with Ka as a food source. In this case Ka was added to KK2MC (KK2 + 2 mM MgSO4, 100 μM CaCl2) to 2 OD600 nm from a 100 OD600 nm stock (these bacteria were grown overnight in 2xTY, pelleted by centrifugation and washed twice in KK2).
Cells were grown axenically in HL5 in conical flasks with shaking at 180 rpm. Media derived from SIH, including SUM, were made in KK2MC pH 6.5.Conditioned medium was made by washing axenically grown Ax2 cells free of HL5, resuspending them to 1x107 cells ml-1 in KK2MC and incubating for 8 hours, 180 rpm, before removing the cells by centrifugation (2400 g, 10 min). Strains are listed in Table S4.
For transformation, cells were harvested from bacteria, resuspended in H40 buffer (40 mM Hepes, 1 mM MgCl2, pH 7.0), mixed with 500 ng vector for a PIP3 reporter (PkgE-PH mCherry), electroporated in ice-cold 2 mm cuvettes (Novagen) using a square wave protocol (2x 350 volts, 8 ms apart), then transferred to 2 ml KK2MC + Ka in a 6 well plate to recover for 5 hours, before G418 selection was added to 10 μg ml-1.
Chemicals were from Sigma unless otherwise indicated. Polyphosphate was obtained from both Spectrum and Merck.
Uptake measurements by fluorimetry
Based on (Rivero and Maniak, 2006): cells at 1x107 ml-1 were shaken at 180 rpm in HL5 with 0.5 mg ml-1 TRITC-dextran and at each time point triplicate 0.8 ml samples centrifugally washed once in ice-cold KK2 and resuspended to 1 ml. Fluorescence was measured in a fluorimeter (Perkin-Elmer LS 50 B with excitation at 544 nm, emission at 574 nm, slit width 10 nm). Background ‘0 minute’ fluorescence was subtracted and uptake volume calculated using standard curves of TRITC-dextran diluted in buffer. Cells loaded in this way were also analysed by flow cytometry (LSR_II flow cytometer, BD Biosciences) to compare the methods.
To measure yeast uptake, cells were resuspended from bacterial plates or 9 cm tissue culture dishes where they had been incubated in growth medium to 5x106 ml-1 in KK2MC in a 5 ml conical flask shaken at 180 rpm, 22oC. TRITC-labelled yeast (sonicated at level 7.0 for 20 seconds on a Misonix sonicator 3000) were added to 1x107 particles ml-1. At 0 and 60 minutes duplicate 200 μl samples were added to 20 μl of Trypan blue quench solution (2 mg ml-1 in 20 mM citrate, 150 mM NaCl, pH 4.5) on ice, shaken for 3 minutes at 2000 rpm, spun down and washed twice with ice-cold KK2 + 10 mM EDTA. The final pellet was resuspended to 1 ml and the fluorescence compared to a standard curve to give the number of yeast per cell.
Uptake measurements by flow cytometry
For high-throughput assays, 50 μl of 1x105 cells ml-1 was inoculated into flat-bottom 96-well plates and incubated at 22oC for the indicated time (usually 24 hours). Then 50 μl of 1 mg ml-1 TRITC-dextran in the same medium was added. After one hour, unless otherwise stated, the medium was thrown off, and the cells washed by ‘dunk-banging’ (the plate was submerged in a container of ice-cold KK2, which was thrown off and the plate patted dry) before 100 μl KK2MC containing 5 mM sodium azide was added to each well to detach the cells and stop exocytosis. Cells were analysed by flow cytometry (LSR-II, BD Biosciences) using the High-Throughput-Sampling attachment, which pipetted them up and down twice, before analysing 65 μl per sample at 3 μl s-1. FlowJo software (https://www.flowjo.com) calculated the median (mean in the case of beads) fluorescence of cells in each well, and then the mean of triplicate wells was calculated. The mean was then taken of all biological replicates. To determine volumes taken up, the same population of cells (loaded with TRITC-dextran in suspension, as above) was analysed by both fluorimetry and flow cytometry. The LSR_II was calibrated through all subsequent experiments using FlowSet fluorospheres calibration beads (Beckman Coulter).
We also used this method to measure uptake of membrane using 10 μM FM1-43 (Invitrogen); phagocytosis of bacteria using 1x108 particles ml-1 Texas-red E. coli bioparticles (Thermo Scientific); or beads of different sizes (YG-beads, Polysciences): 3 μm (2x107 ml-1), 2.0 and 1.75 μm (5x107 ml-1) or 1.5 μl-1 (1x108 ml-1). Particles internalised per cell was calculated by comparing the internalised fluorescence with particles only samples. For time courses, the start time was staggered so that all time-points ended concurrently. When inhibitors were used acutely, they were added with the fluorescent medium to the final indicated concentration. Polyketides were synthesised as described (Morris et al., 1987, Morris et al., 1988, Saito et al., 2006).
To initiate development, axenically growing cells were washed twice, resuspended to 1x107 cells ml-1 in KK2MC and shaken at 180 rpm for one hour before delivering pulses of KK2MC containing cyclic-AMP to give a concentration of 100 nM every 6 minutes using a Watson Marlow 505Di pump. At the indicated times 5x104 cells were diluted into dextran containing KK2MC in 24 well plates for one hour, after which they were washed in-situ using ice-cold KK2 + 10 mM EDTA and detached with KK2MC + 5 mM sodium azide. 100 μl was transferred to duplicate wells in a 96-well plate for flow cytometry analysis.
Development on agar plates was initiated by settling 1.5 ml of washed, axenically grown cells at 2.5x107 cells ml-1 in KK2MC onto fresh 1.8% KK2MC agar in 6 cm plates. After 15 minutes settling, the media was aspirated off, and the plates kept on wet tissues at 22°C. At the indicated times, cells were harvested, resuspended in KK2MC and 1x105 inoculated into KK2MC in a 6-well plate with 0.5 mg ml-1 TRITC-dextran for one hour. Cells were then washed in-situ and resuspended in KK2 + 10 mM EDTA before analysis by low-throughput flow cytometry. The zero hour time-point was of cells taken immediately after washing.
Macropinosome formation rate and diameter
The rate of macropinosome formation was determined in KK2MC by loading cells in a 2-well microscope slide (Nunc) with 2 mg ml-1 FITC-dextran for 1 minute, then washing and fixing with 4% paraformaldehyde for 20 minutes. Fixed cells were washed 5 times and stored in PBS (pH 5.0) at 4oC for imaging. Z-stacks with 0.1 μm steps were taken using a Zeiss 700 series microscope with 2x averaging to reduce noise. Maximum intensity projections were made using FIJI and FITC-positive endosomes counted by eye. The mean of at least 8 cells on a given day was taken as one data-point.
To measure macropinosome diameter at closure, cells in KK2MC expressing a PIP3 reporter (PkgE-PH mCherry) were filmed in their central section at 1 frame per second for 5 minutes on a Zeiss 700 series microscope. The maximum diameter of macropinosomes at closure was measured using the FIJI measure tool. Note that this method will underestimate the diameter of macropinosomes not lying fully within the optical section.
Competing Interests
The authors declare that there are no competing interests.
Author Contributions
Both authors designed the experiments, which were carried out by Thomas Williams.
Funding
We thank the Medical Research Council UK for core funding (U105115237 to RRK).
Data availability
N/A.
Acknowledgements
We thank the rest of the Kay lab for their assistance in moulding this project, particularly Peggy Paschke. Clelia Amato and Robert Insall (Beatson Institute, Glasgow) alerted us to the Rac inhibitor. Jason King (Sheffield University) provided valuable feedback on the macropinosome formation experiments. Miao Pan and Tian Jin (NIAID, Bethesda) kindly sent us the fAR1- strain. The MRC-LMB flow cytometry facility maintained the flow cytometers and provided technical support.