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Integrin-based diffusion barrier separates membrane domains enabling the formation of microbiostatic frustrated phagosomes

  1. Michelle E Maxson
  2. Xenia Naj
  3. Teresa R O'Meara
  4. Jonathan D Plumb
  5. Leah E Cowen
  6. Sergio Grinstein  Is a corresponding author
  1. Hospital for Sick Children, Canada
  2. University Medical Center Hamburg-Eppendorf, Germany
  3. University of Toronto, Canada
  4. St. Michael’s Hospital, Canada
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Cite as: eLife 2018;7:e34798 doi: 10.7554/eLife.34798

Abstract

Candida albicans hyphae can reach enormous lengths, precluding their internalization by phagocytes. Nevertheless, macrophages engulf a portion of the hypha, generating incompletely sealed tubular phagosomes. These frustrated phagosomes are stabilized by a thick cuff of F-actin that polymerizes in response to non-canonical activation of integrins by fungal glycan. Despite their continuity, the surface and invaginating phagosomal membranes retain a strikingly distinct lipid composition. PtdIns(4,5)P2 is present at the plasmalemma but is not detectable in the phagosomal membrane, while PtdIns(3)P and PtdIns(3,4,5)P3 co-exist in the phagosomes yet are absent from the surface membrane. Moreover, endo-lysosomal proteins are present only in the phagosomal membrane. Fluorescence recovery after photobleaching revealed the presence of a diffusion barrier that maintains the identity of the open tubular phagosome separate from the plasmalemma. Formation of this barrier depends on Syk, Pyk2/Fak and formin-dependent actin assembly. Antimicrobial mechanisms can thereby be deployed, limiting the growth of the hyphae.

https://doi.org/10.7554/eLife.34798.001

eLife digest

Billions of microorganisms live on, and in, the human body. Known as the human microbiome, most of these microscopic hitchhikers are harmless. But, for people with a compromised immune system, common species can sometimes cause disease. For example, the yeast Candida albicans, which colonises between 30 and 70% of the population, is normally harmless, but can switch to a disease-causing version that makes branching structures called hyphae. These hyphae grow fast, piercing and damaging the tissues around them.

Immune cells called macrophages usually engulf invading microbes. These cells recognise sugars on the outside of C. albicans, and respond by wrapping their membranes around the yeast, drawing the microorganism in, and sealing it into closed structures called phagosomes. Then, the macrophages fill the phagosomes with acid, enzymes and destructive chemicals, which breaks the yeast down. Yet, C. albicans hyphae grow larger than macrophages, making them difficult to control.

Maxson et al. have now tracked the immune response revealing how macrophages try to control large hyphae. The immune cells were quick to engulf C. albicans in its normal yeast form, but the response slowed down in the presence of hyphae. Electron microscopy revealed that the large structures were only partly taken in. Rather than form a closed phagosome, the macrophages made a cuff around the middle of the hypha, leaving the rest hanging out.

The process starts with a receptor called CR3, which detects sugars on the outside of the hyphae. CR3 is a type of integrin, a molecule that sends signals from the surface to the inside of the immune cell. A network of filaments called actin assemble around the hypha, squeezing the membrane tight. The macrophage then deploys free radicals and other damaging chemicals inside the closed space. The seal is not perfect, and some molecules do leak out, but the effect slows the growth of the yeast. When a phagosome cannot engulf an invading microbe, a state that is referred to as being “frustrated”, the leaking of damaging chemicals can harm healthy tissues and lead to inflammation and disease.

These findings reveal that macrophages do at least try to form a complete seal before releasing their cocktail of chemicals. Understanding how the immune system handles this situation could open the way for new treatments for C. albicans infections, and possibly similar diseases related to “frustrated engulfment” (such as asbestos exposure, where asbestos fibers are also too large to engulf). However, one next step will be to find out what happens to partly engulfed hyphae, and how this differs from the fate of fully engulfed yeast.

https://doi.org/10.7554/eLife.34798.002

Introduction

Candida albicans is a commensal fungus that colonizes the epithelial surfaces of 30–70% of healthy individuals (Perlroth et al., 2007). However, in immune-compromised individuals, C. albicans can cause invasive, life-threatening disease. The mortality rate for infected patients is 46–75%, with candidiasis classified as the fourth most common nosocomial bloodstream infection (Brown et al., 2012). Invasive candidiasis is correlated with a switch of C. albicans from its yeast form to a hyphal form, a shift that can be induced in vitro by nutrient deprivation among other cues (reviewed in Sudbery, 2011). In vivo, C. albicans hyphae are capable of invading epithelium and endothelium; in addition C. albicans is capable of forming recalcitrant biofilms and inducing inflammation (Sudbery, 2011). These conditions activate host defense mechanisms for the control and clearance of C. albicans, mounted predominantly by phagocytic cells of the innate immune system.

Phagocytes can effectively sense, internalize and kill invasive C. albicans. Accordingly, impairment of the phagocytic response, e.g. by elimination of macrophages and neutrophils, is associated with disseminated candidiasis (reviewed in Netea et al., 2015). Phagocytic cells possess receptors that bind the C. albicans cell wall and trigger uptake of the fungus into a phagosome. The C. albicans cell wall is composed mostly (80–90%) of polysaccharides, containing ≈ 60% β-(1,3) and -(1,6) glucans, and ≈ 40% O- and N-linked mannans (Ruiz-Herrera et al., 2006). As such, the main non-opsonic phagocytic receptors for C. albicans are the C-type lectin family of receptors, including Dectin1, the mannose receptor, and DC-SIGN (reviewed in Hardison and Brown, 2012). The phagosome typically matures rapidly after closure, evolving into an acidic, degradative and microbicidal compartment. Acquisition of antimicrobial properties by this compartment depends on its ability to accumulate and retain toxic compounds, including reactive oxygen species (ROS). Superoxide produced by the NADPH oxidase undergoes dismutation into hydrogen peroxide in the acidic luminal environment generated by the V-ATPase, which additionally favors the catalytic activity of various hydrolases. Transporters such as NRAMP-1, that antagonize microbial growth by depleting the phagosome of nutrients, also depend on phagosomal H+ for the extrusion of metal ions.

Unlike most other microbes, C. albicans presents a distinct problem for phagocytes. The hyphal form of C. albicans can grow at a rate of 18.8 μm hr−1 (GOW and Gooday, 1982), quickly exceeding the size of the phagocytes themselves. The challenge is greatest for macrophages, which migrate to infection sites later than the polymorphonuclear cells, and thus encounter growing hyphae (reviewed in Erwig and Gow, 2016). Despite being remarkably plastic, macrophages have difficulty engulfing the much larger C. albicans hyphae, an impasse that no doubt contributes to the pathogenesis of candidiasis.

The aim of the current study was to examine the dynamic and complex process of C. albicans phagocytosis by macrophages. We found that attempts to engulf large hyphae result in the formation of incomplete (frustrated) phagosomes, which nevertheless segregate a section of the hypha, preferentially exposing it to microbiostatic products. The mechanism and fungal components underlying the formation of the diffusion barrier established by the phagocyte when generating the frustrated phagosome was analyzed using a combination of imaging, pharmacological and genetic approaches.

Results

Phagocytosis of C. albicans hyphae

To optimize the phagocytosis of C. albicans, which has a cell wall rich in β-glucans (Gow et al., 2011), we used RAW 264.7 macrophages stably expressing the Dectin1 receptor (RAW-Dectin1; Esteban et al., 2011). Yeast or hyphal forms of C. albicans expressing BFP (Candida-BFP; Strijbis et al., 2013) were used as targets to facilitate their visualization. Under the conditions used to generate them, C. albicans hyphae were considerably longer (>15 μm) than the macrophages (8–10 μm in diameter). After 1 hr of co-incubation with the macrophages the yeast form was fully engulfed (Figure 1A), while a significant number of hyphal C. albicans were only partially internalized (68.5% ± 4.5, while 31.5% ± 4.6 were fully internalized; 1019 events from 12 independent experiments), which was verified using fluorescent concanavalin A to label exposed hyphae (Figure 1B). This was similar to the frustrated engulfment of >20 μm C. albicans hyphae reported earlier (Lewis et al., 2012). Transmission electron microscopy confirmed that most hyphae were only partially internalized (Figure 1C) and, in addition, revealed the existence around the neck of the frustrated phagosome of a low-contrast structure seemingly devoid of membrane-bound organelles (Figure 1C, inset), previously interpreted by Strijbis et al., 2013 as accumulated actin. Indeed, this region corresponded to an actin-rich cuff-like structure (Figure 1D); F-actin was so highly accumulated at the cuff that the remainder of the cellular actin could only be visualized when images were overexposed (Figure 1D, inset). Note that the remainder (i.e. the base) of the frustrated phagocytic cup was virtually devoid of F-actin. 3D visualization verified the continuous accumulation of F-actin around the neck of the tubular phagosomes lining individual hyphae and its sharp delineation of the intracellular and extracellular portions of the fungus (Figure 1E,F,G,H and Video 1). This actin cuff was observed for RAW-Dectin1 cells engulfing C. albicans hyphae up to 100 μm in size (data not shown), and occurred in 96.3% ± 1.9 of the partially internalized hyphae (674 events analyzed in 12 independent experiments). These data support published accounts of actin cuff-like structures seen during the phagocytosis of various filamentous targets (García-Rodas et al., 2011; Gerisch et al., 2009; Heinsbroek et al., 2009; Prashar et al., 2013; Strijbis et al., 2013). The occurrence of frustrated phagocytosis with formation of a pronounced actin cuff was not unique to the RAW-Dectin1 cell line; similar features were seen when murine or human primary macrophages were confronted with C. albicans hyphae (Figure 1—figure supplement 1A and B, respectively). The actin cuff was remarkably stable, lasting for at least 90 min without contracting (Figure 1I). Nevertheless, the actin composing these structures undergoes measurable turnover (treadmilling), since the cuffs underwent gradual disassembly when the cells were treated with latrunculin A, which scavenges actin monomers (last two panels, Figure 1I). These long-lasting yet dynamic cuffs identify the frustrated phagocytic cups generated by macrophages attempting to eliminate C. albicans hyphae.

Figure 1 with 1 supplement see all
Partial phagocytosis of C. albicans hyphae is associated with formation of an actin cuff.

Phagocytosis of C. albicans yeast (A) or hypha (B) by RAW-Dectin1 cells. After incubation with Candida-BFP, RAW-Dectin1 cells were fixed and extracellular C. albicans stained using Alexa594-conjugated concanavalin A (red). The fluorescence of the BFP is shown in white here and elsewhere to reveal the location of the Candida-BFP. Inset in (B): overexposure of the concanavalin A signal to show less intense, staining of the macrophage membrane (as in A). Scale bars: 5 μm and 10 μm, respectively. (C) Transmission electron micrograph of a RAW-Dectin1 cell with a partially internalized C. albicans hypha. Area of organelle clearance corresponding to the cuff structure is indicated in inset by arrows. Scale bar: 5 μm. (D) F-actin enrichment at the neck of partial phagosome. RAW-Dectin1 cells were allowed to internalize C. albicans hyphae, fixed and stained with fluorescent phalloidin (green). Actin cuff indicated with a bracket. Inset: overexposure to show the less intense cellular actin. Scale bar: 10 μm. (E–H) 3D rendering of a C. albicans hypha partially internalized by a RAW-Dectin1 cell. After incubation with Candida-BFP (white), RAW-Dectin1 cells were fixed and extracellular portions of the hyphae stained using Alexa647-conjugated concanavalin A (blue). Actin was stained with fluorescent phalloidin (red). Scale bar: 5 μm. (F) 3D rendering sliced near the middle of the tubular phagosome, (G) same as E showing only the hypha (white) and actin (red), and (H) same as E showing only the hypha (white) and concanavalin A (blue). (I) Stability of the actin cuff assessed by live cell imaging. RAW-Dectin1 cells expressing LifeAct-GFP were allowed to internalize C. albicans hyphae and imaged at defined intervals. Where indicated (105 min) 1 µM latrunculin A was added and recording continued. Actin cuff location indicated by bracket. Scale bar: 10 μm. Images are representative of ≥30 fields from ≥3 separate experiments of each type. In this and subsequent figures the outline of the phagocyte (when not readily apparent) is indicated by a dotted grey line.

https://doi.org/10.7554/eLife.34798.003
Video 1
3D rendering of a RAW-Dectin1 cell with a partially internalized Candida-BFP hypha (white), showing the demarcation of concanavalinA (blue) by the actin cuff (red).

See Figure 1 for additional information.

https://doi.org/10.7554/eLife.34798.005

Dectin1 and cadherins do not localize to the actin cuff

We proceeded to probe the receptors whose signaling could potentiate the formation of the actin cuff. Because C-type lectin signaling contributes importantly to C. albicans phagocytosis (de Turris et al., 2015; Tafesse et al., 2015; Xu et al., 2009), we analyzed whether Dectin1 accumulated in the membrane at sites where cuffs were evident. Remarkably, while Dectin1 was clearly concentrated in patches elsewhere along the frustrated phagocytic cup, it was poorly detectable by immunostaining near the actin cuff (ratio cuff: cup 0.60 ± 0.04; n = 30 p<0.0001; Figure 2A and inset). The failure to detect accumulation of Dectin1 at these sites was not attributable to masking of the exofacial epitope, possibly resulting from tight apposition to the hyphae, because similar results were obtained when the receptors were tagged with emerald fluorescent protein and visualized directly in live cells (ratio cuff: cup 0.56 ± 0.04; n = 15, p<0.0001; Figure 2B and inset).

Figure 2 with 1 supplement see all
Assessing the contribution of Dectin1 and cadherin/catenin to the formation of the actin cuff.

After incubation with Candida-BFP hyphae, RAW-Dectin1 cells were fixed and monolayers stained and visualized as follows. (A) The distribution of Dectin1-HA was detected by immunostaining (red). Actin was stained using fluorescent phalloidin (green); concanavalin A (blue). Inset: actin cuff shows little colocalization (yellow) with Dectin1-HA. (B) Visualization of Emerald-Dectin1 (green). Actin was stained using fluorescent phalloidin (blue); concanavalin A (red). Inset: poor colocalization of actin cuff with Emerald-Dectin1, in yellow. (C) The expression of E-cadherin (top panel) and β-catenin (bottom panel) was assessed by immunoblotting in human macrophages, A431 and RAW-Dectin1 cells; GAPDH was used as loading control. Visualization of: (D) E-cadherin-GFP or (E) β-catenin-GFP transiently transfected into RAW-Dectin1 cells. For both (D) and (E), after phagocytosis and fixation, extracellular C. albicans was stained using Alexa594-conjugated concanavalin A (red), and actin stained using fluorescent phalloidin (blue). Scale bars: 5 μm. (F) RAW-Dectin1 cells were allowed to internalize C. albicans-hyphae in the presence or absence of 4 mM EDTA. Following phagocytosis, extracellular C. albicans was stained using concanavalin A, and actin stained with phalloidin. The number of C. albicans hyphae that were fully internalized or partially internalized with actin cuffs per 37.5x field was counted by confocal microscopy, and the average number per field calculated. Average number of C. albicans per field was 12.7 ± 1.0. For each condition, three independent experiments were quantified, with ≥15 fields counted per replicate. p value was calculated using the unpaired, 2-tailed students t-test. Data are means ±SEM.

https://doi.org/10.7554/eLife.34798.006

In epithelial and endothelial cells, host E- or N-cadherin, respectively, have been reported to contribute to C. albicans internalization (Moreno-Ruiz et al., 2009). This process involved the recruitment of α- and β-catenins and activation of the Arp2/3 pathway for actin nucleation. In agreement with these reports, we observed E-cadherin and β-catenin accumulation at sites of where C. albicans hyphae were being internalized by epithelial A431 cells, with particular accumulation at sites where actin polymerized (Figure 2—figure supplement 1). We considered whether a similar mechanism was responsible for the formation of actin cuffs by macrophages. However, neither E-cadherin nor β-catenin was detectable in RAW-Dectin1 cells or in primary human macrophages by immunoblotting (Figure 2C) or by immunofluorescence (not illustrated). Under comparable conditions, robust signals were obtained when probing A431 cells (Figure 2C). When expressed heterologously in macrophages E-cadherin-GFP was found to line the surface membrane, but was absent from the phagocytic cup (Figure 2D), while β-catenin-GFP was largely soluble and did not accumulate at the cuff (Figure 2E). Thus, E-cadherin and β-catenin are unlikely to mediate phagocytosis of C. albicans in macrophages. Nevertheless, low levels of expression of these proteins (below the level of detection of our assays) or other cadherins may have mediated the internalization. This possibility was assessed by treating the cells with EDTA, which chelates the Ca2+ known to be required for ligand binding by cadherins (reviewed in Brasch et al., 2012). As shown in Figure 2F, omission of Ca2+ had no effect on actin cuff formation in C. albicans-infected RAW-Dectin1 cells.

Integrin αM β2 is involved in the formation of the actin cuff

Actin can also be tethered to the phagocytic cup via integrins (Freeman et al., 2016). Integrins can be directly or indirectly involved in the phagocytosis of opsonized particles, apoptotic cells and a variety of other targets (reviewed in Dupuy and Caron, 2008) and link with actin filaments via talin and vinculin (reviewed in Shattil et al., 2010). However, canonical integrin activation and ligand binding require divalent cations (reviewed in Leitinger et al., 2000), and would therefore be inhibited by their chelation with EDTA. Moreover, actin cuffs formed normally in CALDAG-GEF1−/− macrophages (Figure 3—figure supplement 1), consistent with the notion that cuff formation was independent of canonical activation of integrins, which involves Rap1 (reviewed in Hogg et al., 2011). There is, however, one atypical instance where integrin activation can occur in the absence of divalent cations. The α chain of the integrin complement receptor 3 (CR3, also referred to as Mac1), is unique in that it contains a lectin-like domain (LLD) that binds carbohydrates in a divalent cation-independent manner (Thornton et al., 1996). The LLD is separate from the I-domain –the conventional ligand-binding domain of integrins (reviewed in Ross, 2002)– and, interestingly, binds fungal β-glucan (Ross et al., 1985; Vetvicka et al., 1996). We therefore proceeded to test whether CR3, which consists of αM (CD11b) and β2 (CD18) subunits, is present in the region of the actin cuff. As illustrated in Figure 3, both CD11b and CD18 accumulated in the region of the actin cuff in RAW-Dectin1 cells that had partially internalized C. albicans hyphae (CD11b ratio cuff: cup 4.75 ± 0.29; n = 30, p<0.0001; CD18 ratio cuff: cup 4.79 ± 0.28; n = 30, p<0.0001; Figure 3A,B and insets). Moreover, talin, vinculin and paxillin were also localized to the cuff (Figure 3C,D and insets; Figure 3—figure supplement 1E and inset), as was HS1, the homologue of cortactin in leukocytes (Figure 3E and inset). Like cortactin, HS1 is thought to regulate actin nucleation and branching (Daly, 2004).

Figure 3 with 1 supplement see all
Engagement of integrin αMβ2 (CD11b/CD18) is necessary for formation of the actin cuff.

After incubation with Candida-BFP hyphae, RAW-Dectin1 cells were fixed and extracellular C. albicans stained using Alexa594-conjugated concanavalin A (red). For panels (A–G) F-actin was stained using fluorescent phalloidin (blue), and actin cuff location indicated with a dashed box or bracket. (A) Anti-CD11b immunostaining (green). Inset: Colocalization of actin cuff with CD11b, in yellow. Scale bar: 5 μm. (B) Anti-CD18 immunostaining (green). Inset: Colocalization of actin cuff with CD18, in yellow. Scale bar: 5 μm. (C) Visualization of transfected Talin-GFP. Inset: Colocalization of actin cuff with talin, in yellow. Scale bar: 10 μm. (D) Immunostaining of endogenous vinculin (green). Inset: Colocalization of actin cuff with vinculin, overlaid in yellow. Scale bar: 10 μm. (E) Immunostaining of endogenous HS1 (green). Scale bar: 10 μm. (F–H) Internalization of Candida-BFP hyphae was allowed to proceed in the presence of the CD11b blocking antibody M1/70 or an isotype-matched (rat IgG2b) control antibody. Following phagocytosis, extracellular C. albicans was stained using Alexa594-conjugated concanavalin A (red), and actin stained using fluorescent phalloidin (blue). Immunostaining (green) for rat IgG2b isotype control (F, left panel) or M1/70 (G, left panel). Scale bars: 5 μm. Images shown are representative of at least 3 experiments of each kind. (H) The number of C. albicans hyphae that were fully internalized or partially internalized with actin cuffs per 37.5x field was counted by confocal microscopy. Average number of C. albicans per field was 11.7 ± 0.5. For each condition, four independent experiments were quantified, with ≥15 fields counted per replicate. p value was calculated using the unpaired, 2-tailed students t-test. Data are means ±SEM.

https://doi.org/10.7554/eLife.34798.009

The preceding findings support a model whereby ligation of β-glucan by the LLD causes outside-in activation of CR3 directly (O'Brien et al., 2012; Vetvicka et al., 1996), or in conjunction with Dectin1 signaling (Huang et al., 2015; Li et al., 2011), resulting in Arp2/3-dependent actin nucleation. This model was tested using the M1/70 antibody, which binds to CD11b between its β-propeller and thigh domains (residues 614–682; Osicka et al., 2015) and effectively blocks the binding of CR3 to β-glucan (Xia et al., 1999). Cells pretreated with M1/70 failed to show accumulation of CR3 around partially internalized C. albicans hyphae, and their ability to form actin cuffs was markedly impaired (Figure 3H); actin cuffs were much less prominent or missing altogether when CR3 was blocked (Figure 3F versus G). The number of fully internalized C. albicans did not differ between conditions (Figure 3H). We concluded that binding of the CR3 integrin to C. albicans was critical for the establishment of long-enduring actin cuffs observed during frustrated phagocytosis of the hyphae.

Role of receptor cooperativity in actin cuff formation

Dectin1 and CR3 both bind β-glucans (Brown and Gordon, 2001; Brown et al., 2002; Ross et al., 1985; Vetvicka et al., 1996), and have been reported to cooperate during phagocyte responses to fungal pathogens (Huang et al., 2015; Li et al., 2011). Dectin1 has also been reported to cooperate with TLR2, TLR4 (Ferwerda et al., 2008; Netea et al., 2006; Netea et al., 2002) and mannose receptors (Astarie-Dequeker et al., 1999; Bain et al., 2014; Lewis et al., 2012; McKenzie et al., 2010; Netea et al., 2006) in the recognition of C. albicans. We therefore sought to clarify the receptors and ligands involved in actin cuff formation.

Untransfected RAW 264.7 cells express negligible levels of Dectin1 (Brown et al., 2003; Esteban et al., 2011; Taylor et al., 2004), providing a means to assess the contribution of this receptor to actin cuff formation. As shown in Figure 4A, RAW 264.7 cells rarely formed actin cuffs compared to RAW-Dectin1 cells, suggesting that initial engagement of the hyphae by Dectin1 was essential. The requirement for Dectin1 in C. albicans phagocytosis (Marakalala et al., 2013; Taylor et al., 2007) could be bypassed when the hyphae were serum-opsonized, enabling opsonin receptors to establish the initial contact with the fungus (Figure 4A). Thus, while not accumulating in the region of the cuff, Dectin1 binding to the hyphae (which is evident by its accumulation in the frustrated phagocytic cup; Figure 2A and B) is required for the subsequent activation of F-actin polymerization by CR3.

Figure 4 with 1 supplement see all
Assessing the contribution of C. albicans cell wall components to actin cuff formation.

(A) RAW or RAW-Dectin1 cells were incubated with Candida-BFP hyphae that had been either untreated or serum-opsonized. Following phagocytosis, extracellular C. albicans was stained using concanavalin A, and actin stained with phalloidin. The number of C. albicans hyphae that were fully internalized or partially internalized with actin cuffs per 37.5x field was counted by confocal microscopy. Average number of C. albicans per field was 15.7 ± 1.3. For each condition, three independent experiments were quantified, with ≥4 fields counted per replicate. p value was calculated using the unpaired, 2-tailed students t-test. Data are means ±SEM. (B) RAW-Dectin1 cells were allowed to internalize Candida-BFP hyphae in the presence or absence of laminarin or mannan. For laminarin, RAW-Dectin1 cells were also allowed to adhere C. albicans 15 min prior to the addition of laminarin, as indicated. Other details as in A. Average number of C. albicans per field was 12.9 ± 0.7. (C–D) Evaluation of C. albicans GRACE strain cell wall mutants for actin cuff formation. GRACE strains were induced to form hyphae in the absence or presence of doxycycline (DOX) to repress target gene expression, and incubated with RAW-Dectin1 cells. Following phagocytosis, monolayers were fixed and C. albicans stained with 10 μg mL−1 calcofluor white (white), extracellular C. albicans stained using concanavalin A (red), and actin stained with phalloidin (green). Image in C is representative of ≥30 fields from ≥3 separate experiments of each type. Scale bar: 5 μm. (D) The number of C. albicans hyphae that were fully internalized or partially internalized with actin cuffs per 37.5x field was counted by confocal microscopy, and the average number per field calculated. Average number of C. albicans per field was 20.6 ± 0.6. For each condition, three independent experiments were quantified, with ≥4 fields counted per replicate. p value was calculated using the unpaired, 2-tailed students t-test. Data are means ±SEM. (E) Role of C. albicans β-(1,3)-glucan in actin cuff formation. The GRACE wild-type strain was incubated and induced to form hyphae in the presence or absence of 5 ng mL−1 caspofungin and incubated with RAW-Dectin1 cells for phagocytosis. Following phagocytosis, cells were fixed and C. albicans stained with 10 μg mL−1 calcofluor white (white), extracellular C. albicans stained using fluorescent concanavalin A (red), and actin stained with fluorescent phalloidin (green). Image is representative of ≥30 fields from ≥3 separate experiments. Scale bar: 5 μm. (F) The effect of β-(1,3)-glucan synthase inhibition on actin cuff formation. Hyphae were prepared as in (E), with varying concentrations of caspofungin, as indicated. Phagocytosis, fixation and staining as in (E). Other details as in (A). Average number of C. albicans per field was 19.7 ± 0.8. (G) Actin cuffs are observed during phagocytosis of A. fumigatus hyphae. After incubation with hyphae, monolayers were fixed and A. fumigatus stained with 10 μg mL−1 calcofluor white (white). Actin stained with phalloidin (green). Image representative of ≥30 fields from ≥2 separate experiments. Scale bar: 5 μm.

https://doi.org/10.7554/eLife.34798.012

We also studied cooperativity by using soluble ligands to competitively block defined receptors, and scoring the frequency of actin cuff formation (Figure 4B). Soluble mannan, a ligand for mannose receptor, had no effect on actin cuff formation by RAW-Dectin1 cells. Accordingly, we did not find mannose receptors in the membrane lining the actin cuff (data not shown). Laminarin, a soluble β-glucan ligand for Dectin1 (Brown and Gordon, 2001; Brown et al., 2002) impaired phagocytosis and actin cuff formation when present prior to and during phagocytosis, but not if added after the hyphae had adhered to the RAW-Dectin1 cells (Figure 4B). These findings support the notion that Dectin1, but not mannan receptors, cooperate with CR3 to generate the actin cuffs.

Fungal cell wall components that contribute to actin cuff formation

C. albicans cell wall components include β-(1,3)-glucans, β-(1,6) glucans, O- and N-linked mannans and chitin (Netea et al., 2008; Ruiz-Herrera et al., 2006). These can contribute to the recognition of C. albicans by phagocytes (reviewed in Netea et al., 2008), and potentially also to actin cuff formation. To clarify the contribution of individual wall components we used gene replacement and conditional expression (GRACE) strains (Roemer et al., 2003) with specific depletion targeting chitin, mannan, and β(1,6)-glucan biosynthetic pathways upon incubation with doxycycline (Table 1; O'Meara et al., 2015). Repression of pathways involved in chitin, mannan and β(1,6)-glucan synthesis using doxycycline did not affect actin cuff formation (Figure 4C,D and data not shown), implying that these components are dispensable. We next assessed the role of β(1,3)-glucan through pharmacological inhibition of Fks1 with caspofungin (Douglas et al., 1997), as genetic depletion of Fks1 results in defects in hyphae formation (Ben-Ami et al., 2011). Remarkably, the ability to form actin cuffs was greatly reduced in C. albicans grown and allowed to form hyphae in the presence of caspofungin (Figure 4E and F). The inhibitory effect of caspofungin on actin cuff formation was dose-dependent (Figure 4F), reaching ≈ 80% at 5 ng mL−1 caspofungin, a dose that reduced the β(1,3)-glucan content of the wall by 55.3%, as assessed by aniline blue staining. Actin cuff formation around caspofungin-treated hyphae could not be rescued by serum opsonization (Figure 4—figure supplement 1), suggesting that β(1,3)-glucan is the ligand that promotes actin cuff assembly via CR3. Interestingly, Aspergillus fumigatus hyphae (routinely exceeding 80 µm in length) were also able to illicit actin cuff formation by RAW-Dectin1 cells (Figure 4G). A. fumigatus hyphae, while displaying some unique cell wall components compared to C. albicans hyphae, also have cell wall-associated β(1,3)-glucan (Erwig and Gow, 2016). We concluded that ligation of fungal β(1,3)-glucan by CR3 is required for actin cuff formation during frustrated phagocytosis of long hyphae.

Table 1
C. albicans strains used in this study.
https://doi.org/10.7554/eLife.34798.018
StrainParentGenotypeGene functionReference
Candida-BFPSC5314Peno1-TagBFP-NATRN/A(Strijbis et al., 2013)
CaSS1CAI4ura3::imm434 / ura3::imm434 his3::hisG / his3::hisG leu2::tetR-GAL4AD-URA / LEU2N/A(Roemer et al., 2003)
CHT1CaSS1tetO-CHT1 / cht1∆chitinase(O'Meara et al., 2015)
CDA2CaSS1tetO-CDA2 / cda2∆chitin deacetylase(O'Meara et al., 2015)
MNT2CaSS1tetO-MNT2 / mnt2∆α-(1,2)-mannosyl transferas(O'Meara et al., 2015)
VRG4CaSS1tetO-VRG4 / vrg4∆GDP-mannose transporte(O'Meara et al., 2015)
KRE1CaSS1tetO-KRE1 / kre1∆cell wall glycoprotein, β-(1,6)-glucan synthesis(O'Meara et al., 2015)
KRE6CaSS1tetO-KRE6 / kre6∆β-(1,6)-glucan synthase subunit(O'Meara et al., 2015)
KRE62CaSS1tetO-KRE62 / kre62∆β-(1,6)-glucan synthase subunit(O'Meara et al., 2015)
KEG1CaSS1tetO-KEG1 / keg1∆integral membrane ER protein, β-(1,6)-glucan synthesis(O'Meara et al., 2015)
ENG1CaSS1tetO-ENG1 / eng1∆endo-(1,3)-β-glucanase(O'Meara et al., 2015)
ACF2CaSS1tetO-ACF2 / acf2∆endo-(1,3)-β-glucanase(O'Meara et al., 2015)
BGL22CaSS1tetO-BGL22 / bgl22∆putative β-glucanase(O'Meara et al., 2015)

Signals driving actin cuff formation

Despite the paucity of Dectin1 and mannose receptors (Figure 2A and B), phosphotyrosine was markedly concentrated at the cuff (Figure 5A), possibly as a consequence of CR3 activation. While there is disagreement over the requirement of Src-family kinases (SFKs) for the interaction of phagocytes with fungal targets (Elsori et al., 2011; Herre et al., 2004; Le Cabec et al., 2002; Mansour et al., 2013; Underhill et al., 2005), there is evidence that Syk, as well as Pyk2 and Fak, two related tyrosine kinases, participate in CR3-mediated phagocytosis (Li et al., 2006; Paone et al., 2016; Zhao et al., 2016). The contribution of individual kinases to the tyrosine phosphorylation was explored next.

Figure 5 with 1 supplement see all
Signaling associated with actin polymerization at the phagocytic cup formed around C. albicans hyphae.

After incubation with Candida-BFP hyphae, RAW-Dectin1 cells were fixed and extracellular C. albicans stained using fluorescent concanavalin A. (A) Phosphotyrosine (pTyrosine) was detected by immunostaining (green). F-actin was visualized using TdTom-F-Tractin (red); concanavalin A (blue). Inset: Colocalization of actin cuff with pTyrosine, in yellow. Image is representative of ≥30 fields from ≥3 separate experiments. (B) Phospho-SFK (Y418) was detected by immunostaining (green); concanavalin A (red). Inset: Colocalization of actin cuff with pSFK, in yellow. (C) Phospho-PYK2 (Y402) was detected by immunostaining (green); concanavalin A (red). Inset: Colocalization of actin cuff with pPYK2, in yellow. (D) Phospho-FAK (Y397) was detected by immunostaining (green); concanavalin A (red). Inset: Colocalization of actin cuff with pFAK, in yellow. Images in B, C and D are representative of ≥30 fields from ≥2 separate experiments of each type. (E) Effect of tyrosine kinase inhibitors on actin cuff formation. RAW-Dectin1 cells were allowed to adhere Candida-BFP hyphae for 15 min and then incubated 45 min in the presence of vehicle, PP2, piceatannol or PF573228. Following phagocytosis, extracellular C. albicans was stained using concanavalin A, and actin stained with phalloidin. The number of C. albicans hyphae that were fully internalized or partially internalized with actin cuffs per 94.5x field was counted by confocal microscopy. Average number of C. albicans per field was 3.4 ± 0.6. For each condition, three independent experiments were quantified, with ≥4 fields counted per replicate. p value was calculated using unpaired, 2-tailed students t-test. Data are means ±SEM. (F) Active Rac/Cdc42 were visualized using PAK(PBD)-YFP as a probe (green). Actin was stained using fluorescent phalloidin (blue); concanavalin A (red). Inset: Colocalization of actin cuff with PAK(PBD), in yellow. Image is representative of ≥30 fields from ≥3 separate experiments. Scale bars: 10 μm. (G) Effect of actin assembly inhibitors on actin cuff formation. RAW-Dectin1 cells were allowed to adhere Candida-BFP hyphae for 15 min, then incubated 45 min in the presence of vehicle, CK-666 or SMI-FH2. Following phagocytosis, extracellular C. albicans was stained using concanavalin A, and actin stained with phalloidin. The number of C. albicans hyphae that were fully internalized or partially internalized with actin cuffs per 94.5x field was counted by confocal microscopy. Average number of C. albicans per field as in (E). For each condition, three independent experiments were quantified, with ≥5 fields counted per replicate. p value was calculated using the unpaired, 2-tailed students t-test. Data are means ±SEM.

https://doi.org/10.7554/eLife.34798.019

Phosphorylated SFKs accumulated along the frustrated phagocytic cup (Figure 5A) where Dectin1 was also found (Figure 2A and B), but were not particularly enriched in the region of the actin cuff (ratio cuff: cup 0.98 ± 0.03; n = 17, p=0.61). SFK inhibition by PP2 following adherence of the hyphae to RAW-Dectin1 cells had no effect on actin cuff formation (Figure 5E). In contrast, the phosphorylated (active) forms of Pyk2 and Fak were enriched solely at the actin cuff (pPyk2 ratio cuff: cup 23.69 ± 1.20; n = 46, p<0.0001, pFak ratio cuff: cup 22.56 ± 1.01; n = 34, p<0.0001; Figure 5C and D). Moreover, inhibition of Pyk2/Fak activity by PF573228 following adherence of the hyphae to the cells abolished actin cuff formation, with no effect on internalization (Figure 5E). Also, as reported by Strijbis et al., 2013, we observed phosphorylation of Syk with accumulation at the actin cuff (ratio cuff: cup 21.55 ± 1.75; n = 22, p<0.0001; Figure 5—figure supplement 1). As expected, inhibition of Syk by piceatannol after C. albicans adherence blocked actin cuff formation (Figure 5E). These data provide evidence that, along with Syk, Pyk2/Fak play a role in the interaction between macrophages and C. albicans, and are important for actin cuff formation during frustrated phagocytosis of hyphae.

Interestingly, the interaction of Pyk2 with β2 integrins activates Vav1 (Gakidis et al., 2004; Kamen et al., 2011), a GEF for Rho-family GTPases that is also essential for the phagocytosis and control of C. albicans by macrophages (Strijbis et al., 2013). Accordingly, Rac1 and/or Cdc42 were seemingly involved in the marked polymerization of actin at the cuff. This was indicated by the recruitment of PAK(PBD), a biosensor of the active (GTP-bound) form of these GTPases (Benard et al., 1999), that accumulated at the cuff to levels ≥4 fold higher than along the cup. F-actin accumulation at the cuff was sensitive to the formin inhibitor SMI-FH2, but not to the Arp2/3 inhibitor CK-666 (Figure 5G). Together, these data suggest that activation of Syk and Pyk2/Fak by CR3 leads to activation of Rho-family GTPases, culminating in formin-mediated actin assembly, a process akin to focal adhesion formation (reviewed in Vicente-Manzanares et al., 2005).

Phospholipid segregation between the plasma membrane and the cuff-delimited phagosomal cup

Phospholipids undergo striking changes during the course of conventional phagocytosis. PtdIns(4,5)P2 that is normally found in the plasma membrane is converted to PtdIns(3,4,5)P3 at sites of receptor engagement, and is subsequently degraded by lipases and phosphatases, becoming undetectable in sealed phagosomes. PtdIns(3,4,5)P3 can be detected for up to a minute following sealing, but then disappears abruptly as PtdIns(3)P appears; the latter is detectable on early phagosomes for about 10–15 min (reviewed in Levin et al., 2015). These drastic switches are thought to reflect and possibly dictate the identity and developmental stage of the maturing phagosome. It has been observed that the frustrated tubular phagosomes of heat-killed filamentous Legionella pneumophila are accompanied by a sharp separation of plasmalemmal and phagosomal phosphoinositide species (Naufer et al., 2018; Prashar et al., 2013). Additionally, atypical phosphoinositide dynamics can occur in sealed phagosomes containing filamenting C. albicans (Heinsbroek et al., 2009) or during CR3-mediated phagocytosis of opsonized targets (Bohdanowicz et al., 2010). Therefore we analyzed the phosphoinositides in frustrated phagosomes of C. albicans hyphae. We used the genetically-encoded fluorescent biosensor PLCδ-PH-GFP to monitor the distribution of PtdIns(4,5)P2. Remarkably, while PtdIns(4,5)P2 was present as expected in the surface membrane facing the extracellular milieu, it was undetectable in the invaginated section that constituted the frustrated phagosome (Figure 6A). In stark contrast, PtdIns(3,4,5)P3 –which was visualized using AKT-PH-GFP– was found solely in the open phagosomal cup (Figure 6B), where it co-existed with PtdIns(3)P, detected using the PX-GFP sensor (Figure 6C). In addition to the localization of PtdIns(3,4,5)P3 in the cup reported in a previous collaborative study (Strijbis et al., 2013), we detected additional enrichment of PtdIns(3,4,5)P3 in the actin cuff region (ratio cuff: cup 1.39 ± 0.10; n = 30, p=0.0006). In contrast, PtdIns(3)P was comparatively excluded from the actin cuff (ratio cuff: cup 0.823 ± 0.05; n = 30, p=0.0025). The segregation of these phosphoinositides persisted for the duration of our observations (up to 90 min after frustrated phagosome formation; not illustrated).

Distribution of phosphoinositides and endo-lysosomal markers.

After incubation with Candida-BFP hyphae, RAW-Dectin1 cells were fixed and extracellular C. albicans stained using Alexa594-conjugated concanavalin A (red). Actin was stained using fluorescent phalloidin (blue), and the location of the actin cuff is indicated by the dashed square. Visualization of: (A) PtdIns(4,5)P2 using PLCδ-PH-GFP; (B) PtdIns(3,4,5)P3/PtdIns(3,4)P2 using AKT-PH-GFP, inset: colocalization of actin cuff with AKT-PH, in yellow; (C) PtdIns(3)P using PX-GFP, inset: colocalization of actin cuff with PX, in yellow; (D) LC3-GFP, inset: colocalization of actin cuff with LC3, in yellow; (E) Rab7-GFP, inset: colocalization of actin cuff with Rab7, in yellow; (F) immunostained LAMP1 (green), inset: colocalization of actin cuff with LAMP1, in yellow. Scale bars: 10 μm. Images are representative of ≥30 fields from ≥3 separate experiments of each type. (G–H) 3D rendering of a RAW-Dectin1 cell with a partially internalized C. albicans hypha. After incubation with Candida-BFP, RAW-Dectin1 cells were fixed and extracellular portions of the hyphae were stained using concanavalin A (blue). (G) C. albicans (white) visualized with actin immunostaining (red). (H) Same 3D rendering as in (G), visualizing LAMP1 immunostaining (green), actin (red) and concanavalin A (blue). Scale bar: 5 μm.

https://doi.org/10.7554/eLife.34798.023

The actin cuff forms a diffusional barrier to the movement of proteins and lipids

The sharp boundary between the PtdIns(4,5)P2-rich surface membrane and the tubular membrane endowed with PtdIns(3,4,5)P3 and PtdIns(3)P coincided with the location of the actin cuff, suggesting that the latter may function as a diffusion barrier. However, the restricted localization of the phosphoinositides may have resulted from the strategic positioning of synthetic (i.e. kinases) and degradative (i.e. phosphatases or lipases) enzymes. To more definitively assess the existence of a diffusion barrier, we analyzed the distribution and dynamics of molecules that do not undergo rapid metabolic transformation, including lipid-anchored and transmembrane proteins, which had been reported to segregate in frustrated phagosomes. As shown in Figure 6D, LC3 –a small protein covalently linked to PtdEth– was found in the frustrated phagosome (Kanayama and Shinohara, 2016; Martinez et al., 2015; Sprenkeler et al., 2016; Tam et al., 2016), yet did not reach the surface membrane. Similarly, both wild-type Rab7 (Figure 6E) and constitutively-active Rab7 (not illustrated) are confined to the frustrated phagosomal tube and partially excluded from the actin cuff (Rab7 ratio cuff: cup 0.68 ± 0.05; n = 30, p<0.0001), as was LAMP1 (ratio cuff: cup 0.59 ± 0.03; n = 30, p<0.0001; Figure 6F), a late-endosomal/lysosomal membrane-spanning glycoprotein. The exclusion from the actin cuff was better appreciated by 3D visualization of LAMP1 (Figure 6G,H and Video 2). Because metabolic conversion to other species could not account for the segregation of the latter probes to the invaginated section of the membrane, we considered it more likely that restricted diffusion accounted for the observations.

Video 2
3D rendering of a RAW-Dectin1 cell with a partially internalized Candida-BFP hypha (white), showing the demarcation of phagosomal LAMP1 (green) and concanavalin A (blue) by the actin cuff (red).

See Figure 6 for additional information.

https://doi.org/10.7554/eLife.34798.024

It was nevertheless possible that molecules like LC3, Rab7 or LAMP were inserted through fusion into the tubular part of the membrane, where they could conceivably remain immobile. To exclude this possibility, we assessed their mobility measuring fluorescence recovery after photobleaching (FRAP). The constitutively-active form of Rab7, Rab7(Q67L), was used for these experiments; because this variant is unable to exchange nucleotides, it does not associate stably with GDI and remains membrane associated (Méresse et al., 1995), eliminating the confounding effects of fluorescence recovery from a cytosolic pool. Rapid recovery was observed following photobleaching of a ≈3 µm spot within the phagosomal cup. In four independent experiments, half-maximal recovery was attained after 3.3 s (Figure 7B). Similar analyses were performed using GFP-tagged LAMP1 (Figure 7A,B), which also recovered within seconds (t1/2 = 7.9 sec). Between 75–80% of the fluorescence was recovered in both instances, implying that the majority of the Rab7(Q67L) and LAMP1 molecules were mobile.

Figure 7 with 1 supplement see all
Formation of the actin cuff is associated with the establishment of a diffusional barrier.

RAW-Dectin1 cells were transfected with the indicated constructs, exposed to Candida-BFP, and used for FRAP determinations. F-actin was visualized with LifeAct-RFP (orange). (A) A region of interest (denoted by dotted circle) of LAMP1-GFP in the frustrated phagosome was selected (left panel, −0.5"), photobleached (middle panel, 0"), and allowed to recover for 60 s (right panel, 60"). Scale bar: 5 μm. Images in A, C and D are representative of ≥30 fields from ≥3 separate experiments of each type. (B) Quantitation of fractional recovery of fluorescence after photobleaching LAMP1 (green) or Rab7(Q67L) (blue). In both cases, data were normalized to fluorescence in unbleached regions of the C. albicans phagosomal cup. For either condition, four biological replicates, with a total of ≥30 cells, were quantified. (C–D) A region of interest in the frustrated phagosome (C) or in the plasma membrane (D) of cells expressing Lyn11-GFP was selected (left panel, −0.5"), photobleached (middle panel, 0"), and allowed to recover for 60 s (right panel, 60"). Scale bars: 5 μm. (E) Quantitation of fractional recovery of fluorescence of photobleached Lyn11-GFP in the plasma membrane (blue) or the frustrated C. albicans phagosomal cup (orange). In both cases, FRAP data was normalized to fluorescence in the plasma membrane. For either condition, three biological replicates, with a total of ≥35 cells, were quantified.

https://doi.org/10.7554/eLife.34798.025

The retention of Rab7(Q67L) and LAMP1 in the cup for many minutes despite their ability to move laterally in the plane of the membrane implies that they are unable to cross the junction with the surface membrane. The existence of a diffusion barrier was confirmed by expressing the N-terminal domain of Lyn (Lyn11) tagged with GFP. This region of the protein becomes myristoylated and palmitoylated, targeting it to the plasma membrane and, to a lesser extent, to early endosomes. Following frustrated phagocytosis of hyphae, Lyn11-GFP is found both at the membrane and in the phagosomal cup, where its density is lower, likely because of dilution caused by insertion of unlabeled endomembranes. We analyzed comparatively small phagosomes to enable photobleaching of Lyn11-GFP in the entire cup (Figure 7C). Strikingly, the fluorescence of the cup failed to recover, despite the persistence of abundant Lyn11-GFP in the adjacent plasmalemma. In three independent experiments only 19% of the original fluorescence reappeared, possibly via fusion with Lyn11-GFP-containing early endosomes. Failure to recover was not attributable to immobility of Lyn11-GFP in the membrane, which displayed very fast and nearly complete recovery following photobleaching (Figure 7D,E). These data confirm that the region of the actin cuff acts as a lateral diffusion barrier, separating the inner leaflet of the plasma membrane from that of the open phagocytic cup.

It is noteworthy that while the barrier curtails the diffusion of lipids and proteins anchored to lipids on the inner leaflet of the membrane, exofacial lipids and lipid-associated proteins readily traverse the junction between the membrane and the tubular phagosome. This was demonstrated by incorporation of rhodamine-labeled PtdEth to the surface membrane following stabilization of the frustrated phagosome. The labeled lipid, which inserts into the outer leaflet of the plasmalemma, reached the entire membrane of the frustrated phagocytic cup within ≈5 min (Figure 7—figure supplement 1A). Similarly, fluorescent cholera toxin B subunit, which binds to exofacial ganglioside GM1, promptly entered the phagocytic cup (Figure 7—figure supplement 1B). Thus, the actin-dependent diffusion barrier selectively restricted the mobility of components of the inner leaflet, including transmembrane proteins, while exofacial lipids remained able to traverse the junction.

Examining the role of CR3 and actin in the maintenance of diffusional barriers

How is the diffusion barrier generated? We speculated that the molecular crowding resulting from tight clustering of integrins and their ancillary proteins could restrict the diffusion of membrane-associated components across the cuff. To test this possibility, we investigated whether sufficient molecular crowding could be generated to exclude other membrane components from regions of integrin clustering. To this end, we used antibody-induced cross-linking, a strategy shown earlier to induce the formation of CR3 patches on the plasma membrane (Fukushima et al., 1996; Pavan et al., 1992; Zhou et al., 1993). Whether exclusion could be induced by molecular crowding was assessed analyzing the distribution of CD2-CD45-GFP (Figure 8B), a transmembrane protein having a short, 7 nm ectodomain (Cordoba et al., 2013). As shown in Figure 8A, prior to cross-linking both CD2-CD45-GFP and CR3 were distributed diffusely throughout the membrane, overlapping extensively at the resolution of the confocal microscope. The CD2-CD45-GFP fluorescence intensity in CR3-positive regions compared to the average CD2-CD45-GFP fluorescence intensity of the entire plasma membrane averaged 0.69 ± 0.01 (585 CR3-positive regions in 20 cells from three different experiments). After antibody treatment, CR3 clustered into large, dense patches. Strikingly, CD2-CD45-GFP was largely (81%) and significantly (p>0.0001) excluded from such patches, where the fluorescence was only 0.13 ± 0.01 of the plasmalemmal average (measured in 472 CR3 patches in 15 cells from three experiments). Importantly, the exclusion was not alleviated by treatment with latrunculin A, the fluorescence of the patches averaging 0.12 ± 0.01 of the plasmalemmal average (measured in 445 patches in 18 cells from three experiments), implying that the actin cytoskeleton is not involved in the domain segregation. CD2-CD45-GFP exclusion did not differ between these two conditions (p=0.66). We concluded that integrins could be sufficiently clustered to exclude other membrane components. By forming a continuous and thick ring around the neck of the frustrated phagosome, the molecular crowding of clustered integrins could generate a diffusional barrier.

Clustering and patching of CR3 forms a diffusional barrier that excludes transmembrane proteins.

(A) Raw-Dectin1 cells transiently expressing CD2-CD45-GFP (left panel, green) and stained for external CD18 (middle panel, red). Right panel shows the colocalization of CD2-CD45 and CD18, in yellow. Panel insets: 2.1x magnification. Scale bar: 5 μm. (B) Diagram illustrating the method used to cluster CR3 in CD2-CD45-GFP-expressing Raw-Dectin1 cells, using M18/2 antibody to CD18, followed by secondary and tertiary antibodies. See Materials and methods for details. (C) Effect of CR3 patching and actin depolymerization. After clustering CR3 as in (B), cells were incubated 10 min in the absence (left) or presence (right) of 1 μM latrunculin A, and visualized for CD2-CD45-GFP (green) and extracellular CD18 (red). Colocalization of CD2-CD45 and CD18 channels shown in yellow. Panel insets: 2x magnification of both channels (left inset) and CD2-CD45 channel only (right inset). Scale bars: 5 μm.

https://doi.org/10.7554/eLife.34798.029

While actin is not essential to constrain the diffusion across patches of antibody-aggregated integrins, it is nevertheless required to maintain the integrins clustered in response to the glucan during frustrated phagocytosis. As such, an intact actin cuff is required to establish and maintain the barrier to phosphoinositides or transmembrane proteins. This was validated in cells that had formed a stable frustrated phagosome around C. albicans hyphae and were then treated with latrunculin A, which was shown earlier (Figure 1I) to cause gradual disassembly of the cuff. PtdIns(4,5)P2 –which in untreated cells is excluded from the phagocytic cup (Figures 6A and 9A)– gained access to the entire cup when actin was disassembled by latrunculin (Figure 9B). The PtdIns(4,5)P2 present in the cup, expressed relative to the plasmalemma, increased 4.88 times after latrunculin treatment (Figure 9C). Conversely, LAMP1 –that is restricted to the cup in untreated cells (Figures 6F and 9D)– was able to reach the surface membrane following treatment with latrunculin (Figure 9E). After latrunculin treatment, the ratio of LAMP1 present in the cup decreased 3.99 times (Figure 9F). Clearly, while clustering of CR3 is sufficient to form a diffusional barrier (Figure 8), the actin cuff formed during phagocytosis of C. albicans hyphae likely contributes to the stability of the barrier between CR3 and C. albicans β(1,3)-glucans, presumably by maintaining integrins in their active conformation (Kaizuka et al., 2007; Lavi et al., 2007; Lavi et al., 2012) during frustrated phagocytosis.

Actin depolymerization abolishes the diffusional barrier around C. albicans hyphae.

RAW-Dectin1 cells were transfected with the indicated constructs, exposed to Candida-BFP hyphae and incubated for 30 min in the presence (B and E) or absence (A and D) of latrunculin A. After treatment, cells were fixed and extracellular C. albicans stained using Alexa594-conjugated concanavalin A (red), and actin stained using fluorescent phalloidin (blue). (A and B) Cells transfected with PLCδ-PH-GFP. (C) Effect of latrunculin A on actin cuff-mediated segregation of PLCδ-PH-GFP to the plasma membrane, quantitated as the ratio of the fluorescence intensity of GFP in the phagocytic cup over the plasma membrane. (D and E) Cells transfected LAMP1-GFP. (F) Effect of latrunculin A on actin cuff-mediated segregation of LAMP1 to the frustrated phagocytic cup, quantitated as the ratio of the fluorescence intensity in the phagocytic cup over the plasma membrane. For A and D, location of the actin cuff is indicated with a bracket. Scale bars: 10 μm. Images are representative of ≥30 fields from ≥3 separate experiments of each type. For each condition in C and F, three independent experiments were quantified, with ≥10 fields counted per replicate. p value was calculated using unpaired, 2-tailed students t-test. Data are means ±SEM.

https://doi.org/10.7554/eLife.34798.030

Functional properties of the frustrated phagosome

Despite remaining unsealed, frustrated phagosomes acquired markers of endosomes and lysosomes, implying that they had undergone at least partial maturation. It was therefore conceivable that the cells established the diffusion barrier in an effort to generate a microbicidal compartment, despite their inability to form a sealed vacuole. Acidification of the lumen, secretion of antimicrobial enzymes and peptides and deployment of the NADPH oxidase are among the principal mechanisms used by leukocytes to eliminate pathogens. We first tested the ability of frustrated phagosomes to generate and maintain an acidic lumen, using the fluorescent acidotropic dye LysoBrite Red dye. As expected, the dye accumulated in lysosomes; however, it was never found to concentrate inside the frustrated phagosome (Figure 10A), suggesting that vacuolar ATPases are not functional on its membrane and/or that the junction separating the lumen from the extracellular milieu is permeable to H+. That the latter interpretation is correct was suggested by determinations of permeability of the junction using dextrans of varying size. For these experiments lysosomes were loaded with either 10 kDa or 70 kDa fluorescent dextran and then exposed to C. albicans hyphae. The dextrans were delivered into fully formed (sealed) phagosomes, where they were clearly retained (Figure 10—figure supplement 1A). The smaller (10 kDa) dextran, however, was not detectable inside frustrated phagosomes; the reduced overall staining of the cells (cf. main panel and inset in Figure 10B) suggests that secretion of lysosomes did occur, but that the dextran must have escaped the confines of the frustrated phagosome. In contrast, the 70 kDa dextran was readily visible along the frustrated phagosome, implying that its diffusion into the external medium was limited. Thus, a size-selective filter determined the extent to which solutes were retained within the frustrated phagosome. The cut-off of this filter must be greater than ≈50 kDa, because cathepsin D, a globular protein of ≈28 kDa, managed to escape the frustrated phagosome (Figure 10D), yet was routinely detected in sealed phagosomes (Figure 10—figure supplement 1B). Therefore, the incomplete phagocytic cup formed around partially internalized hyphae would be expected to have limited degradative capacity towards C. albicans. These data are in accord with the findings of (Prashar et al., 2013) that showed frustrated L. pneumophila phagosomes to retain large molecular weight dextrans, but not protons or lysosomal enzymes, despite acquisition of the V-ATPase and fusion with lysosomes.

Figure 10 with 2 supplements see all
Analysis of the antimicrobial environment within the frustrated phagosome.

(A) After phagocytosis of Candida-BFP hyphae by RAW-Dectin1 cells, acidic compartments were labeled with LysoBrite (red). The open hyphal phagocytic cup is marked with a dotted outline. Actin was visualized using transfected LifeAct-GFP. Scale bar: 5 μm. (B and C) Prior to phagocytosis of C. albicans hyphae, the lysosomes of RAW-Dectin1 cells were loaded with (B) 10 kDa or (C) 70 kDa fluorescent dextran (red). After phagocytosis, retention of dextran in the frustrated hyphal phagocytic cup was assessed by live cell microscopy. Actin was visualized using transfected LifeAct-GFP. Scale bars: 10 μm. (D) Retention of lysosomal hydrolases was assessed using transfected cathepsin D-RFP as a marker (red). Following phagocytosis and fixation, extracellular C. albicans was stained using Alexa647-conjugated concanavalin A (blue). Actin was visualized using transfected F-Tractin-GFP. Frustrated hyphal phagocytic cup marked with a dotted outline. Scale bar: 5 μm. (E) Generation of superoxide within the frustrated hyphal cup was detected using NBT. Following phagocytosis and fixation, extracellular C. albicans was stained using Alexa594 conjugated concanavalin A (red). Actin was stained using fluorescent phalloidin (blue). Inset shows merged image of formazan precipitate and the actin cuff. Scale bar: 5 μm. Images are representative of ≥30 fields from ≥3 separate experiments of each type. (F) Effect of the frustrated phagosome on C. albicans hyphal extension rate. After incubation with Candida-BFP hyphae for 10 min, RAW-Dectin1 cells transiently expressing F-Tractin-GFP were fixed at 10 min intervals, and extracellular C. albicans stained using fluorescent concanavalin A and visualized by confocal microscopy. The length of C. albicans hyphae (see (F), top) was measured for hyphae identified as extracellular (Δ), fully internalized (□) or partially internalized with actin cuffs (•), and the average hyphal length at each time-point and hyphal extension rate calculated. Average number of C. albicans per time-point was 55.0 ± 3.1. For each condition, four independent experiments were quantified, with ≥10 fields (37.5x) counted per replicate. p values calculated using the unpaired, 2-tailed students t-test. Data are means ±SEM.

https://doi.org/10.7554/eLife.34798.033

Though unable to retain luminal macromolecules over extended periods of time, the partial barrier to diffusion at the mouth of the frustrated phagosome, together with the geometrical constraint posed by the length and narrowness of the luminal space, are expected to delay the exit of molecules secreted into the phagosome. Rapidly reacting molecules may therefore be able to exert microbicidal/microbiostatic effects under these circumstances. Such is the case of reactive oxygen species produced by the NADPH oxidase. Indeed, we were able to detect preferential deposition of formazan, a product of the reaction of superoxide with nitroblue tetrazolium (NBT), inside frustrated phagosomes (Figure 10E). Heat-killed or paraformaldehyde-killed C. albicans hyphae were utilized for these experiments, eliminating the need to account for superoxide production by live C. albicans (see Materials and methods).

Because the frustrated phagosome appeared to retain some antimicrobial function, we assessed the effect of the frustrated phagosome environment on the fate of partially internalized hyphae. We could not detect significant loss of viability of the partially internalized C. albicans, as assessed by propidium iodide staining. We reasoned that the antimicrobial effectors may not suffice to kill the fungus, yet their effects may manifest as an observable change in the rate of hyphal extension, which can average 0.31 µm min−1 on serum agar (GOW and Gooday, 1982). When measured in RPMI medium without serum (wthout macrophages present) C. albicans hyphae grew at a rate of 0.22 µm min−1 ±0.03. Remarkably, the extension rate of partially internalized hyphae, which displayed an actin cuff, was significantly reduced (0.11 µm min−1 ±0.01). This reduced growth rate was not different (p=0.742) to that of fully internalized hyphae (0.108 µm min−1 ±0.011). In the same experiments, neighboring C. albicans hyphae not in contact with macrophages grew at a rate of 0.198 µm min−1 ±0.014, indistinguishable from that measured in the absence of macrophages. Therefore, while small molecular weight contents can eventually diffuse out of the frustrated phagosome, they are nevertheless retained sufficiently to limit the growth of partially internalized C. albicans. This microbiostatic effect on partially internalized C. albicans hyphae could be ablated by blocking macrophage CR3 with the M1/70 antibody (see Figure 3F,G and H) before phagocytosis (Figure 10—figure supplement 2), reiterating the importance of CR3 ligation to β(1,3)-glucan for the generation and maintenance of this atypical phagocytic environment.

Discussion

Most of the microbicidal and degradative properties of the phagosome depend on the release and containment of lysosomal hydrolases, antimicrobial peptides and reactive oxygen species in close proximity to the internalized microorganism. However, when phagocytes are faced with exceptionally large targets, their internalization can become retarded or frustrated altogether. The inability to complete phagocytosis, as in the case of long asbestos fibers (Donaldson et al., 2010) or bacterial biofilms (Costerton et al., 1999; Scherr et al., 2014; Thurlow et al., 2011) can potentiate harmful inflammation.

C. albicans hyphae can attain lengths of ≥50 μm (GOW and Gooday, 1982), overwhelming the comparatively diminutive phagocytes that are unable to ingest them whole. Accordingly, attempts to internalize such hyphae are frustrated and inflammatory in nature (Branzk et al., 2014; Goodridge et al., 2011; Lewis et al., 2012; Rosas et al., 2008). Nevertheless, our study shows that macrophages endeavor to seal the frustrated phagocytic compartment, in an effort to maximize their antimicrobial effect and minimize the release of inflammatory agents. To this end, they generate de novo a strikingly effective diffusion barrier by a process that involves activation of integrins that induce the formation of a thick F-actin cuff at the neck of the tubular phagosomes. The formation of actin-rich structures was reported previously during infection of macrophages with C. albicans (García-Rodas et al., 2011; Heinsbroek et al., 2009; Strijbis et al., 2013) and other rod/filament shaped microbes (Gerisch et al., 2009; Prashar et al., 2013), but neither their mechanism of assembly nor their functional significance were fully understood, which motivated our studies.

As expected, Dectin1 –the major phagocytic receptor for fungal β-glucan (Brown and Gordon, 2001; Brown et al., 2002; Taylor et al., 2007)– was present along the phagocytic cup, lining the internalized portion of the hyphae. Indeed, the RAW-Dectin1 cell line used for some of our studies was created to allow efficient internalization of fungal zymosan (Esteban et al., 2011), and has been used to study C. albicans-macrophage interactions (Strijbis et al., 2013). Dectin1 signaling leads to robust production of reactive oxygen species by the NADPH oxidase in response to fungal ligands (Brown et al., 2002; Goodridge et al., 2011; Underhill et al., 2005), accounting for our observation that superoxide is detected within the frustrated phagocytic cup. Remarkably, however, Dectin1 did not concentrate in the region of the membrane adjacent to the actin cuff where the diffusion barrier was established. Instead, integrin αMβ2 (CR3, or CD11b/CD18) was found to accumulate in this region.

Engagement of CR3 at the cuff is required for the formation of the underlying actin cuff, a process likely mediated by talin and vinculin, which were also found accumulated at the site. The entire assembly appears central to the establishment of the diffusion barrier, which is lost when blocking CR3 binding with the M1/70 antibody and also when latrunculin is used to disassemble actin filaments. CR3 is unique amongst integrins in that its α domain contains a lectin-like domain (LLD) capable of binding fungal β-glucan (Ross et al., 1985; Vetvicka et al., 1996). This LLD, located at the membrane-proximal C terminus (between residues 943–1047; Lu et al., 1998) is distinct from the traditional ligand-binding I domain. Importantly, the LLD can bind β-glucan in a Ca2+-independent manner while the integrin is in the inactive, bent conformation (Thornton et al., 1996). Binding of glucan has been shown to induce a semi-active conformation of the integrin that is predicted to facilitate outside-in signaling (O'Brien et al., 2012; Vetvicka et al., 1996). We found that Dectin1 did not have a direct role in actin cuff formation, being instead required for adhesion and the initiation of phagocytosis. Interestingly, when Dectin1 expression is low, the deposition of opsonins contained in serum –including complement and possibly also anti-Candida antibodies– suffice to engage CR3 and promote actin cuff formation directly activating this process, as predicted from earlier observations (Boxx et al., 2010; Kozel et al., 1987; Vetvicka et al., 1996). Dectin1 signaling can initiate inside-out activation of CR3 (Li et al., 2011). However, conventional Rap1-dependent inside-out signaling mediated by CalDAG-GEF1 was dispensable for actin cuff formation, as were divalent cations. Therefore, it is likely that CR3 binds fungal β-glucan in a manner that does not require inside-out activation by Dectin1.

We showed that mannan, a ligand for the mannose receptor, had no effect on actin cuff formation. The utilization of a curated set of C. albicans GRACE strains confirmed that mannan was dispensable and further excluded chitin and β(1,6)-glucan as ligands for cuff formation. Importantly, caspofungin inhibition of β(1,3)-glucan synthesis blocked formation of the cuff, in a dose dependent manner. These observations are in accord with involvement of the LLD of CR3, which was previously demonstrated to ligate β-glucan (Mueller et al., 2000; Thornton et al., 1996; Vetvicka et al., 1996). Clearly, β-glucan is also a ligand for Dectin 1 (Brown and Gordon, 2001; Brown et al., 2002; Palma et al., 2006), and initial engagement of this receptor is required for formation of actin cuffs around unopsonized hyphae. However, β(1,3)-glucan appears to play a distinct role in CR3-mediated actin cuff formation, as the effects of caspofungin could not be rescued by serum opsonization, with the caveat that caspofungin treatment may have affected complement deposition on C. albicans (Boxx et al., 2010; Kozel et al., 1987), although we regard this as unlikely because the fungal cell wall is rich in other polysaccharides and proteins that can serve to attach complement.

Our observations implicate clustered CR3 as an initiator of actin polymerization and a key constituent of the diffusion barrier. The signals mediating this effect include activation of Syk, which had been reported earlier (Strijbis et al., 2013), and also of Pyk2 and Fak. The latter related kinases were enriched at the cuff and dual inhibition by PF573228 blocked cuff formation. Along with activated Syk, Pyk2 and Fak have been shown to interact with β2 integrins, including CR3 (Duong and Rodan, 2000; Fernandez and Suchard, 1998; Han et al., 2010; Hildebrand et al., 1995; Kamen et al., 2011; Mócsai et al., 2002; Raab et al., 2017; Rubel et al., 2002; Wang et al., 2010; Yan and Novak, 1999). Pyk2, in particular, is required for paxillin and Vav1 activation during integrin engagement and CR3-mediated phagocytosis (Kamen et al., 2011). Paxillin was proposed to act as a scaffold, bridging integrin-initiated complexes with Rho-GTPases (Deakin and Turner, 2008), and Vav1, previously identified as important for the phagocytosis of C. albicans (Strijbis et al., 2013), also links β2 integrins to the activation of Cdc42, Rac1 and RhoA (Gakidis et al., 2004). These signaling events appear conserved during the frustrated phagocytosis of C. albicans hyphae, as we detected paxillin and active Rac1/Cdc42 at the actin cuff, and Vav1 was found to be enriched in actin cuff-like structures around C. albicans (Strijbis et al., 2013).

The activation of the Rho family GTPases is linked to both formin- and Arp2/3- mediated actin dynamics. The actin cuffs formed around C. albicans hyphae were singularly sensitive to SM1-FH2 –and therefore dependent on formin-mediated linear actin polymerization– and not to inhibitors of the Arp2/3 complex that promotes branched actin polymerization. Interestingly, Rac1 and Cdc42 interact with actin-nucleating formins of the mDia family (Lammers et al., 2008). Collectively, our findings support a model whereby CR3 initiates signaling through Syk and Pyk2/Fak, leading to activation of Vav1 and Rho GTPases, culminating in formin-dependent actin nucleation.

Our studies show that the integrin/actin cuff separates the open phagocytic cup from the plasma membrane, appearing to act as a boundary that segregates distinct and mobile membrane domains. There was a clear segregation of phosphoinositides between the plasma membrane (PtdIns(4,5)P2) and the open cup (PtdIns(3,4,5)P3 and PtdIns(3)P). In principle, such separation could stem from the differential and strategic localization of kinases and/or phosphatases in the two membranes and in the junctional complex. However, we also observed slowly convertible (LC3) or non-convertible lipid-anchored proteins (constitutively-active Rab7) and transmembrane proteins (LAMP1) to be retained in the phagocytic cup, unable to reach the surface membrane. FRAP studies confirmed that these molecules were freely mobile within the phagocytic cup, pointing to the integrin/actin cuff as the diffusion barrier.

While antibody-induced clustering of CR3 was sufficient to form an actin-independent diffusional barrier, actin was required to maintain the barrier function of the cuff during phagocytosis of hyphae. Actin can stabilize integrins in their active conformation (Kaizuka et al., 2007; Lavi et al., 2007; Lavi et al., 2012), and such stabilization is likely required to maintain the cuff during extended frustrated phagocytosis. Actin-dependent diffusional barriers have been invoked in other systems (Golebiewska et al., 2011; Nakada et al., 2003; Prashar et al., 2013), although the pickets that anchor the cytoskeletal fence and restrict the diffusion of membrane components had not been previously identified.

We also analyzed the functional consequences of the establishment of the cuff and diffusion barrier. By segregating the two domains, the barrier enabled the open phagocytic cup to undergo an atypical maturation, despite the fact that scission from the surface membrane never occurred. This enabled targeting and activation within the frustrated phagosome of the NADPH oxidase, which generated toxic superoxide in the immediate vicinity of the portion of the hypha that had been engulfed. Thus, a crucial means for C. albicans control during infection (Sasada and Johnston, 1980; Brothers et al., 2013) remains operational in the frustrated phagosomes.

The observation that the tubular phagosomes were rich in LAMP1, a prototypic lysosomal marker, suggests that lysosomal hydrolases must have been secreted also into the phagosomal lumen. These were, however, not well retained by the phagosomes because, despite the tight seal that separated the inner leaflet of the membrane, the junction separating the aqueous compartments (i.e. the lumen from the extracellular space) was not perfectly tight. While 70 kDa dextran was retained within the phagosome, 10 kDa dextran was not, resembling the findings in frustrated L. pneumophila phagosomes (Prashar et al., 2013) and indicating the establishment of a sieve that excluded molecules with a hydrodynamic radius greater than ≈ 6–8 nm (Nicholson and Tao, 1993). Cathepsin family members (radius ≈ 2.4 nm; Fazili and Qasim, 1986) and similarly-sized hydrolases would therefore eventually escape the lumen. Nevertheless, because a partial seal is formed, their rate of loss might be slowed, allowing hydrolases and other antimicrobial molecules to act on the partially internalized C. albicans hyphae before exiting the open cup. Fast-acting antimicrobial agents, like ROS, released in close proximity to the target would be expected to be at least partially effective. Consistent with this hypothesis, partially internalized C. albicans hyphae exhibited a reduced growth rate compared to external hyphae. Importantly, this growth restriction was abolished upon antibody blockade of CR3 and loss of actin cuff formation, presumably a result of increased leakage of phagosome contents. In this case, agents such as ROS, would not reach sufficient concentration to manifest the microbiostatic effect. However, leakage of phagosomal contents or ROS does not explain the failure of the frustrated phagosomes to kill the fungus, because C. albicans yeast and hyphae survive also within fully sealed phagosomes.

Based on the preceding considerations, we hypothesize that the integrin/actin cuff is generated and maintained by macrophages as a means to sustain antimicrobial functions in open tubular phagosomes formed around C. albicans hyphae and possibly other targets. It is tempting to speculate that the unique conditions established by the diffusion barrier might provide additional benefits to the phagocyte. In dendritic cells, decreased phagosomal proteolysis associated with reduced phagosome acidification protects antigens for enhanced presentation (Mantegazza et al., 2008; Rybicka et al., 2012; Savina et al., 2006). In addition, the frustrated yet maturing phagosome may enable activation of endomembrane Toll-like receptors (TLRs). TLR3 and TLR9 both localize to intracellular compartments and recognize C. albicans nucleic acids and chitin, respectively, contributing to a protective cytokine response to the fungus (Nahum et al., 2011; Wagener et al., 2014). Interestingly, Dectin1 can collaborate with plasmalemmal TLRs (TLR2 or TRL4) to enhance signaling and augment cytokine production (Ferwerda et al., 2008; Netea et al., 2006; Underhill, 2007) and a similar synergy may apply to endomembrane TLRs. Indeed, Dectin1 recognition is required for TLR9 localization to the C. albicans phagosome and TLR9-dependent gene expression (Khan et al., 2016). Thus, the unique structure described here may play an important role in the control of fungal infection and possibly also in the management of biofilms and other large targets by phagocytes.

Materials and methods

Reagents

Mammalian expression vectors were obtained from the following sources: Emerald-Dectin1 (plasmid #56291; Addgene, Cambridge, MA), PAK-PBD-YFP (Srinivasan et al., 2003), E-cadherin-GFP (plasmid #67937; Addgene), β-catenin-GFP(plasmid #16071; Addgene), Talin-GFP (Franco et al., 2004), AKT-PH-GFP (Marshall et al., 2001), PLCδ-PH-GFP (Botelho et al., 2000), PX-GFP (Kanai et al., 2001), LC3-GFP (Kabeya et al., 2000), Rab7-GFP (Bucci et al., 2000), Rab7(Q67L)-RFP (D’'Costa et al., 2015), Lamp1-GFP (Martinez et al., 2000), Lyn11-GFP (Teruel et al., 1999), cathepsin D-RFP (Yuseff et al., 2011), LifeAct-RFP or -GFP (Riedl et al., 2008), F-tractin-GFP (Belin et al., 2014), CD2-CD45-GFP (Cordoba et al., 2013).

Primary antibodies were purchased from the following vendors: HA (catalogue #MMS-101P; Covance, Princeton, NJ), pTyrosine (catalogue #05–321; EMD Millipore, Billerica, MA), pFAK-Y397 (catalogue #3283S; Cell Signaling, Berverly, MA), pPYK2-Y402 (catalogue #3291S; Cell Signaling), pSFK-Y418 (catalogue #44660G; Invitrogen, Carlsbad, CA), pSYK-Y525/526 (catalogue #2771S; Cell Signaling), Talin (catalogue #T3287; Sigma-Aldrich, St. Louis, MO), Vinculin (catalogue #MAB3574; EMD Millipore), HS1 (catalogue #4557S; Cell Signaling), LAMP1 (catalogue # 1D4B-s, Developmental Studies Hybridoma Bank, Iowa City, IA), actin (catalogue #A4700; Sigma-Aldrich), CD11b (catalogue #557394; BD Biosciences, Franklin Lakes, NJ), CD18 (catalogue #557437; BD Biosciences), rat IgG2B isotype control (catalogue #MAB0061; R and D systems, Minneapolis, MN), paxillin (catalogue #P13520; Transduction Laboratories, Lexington, KY), GAPDH (catalogue #MAB374; EMD Millipore), E-cadherin (catalogue #610181; BD Biosciences), β-catenin (catalogue #610153; BD Biosciences). Unconjugated and Alexa488, Cy3, Cy5, HRP-conjugated secondary antibodies against mouse, goat, rat, rabbit IgGs were obtained from Jackson ImmunoResearch Labs (West Grove, PA).

Fungal strains and culture conditions

A list of all C. albicans strains tested is provided in Table 1. C. albicans strain SC5314 expressing BFP (Candida-BFP; Strijbis et al., 2013) was grown at 30°C in YPD (BD Biosciences). C. albicans cell wall mutant strains were obtained from the GRACE collection of tetracycline-repressible mutant strains (O'Meara et al., 2015; Roemer et al., 2003). Depletion of target gene expression was achieved by adding 0.5 μg mL−1 doxycycline (DOX) to the growth medium. To induce hyphae of C. albicans, overnight cultures were subcultured 1:1000 in RPMI-1640 medium and incubated at 37°C for 1–3 hr, as indicated in the text. In some cases, caspofungin (Sigma-Aldrich) was used to pharmacologically inhibit β(1,3)-glucan synthesis. C. albicans overnight cultures were subcultured into RPMI-1640 containing 10, 5, 2.5, 1.25 and 0 ng mL−1 caspofungin for 2 hr at 30°C. Cultures were then moved to 37°C for 1 hr to induce hyphae in the presence of caspofungin. To measure the effect of caspofungin on β(1,3)-glucan levels, C. albicans hyphae-infected wells were stained with 0.05% aniline blue (EVANS et al., 1984; Lee et al., 2016) overnight.

A. fumigatus strain AF293 (clinical isolate) was grown on YPD agar (Bioshop, Burlington, ON) plates at 30°C. Conidia were harvested in PBS containing 0.01% Tween-80. For experiments, resuspended conidia were diluted 1:10 in RPMI-1640 containing 0.01% Tween-80, and allowed to form hyphae at 30°C overnight. Hyphae were then washed twice with PBS 0.01% Tween-80, and diluted 1:10 or 1:100 into RPMI-1640 containing 0.01% Tween-80.

Mammalian cells and culture conditions

The RAW 264.7 cell line was obtained from and authenticated by the American Type Culture Collection (ATCC, Manassas, VA). The RAW-Dectin1-LPETG-3xHA cell line (RAW-Dectin1) was provided by Dr. Karin Strijbis and authenticated for Dectin1-HA expression and Dectin1-mediated phagocytic ability by flow cytometry (Esteban et al., 2011). Prior to experimentation, these cell lines were validated in our laboratory by assessing their morphology, phagocytic ability and expression of plasma membrane markers. RAW 264.7 and RAW-Dectin1 cells were grown in RPMI-1640 medium containing L-glutamine (MultiCell, Wisent, St. Bruno, QC) and 10% heat-inactivated fetal calf serum (FCS; MultiCell, Wisent), at 37°C under 5% CO2. The A431 cell line was obtained from and authenticated by the American Type Culture Collection (ATCC). Prior to experimentation, this cell line was revalidated by assessing its expression of plasma membrane markers, and responsiveness to epidermal growth factor (EGF). A431 cells were grown in DMEM medium containing L-glutamine (MultiCell, Wisent) and 10% heat-inactivated FCS, at 37°C under 5% CO2. All cell lines tested negative for mycoplasma contamination by DAPI staining.

Bone marrow-derived macrophages (BMDM) were obtained from the femoral bones of CALDAG-GEF1−/− (Bergmeier et al., 2007) or +/+ (C57BL/6) mice, and differentiated for 5–7 days in DMEM containing L-glutamine, 10% heat-inactivated FCS, 100 U mL−1 penicillin, 100 μg mL−1 streptomycin, 250 ng mL−1 amphotericin B (MultiCell, Wisent) and 10 ng mL−1 mM-CSF (PeproTech, Rocky Hill, NJ), at 37°C and 5% CO2. To obtain M2 human monocyte-derived macrophages, peripheral blood mononuclear cells were isolated from the blood of healthy donors by density-gradient separation with Lympholyte-H (Cedarlane, Burlington, ON). Human monocytes were then separated by adherence, and incubated in RPMI-1640 containing L-glutamine, 10% heat-inactivated FCS, 100 U mL−1 penicillin, 100 μg mL−1 streptomycin, 250 ng mL−1 amphotericin B and 25 ng mL−1 hM-CSF (PeproTech) for 7 days.

Phagocytosis

Mammalian cell lines or primary cells were seeded on 18 mm coverslips in 12-well plates at 2 × 105 cells mL−1. For infections with C. albicans, the medium was aspirated from the wells and replaced with 1 mL of C. albicans that had been induced to form hyphae. Plates were centrifuged for 1 min at 1500 rpm, then incubated with the following cell types at 37°C and 5% CO2 for phagocytosis to proceed: RAW-Dectin1: 1 hr; BMDM: 15 min; human M2 macrophages: 20 min; A431 cells: 3 hr. In some cases, 30 min prior to infection, C. albicans hyphae were opsonized in human serum to promote the deposition of complement, although deposition of donor-specific anti-Candida antibodies may have also occurred, further favoring phagocytosis.

Where indicated, cells were treated with either vehicle or 1 μM Latrunculin A (Sigma-Aldrich) after 1 hr Candida-BFP infection, or pretreated 30 min with 4 mM EDTA (Bioshop), followed by infection with Candida-BFP in the presence of EDTA. For inhibition of actin polymerization or kinases, after 30 min incubation with Candida-BFP, monolayers were treated with either vehicle, 50 μM CK-666 (Calbiochem, La Jolla, CA), 10 μM SMI-FH2 (Calbiochem), 10 μM PP2 (Calbiochem), 50 μM piceatannol (Sigma Aldrich), or 50 μM PF573228 (Tocris, Oakville, ON), for 30 min. Following infection, monolayers were washed three times with PBS and fixed with 4% paraformaldehyde (PFA). In some cases, wells were treated with various fluorescent reagents before or after phagocytosis, as described below.

For infections with A. fumigatus, RAW-Dectin1 cells were incubated with 1 mL diluted A. fumigatus hyphae, and plates centrifuged for 5 min at 1500 rpm. Plates were incubated for 2 hr at 37°C and 5% CO2 for phagocytosis to proceed. RAW-Dectin1 cells were pretreated 10 min and infected in the presence of 100 mM L-cysteine (Sigma-Aldrich) to prevent gliotoxin-mediated inhibition of phagocytosis (Schlam et al., 2016). Following infection, monolayers were washed three times with PBS and fixed with 4% paraformaldehyde (PFA). In some cases, wells were treated with various fluorescent reagents before or after phagocytosis, as described below.

After phagocytosis, external C.albicans were labeled for 20 min at room temperature using a solution of 5 μg mL−1 fluorescent conjugated concanavalin A (ThermoFisher Scientific, Waltham, MA). To stain actin filaments, cells were permeabilized 5 min with 0.1% Triton X-100 and incubated 30 min with a 1:1000 dilution of fluorescent phalloidin (Thermofisher Scientific) or acti-stain (Cytoskeleton, Inc., Denver, CO). In some cases, C. albicans and A. fumigatus were stained with 10 μg mL−1 calcofluor white (Fluorescent Brightener 28; Sigma-Aldrich).

DNA transfection

For transient transfection, RAW-Dectin1 cells were plated on 18 mm glass coverslips at a concentration of 2 × 105 cells mL−1 16–24 hr prior to experiments. FuGENE HD (Promega, Madison, WI) transfection reagent was used according to the manufacturer’s instructions. RAW-Dectin1 cells were transfected at a 3:1 ratio using 1.5 μL FuGENE HD and 0.5 μg DNA per well, and used for experiments 16 hr after transfection.

In some cases, DNA transfections were performed using the Neon transfection system (Life Technologies, Carlsband, CA) according to the manufacturer’s protocol. RAW-Dectin1 cells were lifted, washed and resuspended to a concentration of 4 × 106 cells mL−1 and 100 μL of the suspension were mixed with 5 μg DNA. Electroporation was done using a single 20 ms pulse of 1750 V. Cells were then immediately transferred to RPMI-1640 containing L-glutamine and 10% heat-inactivated FCS, before seeding on coverslips at concentration of 2 × 105 cells mL−1. Cells were used for experiments 16 hr after electroporation.

Immunofluorescence

After phagocytosis, fixation and concanavalin A staining (as indicated), monolayers were permeabilized in PBS containing 0.1% Triton X-100 for 5 min and blocked in PBS containing 5% skim milk and 0.1% Triton X-100 for 30 min at room temperature. Samples were incubated with primary antibodies for 30 min at room temperature. Primary antibody dilutions were: HA (1:1000), pTyrosine (1:100), pFAK-Y397 (1:100), pPYK2-Y402 (1:100), pSRC-Y418 (1:100), pSYK-Y525/526 (1:100), talin (1:500), vinculin (1:500), HS1 (1:250), LAMP1 (1:20), actin (1:100), E-cadherin (1:100), β-catenin (1:100), CD11b (1:100), CD18 (1:100), paxillin (1:100). After rinsing with PBS, samples were incubated 30 min at room temperature with Alexa488, Cy3 or Cy5-conjugated secondary antibodies at a 1:10,000 dilution. Where indicated, fluorescent phalloidin at a 1:1000 dilution was included with secondary antibodies. Samples were rinsed and viewed in PBS by confocal microscopy.

Immunoblotting

Cells were grown in six well plates at a concentration of 4 × 105 cells per well. After infections, wells were lysed in Laemmli buffer (Bio-Rad, Mississauga, ON). Samples were run on a 7% SDS-PAGE gel for separation and the gel was transferred to a polyvinylidene difluoride (PVDF) membrane. Membrane was blocked in PBS containing 5% skim milk and 0.05% Tween-20 for 30 min at room temperature, followed by primary antibody staining for 1 hr at room temperature, in blocking buffer. Primary antibodies dilutions: E-cadherin (1:10,000), β-catenin (1:1000), GAPDH (1:20,000; loading control). After washing membrane in PBS containing 0.05% Tween-20, samples were incubated 30 min at room temperature with HRP-conjugated secondary antibodies at a 1:3000 dilution. Blots were visualized using the ECL Prime Western Blot detection reagent (GE Healthcare, Mississauga, ON) on an Odyssey Fc (LI-COR, Lincoln, NE).

Rhodamine-PtdEth labeling

5 μg Rhodamine-PtdEth (L-α-phosphatidylethanolamine-N-(lissamine rhodamine B sulfonyl) ammonium salt; Avanti Polar Lipids, Inc., Alabaster, AL) was dried under N2 and resuspended in 10 μL methanol. After vortexing, 900 μL 3 mg mL−1 bovine serum albumin (BSA) was added. This was then diluted 1:1 in cold serum-free RPMI-1640 containing 25 mM HEPES (HPMI; MultiCell, Wisent). Following phagocytosis, the medium was aspirated and replaced with 500 μL of the prepared Rhodamine-PtdEth/HPMI, and incubated at 4°C for 10 min. The adherent cells were rinsed three times with cold HPMI, heated to 37°C 10 min and imaged live.

Cholera toxin B labeling

Following phagocytosis, cells were rinsed three times with cold HPMI. Cholera toxin subunit B, Alexa488 conjugate (Thermofisher Scientific) was added to a final concentration of 1 μg mL−1 and cells incubated at 4°C for 10 min. The adherent cells were rinsed three times with cold HPMI, heated to 37°C 10 min and viewed live.

LysoBrite red staining

Following phagocytosis, acidic intracellular compartments were stained using a 1:5000 dilution of the acidotropic LysoBrite Red dye (AAT Bioquest, Sunnyvale, CA) for 5 min at 37°C. Monolayers were rinsed three times with PBS, placed in HPMI, and imaged live.

Dextran loading

RAW-Dectin1 cells were pulsed overnight with 20 μg mL−1 Alexa647-conjugated 10 kDa dextran or 25 μg mL−1 tetramethylrhodamine-conjugated 70 kDa dextran. Cells were washed and incubated with Candida-BFP, as described above, for 1 hr. Following phagocytosis, monolayers were rinsed three times with PBS, placed in HPMI, and viewed live.

Nitroblue tetrazolium assay

This assay required PFA-inactivated Candida-BFP hyphae, as metabolically active C.albicans reduced nitroblue tetrazolium (NBT) to formazan, confounding the results. Candida-BFP hyphae were made as described above, followed by fixation in 8% PFA for 20 min. PFA-inactivated hyphae were rinsed and used to infect RAW-Dectin1 cells for 1 hr, in the presence of 0.5 mg mL−1 NBT (Sigma-Aldrich). Following infection, cells were washed three times with PBS and fixed with 4% PFA. Formazan precipitate, created in response to superoxide anion produced by phagosomal NADPH oxidase, was visualized by bright field microscopy.

Blocking experiments

To block CR3, adherent RAW-Dectin1 cells were incubated with 10 μg blocking antibody to CD11b (monoclonal M1/70) or rat IgG2B isotype control (MAB0061; R and D systems) for 30 min at room temperature. After warming to 37°C, cells were incubated with Candida-BFP hyphae as described above. Following phagocytosis, monolayers were rinsed and fixed in 3% PFA as mentioned above. After permeabilization and blocking, described above, blocking antibody was detected using a Alexa488-conjugated secondary antibody against rat IgG (1:10,000; Jackson ImmunoResearch Labs), and viewed by confocal microscopy.

For sugar blocking experiments, adherent RAW-Dectin1 cells were pretreated 30 min with 100 μg mL−1 mannan (Sigma Aldrich) or laminarin (Sigma Aldrich), followed by infection with C.albicans in the continued presence of each sugar. In the case of laminarin, Candida-BFP hyphae were allowed to adhere to RAW-Dectin1 cells 10 min, followed by the addition of 100 μg mL−1 laminarin for the remainder of the 1 hr infection.

CR3 patching

RAW-Dectin1 cells transiently transfected with CD2-CD45-GFP were washed with PBS and cooled to 10°C in 1X HBSS (MultiCell, Wisent). To crosslink surface CR3, cells were sequentially incubated with: (1) 5 μg mL−1 anti-CD18 primary (monoclonal M18/2); (2) goat anti-rat secondary and (3) donkey anti-goat tertiary antibodies, for 30 min each at 4°C, in PBS containing Ca2+, Mg2+ and 0.1% glucose. Labeled cells were then incubated 30 min at 37°C in the presence of 30 μM DYNGO 4a, to patch crosslinked CR3 without its internalization. Following this treatment, cells were treated 10 min with or without 1 μM latrunculin A, in the presence of 1 μM N-ethylmaleimide to prevent internalization of crosslinked CR3 following actin depolymerization. Monolayers were fixed in 3% PFA, and extracellular CR3 was labeled using a fluorescent anti-goat antibody. Plasma membrane clusters of CR3 were analyzed in Volocity software for exclusion of CD2-CD45-GFP. Exclusion was calculated from confocal slices as the ratio of the GFP intensity per pixel in the CR3-positive patches to the average intensity in the plasma membrane.

Hyphal growth rate of C. albicans during infection

RAW-Dectin1 cells were incubated with Candida-BFP hyphae for 10 min to allow adherence, then fixed in 4% PFA after 0, 10, 20, 30, 40, 50 or 60 min. Following phagocytosis, the cells were fixed in 3% PFA as above, external Candida-BFP were labeled with fluorescent concanavalin A, and permeabilized and stained with fluorescent phalloidin, as described above. Samples were imaged by confocal microscopy, and the length of individual Candida-BFP hypha measured in μm. Hyphal extension rate was calculated by linear regression analysis in GraphPad Prism software (GraphPad Software, Inc., La Jolla, CA).

Confocal microscopy

Confocal images were acquired using a Yokogawa CSU10 spinning disk system (Quorum Technologies Inc., Guelph, ON) or a Leica SP8 laser scanning system (Leica). Images were acquired using a 63x/1.4 NA oil objectives or 25x/0.8 NA water objective (ZEISS, Germany), as indicated, with an additional 1.5x magnifying lens. For live experiments, cells were maintained at 37°C using an environmental chamber (Live Cell Instruments, Korea). Routine analyses and colocalizations were done using Volocity software (Perkin Elmer, Woodbridge, ON). 3D data visualization was done using Imaris (Bitplane, Concord, MA) software.

For colocalization analyses, Volocity software was used to calculate positive product of the differences of the mean (Li et al., 2004) channels, which were then overlayed on merged images for visualization.

For fluorescent intensity calculations, background subtracted intensities per unit area for expressed fluorescent protein constructs or endogenous proteins (immunofluorescence) were measured in Volocity software. Ratios were calculated comparing relative intensities in the actin cuff compared to phagocytic cup, or phagocytic cup compared to membrane, as indicated in the text.

Fluorescence recovery after photobleaching (FRAP)

FRAP experiments of GFP or RFP-tagged proteins, transiently expressed in RAW-Dectin1 cells, were conducted on an A1R point-scanning confocal system (Nikon Instruments, Japan). For FRAP, Candida-BFP hyphae-infected cells were imaged in HPMI at 37°C. Images were acquired using a 60x/1.4 NA oil objective (Nikon), 1.2-AU pinhole, resonant scanning mode, and 16x line averaging. For a complete 2 min FRAP acquisition at 1.9 fps, after 5 s of initial imaging, a region of interest 3 μm in diameter was bleached for 1.06 s using the 405 laser at 100% power, followed by imaging for fluorescence recovery. Images were exported and analyzed for fluorescence intensity using Volocity software.

After background subtraction, fluorescence intensity units were normalized (see Figure 6 legend) using Microsoft Excel software, and transformed to a 0–1 scale, to correct for differences in bleaching depth and allow for comparison of up to 30 individual FRAP curves per condition. Graphpad Prism software was used to fit the FRAP curves to a single exponential, plotted as fractional recovery over time.

Transmission electron microscopy (TEM)

Candida-BFP hyphae-infected RAW-Dectin1 cells were washed with cold PBS, and fixed with 2% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.3. For improved specimen preparation, samples were then subjected to a zymolyase digestion protocol (Bauer et al., 2001) designed to weaken the fungal cell wall and allow for adequate structural preservation. Samples were then post-fixed in 1% osmium tetroxide in 0.1 M sodium cacodylate buffer, pH 7.3, dehydrated in a graded ethanol series followed by propylene oxide, and embedded in Quetol-Spurr resin. Ninety nm sections were cut on a Leica Ultracut ultramicrotome and stained with uranyl acetate and lead citrate. Samples were imaged on a FEI Tecnai 20 transmission electron microscope, equipped with an AMT 16000 digital camera.

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Decision letter

  1. Dominique Soldati-Favre
    Reviewing Editor; University of Geneva, Switzerland

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 "An integrin-based diffusion barrier separates membrane domains enabling formation of microbicidal frustrated phagosomes" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Neil Gow (Reviewer #1).

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

As you will see from their individual comments below, the reviewers concur that this study reports some advances the field of host-fungus interactions and add supporting data on role for integrin in the establishment or maintenance of the actin cuff. The manuscript is clearly written, and the work is well executed and clearly illustrated with beautiful imaging data.

However, the originality of the manuscript is in part compromised because some of the concepts were preceded by a study from Dr. Terebiznik's lab. While the current work needs to be more cross-referral to this, there is still novelty in the current submission leaning on the identification of Mac-1-dependent diffusion barriers formed between the hyphae cell wall and the plasma membrane of the macrophage. The experimental data supporting this model requires important revisions in order for it to be convincing (more quantitative data). Furthermore, data in support of the microbicidal properties of these frustrated phagosomes is missing. The roles of Dectin-1 and Integrin receptors in the formation of actin cuffs and barriers should investigated in more detail whereas it is optional to expand the study into inflammatory response.

While we are rejecting the paper as a result of the need for a substantial revision (longer than 2 months), we are broadly supportive of this manuscript and would consider a newly submitted form of this paper that would be treated as a revised manuscript.

Reviewer #1:

This paper addresses a well appreciated but univestigated conundrum. How do immune phagocytes interact with target cells that are much larger than they can envelop? The authors show that macrophages that are challenged with elongated hyphal cells of the fungus Candida albicans wrap around the end of a hypha and form an actin cuff that partially seals the phagocytic cup at the mouth of the tubular phagosome. This structure retains some of the ability to establish an antimicrobial environment within the unsealed tubular phagosome – for example allowing NAPPH oxidase activation, but not phagosome acidification. The actin cuff is shown to represent a diffusion barrier that segregates phosphoinositides between the phagocytes cell membrane and the open cup. Some elegant experiments investigate how this is segregation is achieved and regulated. Overall this is an interesting, novel and well executed study that clearly advances the field of host-fungus interactions.

I have a number of questions, suggestions and presentational issues.

1) The fungus studied is Candida albicans but is most commonly referred to as Candida. It would not be simply pedantic to request that it be referred to as C. albicans throughout since most Candida species cannot form elongated hyphae.

2) It would be interesting to know if similar actin cuff seals are formed when phagocytes encounter hyphae of Aspergillus fumigatus (interesting because this fungus has quite marked differences in cell wall composition).

3) The hyphae used are not very long (1 h or 2 h hyphae were used) and the macrophages shown seem to be invaginating one end. Macrophages will also wrap around the trunk of a very large hypha. Are actin cuffs formed here- and what is their architecture? It would be useful to include some images of longer hyphae of 4-6 hours incubation in RPMI.

4) Various C. albicans mutants are available with alterations in the cell wall that would help determine what surface components of the fungus are important for inducing the cuffs.

5) In many of the figures it is not possible to clearly evaluate the shape and position of the phagocyte relative to the fungus. It would be very helpful to show a DIC or phase image of the same interacting cells in order to really see the orientation of the hypha and phagocyte. I have no doubt that the descriptions in the text are accurate, but I have to somewhat take it on trust since I cannot see the outline of the two interacting cells clearly in all cases. (For example see Figure 3B,C; Figure 4B,E,F; Figure 5B,C,F; Figure 6E; Figure 8A,D and others).

6) The authors show that the CTL Dectin-1 is not present at the actin cuff, but phosphotyrosine accumulated at this site suggesting that some other pattern recognition receptor may drive actin assembly. It would be useful to verify that glucan phosphate or laminarin did not block actin cuff formation.

7) Since the mannose receptor is likely to engage with hypha mannan, (which is located in the outer cell wall) it would be interesting to know if this is located at this site.

8) I'm not sure that the TEM shown in Figure 1C can be seen as evidence of showing the actin cuff unless the actin is perhaps stained using colloidal gold.

9) My88-/- cells would be useful in assessing the possible involvement of TLRs.

10) The actin cuffs appear to be elongated in some images and much shorter or almost like spots in others. Do they retain their shape over time or contract to a spot?

Reviewer #2:

In this manuscript M Maxson and collaborators have characterized the phagocytosis of Candida albicans hyphae, which for long hyphae, cannot progress beyond the stage of phagocytic cup, leading to the formation of a frustrated phagosome. The manuscript is clearly written, and the findings illustrated with beautiful imaging data. The authors propose that actin cuffs in the frustrated phagosomes, which have been described in previous reports on the phagocytosis in Candida albicans (Bain et al., 2014 and Lewis et al., 2012), may act as diffusion barriers and contribute to membrane fences that allow the accumulation of ROS in the frustrated phagosomes. They also propose that these actin cuffs are the product of the atypical activation of MAC1receptors in the macrophages by ligands expressed on the cell wall of Candida and that MAC1 may restrict the free diffusion of lipids and membrane proteins along the membrane. The proposed model is sound, and it may explain possible microbicidal actions phagocytes may attempt when pathogens cannot be completely engulfed. However, there are many important points in the manuscript that require clarification and revision to better support the authors' model.

The authors claim that the frustrated phagosomes are microbicidal. Yet, there is no data on fungal survival to support this conclusion. Are the macrophage-trapped hyphae growing longer or dying as time passes?

The contribution of Candida hyphae to the formation of the frustrated phagosomes and actin cuffs must be assessed. Changes in Candida cell wall composition have been previously reported to affect macrophage actin dynamics (Bain et al., 2014). However, the authors are not considering or discarding a role for the yeast in the proposed model.

For the imaging data, even though the regions of interest are nicely indicated with brackets, since different markers don't always localize to same regions along the frustrated phagosome, having additional panels with channels merged would make it easier to visualize what the authors are illustrating.

In subsection “Signals driving actin cuff formation”, referring to Figure 2AB, the authors indicate, a lack of Dectin-1 at the actin cuffs. However Emerald-Dectin-1 can be seen in the cuff in Figure 2B, which is in good agreement with Strijbis et al., (2013) that also showed that Dectin-1 is enriched in the cuff region of the phagocytic cup, as well as in areas of enrichment outside the cuff region. To support their claim, the authors must present quantitative data on Dectin-1 distribution along the frustrated phagosome. Also, it is not indicated, but the imaging shown in Figure 2, and other figures, seems to be a single confocal z-plane. It may also be appropriate to include extended focus projections, considering the size of the hyphae, which could be included as supplementary files.

Clarifying the distribution of Dectin1 in the frustrated phagosomes, is critical because it affects the interpretation of data in Figure 2D and other results in the manuscript.

Figure 2D showing PAK-PBD and tyrosine phosphorylation in the actin cuffs are likely expected results since Strijbis et al., 2013, have shown that the GEF for CDC42 and RAC1, VAV1, as well as the tyrosine kinases Syk and BTK, are all recruited to cuffs. Quantitative data for the recruitment of these molecules to the frustrated phagosomes and data proving that their activities are involved in the formation or sustaining the cuffs will certainly help the proposed model.

Figure 3D shows that on average, for a given field, 6 hyphae are observed in frustrated phagosomes with actin cuffs and this is consistent throughout the manuscript. However, it is not clear how many individual events were analyzed. How frequently are these frustrated phagosomes observed? Do all hyphae engaged by macrophages have these structures associated with them?

Figure 4. For accumulation of receptors provide quantitative data. According to the authors, MAC1 activation signals for the formation of the actin cuffs. However, in Figure 4A, CD11b did not localize to the thicker section of the cuffs. Please clarify and indicate the frequency of this phenotype.

Figure 4G shows a shorter cup in the presence of CD11b blocking antibodies. The treatment could cause an inhibition of phagocytosis. Authors must demonstrate that phagocytosis is not affected by the treatment.

Figure 4G/H In cells treated with the M1/70 antibody, actin is still present in the frustrated phagosome. Is this a cuff? In fact, Concavalin A labeling is limited to the phalloidin boundary. This suggests the presence of a cuff/ barrier, even after CD11b treatment.

Is dectin1 distribution along the frustrated phagosome affected after CD11b is blocked? Blocking CD11b in dectin1 knockout cells will help address whether dectin1 contributes to this barrier.

Figure 5, subsection “Phospholipid segregation between the plasma membrane and the cuff-delimited phagosomal cup” states "Astonishingly, while PtdIns(4,5)P2 was present as expected in the surface membrane facing the extracellular milieu, it was undetectable in the invaginated section that constituted the frustrated phagosome (Figure 5A)". This and other characteristics of the cuff and the frustrated phagosomes reported in Figure 5 and Figure 8 are similar to those reported by Prashar et al., (2013). This must be acknowledged in the manuscript. It must be acknowledged that PtdIns(3,4,5)P3 distribution in the frustrated phagosome has been previously described by Strijbis et al. In Figure 5D, how do the authors explain the presence of LC3 in the actin cuff?

Subsection “The actin cuff forms a diffusional barrier to the movement of proteins and lipids” "The sharp boundary between the PtdIns(4,5)P2-rich surface membrane and the tubular membrane endowed with PtdIns(3,4,5)P3 and PtdIns(Bauer et al., 2001)P coincided with the location of the actin cuff, suggesting that the latter may function as a diffusion barrier".

Subsection “The actin cuff forms a diffusional barrier to the movement of proteins and lipids” Similarly, both wild-type Rab7 (Figure 5E) and constitutively-active Rab7 (not illustrated) are confined to the frustrated phagosomal tube, as was LAMP1 (Figure 5F)……..we considered it more likely that restricted diffusion accounted for the observations"

Differently to what the authors describe in the paragraph from above, the phagosomal markers are penetrating the barrier as in the case of Figure 5B and E and surpassing the cuffs in Figure 5C, E and F. As indicated before for other results the authors must present quantitative data on the distribution of the markers and extended focus projections to support the proposed model.

Figure 7 strongly supports authors' claim However, if latranculin A treatment allowed Lamp-1 and PtdIns(4,5)P2 to migrate to previously excluded zones, why concanavalin A is not following and labels the full extension of the hyphae? That will be the expected if the diffusion barriers in the cuffs are dismantled. Quantitative data is missing, and full cell imaging would be preferable to better illustrate the authors' claim.

Figure 8 The characteristics of the frustrated phagosome are remarkably similar to long cups described by Prasher et al., 2013 and Strijbis et al., 2003. This must be cited in the text. The accumulation of NBT in Figure 8E is puzzling since small molecules can diffuse out across the actin cuff barrier. The authors must address the possibility that the NBT is accumulating inside the hyphae. NBT can accumulate in fungi Camile et al., 2008. Thus, authors must prove a link between ROS and microbicidal conditions in the long open phagosomes.

Reviewer #3:

The manuscript by Maxson et al., describes and analyzes partially sealed phagocytic cups in macrophages which form around Candida albicans hyphae. It demonstrates that an actin-rich cuff at the distal margin of the cup supports a lateral segregation between the components of the plasma membrane and the contiguous inner leaflet of the phagocytic cup. This lateral segregation excludes the PI(4,5)P2 from the cup and confines the PI(Bauer et al., 2001)P and PIP3 to the cup. The actin cuff contains the integrin CR3 and associated integrin signaling molecules. The barrier effectively limits the movement of inner leaflet probe molecules into or out of the cup, without significantly diminishing diffusion within the plasma membrane or the cup. The actin cuff can be disrupted by the actin depolymerizing drug latrunculin A. The barrier limits escape by diffusion of large macromolecules delivered into the lumen of the cup, but not the diffusive loss of smaller molecules, including dextrans and protons (pH). Nonetheless, reactive oxygen species can be delivered into the unclosed cups. The experimental work is carefully done and the morphological evidence in support of the claims is beautiful. However, much if not most of the conclusions reported here were first described in a large study by Prashar et al., (2013), using an analogous experimental model. Examining the interactions between macrophages and filamentous bacteria, that study demonstrated (Astarie-Dequeker et al., 1999) the actin cuff (called a "jacket") that segregates two domains of contiguous membrane (plasma membrane and phagocytic cup), (Bain et al., 2014) the effect of actin depolymerization on the maintenance of that segregation, (Bauer et al., 2001) the nature of the diffusion barrier with respect to extracellular probes and (Belin et al., 2014) the retention of diffusible molecules in the cup lumen (different sizes of dextrans, pH, hydrolytic enzymes). Moreover, unlike the present manuscript which shows that ROS can be generated in the unclosed cups, the previous paper measured the effect of partial phagocytosis on microbicidal activities against filamentous bacteria (albeit somewhat incompletely). The present manuscript but does not measure microbicidal activity against C. albicans.

Assuming this manuscript can be rewritten to acknowledge appropriately the demonstrated precedents and concepts of that earlier work, the present study does add some interesting new data which supports a role for integrin in the establishment or maintenance of the actin cuff. The role of integrin in the maintenance of the barrier remains indirect, however. Perhaps the actin ring is the main ingredient of the barrier, and anything which organizes actin into a ring can support the formation of diffusion barriers such as described here, and elsewhere for analogous structures (Golebiewska et al., 2011; Welliver et al., 2011. To better define the nature of the diffusion barrier, the diffusion measurements (e.g. Figure 6) or probe localization studies (e.g. Figure 5) described in the manuscript should be extended to analyze the roles of actin (latrunculin A) and integrin (M1/70) to barrier maintenance. This could provide a mechanistic underpinning to the diffusion measurements.

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

Thank you for resubmitting your work entitled "Integrin-based diffusion barrier separates membrane domains enabling formation of microbiostatic frustrated phagosomes" for further consideration at eLife. Your revised article has been favorably evaluated by Ivan Dikic (Senior editor), a Reviewing editor, and three reviewers.

This revised manuscript contains extensive additional experimental work and text that address concerns noted in earlier reviews. The new work is thorough and adequately supportive of the conclusions. The manuscript now establishes a diffusion barrier in the inner leaflet of incompletely closed phagocytic cup membranes, comprised of CR3 integrins and maintained by an actin-rich cuff. Further, it identifies signaling molecules that contribute to cuff formation, and establishes the incompletely closed cups as microbiostatic. Related studies are adequately cited. The discussion provides a thoughtful analysis of the biology and its implications. Yet there are some minor remaining issues that need to be addressed in the text before acceptance, as outlined below:

The authors addressed all the critiques from the reviewers.

They reported additional results on the phagocytosis of C. albicans hyphae, improved the quality of the imaging, and provided numerical and statistical data to support their most relevant experiments.

However, below there are several points that we believe could be discussed in the manuscript:

1) In the abstract it is stated “[…]in response to non-canonical activation of integrins by fungal β(1,3)-glucans." We suggest using the singular "glycan". There is more than one form of fungal β(1,3)- glucan, but this is not relevant to this article.

2) B-glucans are involved in the activation of complement and the deposition of C3b fragments on the surface of C. albicans. (Boxx et al., 2010., Kozel et al., 1987, Vetvicka et al., 1996). What could be the contribution of C3b opsonization on the formation of the actin cuffs and the properties of the membrane and luminal diffusion barriers?

3) Previous reports on the phagocytosis of C. albicans showed the formation of PIPs in phagosomes containing this yeast (Heinsbroek, 2009). This was attributed to pathogenic mechanisms. On the other hand, and coinciding with the authors' interpretation, Naufer et al., 2018 recently reported that PI(Bauer et al., 2001)P co-exists with phagolysosomal markers in the phagocytic cup of heat-killed filamentous bacteria. Maybe the authors could mention this in their Discussion section.

4) Please clarify: In Figure 4, human serum was used to complement opsonize hyphae. How did the authors account for the presence of antibodies against C. albincans in the serum? Antibodies could be expected as this yeast is part of the human microbiome

5) Caspofungin treatment could prevent the deposition of complement in hyphae, and its recognition by antibodies and Dectin-1. This could explain the results in Figure 4—figure supplement 1.

6) The authors showed that ROS production in the phagocytic cup retarded the elongation of the hyphae. This was alleviated by blocking CR3 with monoclonal antibodies to impede the formation of cuffs, a procedure that favours a ROS leaching. Since ROS are generally considered microbicidal, perhaps, ROS failing to reach lethal concentrations in the open cup could be the cause of the effect reported by the authors. Therefore, assuming a microbiostatic mechanism is probably incorrect.

https://doi.org/10.7554/eLife.34798.041

Author response

[Editors’ note: the author responses to the first round of peer review follow.]

Reviewer #1:

I have a number of questions, suggestions and presentational issues.

1) The fungus studied is Candida albicans but is most commonly referred to as Candida. It would not be simply pedantic to request that it be referred to as C. albicans throughout since most Candida species cannot form elongated hyphae.

As requested, we have changed our notation to C. albicans throughout the text, and only refer to the BFP-expressing C. albicans as Candida-BFP to preserve the original nomenclature coined by the authors that generated this strain (Strijbis, 2013).

2) It would be interesting to know if similar actin cuff seals are formed when phagocytes encounter hyphae of Aspergillus fumigatus (interesting because this fungus has quite marked differences in cell wall composition).

As suggested by the reviewer, we performed new experiments in collaboration with the laboratory of Dr. Leah Cowen (who is now included as a co-author) using Aspergillus fumigatus. The new data are shown in the revised Figure 4G and discussed in the Results section. Like others before, we observed that A. fumigatus is not effectively internalized by macrophages, due in part to the presence of cell wall glycosaminoglycans and also because it produces gliotoxin, an effective inhibitor of phagocytosis. We minimized the effects of gliotoxin (that reacts with sulfhydryl groups, forming mixed disulfides) by exposing the cells to Aspergillus fumigatus in the presence of L-cysteine (see Materials and methods section). Under these conditions, when the macrophages engage the hyphae near the end, we were able to observe the formation of distinct actin cuffs, resembling those formed around C. albicans hyphae, consistent with the fact that the cell wall of A. fumigatus contains β(1,3)-glucans.

3) The hyphae used are not very long (1 h or 2 h hyphae were used) and the macrophages shown seem to be invaginating one end. Macrophages will also wrap around the trunk of a very large hypha. Are actin cuffs formed here- and what is their architecture? It would be useful to include some images of longer hyphae of 4-6 hours incubation in RPMI.

Based on this suggestion, we conducted experiments using longer C. albicans hyphae (allowed to grow for 4 hours), and also observed the formation of actin cuffs when these were engaged near the end. A representative image is shown in Author response image 1 (top row). In the case of the longer hyphae, as was also the case with the very long Aspergillus hyphae, the macrophages often interacted with intermediate regions of the hyphae. In these instances, the macrophages partially wrapped themselves around the hyphae, forming dense actin structures that did not seem to surround the hyphae in their entirety (we call them “tacos”; see Author response image 1, bottom row, for representative image). Because these seemingly unsealed structures would not function as diffusion barriers –the main topic of the paper– we have opted not to illustrate or discuss them but would be happy to do so if the reviewer felt this is important.

4) Various C. albicans mutants are available with alterations in the cell wall that would help determine what surface components of the fungus are important for inducing the cuffs.

As recommended by the reviewer we have conducted an extensive series of experiments in collaboration with Drs. L. Cowen and T. O’Meara to define the C. albicans cell wall components required for actin cuff formation. To this end we utilized a curated set of C. albicans GRACE strain mutants generously made available by Merck and Genome Canada, which filament normally but have mutations affecting biosynthetic pathways for chitin, mannans or β(1,6)glucans (see O’Meara et al., 2015). The new results are now illustrated in Figure 4C and D and described in the Results section. To assess the role of β(1,3)-glucans, we pharmacologically inhibited β(1,3)-glucan synthesis using caspofungin. These results are presented in the new Figures 4E and F and discussed in the Results section. In a nutshell, we found that β(1,3)-glucan is required for actin cuff formation, while chitin, mannans and β(1,6)-glucans were dispensable. Please note that we also utilized caspofungin in Figure 4—Figure supplement 1, discussed in the Results section, to distinguish the role of β(1,3)-glucan in actin cuff formation from its role in Dectin1-mediated phagocytosis.

5) In many of the figures it is not possible to clearly evaluate the shape and position of the phagocyte relative to the fungus. It would be very helpful to show a DIC or phase image of the same interacting cells in order to really see the orientation of the hypha and phagocyte. I have no doubt that the descriptions in the text are accurate, but I have to somewhat take it on trust since I cannot see the outline of the two interacting cells clearly in all cases. (For example see Figure 3B,C; Figure 4B,E,F; Figure 5B,C,F; Figure 6E; Figure 8A,D and others).

As requested by the reviewer, to enable the reader to locate the phagocytes with respect to the hyphae, we have outlined the macrophages using grey dotted lines (see Figure 1, Figure 1—figure supplement 1, Figure 2, Figure 3, Figure 3—figure supplement 1, Figure 4, Figure 5, Figure 5—figure supplement 1, Figure 6 and Figure 10), except in those cases where the cell outline is readily discernible as background fluorescence.

6) The authors show that the CTL Dectin-1 is not present at the actin cuff, but phosphotyrosine accumulated at this site suggesting that some other pattern recognition receptor may drive actin assembly. It would be useful to verify that glucan phosphate or laminarin did not block actin cuff formation.

Considering this comment, together with comment #7 below, we performed additional experiments to characterize the properties of the receptor-ligand interaction involved. As suggested, we used laminarin to block Dectin1 and the new results have been included in the revised Figure 4B. When added before the addition of C. albicans, laminarin impaired phagocytosis, consistent with the involvement of Dectin1 in the initial stages of the interaction, which was validated by comparing RAW cells (that express little Dectin1) with RAW-Dectin1 cells, which are stably transfected to express higher levels of Dectin1 (new Figure 4A). However, when added after C. albicans adherence, laminarin had no effect on actin cuff formation (Figure 4B). As discussed in the manuscript, we believe that at this later stage the integrins, specifically CR3, have been activated and become the main drivers of the association and of the formation of the actin cuff.

7) Since the mannose receptor is likely to engage with hypha mannan, (which is located in the outer cell wall) it would be interesting to know if this is located at this site.

The possible involvement of mannan receptors was tested in several ways. First, as mentioned above, we used mutant strains that are mannan-deficient. Secondly, we tested the effects soluble mannans on the internalization of C. albicans and the formation of actin cuffs and have included these data in the new Figure 4B. In accordance with the mutant data, we found soluble mannan to be without effect. Lastly, we immunostained for mannose receptors using antibody MR5D3 (Bio-Rad; catalogue # MCA2235GA) and found no evidence of accumulation at the cuff. These new results are described in subsection “Role of receptor cooperativity in actin cuff formation” of the revised text.

8) I'm not sure that the TEM shown in Figure 1C can be seen as evidence of showing the actin cuff unless the actin is perhaps stained using colloidal gold.

We agree that the electron micrographs do not provide direct evidence that actin is accumulated in the region of cuff, a case much better made by the phalloidin staining that is profusely illustrated throughout the paper. We clearly overstated the case in the original version and have toned down the interpretation of the images in the revised text. Instead, we refer to the similar exclusion area seen by Strijbis et al., (2013), who used tannic acid negative staining to better visualize actin (see subsection “Phagocytosis of C. albicans hyphae” of revised text).

9) My88-/- cells would be useful in assessing the possible involvement of TLRs.

The involvement of TLRs/MyD88 in the recognition of and control of C. albicans by macrophages is well documented. Indeed, Marr et al., (2003) showed that MyD88−/− cells have a pronounced phagocytic defect. While reflecting the importance of the cooperation between MyD88/TLR and Dectin1, this phagocytic defect would complicate performing the suggested experiments using MyD88−/− cells, as well as their interpretation, since it would be difficult to separate the contribution of MyD88 to actin cuff formation from that to Dectin1-mediated phagocytosis. While we have included additional experiments to dissect the contributions of Dectin1 and CR3, we feel that additional studies of MyD88 and TLRs would be outside the scope of this manuscript.

10) The actin cuffs appear to be elongated in some images and much shorter or almost like spots in others. Do they retain their shape over time or contract to a spot?

It is our observation that the actin cuffs can vary somewhat in length and continuity from cell to cell, sometimes fragmented into two or more smaller cuffs. However, once established in any single cell, the actin cuff does not change shape or contract significantly after formation for up to 90 min (the duration of our experiments), despite the fact that actin treadmilling is ongoing at the cuff. We refer the reviewer to subsection “Phagocytosis of C. albicans hyphae” of the revised text, where these observations are described.

Reviewer #2:

The authors claim that the frustrated phagosomes are microbicidal. Yet, there is no data on fungal survival to support this conclusion. Are the macrophage-trapped hyphae growing longer or dying as time passes?

This point was also raised, equally aptly, by reviewer 3. To address this issue, we conducted new experiments to assess the antimicrobial effects of the actin cuff on C. albicans hyphae. We initially measured the viability of the hyphae that had been partially internalized using propidium iodide. As is now described in the Results section, we did not see any change in fungal viability in the partially internalized C. albicans hypha (nor in fully internalized hyphae, for that matter; data not shown) for at least 60 min, the normal duration of our experiments. We reasoned that the antimicrobial effectors may not suffice to kill the fungus yet may curtail hyphal growth. The remarkable extension rate of C. albicans hyphae, which has been well-documented (18.8 µm/hr = 0.31 µm/min, Gow, 1985), enables accurate quantitation during the course of our experiments. We therefore, compared the rate of growth of partially internalized hyphae with an actin cuff with that of hyphae that were fully internalized or that were not engaged by macrophages (i.e. remained extracellular) throughout. These results are now summarized in the new Figure 10F and described in subsection “Functional properties of the frustrated phagosome” of the revised text. Briefly, we found that sequestration of a part of the hyphae within the frustrated phagosomes delimited by the cuff reduced the rate of growth considerably, to the same extent as seen for hyphae fully internalized within a sealed phagosome. Importantly, blocking of CD11b with the M1/70 antibody abolished this microbiostatic effect (Figure 10—figure supplement 2). In light of this newly characterized effect on hyphal growth, we have changed the title from “microbicidal” to “microbiostatic”.

The contribution of Candida hyphae to the formation of the frustrated phagosomes and actin cuffs must be assessed. Changes in Candida cell wall composition have been previously reported to affect macrophage actin dynamics (Bain et al., 2014). However, the authors are not considering or discarding a role for the yeast in the proposed model.

As described above when addressing comment #4 of reviewer 1, we performed a new series of experiments to define the cell wall components required for actin cuff formation, using a curated set of C. albicans GRACE strain mutants obtained from Merck and from Genome Canada, which have mutations affecting biosynthetic pathways for chitin, mannans or β(1,6)glucans (see O’Meara et al., 2015). We also conducted pharmacological experiments targeting the formation of the β(1,3)-glucan bond, using caspofungin. In a nutshell, we found that β(1,3)glucan is required for actin cuff formation, while chitin, mannans and β(1,6)-glucans were dispensable. Please see the new Figure 4 (Figure 4C, D, E and F, Figure 4—figure supplement 1) and the respective text in subsection “Fungal cell wall components that contribute to actin cuff formation” of the revised manuscript for details.

For the imaging data, even though the regions of interest are nicely indicated with brackets, since different markers don't always localize to same regions along the frustrated phagosome, having additional panels with channels merged would make it easier to visualize what the authors are illustrating.

As suggested, to ease interpretation of the images, we have added ~ 2x merged insets of the actin cuff region, where the colocalization of various markers is indicated as yellow. See revised Figure 2, Figure 3, Figure 5, Figure 6 and Figure legends. In addition, we now provide quantitation of specific marker enrichment/exclusion in the actin cuff, calculated as the ratio of fluorescence in the cuff vs. the phagocytic cup (Results section, and Materials and methods section of the revised manuscript). Finally, we have added new 3D reconstructions to aid the viewer in visualizing the shape of the cuff and its relationship to the extracellular portions of the hyphae and to the phagosomal marker LAMP1 (see Figure 6G and H). Rotational views and progressive deconstruction of the images are provided in accompanying Video 2.

In subsection “Signals driving actin cuff formation”, referring to Figure 2AB, the authors indicate, a lack of Dectin-1 at the actin cuffs. However Emerald-Dectin-1 can be seen in the cuff in Figure 2B, which is in good agreement with Strijbis et al., (2013) that also showed that Dectin-1 is enriched in the cuff region of the phagocytic cup, as well as in areas of enrichment outside the cuff region. To support their claim, the authors must present quantitative data on Dectin-1 distribution along the frustrated phagosome.

As requested by the reviewer, we have quantitated the ratio of Dectin1-HA and Emerald-Dectin1 in the cuff compared to the cup; this quantitation shows that the receptor is indeed partially excluded from the cuff (see Results section). Moreover, we now provide insets to Figure 2A and B showing the localization of Dectin1 and merged images showing also the actin cuff. Strijbis et al., (2013) did assess Dectin1 localization and found that “Dectin1 was enriched in the cuff region of the phagocytic cuff….but also showed areas of enrichment outside the cuff region”. We believe that this is consistent with our observations that Dectin1 is present throughout the phagocytic cup but is not enriched in the area of the actin cuff, relative to the rest of the frustrated phagocytic cup. Additionally, please see point 5 below for further discussion of the localization of Dectin1.

Also, it is not indicated, but the imaging shown in Figure 2, and other figures, seems to be a single confocal z-plane. It may also be appropriate to include extended focus projections, considering the size of the hyphae, which could be included as supplementary files.

Clarifying the distribution of Dectin1 in the frustrated phagosomes, is critical because it affects the interpretation of data in Figure 2D and other results in the manuscript.

We agree that clarifying the role of Dectin1 is important. To this end, we have added to Figure 2A and B insets showing the merged images and dotted outlines of the cells and have included the quantitation of Dectin1 density described above (see Results section). Moreover, as described under comment #3, we now include 3-dimensional reconstructions and videos of the cuff and its relationship to other components. We also attach Author response image 2, the extended focus projections of the confocal images illustrated as Figures 2A and B, for the reviewer’s perusal. We do not feel that inclusion of such extended focus images would be more informative but would certainly include them as part of Figure 2—figure supplement 1 if the reviewer feels this is useful.

Figure 2D showing PAK-PBD and tyrosine phosphorylation in the actin cuffs are likely expected results since Strijbis et al., 2013, have shown that the GEF for CDC42 and RAC1, VAV1, as well as the tyrosine kinases Syk and BTK, are all recruited to cuffs. Quantitative data for the recruitment of these molecules to the frustrated phagosomes and data proving that their activities are involved in the formation or sustaining the cuffs will certainly help the proposed model.

As suggested by the reviewer, we have quantitated the enrichment of PAK(PBD) and phosphotyrosine in the actin cuff region (see Results section and Materials and methods section). Additionally, we present a new Figure 5 where the contribution of distinct tyrosine kinases to actin cuff formation was assessed in the context of the frustrated phagosome anchored by CR3.

This extends and complements the work of Strijbis et al., (2013), who first described the role of Syk and BTK in the context of Dectin1-mediated phagocytosis. To this end, we used immunostaining to assess the activation of several candidate tyrosine kinases. Specifically, we have detected activation (phosphorylation) of Syk and, interestingly, PYK2/FAK kinases at the actin cuff (see Figure 5, and subsection “Signals driving actin cuff formation” of the text for quantitation). To our knowledge, PYK2 and FAK were previously not known to participate in the host response to C. albicans. These observations prompted us to assess the functional requirement of these kinases for cuff formation and maintenance. As illustrated in the new Figure 5E, inhibition of PYK2/FAK obliterated the formation of actin cuffs, as did inhibition of Syk. It is noteworthy that, by contrast, Src-family kinase inhibitors were without effect.

To extend the findings of Strijbis et al., (2013) regarding actin polymerization, we performed new experiments assessing the contribution of Arp2/3- vs. formin-dependent actin assembly to actin cuff formation, since both can be driven by active Rac1 and Cdc42 GTPases. As shown in the new Figure 5G we found that formins are the primary driver of actin cuff formation. These data provide the basis of a more complete model of cuff formation and are consistent with known pathways driving actin polymerization after engagement of CR3 (see Results section and Discussion section).

Figure 3D shows that on average, for a given field, 6 hyphae are observed in frustrated phagosomes with actin cuffs and this is consistent throughout the manuscript. However, it is not clear how many individual events were analyzed. How frequently are these frustrated phagosomes observed? Do all hyphae engaged by macrophages have these structures associated with them?

In accordance with the reviewer’s request we have included quantitation of the frequency of actin cuff occurrences in the text (see Results section). As mentioned in the revised text, under the conditions used, partially internalized hyphae occur at a frequency of 68.5%, and of those 96.3% have actin cuffs. In addition, we now provide the average number of C. albicans hyphae visualized per field, to accompany the graphical data (see Figure legends), and have amended all graphical figures to show the number of C. albicans with an actin cuff, along with the number of fully internalized hyphae, to better reflect the total number of C. albicans events analyzed per field (see Figure 2, Figure 3, Figure 4 and Figure 5).

Figure 4. For accumulation of receptors provide quantitative data. According to the authors, MAC1 activation signals for the formation of the actin cuffs. However, in Figure 4A, CD11b did not localize to the thicker section of the cuffs. Please clarify and indicate the frequency of this phenotype.

To provide further clarity as to the localization of CD11b and CD18, we now provide merged ~2x insets showing the colocalization (in yellow) of CD11b and CD18 with the actin cuff (see Figure 3A and B), and we have quantitated the enrichment of CD11b and CD18 in the actin cuff region (see Results section and Materials and methods section for calculations). Note that, in other integrin-induced structures such as stress fibres and podosomes, the regions of greatest actin accumulation are located at some distance from the active integrins themselves. It is also possible that access to the epitope (which is exofacial in both instances) may be restricted by the close apposition of the host and pathogen surfaces, particularly in those areas where the actin cuff may constrict the neck.

Figure 4G shows a shorter cup in the presence of CD11b blocking antibodies. The treatment could cause an inhibition of phagocytosis. Authors must demonstrate that phagocytosis is not affected by the treatment.

As the reviewer has requested, we have amended the former Figure 4H (now Figure 3H) to include the number of fully internalized C. albicans, in addition to those that are partially internalized and therefore display actin cuffs. As the new figure indicates, the number of fully internalized hyphae, a process largely mediated by Dectin1, was not significantly affected. The decrease in the number of actin cuffs seen upon inhibition of CD11b was accompanied by a reciprocal increase in the number of partially internalized hyphae that did not display actin cuffs, again implying that engagement of the hyphae and initiation of phagocytosis did not require CR3. To simplify the graph in Figure 3H, the number partially internalized without actin cuffs has not been included but is referred to in the text (see Results section).

Figure 4G/H In cells treated with the M1/70 antibody, actin is still present in the frustrated phagosome. Is this a cuff? In fact, Concavalin A labeling is limited to the phalloidin boundary. This suggests the presence of a cuff/ barrier, even after CD11b treatment.

The original Figure 4G (now Figure 3G) is representative of the observed disruption of the actin cuff around partially internalized C. albicans hyphae. As should be apparent, the amount of actin is greatly reduced and, more importantly, the entire cup is lined by actin, unlike the discrete distribution at the cuff, which is now clearly illustrated in the 3-dimensional images shown in Figures 1E-H and Video 1. It is important to note that preferential labeling of the extracellular portions of the hyphae by Concanavalin A (ConcA) is not predicated on the barrier function of the integrin/actin cuff. Indeed, the Stokes radius of ConcA, reported as 3.0 nm (Sawyer et al., 1975; Ahmad et al., 2007) is similar that of 10 kDa dextran (2.3 nm), and considerably smaller than that of the 100 kDa dextran (>6 nm), which was shown not to traverse the junction. It is the result of the comparatively brief (20 min) incubation at reduced (room) temperature with the lectin, which favors readily detectable binding to exposed carbohydrates, while minimally staining those inside the cup, where diffusion is limited by the narrow spacing between the membrane and hyphal wall. Moreover, ConcA staining is done after fixation with paraformaldehyde. Paraformaldehyde crosslinking likely restricts further the entry of Concanavalin A to the regions trapped in the cup. Nevertheless, to address the effectiveness of the block exerted by antibody M1/70, we took advantage of the observation that formation of the cuff impaired the growth of the hyphae. We performed new experiments where the rate of growth was compared in hyphae that were partially internalized by cells in the presence and absence of antibody M1/70. The new data, which are presented in Figure 10—figure supplement 2, show that blocking the engagement of CR3 eliminated the microbiostatic effect of the macrophages, i.e. the hyphae grew at normal rates in M1/70-treated cells. See response to comment #2 by reviewer 3 for further description of these data.

Is dectin1 distribution along the frustrated phagosome affected after CD11b is blocked? Blocking CD11b in dectin1 knockout cells will help address whether dectin1 contributes to this barrier.

As requested by this reviewer, as well as by reviewer 1, we performed additional experiments to assess the role of Dectin1 in actin cuff formation. We took advantage of the earlier observation that Dectin1 is essential for C. albicans internalization(Taylor et al., 2007, Marakalala et al., 2013 –these references have now been cited in subsection “Role of receptor cooperativity in actin cuff formation”). Based on this knowledge we designed experiments comparing the Dectin1-deficient (parental) RAW 264.7 cell line with the RAW-Dectin1 stable transfectants. Importantly, while cuff formation was minimal in the Dectin1 deficient cells, we could bypass the need for the glucan receptor using complement opsonization (see new Figure 4A, discussed in subsection “Role of receptor cooperativity in actin cuff formation” of the revised text). We interpret these results to mean that Dectin1 is required for initial engagement and initiation of phagocytosis, a role that can be alternatively fulfilled by the traditional integrin-binding domain of CR3 yet is not essential for cuff formation. Because in the absence of complement phagocytosis will not occur in Dectin1 knockout cells, the blocking experiment suggested by the reviewer cannot be performed. Instead, we timed the inhibition of Dectin1 using laminarin, as recommended by reviewer 1. When added before phagocytosis, laminarin greatly inhibited cuff formation, as expected, but it was ineffective when added after the integrin was already engaged (Figure 4B). Note that size of the laminarin used (≈500 Da) was sufficiently small for it to penetrate the junction between the macrophage and the hyphae, based on the data of Figure 10.

Figure 5, subsection “Phospholipid segregation between the plasma membrane and the cuff-delimited phagosomal cup” states "Astonishingly, while PtdIns(4,5)P2 was present as expected in the surface membrane facing the extracellular milieu, it was undetectable in the invaginated section that constituted the frustrated phagosome (Figure 5A)". This and other characteristics of the cuff and the frustrated phagosomes reported in Figure 5 and Figure 8 are similar to those reported by Prashar et al., (2013). This must be acknowledged in the manuscript. It must be acknowledged that PtdIns(3,4,5)P3 distribution in the frustrated phagosome has been previously described by Strijbis et al.

At the reviewer’s request, we have cited these references in the appropriate context in several places in the text. Please see Results section and Discussion section).

In Figure 5D, how do the authors explain the presence of LC3 in the actin cuff?

LC3 has been reported to insert into the phagocytic cup during LC3-assisted phagocytosis (see Martinez et al., 2015 reference in Results section), a maturation pathway that has been implicated as important for the control of fungus, including C. albicans (Kanayama et al., 2016, Tam et al., 2016, Sprenkeler et al., 2016 –references now added to the text, see Results section). We chose to analyze its distribution because it is lipid-anchored yet can be covalently labeled by a fluorescent protein. The precise mechanism that directs LC3 to phagosomes is not known and, most importantly, we do not know whether these determinants are deployed in the area of the membrane where the integrins generate the cuff, which would readily explain its presence there. Most importantly, as discussed in more detail below, the diffusion barrier may slow down the movement of lipids, lipid-anchored proteins and proteins markedly, without necessarily preventing their entry into the cuff altogether. In this instance, the much faster diffusion of these molecules after they exit the cuff would make them undetectable outside the cup.

Subsection “The actin cuff forms a diffusional barrier to the movement of proteins and lipids” "The sharp boundary between the PtdIns(4,5)P2-rich surface membrane and the tubular membrane endowed with PtdIns(3,4,5)P3 and PtdIns(Bauer et al., 2001)P coincided with the location of the actin cuff, suggesting that the latter may function as a diffusion barrier".

Subsection “The actin cuff forms a diffusional barrier to the movement of proteins and lipids” Similarly, both wild-type Rab7 (Figure 5E) and constitutively-active Rab7 (not illustrated) are confined to the frustrated phagosomal tube, as was LAMP1 (Figure 5F)[…]we considered it more likely that restricted diffusion accounted for the observations"

Differently to what the authors describe in the paragraph from above, the phagosomal markers are penetrating the barrier as in the case of Figure 5B and E and surpassing the cuffs in Figure 5C, E and F. As indicated before for other results the authors must present quantitative data on the distribution of the markers and extended focus projections to support the proposed model.

This is an important issue that indeed required clarification. First, to more clearly illustrate the relative disposition of the markers vis-a-vis the actin cuff, we have added ~ 2x merged insets of the cuff region where the colocalization of various markers with actin is indicated in yellow (see revised Figure 6). As we described in the original version, the boundary of the lipids coincided with the location of the actin band, although in the case of PtdIns(4,5)P2 the inositide is restricted to the plasmalemma, while PtdIns(3,4,5)P3 is both in the cup and throughout the cuff. It is quite likely that PI3-kinase is active in the region where the integrins and active Syk and PYK2/FAK accumulate, so that PtdIns(3,4,5)P3 could be generated within the cuff itself. As stated above, in the case of the lipids in particular, it is likely that the barrier reduces the diffusion rate of the lipids considerably, without necessarily preventing their entrance into the region of the cuff altogether. The limited amounts of PtdIns(4,5)P2 entering the cuff would be converted to PtdIns(3,4,5)P3 and/or hydrolyzed by PLC. This, we believe, accounts for the differential appearance of the two inositides.

Also, as requested, we have now provided quantitation of marker enrichment/exclusion in the actin cuff, calculated as the ratio of fluorescence in the cuff vs. phagocytic cup (see Results section and Materials and methods section). Based on these measurements we have modified the wording of that section as follows:

“Similarly, both wild-type Rab7 (Figure 6E) and constitutively-active Rab7 (not illustrated) are confined to the frustrated phagosomal tube and partially excluded from the actin cuff (Rab7 ratio cuff: cup 0.68 ± 0.05; n=30, p<0.0001), as was LAMP1 (ratio cuff: cup 0.59 ± 0.03; n=30, p<0.0001; Figure 6F)”.

The partial exclusion of the marker proteins is consistent with molecular crowding caused by integrin accumulation at the cuff, which we propose causes reduced penetration and diffusion within the area of the cuff.

Figure 7 strongly supports authors' claim However, if latranculin A treatment allowed Lamp-1 and PtdIns(4,5)P2 to migrate to previously excluded zones, why concanavalin A is not following and labels the full extension of the hyphae? That will be the expected if the diffusion barriers in the cuffs are dismantled. Quantitative data is missing, and full cell imaging would be preferable to better illustrate the authors' claim.

As we described above in response to comment #9 by reviewer 2, preferential labeling of the extracellular portions of the hyphae by Concanavalin A (ConcA) is not predicated on the barrier function of the integrin/actin cuff. It is the result of the comparatively brief (20 min) incubation at reduced (room) temperature with the lectin, which favors readily detectable binding to exposed carbohydrates, while minimally staining those inside the cup, where diffusion is limited by the narrow spacing between the membrane and hyphal wall. Moreover, ConcA staining is done after fixation with paraformaldehyde. Paraformaldehyde crosslinking likely restricts further the entry of Concanavalin A to the regions trapped in the cup.

As requested, we have included quantitative data showing the effects of latrunculin A on the distribution of PLCδ-PH-GFP and Lamp1-GFP (new Figures 9C and F), supporting our observations depicted in the representative figures (Figures 9A, B, D, E).

Figure 8 The characteristics of the frustrated phagosome are remarkably similar to long cups described by Prasher et al., 2013 and Strijbis et al., 2003. This must be cited in the text.

In accordance with the reviewer’s request, we have cites these references in several places in the revised manuscript (see Results section and Discussion section).

The accumulation of NBT in Figure 8E is puzzling since small molecules can diffuse out across the actin cuff barrier. The authors must address the possibility that the NBT is accumulating inside the hyphae. NBT can accumulate in fungi Camile et al., 2008. Thus, authors must prove a link between ROS and microbicidal conditions in the long open phagosomes.

We were aware that the C. albicans can metabolize NBT, producing formazan. Bearing this in mind we performed the experiments reported in the original Figure 8 (now Figure 10 of the revised manuscript) using killed bacteria. Note that we tested both heat-killed and paraformaldehyde-killed C. albicans hyphae to ensure that the production of formazan was not dependent on the structure of molecular patterns that might have been affected by the method used. While these details were described in the Materials and methods section of the original version, we have now mentioned them also in the Results section to avoid confusion (see Results section and also Materials and methods section)

Reviewer #3:

The manuscript by Maxson et al., describes and analyzes partially sealed phagocytic cups in macrophages which form around Candida albicans hyphae. It demonstrates that an actin-rich cuff at the distal margin of the cup supports a lateral segregation between the components of the plasma membrane and the contiguous inner leaflet of the phagocytic cup. This lateral segregation excludes the PI(4,5)P2 from the cup and confines the PI(Bauer et al., 2001)P and PIP3 to the cup. The actin cuff contains the integrin CR3 and associated integrin signaling molecules. The barrier effectively limits the movement of inner leaflet probe molecules into or out of the cup, without significantly diminishing diffusion within the plasma membrane or the cup. The actin cuff can be disrupted by the actin depolymerizing drug latrunculin A. The barrier limits escape by diffusion of large macromolecules delivered into the lumen of the cup, but not the diffusive loss of smaller molecules, including dextrans and protons (pH). Nonetheless, reactive oxygen species can be delivered into the unclosed cups. The experimental work is carefully done and the morphological evidence in support of the claims is beautiful. However, much if not most of the conclusions reported here were first described in a large study by Prashar et al., (2013), using an analogous experimental model. Examining the interactions between macrophages and filamentous bacteria, that study demonstrated (Astarie-Dequeker et al., 1999) the actin cuff (called a "jacket") that segregates two domains of contiguous membrane (plasma membrane and phagocytic cup), (Bain et al., 2014) the effect of actin depolymerization on the maintenance of that segregation, (Bauer et al., 2001) the nature of the diffusion barrier with respect to extracellular probes and (Belin et al., 2014) the retention of diffusible molecules in the cup lumen (different sizes of dextrans, pH, hydrolytic enzymes).

In accordance with the reviewer, we have cited this reference repeatedly throughout the text to more accurately lay the ground for our studies (see Results section and Discussion section). It is important to point out that our study extends and complements the Prashar et al., study in several ways: (Astarie-Dequeker et al., 1999) we identify the receptors involved in generating the actin cuff; (Bain et al., 2014) using a variety of mutants and pharmacological agents, we now identify the ligand recognized by the lectin domain of the integrin, namely the β(1,3)-glucan bond (see new Figure 4); (Bauer et al., 2001) we describe experiments indicating that molecular crowding is responsible for the diffusion barrier (see new Figure 8); (Belin et al., 2014) we describe the activation and functional role of PYK2/FAK in the phagocytosis of C. albicans hyphae (see new Figure 5); (Ben-Ami et al., 2011) we document actin turnover (treadmilling) at the cuff, despite the maintenance of the net amount of actin over extended periods; (Benard et al., 1999) we assign a primary role to formins, as opposed to Arp2/3 in the process (see new Figure 5G); (Bergmeier et al., 2007) we demonstrate that molecules trapped in the phagocytic cup remain mobile; (8) we demonstrate that generation of the cuff has a microbiostatic effect, limiting the growth of the hyphae and (9) we highlight the differential behaviour of lipid/lipid-anchored molecules located in the inner and outer leaflets of the membrane.

Moreover, unlike the present manuscript which shows that ROS can be generated in the unclosed cups, the previous paper measured the effect of partial phagocytosis on microbicidal activities against filamentous bacteria (albeit somewhat incompletely). The present manuscript but does not measure microbicidal activity against C. albicans.

This point was also raised, equally aptly, by reviewer 2. To address this issue, we conducted new experiments to assess the antimicrobial effects of the actin cuff on C. albicans hyphae. We initially measured the viability of the hyphae that had been partially internalized using propidium iodide. As is now described in the Results section, we did not see any change in fungal viability in the partially internalized C. albicans hypha (nor in fully internalized hyphae, for that matter; data not shown) for at least 60 min, the normal duration of our experiments. We reasoned that the antimicrobial effectors may not suffice to kill the fungus yet may curtail hyphal growth. The remarkable extension rate of C. albicans hyphae, which has been well-documented rate (18.8 µm/hr = 0.31 µm/min, Gow, 1985), enables accurate quantitation during the course of our experiments. We therefore, compared the rate of growth of partially internalized hyphae with an actin cuff with that of hyphae that were fully internalized or that were not engaged by macrophages (i.e. remained extracellular) throughout. These results are now summarized in the new Figure 10F and described in subsection “Functional properties of the frustrated phagosome” of the revised text. Briefly, we found that sequestration of a part of the hyphae within the frustrated phagosomes delimited by the cuff reduced the rate of growth considerably, to the same extent as seen for hyphae fully internalized within a sealed phagosome. Importantly, blocking of CD11b with the M1/70 antibody abolished this microbiostatic effect (Figure 10—figure supplement 2, and Results section). In light of this newly characterized effect on hyphal growth, we have changed the title from “microbicidal” to “microbiostatic”.

Assuming this manuscript can be rewritten to acknowledge appropriately the demonstrated precedents and concepts of that earlier work, the present study does add some interesting new data which supports a role for integrin in the establishment or maintenance of the actin cuff.

As mentioned above, we have rewritten the text to better assign credit to earlier reports, while highlighting our novel findings. The most salient novel contributions are itemized in the response to comment #1 by this reviewer. We feel that the new data and insights presented represent a significant advance to the study of phagocytosis and host-fungus interactions.

The role of integrin in the maintenance of the barrier remains indirect, however. Perhaps the actin ring is the main ingredient of the barrier, and anything which organizes actin into a ring can support the formation of diffusion barriers such as described here, and elsewhere for analogous structures Golebiewska et al., 2011; Welliver et al., 2011. To better define the nature of the diffusion barrier, the diffusion measurements (e.g. Figure 6) or probe localization studies (e.g. Figure 5) described in the manuscript should be extended to analyze the roles of actin (latrunculin A) and integrin (M1/70) to barrier maintenance. This could provide a mechanistic underpinning to the diffusion measurements.

The reviewer raises an important and valid point, which we have tried to address by performing a series of new experiments. As intimated (but not demonstrated) in the original version, we hypothesized that the diffusional barrier is the consequence of molecular crowding, generated by clustering of CR3 and associated molecules to a high density. Dense clustering was envisaged to occur upon exposure to a profusion of β(1,3)-glucan bonds on the hyphae. To test this hypothesis, we induced crowding of CR3 by crosslinking the integrins with antibodies (in the absence of C. albicans hyphae), adapting the traditional technique used previously to patch/cap surface receptors (see new Figure 8B for diagrammatic illustration). Having successfully clustered CR3, we analyzed whether this sufficed to exclude other proteins.

To this end, we transfected the cells with a fluorescent transmembrane protein (CD2-CD45GFP) and quantified the relative distribution of the two proteins before and after cross-linking CR3. As shown in the new Figure 8, patches formed by clustering CR3 excluded CD2-CD45-GFP, implying that a diffusion barrier had been generated by increasing the local density (crowding) the integrin. Importantly, neither formation of the patches, nor the exclusion of CD2CD45-GFP required actin, since latrunculin was without effect. These observations imply that clustering of the transmembrane protein CR3, as opposed to its effects on actin recruitment, are the primary determinant of the diffusion barrier. This conclusion is in good agreement with the realization that transmembrane “pickets”, rather than the underlying actin “fence”, are the primary obstacles to the diffusion of membrane-associated proteins and lipids (Freeman, et al., 2018, in press).

It is important to bear in mind that, while actin itself may not be required to exclude mobile proteins and lipids form areas of CR3 crowding, it is nevertheless required to establish and maintain the clusters, ostensibly by stabilizing the active form of CR3 via talin and vinculin, as described for other integrins. This explains why the diffusional barrier generated by interaction with hyphae disassembles when the cells are treated with latrunculin A (Figure 9). The new experiments are now described in subsection “Examining the role of CR3 and actin in the maintenance of diffusional barriers” of the revised paper.

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

1) In the abstract it is stated “[…]in response to non-canonical activation of integrins by fungal β(1,3)-glucans." We suggest using the singular "glycan". There is more than one form of fungal β(1,3)- glucan, but this is not relevant to this article.

We have changed the text as requested; please see the Abstract of the revised version.

2) B-glucans are involved in the activation of complement and the deposition of C3b fragments on the surface of C. albicans. (Boxx et al., 2010., Kozel et al., 1987, Vetvicka et al., 1996). What could be the contribution of C3b opsonization on the formation of the actin cuffs and the properties of the membrane and luminal diffusion barriers?

In the absence of serum opsonization, the observed phenomena are dependent on engagement of Dectin1, which secondarily enables the activation of integrins that bind to glucan. However, when Dectin1 expression is low, complement deposition suffices to engage the integrins and directly activate this process. We have observed that the actin cuffs created via C3b opsonization occur with similar frequency and appearance, therefore we assume that in our experiments and in the host, C3b deposition has a parallel role to Dectin1. We have included text in the Discussion section to elaborate on this point.

3) Previous reports on the phagocytosis of C. albicans showed the formation of PIPs in phagosomes containing this yeast (Heinsbroek, 2009). This was attributed to pathogenic mechanisms. On the other hand, and coinciding with the authors' interpretation, Naufer et al., 2018 recently reported that PI(Bauer et al., 2001)P co-exists with phagolysosomal markers in the phagocytic cup of heat-killed filamentous bacteria. Maybe the authors could mention this in their Discussion section.

As reported by Heinsbroek et al., 2009, we have also seen secondary waves of actin polymerization as a result of reacquisition of PI(4,5)P2 and PI(3,4)P2/PI(3,4,5)P3 on the phagosomes of serum opsonized sheep red blood cells (Bohdanowicz et al., 2010), internalized via CR3. We have added additional text and references to both papers in subsection “Phospholipid segregation between the plasma membrane and the cuff-delimited phagosomal cup” of the revised manuscript. We have also mentioned the work of Naufer et al., 2018 to indicate that live C. albicans is not required for actin cuff formation (as we see in Figure 10E). This new paper is also now referenced in subsection “Phospholipid segregation between the plasma membrane and the cuff-delimited phagosomal cup”. However, we feel that extensive discussion of the similarities/differences between these findings and our work is outside the scope of this manuscript.

4) Please clarify: In Figure 4, human serum was used to complement opsonize hyphae. How did the authors account for the presence of antibodies against C. albincans in the serum? Antibodies could be expected as this yeast is part of the human microbiome

The reviewers raise a valid point; in the text and figures we have purposefully used the general term “serum opsonization” and not complement opsonization, because additional opsonization by antibodies to C. albicans could indeed occur. This is now explicitly acknowledged in the Discussion sectionof the main text, and also in the Materials and methods section. Although not mentioned in the paper, we believe that immunoglobulins contribute comparatively little to the opsonization because Fc receptor-dependent phagocytosis is dependent on SFKs and Arp2/3, while we find the formation of actin cuffs to be insensitive to the respective inhibitors PP2 and CK666 (Figures 5E and 5G).

5) Caspofungin treatment could prevent the deposition of complement in hyphae, and its recognition by antibodies and Dectin-1. This could explain the results in Figure 4—figure supplement 1.

We thank the reviewers for their astute observation. We have included discussion of this caveat in the Discussion section. It is possible that complement binds less to the caspofungin-treated hyphae, as reports of Boxx et al., 2010 and Kozel et al., 1987 indicate that a portion of the complement binding occurs on fungal glucans. Nevertheless, we regard this as unlikely because the fungal cell wall is rich in other polysaccharides and proteins that can serve to attach complement. In this regard, it is noteworthy that caspofungin-treated C. albicans bind concanavalin A normally (Figure 4E), indicating that mannans are still present.

6) The authors showed that ROS production in the phagocytic cup retarded the elongation of the hyphae. This was alleviated by blocking CR3 with monoclonal antibodies to impede the formation of cuffs, a procedure that favours a ROS leaching. Since ROS are generally considered microbicidal, perhaps, ROS failing to reach lethal concentrations in the open cup could be the cause of the effect reported by the authors. Therefore, assuming a microbiostatic mechanism is probably incorrect.

As pointed out be the reviewers, when CR3 is blocked ROS may exit the open cup more readily, reducing their efficiency. However, leakage of phagosomal contents or ROS does not explain the failure of the frustrated phagosomes to kill the fungus, because C. albicans yeast and hyphae survive also within fully sealed phagosomes. Thus, in the case of C. albicans, ROS is not strictly microbicidal. This is now discussed in the Discussion section of the revised text.

Additionally, after reviewing the changes requested for submission of the revised manuscript, we now ensure that the requested file formats are provided, the title has been reconciled, the requested cell line information is now included in the Materials and methods section, and the source data for graphs in the main figures is provided. Please note, that in lieu of a Key Resources Table and RRIDs, we have comprehensively provided all essential information (suppliers and catalogue numbers for all cell lines, antibodies, and plasmids and reagents) in the Materials and methods section.

https://doi.org/10.7554/eLife.34798.042

Article and author information

Author details

  1. Michelle E Maxson

    Program in Cell Biology, Hospital for Sick Children, Toronto, Canada
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon 0000-0002-1493-490X
  2. Xenia Naj

    Institute for Medical Microbiology, Virology and Hygiene, University Medical Center Hamburg-Eppendorf, Hamburg, Germany
    Contribution
    Formal analysis, Funding acquisition, Investigation, Writing—review and editing
    Competing interests
    No competing interests declared
  3. Teresa R O'Meara

    Department of Molecular Genetics, University of Toronto, Toronto, Canada
    Contribution
    Conceptualization, Funding acquisition, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  4. Jonathan D Plumb

    Program in Cell Biology, Hospital for Sick Children, Toronto, Canada
    Contribution
    Conceptualization, Formal analysis, Validation, Investigation, Visualization, Writing—review and editing
    Competing interests
    No competing interests declared
  5. Leah E Cowen

    Department of Molecular Genetics, University of Toronto, Toronto, Canada
    Contribution
    Conceptualization, Resources, Funding acquisition, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  6. Sergio Grinstein

    1. Program in Cell Biology, Hospital for Sick Children, Toronto, Canada
    2. Keenan Research Centre for Biomedical Science, St. Michael’s Hospital, Toronto, Canada
    3. Department of Biochemistry, University of Toronto, Toronto, Canada
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Methodology, Writing—original draft, Project administration, Writing—review and editing
    For correspondence
    sergio.grinstein@sickkids.ca
    Competing interests
    No competing interests declared
    ORCID icon 0000-0002-0795-4160

Funding

Canadian Institutes of Health Research (FDN‑143202)

  • Sergio Grinstein

Heart and Stroke Foundation of Canada (Heart and Stroke Pfizer Fellowship)

  • Michelle E Maxson

National Institutes of Health (AI115947-01)

  • Teresa R O'Meara

The Research Training Group 1459

  • Xenia Naj

Natural Sciences and Engineering Research Council of Canada (06261)

  • Leah E Cowen

Canadian Institutes of Health Research (FDN-154288)

  • Leah E Cowen

Canadian Institutes of Health Research (PJT-153403)

  • Leah E Cowen

National Institutes of Health (1R01AI127375-01)

  • Leah E Cowen

Natural Sciences and Engineering Research Council of Canada (477598)

  • Leah E Cowen

Natural Sciences and Engineering Research Council of Canada (462167)

  • Leah E Cowen

Canadian Institutes of Health Research (PJT-148548)

  • Leah E Cowen

Canadian Institutes of Health Research (MOP-86452)

  • Leah E Cowen

Canadian Institutes of Health Research (MOP-119520)

  • Leah E Cowen

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

Acknowledgements

We sincerely thank Dr. Karin Strijbis (Utrecht University, The Netherlands), for providing RAW-Dectin1-LPETG-3xHA cells and C. albicans strain SC5314 expressing BFP, and Dr. Wolfgang Bergmeier (University of North Carolina), for providing tissue from CalDAG-GEF1 knockout mice. We thank Merck and Genome Canada for making the C. albicans GRACE collections available. MEM was the recipient of a Heart and Stroke Pfizer Research Fellowship. TRO is supported by a National Institutes of Health (NIH) Ruth L Kirschstein National Research Service Award (NRSA, AI115947-01) from the NIAID. LEC is supported by Canadian Institutes of Health Research Operating Grants (PJT-153403, PJT-148548, MOP-86452, and MOP-119520) and Foundation Grant (FDN-154288), the Natural Sciences and Engineering Council (NSERC) of Canada Discovery Grants (06261 and 462167), an NSERC EWR Steacie Memorial Fellowship (477598), and a National Institutes of Health NIAID R01 (1R01AI127375-01). SG is supported by Canadian Institutes of Health Research grant FDN–143202.

Reviewing Editor

  1. Dominique Soldati-Favre, University of Geneva, Switzerland

Publication history

  1. Received: January 4, 2018
  2. Accepted: March 16, 2018
  3. Accepted Manuscript published: March 19, 2018 (version 1)
  4. Version of Record published: April 12, 2018 (version 2)

Copyright

© 2018, Maxson et al.

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

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