PtdIns(3,4)P₂, Lamellipodin, and VASP coordinate actin dynamics during phagocytosis in macrophages

Phagocytosis, the process whereby cells engulf and dispose of effete cells, microorganisms, and foreign particles, is pivotal for immunity and tissue homeostasis (Stuart and Ezekowitz, 2005; Flannagan et al., 2012). Phagosome formation, the initial step during phagocytosis, entails marked reorganization of the actin cytoskeleton and membrane remodeling events that lead to pseudopod extension and particle engulfment (Levin et al., 2016; Freeman and Grinstein, 2014). Earlier studies have suggested that phosphoinositides are pivotal molecules in the regulation of phagocytosis. Phosphoinositides, which play crucial roles in signaling and membrane traffic (de Craene et al., 2017), reside primarily in the cytosolic leaflet of organelles including the plasma membrane and phagosomes. Phosphatidylinositol 4,5-bisphosphate (PtdIns[4,5]P₂) is enriched in the plasma membrane where, amongst its many roles, it acts as a positive regulator of actin polymerization (Raucher et al., 2000). The conversion of PtdIns(4,5)P₂ to phosphatidylinositol 3,4,5-trisphosphate (PtdIns[3,4,5]P₃) is a potent stimulus resulting in alterations in actin dynamics through the activation/inactivation of Rho-GTPases, as well as promoting the hydrolysis of PtdIns(4,5)P₂ by activating phospholipase Cγ. Indeed, it has been appreciated for nearly two decades that IgG-opsonized particle engagement by the phagocytic Fcγ receptor leads to the activation of the phosphoinositide-3-kinase (PI3K; Gu et al., 2003) that is required for the internalization of large phagocytic prey (Araki et al., 1996; Cox et al., 1999). Remarkably, the dynamic changes in phosphoinositides and the actin cytoskeleton during phagocytosis are restricted to the site of particle engagement and do not propagate to the rest of the plasma membrane. Accordingly, phagosome formation is characterized by the focal generation of PtdIns(3,4,5)P₃ (Dewitt et al., 2006) at the site of contact and extending pseudopods, while a concomitant disappearance of PtdIns(4,5)P₂ is observed at the base of the phagocytic cup prior to sealing of the nascent phagosome (Botelho et al., 2000).

The downstream effects of PI3K activation during phagocytosis have been mostly attributed to the generation of PtdIns(3,4,5)P₃. However, some of these findings should be interpreted with caution, as most were obtained using probes with dual specificity for PtdIns(3,4,5)P₃ and PtdIns(3,4)P₂, such as the pleckstrin-homology (PH) domain of Akt (Ebner et al., 2017; Manna et al., 2007). Recent advances in lipid bio-sensor design have enabled the distinction of the differential roles of PtdIns(3,4,5)P₃ and the related PtdIns(3,4)P₂ (Goulden et al., 2019; Liu et al., 2018). PtdIns(3,4)P₂ is increasingly appreciated as involved in cellular processes such as endocytosis, macropinocytosis, cell migration, and neurite initiation (Posor et al., 2013; Li and Marshall, 2015; Zhang et al., 2017; Gozzelino et al., 2020). To date, however, the role of PtdIns(3,4)P₂ and its effectors during phagocytosis remains largely unknown. Using recently developed specific probes, we demonstrate here that PtdIns(3,4)P₂ accumulates greatly at sites of phagocytosis, where it persists long after PtdIns(3,4,5)P₃ disappears. More importantly, we show that selective depletion of PtdIns(3,4)P₂ attenuates particle internalization, highlighting the importance of this unique phosphoinositide.

The distribution of PtdIns(3,4)P₂ in unstimulated RAW 264.7 macrophages was investigated first. To this end, we employed a recently described genetically encoded biosensor based on tandem carboxy-terminal PH (cPH) domains of TAPP1 (Thomas et al., 2001; Goulden et al., 2019). Using these biosensors, NES-mCh-cPHx3 and NES-EGFP-cPHx2 (cPHx3 and cPHx2, respectively), we detected a discrete plasmalemmal pool of PtdIns(3,4)P₂ in resting cells (Fig. 1 A, left panel). To confirm the specificity and responsiveness of the probes, we co-expressed inositol polyphosphate 4-phosphatase type II (INPP4B), which selectively hydrolyzes the D-4 position of PtdIns(3,4)P₂ (Gewinner et al., 2009; Norris and Majerus, 1994). To ensure plasmalemmal targeting of INPP4B, the phosphatase was attached to a carboxy-terminal CAAX motif (INPP4B-CAAX) that is prenylated and polycationic. As shown in Fig. 1 A (middle and right panels), the biosensors detached from the membrane upon co-expression of INPP4B-CAAX, but not when the catalytically inactive INPP4B(C842A)-CAAX mutant was co-expressed. These observations validate the selectivity of the probes and confirm that modest yet detectable amounts of PtdIns(3,4)P₂ are indeed present in the membrane of resting macrophages. Furthermore, we documented that production of PtdIns(3,4)P₂ is PI3K dependent, since treatment with nanomolar concentrations of the PI3K inhibitor wortmannin completely released the cPHx3 probe from the plasma membrane (Fig. 1, B and C).

Next, we examined the distribution of PtdIns(3,4)P₂ during phagocytosis. Upon exposure to IgG-opsonized sheep erythrocytes (SRBCs), RAW 264.7 cells exhibited a marked accumulation of the PtdIns(3,4)P₂ probe at the site of phagocytosis to levels greatly surpassing those of the neighboring unengaged plasma membrane, an observation consistent with stimulated local production of the lipid (Fig. 2 A, left panel). PtdIns(3,4)P₂ accumulated in the phagosomal cup and remained in the membrane even after scission of the nascent phagosome (Video 1). Of note, there were marked differences in the distribution and dynamics of recruitment of cPHx2 and PH-BTKx2 (a sensor for PtdIns[3,4,5]P₃) during phagocytosis. Specifically, PtdIns(3,4,5)P₃ accumulated primarily—albeit transiently—at the base of the phagocytic cup, whereas PtdIns(3,4)P₂ was acquired slightly later, initially attaining higher levels at the tips of the extending pseudopods (Fig. S1 and Video 2) and later persisting for a few minutes in the sealed phagosome. The accumulation of the PtdIns(3,4)P₂ probe at the phagosomal cup was largely obliterated by co-expression of the INPP4B-CAAX construct, but no depletion was observed when the catalytically inactive INPP4B(C842A)-CAAX construct was used as negative control (Fig. 2, A and B). These findings imply that PtdIns(3,4)P₂ is produced locally at sites of phagocytosis.

We next sought to elucidate the pathway(s) responsible for the biosynthesis of PtdIns(3,4)P₂ during phagocytosis. We considered two possibilities, which are not mutually exclusive: that the PtdIns(3,4)P₂ was being produced by dephosphorylation of PtdIns(3,4,5)P₃ and/or that it was synthesized from PI(4)P by class II PI3Ks. Class I PI3K is known to be activated during phagocytosis and its inhibition arrests engulfment by preventing full extension of the pseudopods around the particle (Araki et al., 1996; Cox et al., 1999). To assess its involvement, we treated cells with nanomolar concentrations of the pan-PI3K inhibitor PI-103 or the class I PI3K-selective inhibitor GDC-0941 and monitored PtdIns(3,4)P₂ formation during phagocytosis (Fig. 2, C and D). In both cases, the recruitment of cPHx2 was virtually eliminated, consistent with a major role for class I PI3Ks in the generation of PtdIns(3,4)P₂. Nevertheless, we examined the possible contribution of Class II PI3Ks. For this, we acutely depleted the plasmalemmal pool of PtdIns(4)P using GSK-A1, a potent and specific inhibitor of the type III phosphatidylinositol 4-kinase PI4KA (PI4KIIIα) that is largely responsible for the generation and maintenance of plasmalemmal PtdIns(4)P; it is noteworthy that selective inhibition of PI4KIIIα by GSK-A1 does not acutely alter the plasmalemmal levels PtdIns(4,5)P₂ (Bojjireddy et al., 2014), the substrate of class I PI3Ks. Upon treatment with GSK-A1, the recruitment of mCherry-cPHx3 to the phagocytic cup remained largely unaffected despite a profound depletion of PtdIns(4)P monitored by the high-avidity biosensor 2xP4M (Fig. 2 E). These findings strongly suggest that dephosphorylation of PtdIns(3,4,5)P₃ is the main source of PtdIns(3,4)P₂ production at the phagocytic cup.

The involvement of PI3Ks that generate PtdIns(3,4,5)P₃ suggests that phosphoinositide 5-phosphatases are also required for the synthesis of PtdIns(3,4)P₂ during phagocytosis. Multiple 5-phosphatases could in theory play a role in the production of PtdIns(3,4)P₂. For instance, Src homology 2 domain-containing inositol-5-phosphatase 2 (SHIP2) is involved in the production of PtdIns(3,4)P₂ at sites of endocytosis (Hak et al., 2018) and at invadopodia (Sharma et al., 2013). Importantly, SHIP2 has also been reported to translocate to sites of phagocytosis (Ai et al., 2006), making it an attractive candidate to mediate the production of PtdIns(3,4)P₂. However, the overexpression of the catalytically inactive mutant of SHIP2 (D607A), which is expected to exert a dominant-negative effect, had no effect over phagosomal levels of PtdIns(3,4)P₂ (Fig. S1, C and D). Similar negative results were observed when the cells were treated with either a SHIP2-selective inhibitor (AS1949490) or with the pan-SHIP1/2 inhibitor (K118; Fig. S1 E). We therefore turned our attention to other 5-phosphatases: there is compelling evidence that SHIP1 (Ming-Lum et al., 2012), INPP5E (Segawa et al., 2014), and OCRL (Bohdanowicz et al., 2012) are all recruited to nascent phagosomes. In addition, we identified synaptojanin-2 (SYNJ2) and INPP5B as being present at the site of phagocytosis (Fig. 2 F). It is therefore conceivable that multiple 5-phosphatases collaborate to dephosphorylate PtdIns(3,4,5)P₃ to PtdIns(3,4)P₂ (Malek et al., 2017). Because of the likelihood of functional redundance, the identity of the specific phosphatases involved was not pursued further.

We next sought to determine whether PtdIns(3,4)P₂ accumulation is necessary for efficient FcγR-mediated particle uptake. To this end, phagocytic efficiency was compared in RAW 264.7 macrophages overexpressing INPP4B-CAAX to deplete plasmalemmal PtdIns(3,4)P₂, or the catalytically inactive version INPP4B(C842A)-CAAX, used as a negative control. We observed a stark decrease in the phagocytic efficiency of cells expressing the INPP4B-CAAX, compared to cells expressing the INPP4B(C842A)-CAAX mutant (Fig. 3, A and B). In addition to the decrease in efficiency of internalization, we noted a modest reduction in the number of SRBCs that were contacted by the INPP4B-CAAX expressing cells. (Fig. S2 C and Fig. 3 A, right panel). Thus, we hypothesized that a defect in pseudopod extension or sealing must be responsible for the observed decrease in phagocytic efficiency.

Actin remodeling, which drives pseudopod extension, is a hallmark of the early stages of the phagocytic process. To assess the role of PtdIns(3,4)P₂ in actin polymerization, we performed live-cell imaging of cells co-expressing LifeAct-GFP and either INPP4B-CAAX or INPP4B(C842A)-CAAX. In cells expressing the inactive phosphatase, focal actin polymerization was followed by pseudopod formation, and progressive particle engulfment in a zipper-like fashion (Fig. 3, C, right; Fig. 3 D, bottom row; and Video 2). In contrast, cells where PtdIns(3,4)P₂ was depleted by expression of INPP4B-CAAX formed pseudopods that failed to progress and wrap around the phagocytic target, despite exhibiting an apparently normal initial actin polymerization (Fig. 3 C left; Fig. 3 D, top row; and Video 3). These findings indicate that PtdIns(3,4)P₂ is necessary for phagocytosis and that it plays a direct role in regulating actin organization and pseudopod progression during the early stages of particle engulfment.

To date, only a handful of proteins have been identified as specific effectors of PtdIns(3,4)P₂ (Li and Marshall, 2015). These include the modular adaptor protein Ras-associated and pleckstrin homology domains containing protein 1, more commonly referred to as Lpd. Through its ability to cluster and tether Ena/vasodilator-stimulated phosphoprotein (VASP) proteins to actin filaments, Lpd is a key regulator of the actin cytoskeleton (Hansen and Mullins, 2015). As such Lpd has roles in lamellipodial formation (Krause et al., 2004), stabilization of actin-dependent cellular protrusions (Dimchev et al., 2019,Preprint), cell migration, and endocytosis (Dimchev et al., 2020; Carmona et al., 2016; Lagarrigue et al., 2015; Vehlow et al., 2013; Hak et al., 2018). Because the PH domain of Lpd has been shown to bind PtdIns(3,4)P₂ (Krause et al., 2004), it appeared a likely candidate for the regulation of pseudopod extension during particle engulfment. To examine this possibility, Lpd was expressed in RAW 264.7 macrophages and its distribution assessed during phagocytosis of IgG-opsonized SRBCs. We observed robust accumulation of GFP-Lpd at the phagocytic cup, where PtdIns(3,4)P₂ was also enriched (Fig. 4 A). Furthermore, upon treatment with wortmannin the levels of both PtdIns(3,4)P₂ and Lpd at the phagocytic cup decreased in parallel fashion (Fig. 4, B and C). Consistent with the notion that Lpd binds to PtdIns(3,4)P₂ via its PH domain, we found that a GFP-tagged tandem PH domain of Lpd (Lpd-2xPH) was recruited to the phagocytic cup in cells expressing the inactive INPP4B(C842A)-CAAX (Fig. 4 D) yet failed to accumulate in cells expressing INPP4B-CAAX. Furthermore, in resting RAW 264.7 cells, Lpd-2xPH showed modest accumulation at the plasma membrane, as we had seen earlier for the cPHx3 and cPHx2 probes, and this limited recruitment was similarly abolished by INPP4B-CAAX, but not by the catalytically inactive INPP4B(C842A)-CAAX (Fig. S2, A and B). From these experiments, we conclude that Lpd is recruited early during phagocytosis at least in part by its PH domain–mediated interaction with PtdIns(3,4)P₂.

We next sought to determine whether Lpd is necessary for FcγR-mediated phagocytosis. The expression of endogenous Lpd in RAW 264.7 macrophages and its susceptibility to shRNA-mediated silencing were first validated by immunostaining (Fig. 5 A and Fig. S3). The enrichment of the endogenous Lpd during phagocytosis was similarly confirmed by immunostaining; its distribution at the phagocytic cup closely resembled that of F-actin stained with phalloidin (Fig. 5 B). Upon silencing of Lpd, we observed an aberrant “flaring” of F-actin around the opsonized particle (Fig. 5 C, top right panel). Lpd-silenced macrophages exhibited “loose” phagocytic cups characterized by multiple protrusions, in contrast to the tightly apposed pseudopods of control cells. In addition, Lpd-shRNA cells exhibited a ∼60% decrease in phagocytic efficiency compared to control cells (Fig. 5, D and E). Taken together, these findings suggest that Lpd is necessary for proper F-actin organization and pseudopod extension during phagocytosis.

Ena/VASP proteins promote actin polymerization by accelerating filament elongation and opposing the action of capping proteins (Barzik et al., 2005; Hansen and Mullins, 2010; Breitsprecher et al., 2011). Ena/VASP proteins regulate the cytoskeleton during TCR signaling (Krause et al., 2000) and also play a role in Fc-mediated phagocytosis (Coppolino et al., 2001) and macroendocytosis in Dictyostelium discoideum (Körber and Faix, 2022). It is relevant that Ena/VASP proteins harbor an EVH1 domain that interacts with the multiple proline-rich regions (FPPPP) present in Lpd (Krause et al., 2004; Hansen and Mullins, 2015; Fig. 6 A) and that they jointly regulate the dynamics of filopodia (Cheng and Mullins, 2020). We sought to determine if Lpd and VASP associate within the phagocytic cup. Co-expression of the Lpd and VASP constructs revealed colocalization of these proteins during phagocytosis. Notably, Lpd, VASP, and F-actin all share a similar distribution within the phagocytic cup (Fig. 6 B). We also confirmed the enrichment of endogenous VASP at the phagocytic cup through immunostaining (Fig. 6 E).

Next, we tested whether VASP was necessary for phagocytosis. To this end, we took advantage of the Listeria monocytogenes effector protein ActA, which binds tightly to VASP. We expressed an N-terminally truncated ActA protein that includes the four repeats of the VASP-binding sequences attached to a C-terminal motif that targets the protein to the cytosolic surface of mitochondria (mRFP-Mito-FP4; Pistor et al., 1994; Bear et al., 2000). Expression of mRFP-Mito-FP4 effectively sequestered the vast majority of VASP to the mitochondria and prevented its accumulation at the phagocytic cup, while a mutant version (mRFP-Mito-AP4) that also targets to mitochondria but is unable to bind VASP was without effect (Fig. 6 C and Fig. S4 A). Tethering of VASP to the mitochondria by means of mRFP-Mito-FP4 reduced the phagocytic efficiency from 79.8 to 31.4% (Fig. 6 D). These results demonstrate that VASP is a positive regulator of phagocytosis and support the notion that Lpd is exerting its effects at least in part by binding to VASP.

Next, we examined whether the interaction between Lpd and VASP was necessary for phagocytosis. We overexpressed a mutant version of Lpd in which all Ena/VASP-binding sites had been inactivated through mutations (Lpd^EVmut). Overexpression of GFP-Lpd^EVmut produced a ∼57% decrease in phagocytosis when compared to RAW 264.7 cells transfected with the wild-type GFP-Lpd construct (Fig. 7, A and B). Furthermore, even though the initial recruitment of GFP-Lpd^EVmut to the phagocytic cup was unaffected, the distribution of the construct became altered during phagocytic cup formation (Fig. S4 B). We noticed that the phagocytic cups formed by this mutant contained multiple filopodia- and ruffle-like projections that were rich in F-actin, as visualized through phalloidin staining (Fig. 7 C). These atypical phagocytic cups and pseudopods closely resembled the ones we had previously observed upon silencing of Lpd (Fig. 5 C). Additionally, these aberrant phagocytic cups were also enriched in PtdIns(3,4)P₂, detected by the cPHx3 probe (Fig. 7 D), demonstrating that the lipid metabolism itself was unperturbed by expression of the Lpd mutant. These findings suggest that interactions between Lpd and VASP are required for proper cytoskeleton organization and coordinated extension of pseudopods at the site of particle engulfment.

We found that comparatively small amounts of PtdIns(3,4)P₂ are present in the plasma membrane of resting RAW 264.7 macrophages and that this phosphoinositide is greatly enriched at the sites of particle engagement during FcγR-mediated phagosome formation. Selective depletion using a highly specific phosphatase revealed that PtdIns(3,4)P₂ has a critical role in supporting phagocytosis. This was previously unrecognized in part due to the use of probes like AKT-PH that have dual specificity and of PI3K inhibitors, rather than targeted enzymatic depletion by INPP4B. We also report that Lpd and its binding partner VASP are jointly required for robust phagocytosis: loss of either from the site of phagocytosis results in an atypical and seemingly uncoordinated actin assembly within the extending pseudopods that hence fail to encircle the target.

The importance of PI3Ks during the process of phagocytosis has been widely documented (Cox et al., 2002; Vieira et al., 2001; Tamura et al., 2009). Genetic manipulation of PI3Ks as well as the use of pharmacological inhibitors impairs phagocytosis, particularly that of larger targets. Additionally, localized production of PtdIns(3,4)P₂ or PtdIns(3,4,5)P₃ at the phagocytic cup has been implicated in phagosomal closure and Ca²⁺ signaling in HL60 neutrophils (Dewitt et al., 2006). Although some of these defects can be directly linked to the role of PtdIns(3,4,5)P₃ cognate effectors, based on this study it is likely that a second, underappreciated effect of PI3K inhibition is the impairment of PtdIns(3,4)P₂ formation, which is itself required for phagocytosis. Curiously, in our studies, we find that PtdIns(3,4)P₂ is found at both the base of the phagocytic cup as well as at the tips of the pseudopods, whereas PtdIns(3,4,5)P₃ is largely restricted to the base of the phagocytic cup (Fig. S1). This raises the possibility that the relevant phosphoinositide 5-phosphatases are particularly active at the tips of the advancing pseudopods. Phenotypically, inhibition of PI3K—and thus attenuation of both PtdIns(3,4)P₂ and PtdIns(3,4,5)P₃ production—results in arrest of pseudopod progression roughly halfway around the opsonized particle (Araki et al., 1996; Cox et al., 1999), while selective depletion of PtdIns(3,4)P₂ prevents pseudopod extension (Fig. 3, C and D, right panel, and Video 3). PtdIns(3,4,5)P₃ is thought to regulate actin dynamics during phagocytosis by locally activating Rho family GEFs and GAPs (Schlam et al., 2015), and by activating phospholipase Cγ, thereby depleting PtdIns(4,5)P₂ from the base of the phagocytic cup. In view of our observations, it seems worthwhile to reconsider whether the purported PtdIns(3,4,5)P₃ effectors are in fact selective for this lipid or are instead responsive to PtdIns(3,4)P₂.

An earlier report suggested that the inositol polyphosphate 4-phosphatase type 1 (INPP4A), which is expressed in RAW macrophages, is a negative regulator of phagocytosis (Nigorikawa et al., 2015). That study documented enhanced ability to internalize particles by RAW cells expressing shINPP4A and by primary macrophages from INPP4A-knockout mice. While the authors did not provide a mechanistic explanation for the observed effects, their findings are consistent with our conclusion that increased PtdIns(3,4)P₂ is essential for optimal phagocytosis, at least partly via Lpd and VASP.

Lpd belongs to the Mig-10, RIAM, Lpd (MRL) family of modular adaptor proteins (Coló et al., 2012). RIAM, despite sharing homology with Lpd (Lafuente et al., 2004), is thought to act downstream of Rap1, and evidence suggests that its PH domain binds preferentially to PtdIns(3)P, PtdIns(5)P (Jenzora et al., 2005), and PtdIns(4,5)P₂ (Patsoukis et al., 2017; Wynne et al., 2012), not PtdIns(3,4)P₂. Previously, RIAM was described to support complement-dependent phagocytosis by relaying integrin signaling (Torres-Gomez et al., 2020). However, RIAM is not required for FcγR-dependent phagocytosis (Medraño-Fernandez et al., 2013), the mode we studied. Furthermore, our results indicate that silencing of Lpd produces severe defects in the phagocytic cups of macrophages where RIAM was left intact. These findings suggest that Lpd, contrary to RIAM, is necessary for Fc-mediated phagocytosis and that these MRL proteins play non-redundant roles during this process.

We found that the PH domain of Lpd is sufficient for its localization at the phagocytic cup. It is possible, however, that other interactions contribute to its recruitment and retention within the forming phagosome. Indeed, we noted that Lpd dissociates from the maturing phagosome prior to the disappearance of the PtdIns(3,4)P₂, suggesting that it may function as a co-incidence detector; other potential-binding partners may well be present at the site of phagocytosis. In this regard, it is noteworthy that Lpd directly binds actin filaments through an unstructured and highly basic region in its carboxyl-terminus (Hansen and Mullins, 2015). Furthermore, Lpd can also bind to active Rac1 (Law et al., 2013; Bae et al., 2014), which is similarly recruited to forming phagocytic cups (Scott et al., 2005). Additionally, the formin-binding protein 17 binds to Lpd and recruits it to the membrane during fast endophilin-mediated endocytosis (Chan Wah Hak et al., 2018). Of note, formin-binding protein 17 is also present within podosomes and phagocytic cups in macrophages (Tsuboi et al., 2009).

Lpd is central to the regulation of cytoskeletal dynamics (Krause et al., 2004) as it interacts with multiple actin regulators such as the Ena/VASP family proteins (Chang et al., 2013; Carmona et al., 2016; Hansen and Mullins, 2015) and the SCAR/WAVE regulatory complex (Carmona et al., 2016; Law et al., 2013). Ena/VASP proteins localize to the phagocytic cup and are indispensable for phagocytosis: Coppolino et al. (2001) reported that upon binding of Ena/VASP to a cytosolic GFP-ActA construct, phagocytosis was inhibited. In the current study, we were able to validate these findings using a more thorough method that entailed mistargeting Ena/VASP proteins to the mitochondria. Additionally, we found that all three Ena/VASP family members (EVL, Mena, VASP) localize to the phagocytic cup when ectopically expressed (Fig. S4 C). Phosphorylation of Lpd by c-Abl kinase reportedly increases its interaction with Ena/VASP (Michael et al., 2010). While we did not explore this possibility in the current study, Abl family kinases have been implicated in the positive regulation of phagocytosis (Greuber and Pendergast, 2012). Therefore, it is possible that a kinase-dependent increase in Lpd–Ena/VASP interaction is one of the mechanisms by which this positive regulation takes place. Furthermore, PI3K activation through Abl kinase was also recently reported to take place during podosome formation in macrophages (Zhou et al., 2020).

In addition to VASP, Lpd can also regulate the actin cytoskeleton through its interactions with the SCAR/WAVE complex. A recent study demonstrated that loss of the WAVE regulatory complex (WRC) impairs phagocytosis (Stahnke et al., 2021). In our study, we compared the effects of Lpd mutants in which all Ena/VASP or all SCAR/WAVE-binding sites had been mutated. The Lpd^EVmut caused the most robust decrease in phagocytic efficiency and aberrant phagocytic cups (Fig. 7). Nevertheless, the SCAR/WAVE binding-deficient mutant also exhibited a smaller, yet significant, decrease in phagocytic efficiency (Fig. S5). Furthermore, Abi1, the component of the WRC that interacts directly with Lpd (Law et al., 2013), was also localized to the phagocytic cup (Fig. S5 A). Jointly, these findings suggest that Lpd is central to the regulation of actin dynamics during phagocytic cup formation not only through its interactions with VASP but probably also through the regulation of the WRC, although additional experiments will be needed to validate the involvement of the latter pathway.

The diversity of phagocytic targets and receptors entails different molecular mechanisms during phagocytic cup formation. Despite these differences, all types of phagocytosis share an inherent dependence on the rearrangement of the actin cytoskeleton and on the dynamic remodeling of the plasma membrane. FcγR-mediated phagocytosis is the best characterized model of phagocytosis and the one used throughout this study. However, complement-mediated phagocytosis is another important type of opsonin-mediated phagocytosis in which the receptors recognize activated complement components, such as iC3b, deposited on the phagocytic target (van Lookeren Campagne et al., 2007). Here, we were able to document that localized synthesis of PtdIns(3,4)P₂ and recruitment of Lpd and VASP to the phagocytic cup also occur during CR3-mediated phagocytosis (Fig. S5, D–F). However, the overall phagocytic efficiency was unaffected in INPP4B-expressing cells (Fig. S5 G), possibly due to the compensation by the RIAM-VASP module described during CR3-mediated phagocytosis (Torres-Gomez et al., 2020). These findings are in agreement with previous reports that have identified PtdIns(3,4)P₂ (Dewitt et al., 2006) and VASP (Torres-Gomez et al., 2020) during complement receptor-mediated phagocytosis. Furthermore, the scaffolding protein SWAP70 was identified as a PtdIns(3,4)P₂ effector during phagocytosis of zymosan particles in dendritic cells (Baranov et al., 2016; Baranov et al., 2019). This demonstrates that PtdIns(3,4)P₂ regulates actin dynamics through multiple effectors in at least a subset of phagocytic receptors.

The plasmids used in this study are summarized in Table 1. Where indicated, plasmids were constructed using In-Fusion HD EcoDry Cloning Kits (Cat# 121416; Takara) and were verified by Sanger sequencing prior to transfection into mammalian cells. The murine Lpd knockdown construct used specific oligonucleotides: forward, 5′-TGC​GTC​AAG​TCA​CAG​AAC​CAT​TCA​AGA​GAT​GGT​TCT​GTG​ACT​TGA​CGC​TTT​TTG​GAA​AGA​ATT​CG-3′; reverse, 5′-TCG​ACG​AAT​TCT​TTC​CAA​AAA​GCG​TCA​AGT​CAC​AGA​ACC​ATC​TCT​TGA​ATG​GTT​CTG​TGA​CTT​GAC​GCA-3′.

The murine macrophage RAW 264.7 cell line was obtained from the American Type Culture Collection and cultured in DMEM (Wisent Bioproducts) supplemented with 10% heat-inactivated fetal bovine serum and incubated at 37°C under 5% CO₂.

Quantitative phagocytosis assays have been described previously (Montaño et al., 2018). Briefly, ∼5 × 10⁴ cells were seeded onto 1.8-cm glass coverslips and grown for 18–24 h. Opsonization of SRBC was performed by incubating 100 μl of a 10% SRBC suspension with 3 μl of rabbit IgG fraction against SRBCs at 37°C for 1 h. SRBCs were then labeled with a fluorescent secondary antibody against rabbit. 10 μl of labeled and opsonized SRBCs was added to individual coverslips containing RAW cells within 12-well plates or directly into imaging chambers for live-cell imaging. Synchronization of phagocytosis was achieved by centrifugation (300× g for 10 s) of the phagocytic targets onto the cell-containing coverslips. Phagocytosis was terminated by replacing the medium with ice-cold PBS. To obtain the differential inside-outside staining used throughout this study, cells were incubated with cold PBS containing a secondary antibody against rabbit to label non-internalized SRBCs, previously internalized labeled-SRBCs are not accessible to this secondary antibody staining and therefore are not additionally labeled.

Rabbit polyclonal Lpd antibody (Krause et al., 2004) was used for immunofluorescence at 1:200. Rabbit monoclonal VASP (9A2) antibody (#3132; Cell Signaling Technology) was used for immunofluorescence at 1:200. SRBCs 10% suspension were purchased from MP Biomedicals. Anti-sheep red blood cell antibodies were purchased from Cedarlane Laboratories and supplied by MP Biomedicals (Cat #0855806). Fluorescent secondary antibodies against mouse (#711-165-150) and rabbit (#711-605-152, 711-545-152, and 711-156-152) were purchased from Jackson ImmunoResearch Labs. Paraformaldehyde (16% wt/vol) was purchased from Electron Microscopy Sciences. Additional reagents: Wortmannin (681675; Sigma-Aldrich), PI-103 (528100; Sigma-Aldrich), GDC-0941 (509226; Millipore Sigma), GSK-A1 (kindly provided by Dr. Tamas Balla, National Institutes of Health, Bethesda, MD [Bojjireddy et al., 2014]), AS1949490, and K118 (Echelon Biosciences), fluorescent phalloidin (Molecular Probes), PBS, and HBSS (Wisent Bioproducts).

RAW 264.7 cells were plated in 18-mm-round coverslips ∼36–40 h before experiments. Subsequently, cells were transfected using Fugene HD (Promega) according to the manufacturer’s protocol. Briefly, 1 μg of plasmid DNA was mixed with 3 μl of Fugene HD in 100 μl of serum-free DMEM. The mix was incubated for 15 min at room temperature, and then distributed into two wells of a 12-well plate containing cells seeded onto glass coverslips. Experiments were performed ∼16–18 h after transfection.

Transgenic, doxycycline-inducible RAW 264.7 stable lines were generated by the Sleeping Beauty transposon system (Kowarz et al., 2015). The open reading frames of TagBFP2-INPP4B(WT)-CAAX or TagBFP2-INPP4B(C842A)-CAAX were inserted into the pSBTet-pur backbone. pSBTet-pur plasmids were co-transfected in a 10:1 ratio to pCMV(CAT)-T7-SB100 in 6-well plates using FuGENE HD as described above. Cell-culture medium was replaced 18 h later, and to select for genomic integration, RAW 264.7 cells were incubated with 2.5 μg/ml puromycin (P8833; Sigma-Aldrich). Puromycin selection was maintained for 5 d prior to expanding the polyclonal population into culture flasks for expansion and validation. For induction of TagBFP2-fusions, 1 μg/ml doxycycline (Hyclate, D9891; Sigma-Aldrich) was added to the culture medium for 48 h. Generally, ∼50% of the polyclonal RAW 264.7 cell population exhibited robust expression of the transgenes.

Paraformaldehyde (4% wt/vol)-fixed cells were permeabilized in 0.1% (vol/vol) Triton X-100 in PBS for 7 min, blocked in 2% (wt/vol) BSA in PBS for 1 h at room temperature or overnight at 4°C. Then, coverslips were incubated for 1 h with primary antibodies (diluted in 2% BSA), washed multiple times in PBS, and subsequently incubated with secondary antibodies (diluted in PBS) for 1 h.

Confocal fluorescence microscopy was performed using spinning-disk confocal microscopes (Quorum Technologies). Our systems use an Axiovert 200M microscope (Carl Zeiss) equipped with a 63× oil immersion objective (NA 1.4) and a ×1.5 magnifying lens. These microscopes carry a motorized XY stage (Applied Scientific Instrumentation), a piezo Z-focus drive and diode-pumped solid-state lasers emitting at 440, 491, 561, 638, and 655 nm (Spectral Applied Research). Images were recorded with back-thinned EM-CCD camera (Hamamatsu ImageEM C9100-13). Acquisition settings and capture are controlled by Volocity software v6.2.1 (PerkinElmer). Selection of regions of interest, fluorescence intensity measurements, and brightness–contrast corrections was performed with Volocity software or with FIJI (National Institutes of Health). Brightness and contrast parameters were adjusted across entire images without altering the linearity of mapped pixel values. For live-cell imaging, cells grown on glass coverslips were mounted in a Chamlide magnetic chamber (Live Cell Instruments), filled with 1 ml of HBSS, and kept at 37°C with a heating stage.

Statistical analyses were performed using GraphPad PRISM. Statistical details for each experiment are also stated in the figure legends. Statistical significance (****P < 0.0001, ***P < 0.001, **P < 0.01, *P < 0.05) is denoted in figures when applicable. All microscopy-based experiments were performed independently at least three times.

Fig. S1 shows that PtdIns(3,4)P₂ and PtdIns(3,4,5)P₃ are differentially distributed within the phagocytic cup, and SHIP1 and SHIP2 are dispensable for PtdIns(3,4)P₂ production at this site. Fig. S2 shows that PtdIns(3,4)P₂ determines the cellular localization of the PH domain of Lpd and the particle association index in PtdIns(3,4)P₂—depleted cells. Fig. S3 shows the shRNA-mediated silencing of Lpd. Fig. S4 shows that MITO-FP4 targets endogenous VASP to the mitochondria, the distribution of the Lpd^EVmut during phagocytosis and that EVL, Mena, and VASP localize at the phagocytic cup. Fig. S5 shows that Lpd–WRC interactions positively regulate phagocytosis and PtdIns(3,4)P₂ and that Lpd and VASP localize to the phagocytic cup during CR3-mediated phagocytosis, but PtdIns(3,4)P₂ is dispensable for CR3-mediated phagocytosis. Video 1 shows the dynamics of PtdIns(3,4)P₂ and PtdIns(3,4,5)P₃ during phagocytosis. Video 2 shows the actin dynamics during phagocytosis. Video 3 shows the actin dynamics in PtdIns(3,4)P₂-depleted macrophages during phagocytosis.

F. Montaño-Rendón was supported by the Mary H. Beatty Fellowship from the University of Toronto. This work was supported by a Canadian Institutes of Health Research Foundation Grant (FDN-143202) to S. Grinstein and a Canadian Institutes of Health Research Project Grant (PJT1655968) to G.D. Fairn. G.D. Fairn is also supported by a Tier 1 Canada Research Chair in Multiomics of Lipids and Innate Immunity.

The authors declare no competing financial interests.

Author contributions: F. Montaño-Rendón, S. Grinstein, and G.D. Fairn conceived the project. F. Montaño-Rendón, S. Grinstein, and G.D. Fairn designed the experiments and analyzed the data. G.F.W. Walpole, M. Krause, and G.R.V. Hammond made and/or provided critical reagents. F. Montaño-Rendón performed the experimental work and wrote the original draft of the manuscript. All authors reviewed and edited the manuscript.