Fungal sensing enhances neutrophil metabolic fitness by regulating antifungal Glut1 activity

Summary

Combating fungal pathogens poses metabolic challenges for neutrophils, key innate cells in anti-Candida albicans immunity, yet how host-pathogen interactions cause remodeling of neutrophil metabolism is unclear. We show that neutrophils mediate renal immunity to disseminated candidiasis by upregulating glucose uptake via selective expression of glucose transporter 1 (Glut1). Mechanistically, dectin-1-mediated recognition of β-glucan leads to activation of PKCδ, which triggers phosphorylation, localization and early glucose transport by a pool of pre-formed Glut1 in neutrophils. These events are followed by increased Glut1 gene transcription leading to more sustained Glut1 accumulation, also dependent on the β-glucan/dectin-1/CARD9-axis. Card9-deficient neutrophils show diminished glucose incorporation in candidiasis. Neutrophil-specific Glut1 ablated mice exhibit increased mortality in candidiasis, caused by compromised neutrophil phagocytosis, ROS and NET formation. In human neutrophils, β-glucan triggers metabolic remodeling and enhances candidacidal function. Our data show that the host-pathogen interface increases glycolytic activity in neutrophils by regulating Glut1 expression, localization and function.

Graphical Abstract

eTOC blurb

Neutrophils protect against fungal infections including candidiasis. Neutrophils kill Candida albicans in an energy demanding process. The mechanisms by which neutrophils meet their metabolic demand are unknown. Li and colleagues show how fungal sensing rewires metabolism in neutrophils via regulating glucose transporter 1 expression and function during renal host defense.

Introduction

Neutrophils are the most abundant, short lived, and terminally differentiated leukocytes in circulation (Mantovani et al., 2011; Mayadas et al., 2014). Neutrophils are the first to arrive at the site of infection where they execute rapid antimicrobial activities to eliminate and prevent pathogen dissemination to sterile sites (Burn et al., 2021). Encountering invading pathogens in diverse tissues exert metabolic challenges for these cells. Despite advances in understanding immunometabolism in T cells and macrophages (Geltink et al., 2018; Makowski et al., 2020; O’Neill et al., 2016; Pearce and Pearce, 2013), surprisingly little is known about the mechanisms by which neutrophils meet the high metabolic demand for effector functions in infectious settings. A comprehensive understanding of the antimicrobial metabolic shift could allow for selective targeting of metabolic pathways to improve neutrophil function without causing collateral tissue damage during host defense.

Unlike most cell types, mitochondria are not required for energy production in neutrophils in the presence of glucose, thus making this innate cell type metabolically unique (Fossati et al., 2003; Maianski et al., 2004; Reiss and Roos, 1978; Zucker-Franklin, 1968). Instead, neutrophils rely on glycolysis and pentose phosphate pathway (PPP) not only to fulfill their energy requirements, but also to support specialized effector functions including degranulation, phagocytosis, reactive oxygen species (ROS) production and neutrophil extracellular trap (NET) formation (Jeon et al., 2020; Kumar and Dikshit, 2019). Individuals with rare genetic deficiency of glucose-6-phopspahte dehydrogenase or glucose-6-phosphate transporter, key enzymes in glycolytic pathway, show impaired ATP and ROS generation (Ardati et al., 1997; Hiraiwa et al., 1999). Hence, neutrophils must tightly regulate glucose uptake, intracellular glucose cycling and glycolysis to execute rapid antimicrobial activities.

Most cells import glucose by facilitative diffusion across the cell membrane, a process mediated by the glucose transporter (Glut) family of membrane transport proteins. There are 14 Glut-family transporters in mammals (Glut 1–14) (Thorens and Mueckler, 2010). Of these, Glut5, Glut6 and Glut13 do not participate in glucose transport and Glut11 and Glut14 are not in the mouse genome (Douard and Ferraris, 2008; Maedera et al., 2019; Scheepers et al., 2005; Uldry et al., 2001; Wu and Freeze, 2002). The range of Glut transporters utilized by neutrophils has not yet been defined. Moreover, the mechanisms that regulate Glut expression in neutrophils are poorly understood. Unlike Glut4, which is insulin-sensitive, expression of Glut1 (encoded by SLC2A1) at the cell membrane is not regulated by blood glucose levels (Leto and Saltiel, 2012). Rather, Glut1 expression is controlled by transcription, translation and/or translocation of the protein from a pre-formed intracellular pool (Buller et al., 2008; Griffin et al., 2004; Hwang and Ismail-Beigi, 2006; Lee et al., 2015; Wang et al., 2019). Glut1 in CD4 T cells and macrophage are essential for activation, expansion and survival of these immune cells (Freemerman et al., 2014; Macintyre et al., 2014). However, our understanding of the role of neutrophil specific Glut1 expression and its regulation in host defense is far from complete.

Neutrophils are critical for controlling disseminated candidiasis, a bloodstream nosocomial infection caused by the dimorphic commensal fungus Candida albicans, that causes high mortality (30–40%) (Horn et al., 2009; Pappas et al., 2018; Pasqualotto and Denning, 2008; Zaoutis et al., 2005). Humans with neutropenia have a higher risk of disseminated candidiasis and mice depleted of neutrophils are susceptible to invasive infection (Desai and Lionakis, 2018; Fortun and Gioia, 2017; Lionakis and Netea, 2013; Uzun et al., 2001; Velasco and Bigni, 2008). Following disseminated candidiasis, neutrophils traffic to the infected organs such as kidney, brain, spleen, and liver, within 2 h (Lionakis et al., 2012). Neutrophils sense fungi by recognizing pathogen associated molecular patterns (PAMPs) via pathogen recognition receptors (PRRs), particularly C-type lectin receptors (CLRs) (Netea et al., 2006; Taylor et al., 2007). The fungal recognition via PRRs authorizes neutrophils to eliminate fungi by oxidative and non-oxidative mechanisms, and also deploy NET to neutralize pathogenic hyphal form of C. albicans (Brown, 2011; Urban et al., 2006). Additionally, the host-pathogen interaction is essential for the development of “trained” immunity in innate cells (Cheng et al., 2014). All these specialized antifungal functions require energy and neutrophils respond to the increasing metabolic demand by adapting to simpler metabolic pathways such as glycolysis (Weerasinghe and Traven, 2020). Indeed, recognition of fungal β-glucan by dectin-1 in monocytes leads to epigenetic changes and increased glycolysis in an Akt/mTOR/HIF1α-dependent manner, which are critical for the development of ‘trained’ immunity. While it is evident that host-pathogen interface is a critical determinant of antifungal functions, the mechanisms by which fungal sensing machinery in neutrophils rewire cell-intrinsic metabolic pathways to meet the increasing metabolic requirement in the kidney is poorly understood.

In this report, we demonstrate that renal immunity to disseminated candidiasis requires Glut1-dependent glucose uptake, which is controlled at multiple levels including Glut1 gene transcription, surface expression and glucose uptake. Recognition of fungal β-glucan via dectin-1 promotes PKCδ-mediated phosphorylation and surface localization of pre-formed Glut1, a process that is essential for early glucose uptake and antifungal activity of neutrophils in the kidney. This is later followed by increased transcription of Glut1 that allows for sustained neutrophil function, which is dependent on the β-glucan/dectin-1/CARD9-axis. Accordingly, mice with neutrophil-specific deletion of Glut1 showed increased susceptibility to DC. Glut1-mediated glucose uptake by neutrophils is indispensable for renal antifungal function including phagocytosis, ROS production and NET formation. Finally, β-glucan metabolically remodeled healthy donor neutrophils by upregulating Glut1 function, which is essential for fungicidal activity. Collectively, these data advance our fundamental understanding of how recognition of fungal pathogens triggers glycolytic activity by selectively regulating Glut1 expression and function in neutrophils during antifungal immunity.

Results

Glut1 transcription and phosphorylation are upregulated in neutrophils during disseminated candidiasis

The mechanisms by which neutrophils meet their high metabolic demand for rapid antimicrobial activities are poorly understood. We induced disseminated candidiasis by intra-venous (i.v.) infection with C. albicans (strain SC5314, 10⁵ CFU) and found that glucose uptake is increased in neutrophils within the infected kidney compared to sham-infected mice at 24 h post-infection (p.i.), as measured by the enhanced incorporation of glucose analog 2-NBDG (Fig 1A, S1A). Moreover, in vitro 2-NBDG incorporation assay revealed increased glucose transport by murine bone marrow (BM)-derived neutrophils at 1.5 h post-C. albicans stimulation (first time point analyzed) (Fig S1B).

Fig 1: Neutrophils upregulate Glut1 transcription and translocation during fungal stimulation.

(A) WT mice (C57BL/6) were infected with 10⁵ CFU of C. albicans (C. a) for 24 h (n=4–5). Glucose uptake by kidney infiltrating neutrophils was detected by injecting 2-NBDG followed by flow cytometry. (B) BM neutrophils from WT mice were stimulated with C. a [multiplicity of infection (MOI)=0.2 (low) or 1 (high)] for 3 h. Gene expression of different glucose transporters was measured by qPCR, normalized to Gapdh. Neutrophils were stimulated with C. a for 3 h or 6 h. Glut1 and Glut3 protein were evaluated by (C) western blot and (D) flow cytometry. (E) WT mice were infected with C. a (n=4–6). Glut1 expression in kidney-infiltrating neutrophils was measured by flow cytometry at indicated days p.i. (F) Neutrophils were stimulated with C. a (MOI=1) and phosphorylation of Glut1 (pGlut1^S226) was assessed by western blot. Phorbol 12-myristate 13-acetate (PMA) (100 ng/ml) was used as a positive control. (G) Neutrophils were stimulated with C. a (MOI=1) for 1.5 h. Surface localization of Glut1 in neutrophils was visualized by ImageStream analysis. The surface translocation of Glut1 was expressed as percentage of surface expressed (Ly6G co-localized) Glut1 out of total Glut1 protein. Data pooled from 2 (A and E) and 3–4 independent experiments (B-D, F and G) and representative images and histogram plots are shown. Statistical analysis by Student’s T test (A, G), One-way ANOVA (D, E) and Two-way ANOVA (B). Data are represented as mean ± SEM (A, B, D, E, F). See also Fig S1.

To define how neutrophils incorporate glucose in disseminated candidiasis, we evaluated expression of multiple glucose transporters in BM neutrophils in the presence or absence of C. albicans (yeast and pseudo-hyphae) for 3 h under in vitro condition. Of 9 glucose transporters tested, only the Slc2a1 (encoding Glut1) and Slc2a3 (encoding Glut3) transcripts were expressed in neutrophils (Fig 1B). Slc2a3 was highly expressed at baseline and remained unchanged during C. albicans stimulation. In contrast, Slc2a1 mRNA was low at baseline but upregulated in neutrophils at 3 h post-C. albicans stimulation in a dose-dependent manner. Congruent with mRNA expression, there was increased Glut1 protein at 6 h but not 3 h post-fungal stimulation (Fig 1C, D). There was no alteration in Glut3 protein levels in C. albicans stimulated neutrophils at these time points. Consistent with these in vitro studies, kidney-infiltrating neutrophils showed increased Glut1 expression during disseminated candidiasis, assessed at 1, 3 and 5 days p.i. (Fig 1E). These data indicate that Glut1 is transcriptionally regulated in neutrophils in systemic fungal infection.

From these results, it was clear that neutrophils incorporate glucose as early as 1.5 h post-C. albicans stimulation, even though there was no evident increase in Glut1 transcript or protein over baseline at this time point. A previous study showed that Glut1 is stored in intracellular vesicles and PKC-dependent phosphorylation of Glut1 is required for surface localization and glucose transport (Lee et al., 2015). We hypothesized that neutrophil activation would lead to rapid phosphorylation and subsequent translocation of a pool of pre-formed Glut1 from cytoplasmic vesicles to the cell membrane, which will aid in early glucose uptake at 1.5 h post-stimulation. Indeed, there was rapid phosphorylation of Glut1 (pGlut1^S226) within 30 minutes of C. albicans stimulation (Fig 1F), followed by Glut1 translocation to the cell membrane at 1.5 h post-stimulation, measured by ImageStream analysis (Fig 1G). Overall, these data indicate that C. albicans stimulation regulates Glut1 transcription, phosphorylation, and cell membrane translocation in neutrophils.

Fungal β-glucan regulates Glut1 transcription and phosphorylation in neutrophils

To define how host-pathogen interaction regulates Glut1 function in neutrophils, we assessed the role of fungal PRRs. Heat-killed (HK) C. albicans has exposed cell wall components such as β-glucans, which bind to the CLR dectin-1. Indeed, HK-C. albicans-stimulated neutrophils induced expression of Slc2a1 mRNA and phosphorylation of Glut1 at levels similar to live fungus at 3 h post-stimulation, suggesting that fungal non-protein and heat- resistant protein ligands control Glut1 function in neutrophils (Fig 2A, B). Accordingly, we activated neutrophils with purified fungal cell wall components including mannan, curdlan and zymosan. Curdlan and zymosan but not mannan increased Slc2a1 transcription and phosphorylation of Glut1 in neutrophils in a dose-dependent manner (Fig 2A, B). Since both curdlan and zymosan are β-glucans, we used curdlan for subsequent experiments. The curdlan-stimulated neutrophils demonstrated increased membrane translocation of Glut1 and in vitro glucose uptake (Fig 2C, D). In keeping with these results, BM neutrophils from Clec7a (Dectin-1)-deficient mice but not Myd88^−/− animals exhibited diminished expression of Slc2a1 mRNA and Glut1 phosphorylation upon C. albicans or curdlan stimulation (Fig 2E, F and Fig S1C, D). These results indicate that dectin-1-dependent recognition of fungal β-glucan regulates Glut1 transcription and phosphorylation in neutrophils.

Fig 2: Fungal β-glucan and dectin-1 interaction controls Glut1 function in neutrophils.

(A) BM neutrophils from WT mice were stimulated with heat-killed C. albicans (C. a) [HKCa, MOI=1 (low) or 5 (high)], mannan [Man: 10 (low) or 100 μg/ml (high)], curdlan [Cur: (10 (low) or 100 μg/ml (high)], or zymosan [Zym: (10 (low) or 100 μg/ml (high)] for 3 h. Gene expression of Slc2a1 was measured by qPCR. (B) Neutrophils were stimulated with C. a (MOI=1), HKCa (MOI=1), mannan (100 μg/ml), curdlan (100 μg/ml) or zymosan (100 μg/ml) for 30 min and Glut1 phosphorylation was assessed by western blot. Neutrophils were stimulated with curdlan (100 μg/ml) for 1.5 h. (C) Localization of Glut1 in neutrophils was visualized by ImageStream analysis and (D) glucose uptake (2-NBDG⁺ cells) was detected by flow cytometry. (E) Neutrophils from WT or Dectin-1^−/− mice were stimulated with C. albicans (MOI=0.2) or curdlan (10 μg/ml) for 3 h. Gene expression of Slc2a1 was measured by qPCR. (F) Neutrophils from WT or Dectin-1^−/− mice were stimulated with C. a (MOI=1) or curdlan (100 μg/ml). Phosphorylation of Glut1 was assessed by western blot at 30 min post-stimulation. Data pooled from at least 3 independent experiments (A-F) and representative images and histogram plots are shown. Statistical analysis by One-way ANOVA (A), Student’s T test (C, D) and Two-way ANOVA (E). Data are represented as mean ± SEM (A, C, D, E). See also Fig S1.

Activation of the dectin-1/Syk pathway controls Glut1 transcription and phosphorylation in neutrophils

The signaling pathways downstream of CLRs include Syk-dependent activation of PKCδ and CARD9, as well as a Syk-independent Raf-1 pathway (Geijtenbeek and Gringhuis, 2009). Both Syk-dependent and Syk-independent pathways activate NF-κB (Fig 3A). Selective blocking of Syk, PKCδ, CARD9 or NF-κB activity with pharmacological signaling inhibitors or gene knock-out mice impaired upregulation of Slc2a1 transcription in neutrophils in response to C. albicans or curdlan, assessed at 3 h (Fig 3B). The inhibitors showed no effect on neutrophil viability at the working concentrations (Fig S1E). In contrast, Raf-1 inhibition did not impact Slc2a1 expression (Fig S2A). These results demonstrate a specific role for the Syk/PKCδ/CARD9/NF-κB pathway in promoting Slc2a1 transcription following fungal recognition.

Fig 3: Glut1 regulation is mediated by dectin-1/PKCδ pathway in neutrophils.

(A) Schematic diagram of downstream signaling pathways of dectin-1. The diagram also shows different inhibitors or gene knockout mice used to sequentially block the signaling intermediates in dectin-1 signaling pathway. (B) BM neutrophils from WT or Card9^−/− mice were stimulated with C. albicans (C. a) (MOI=0.2) or curdlan (Cur) (10 μg/ml) for 3 h with or without indicated inhibitors. Gene expression of Slc2a1 was measured by qPCR. (C-F) BM neutrophils from WT or Card9^−/− mice were stimulated with C. a (MOI=1) or curdlan (100 μg/ml) in the presence or absence of indicated inhibitors. Phosphorylation of Glut1 was assessed by western blot at 30 min post-stimulation. Data pooled from at least 3 independent experiments (B-F) and representative images are shown. Statistical analysis by Two-way ANOVA (B). Data are represented as mean ± SEM (B). See also Fig S2.

We also detected phosphorylation of Glut1 following selective blocking of Syk, PKCδ, CARD9 or NF-κB activity with pharmacological inhibitors or knock-out mice in neutrophils (Fig 3A). Neutrophils stimulated with C. albicans or curdlan showed increased phosphorylation of PKCδ and Glut1 relative to vehicle treated group at 30 min post-stimulation (Fig 3C). Pharmacological inhibition of Syk or PKCδ completely blocked Glut1 phosphorylation (Fig 3C, D). Inhibition of Raf-1 showed no impact on the phosphorylation of Glut1 (Fig S2B). Surprisingly, loss of CARD9 or treatment with an NF-κB inhibitor also had no effect on Glut1 phosphorylation (Fig 3E, F), indicating that PKCδ activation upstream of CARD9 is required and sufficient for Glut1 phosphorylation. Overall, these data highlight that activation of Syk and PKCδ downstream of dectin-1 activation is required for the phosphorylation of Glut1 in neutrophils but CARD9, NF-κB or Raf-1, are not.

Compromised glucose uptake by Card9^−/− neutrophils in disseminated candidiasis

In order to test the role of dectin-1/Syk/CARD9 pathway in regulation of Glut1 function, we next assessed Glut1 translocation and glucose uptake in neutrophils following sequential inhibition of signaling molecules in Syk-dependent pathway, as in Fig 3A. In line with the phosphorylation data (Fig 3C, D), Syk or PKCδ inhibitors blocked the surface translocation of Glut1 at 1.5 h post-curdlan stimulation (Fig 4A, B, Fig S2C, D). Accordingly, glucose uptake by neutrophils was reduced in the presence of Syk or PKCδ inhibitors at 1.5 h post-stimulation (Fig 4E, F, Fig S2G, H). In contrast, CARD9 deficiency or NF-κB inhibition showed no impact on the cell membrane translocation of Glut1 (Fig 4C, D, Fig S2E, F). The glucose incorporation in Card9^−/− neutrophils or NF-κB inhibition was diminished at 6 h but not at 1.5 h post-stimulation (Fig 4G, H, I, Fig S2I, J, K). These results indicate that Glut1 function and glucose uptake in early stage (1.5 h post-stimulation) is mediated by the rapid phosphorylation and membrane translocation of pre-formed and intracellularly stored Glut1 by activation of PKCδ. This is proceeded by a late stage of glucose uptake (6 h post-stimulation), which is regulated by increased Glut1 transcription and protein expression in a Syk/PKCδ/CARD9/NF-κB-axis-dependent manner.

Fig 4: Increased glucose uptake in fungal stimulated neutrophils is mediated by Syk-PKCδ/CARD9/NFκB-axis.

BM neutrophils from WT or Card9^−/− mice were stimulated with curdlan (Cur) (100 μg/ml) for 1.5 h in the presence or absence of indicated inhibitors. (A-D) Glut1 localization in neutrophils was visualized by ImageStream and (E-H) glucose uptake (2-NBDG⁺ cells) by neutrophils was detected by flow cytometry. (I) Neutrophils from WT or Card9^−/− mice were stimulated with curdlan (100 μg/ml) for 6 h, and glucose uptake by neutrophils was detected by flow cytometry. WT and Card9^−/− mice were infected with C. albicans (C. a) for 24 h (n=4–6). (J) Glucose uptake and (K) Glut1 protein level in kidney infiltrating neutrophils were detected by flow cytometry. (L) Glut1 localization in neutrophils was visualized by ImageStream. Data pooled from at least 2–3 independent experiments (A-L) and representative images and flow cytometry plots are shown. Statistical analysis by Two-way ANOVA. Data are represented as mean ± SEM (A-L). See also Fig S2, S3.

To assess the contribution of these pathways to a physiological fungal infection setting, we measured glucose incorporation by Card9^−/− neutrophils in kidney following disseminated candidiasis. WT and Card9-deficient mice showed comparable recruitment of neutrophils to the kidney at 24 h p.i., as described (Drummond et al., 2015) (Fig S3A). However, kidney-infiltrating neutrophils in Card9^−/− mice showed reduced 2-NBDG⁺ cells compared to WT animals (Fig 4J, Fig S3B). Card9^−/− neutrophils also exhibited diminished Glut1 protein expression at this time point (Fig 4K). However, the percentage of total Glut1 protein localized to the cell surface of kidney-infiltrating neutrophils was comparable between groups at 24 h p.i. (Fig 4L). These results indicate that Glut1 transcription but not phosphorylation or translocation in neutrophils is dependent on CARD9 during C. albicans infection and reveal functionally and kinetically distinct mechanisms by which glucose uptake is controlled in neutrophils during fungal encounter.

Glut1 deficiency inhibits glycolytic activity in neutrophils

To interrogate the role of neutrophil-specific Glut1 in disseminated candidiasis, we crossed MRP8-Cre-IRES/GFP and Slc2a1^fl/fl mice to specifically delete Glut1 in this cell type (henceforth referred to as PMN^ΔGLUT1) (Lagasse and Weissman, 1994; Reber et al., 2017). In MRP8Cre mice, Cre recombinase gene, internal ribosomal entry site, and an enhanced green fluorescent protein gene were placed downstream of the human S100 calcium binding protein A8 (calgranulin A) (MRP8 or S100A8) promoter. GFP is conditionally expressed as a bicistronic transcript linked to MRP8 expression in the transgenic mice and reports MPR8Cre positive cells. PMN^ΔGLUT1 mice showed successful Glut1 deletion in neutrophils from blood, BM and spleen, as evidenced by reduced Glut1 protein expression (Fig 5A, B and Fig S4A). The lack of Glut1 did not impact Glut3 mRNA and protein expression in neutrophils at the baseline or after C. albicans stimulation (Fig 5B, Fig S4B). There was no detectable MRP8Cre activity in the kidney-resident macrophages and infiltrating monocytes/macrophages at the baseline (Fig S4C). Furthermore, PMN^ΔGLUT1 mice demonstrated normal neutrophil morphology and granulopoiesis, number of granulocyte-macrophage progenitors (GMPs) and neutrophils in the blood and spleen (Fig 5C, Fig S4D–F). The BM GMPs showed no MRP8Cre activity at the baseline and after infection (Fig S4G). These results indicate that Glut1 is not required for neutrophil development and survival.

Fig 5: Glut1 deficiency suppresses glycolysis in neutrophils during fungal stimulation.

(A) Glut1 expression in mouse peripheral blood neutrophils was measured by flow cytometry. (B) Glut1 and Glut3 protein levels in BM neutrophils were measured by western blot. (C) The number of neutrophils in blood and spleen of 6–7 weeks old male mice was assessed by flow cytometry. (D) PMN^WT and PMN^ΔGLUT1 mice were infected with C. albicans (C. a) for 24 h (n=4–6). Glucose uptake by kidney infiltrating neutrophils (GlucoseCy5⁺ cells) was detected by flow cytometry. (E-I) BM neutrophils from PMN^WT or PMN^ΔGLUT1 mice were stimulated with curdlan (Cur) (10 μg/ml) for 3 h. Cell pellets were subjected to metabolomics by untargeted high-resolution LC-HRMS. Data pooled from at least 3 independent experiments (A-I) and representative images and flow cytometry plots are shown. Statistical analysis by Student’s T test (C) or Two-way ANOVA (D and I). Data are represented as mean ± SEM (C, D, I). See also Fig S4, S5.

We measured glucose uptake by kidney-infiltrating neutrophils from control (PMN^WT) or PMN^ΔGLUT1 mice after systemic infection with C. albicans. The glucose analog 2-NBDG (FITC labeled) is incompatible with the GFP expressed in MRP8-Cre-IRES/GFP mice. So, we used another fluorescent glucose tracer, Cy5-linked 1-amino-glucose (GlucoseCy5), to monitor the in vivo glucose uptake by neutrophils (Watson et al., 2021). While glucose incorporation in kidney infiltrating neutrophils was comparable between the sham-infected groups, PMN^ΔGLUT1 neutrophils showed reduced glucose transport at 24 h p.i. (Fig 5D). These results indicate that while Glut1 is dispensable at the basal level, it is essential for glucose uptake in kidney-infiltrating neutrophils during disseminated candidiasis.

To further dissect the impact of Glut1 deletion on the energy metabolism, we performed metabolomics study on neutrophils from PMN^ΔGLUT1 and PMN^WT mice. Since it is impossible to distinguish between fungal and mammalian metabolites, we used curdlan to stimulate the neutrophils. Metabolomics analysis revealed ~950 metabolites in neutrophils at 3 h post-curdlan stimulation, as assessed by untargeted high-resolution LC-HRMS. Curdlan-stimulated PMN^WT neutrophils showed increased expression of 30% of the measured metabolites (295/950) compared to unstimulated control (Fig 5E, F, G, H). In contrast, very few metabolites (20/950) were changed in curdlan stimulated PMN^ΔGLUT1 neutrophils than control, suggesting impaired metabolic regulation in the absence of Glut1. The PMN^ΔGLUT1 neutrophils showed reduced expression of ~33 metabolites compared to PMN^WT neutrophils after curdlan stimulation. Notably, the metabolites related to glycolysis including hexoses, lactate and hexose-6-phosphate were reduced in stimulated PMN^ΔGLUT1 neutrophils compared to PMN^WT group (Fig 5I, Fig S5A). Interestingly, the effect of Glut1 deficiency on metabolites related to the TCA cycle and glutamate metabolism was minimal in curdlan-stimulated neutrophils (Fig S5B). When subjected to a glycolysis stress test, absence of Glut1 did not impact the basal glycolysis and glycolytic capacity of neutrophils (Fig S5C). However, glycolysis and glycolytic capacity was reduced in PMN^ΔGLUT1 neutrophils following curdlan stimulation. Overall, our data indicate that the absence of Glut1 compromised glucose uptake and glycolysis in neutrophils during fungal stimulation.

Loss of Glut1 in neutrophils increases susceptibility to disseminated candidiasis

We next assessed the susceptibility of PMN^ΔGLUT1 mice to invasive fungal infection. Mice were systemically infected with C. albicans and evaluated for survival over 14 days. PMN^ΔGLUT1 mice showed exacerbated susceptibility to systemic infection compared to PMN^WT mice (Fig 6A). PMN^ΔGLUT1 mice succumbed to infection (~100% mortality) by day 7 p.i., a time point at which majority (~80%) of control mice were alive. Accordingly, PMN^ΔGLUT1 kidneys showed increased fungal burden and hyphal invasion into the renal parenchyma at day 3 p.i. (Fig 6B, C).

Fig 6: Deficiency of Glut1 in neutrophils exacerbates susceptibility of mice to disseminated candidiasis.

PMN^WT and PMN^ΔGLUT1 mice were infected with C. albicans (C. a). (A) Mice were evaluated for survival over a period of 14 days (n=12). (B) Kidney fungal burden (n=8) and (C) renal histopathology (PAS staining) was evaluated at day 3 p.i. Black arrows indicate C. a hyphae in renal parenchyma. Magnification: 20×10. PMN^WT and PMN^ΔGLUT1 mice were sublingually inoculated with a cotton ball saturated in C. a for 75 min (n=8–14). (D) Mice body weight were monitored daily and (E) fungal burden of the tongue was evaluated at day 5 p.i. (F) BM neutrophils from PMN^WT or PMN^ΔGLUT1 mice were incubated with C. a (MOI=0.2) for 3 h. Candidacidal capacity was assessed by in vitro fungal killing assay. (G) PMN^WT and PMN^ΔGLUT1 mice were infected with 5 × 10⁶ CFU of Streptavidin-AF633⁺dTomato⁺ C. a. Mice were sacrificed 2 h p.i. and phagocytosis and intracellular killing by kidney-infiltrating neutrophils were measured by flow cytometry analysis. Live C. a engaged neutrophils emit two fluorescence signals (dTomato⁺AF633⁺) while dead C. a engaged neutrophils just emit only AF633 fluorescence signal (dTomato⁻AF633⁺). (H) Mice were infected with C. a for 24 h. ROS production in kidney infiltrating neutrophils were measured by CellROX^® Deep Red reagent. (I) Neutrophils were incubated with C. a (MOI=0.2) for 3 h and NET formation was visualized by staining with Sytox Orange and anti-neutrophil elastase antibody. Black triangles indicate fungal hyphae. Magnification: 40×10. Frequency of NETosis score was blindly evaluated and expressed as arbitrary units. Data pooled from at least 2–3 independent experiments (A-I) and representative images and FACS plots are shown. Statistical analysis by log-rank test (A), Mann-Whitney test (B, E), Student’s T test (F, G) or Two-way ANOVA (H, I). Data are represented as geometric mean with geometric SD (B, E) or mean ± SEM (D, F, G, H, I). See also Fig S6, S7.

We also subjected PMN^ΔGLUT1 mice to oropharyngeal candidiasis (OPC), a setting where neutrophils play an important role in fungal clearance (Huppler et al., 2014). Following oral infection with C. albicans, PMN^ΔGLUT1 mice lost more body weight than controls (Fig 6D). At day 5 p.i., a time point when WT mice are known to fully clear C. albicans from the oral mucosa, PMN^ΔGLUT1 mice exhibited elevated oral CFU compared to PMN^WT mice (Fig 6E). Collectively, these data highlight a critical role for neutrophil-specific Glut1 expression in antifungal host defense.

Cellular mechanisms of Glut1-dependent immunity to C. albicans

To define the cellular mechanisms for susceptibility of PMN^ΔGLUT1 mice to disseminated candidiasis, we quantified inflammatory cells infiltration in the kidney at 24 h p.i. Surprisingly, PMN^ΔGLUT1 and PMN^WT mice showed comparable numbers of neutrophils (liveCD45⁺Ly6G⁺CD11b⁺ cells), monocytes (liveCD45⁺Ly6G⁻CD11c⁺CD11b^loF4/80^lo cells) and macrophages (liveCD45⁺Ly6G⁻CD11c⁺CD11b^hiF4/80^hi cells) in the infected kidney (Fig S6A). Similar results were obtained when neutrophil infiltration in the kidney was evaluated at an earlier time point (2 h p.i) (Fig S6B). Moreover, PMN^ΔGLUT1 neutrophils showed no defect in migration in the presence of the neutrophil-attracting chemokine CXCL2, assessed in a transwell migration assay (Fig S6C). There were also no obvious changes in patterns of cell death in these mice, with similar numbers of early (AnnexinV⁺7-AAD⁻) and late (AnnexinV⁺7-AAD⁺) apoptotic neutrophils in the infected kidney between groups (Fig S6D). These results indicate that the defect in renal fungal clearance in PMN^ΔGLUT1 mice cannot be attributed to impaired migration or increased apoptosis of neutrophils.

Neutrophils kill C. albicans mainly by phagocytosis, ROS formation and NETosis (Desai and Lionakis, 2018; Lionakis and Netea, 2013; Netea MG, 2015; Urban et al., 2006). An in vitro C. albicans killing assay demonstrated that Glut1 deficiency led to impaired fungicidal activity of neutrophils at 3 h post-incubation (Fig 6F). To determine the fungicidal capacity of neutrophils lacking Glut1 in vivo, we infected PMN^ΔGLUT1 and PMN^WT mice with a streptavidin AF633-dTomato⁺ C. albicans reporter strain that enables simultaneous fluorescent visualization of phagocytosis and fungal killing in the same cells: AF633⁺dTomato⁺ signal report intracellular live fungi and AF633⁺dTomato⁻ signal denote killed fungal cells within the neutrophils. As shown, kidney-infiltrating neutrophils in PMN^ΔGLUT1 mice had compromised phagocytosis and intracellular fungal killing activity compared to PMN^WT mice (Fig 6G, S7A, B).

Our data show that PMN^ΔGLUT1 neutrophils are defective in glycolysis, a pre-requisite for generating NADPH oxidase-mediated ROS production (Fig 5) (Mayadas et al., 2014). Hence, we assessed ROS generation by kidney-infiltrating neutrophils during candidiasis. As expected, PMN^ΔGLUT1 neutrophils produced diminished ROS compared to control neutrophils in the infected kidney, as measured by CellROX^® Deep Red at 24 p.i. (Fig 6H). Since NET formation is partially dependent on ROS generation (DeSouza-Vieira et al., 2016), we evaluated NETosis by PMN^ΔGLUT1 neutrophils in the presence of fungi. To that end, neutrophils were incubated with C. albicans for 3 h and NET formation was visualized by co-staining with Sytox Orange and an anti-neutrophil elastase antibody. Indeed, PMN^ΔGLUT1 neutrophils demonstrated reduced NET formation around fungal hyphae than PMN^WT neutrophils (Fig 6I). Overall, these results demonstrate that Glut1-mediated glucose uptake in neutrophils is critical for physiological fungicidal activity and control of systemic C. albicans infection.

Glut1 in human neutrophils is essential for fungicidal activity

There are a number of differences between mouse and human neutrophils, including the absolute number in the blood, composition of granules, expression of neutrophil-derived antimicrobial peptides and ability to kill C. albicans (Zschaler et al., 2014). To assess the role of Glut1 in human PMNs, we isolated neutrophils from peripheral blood from 10 healthy volunteers and activated the cells with C. albicans or curdlan. As in murine neutrophils, SLC2A1 mRNA was upregulated 3 h after C. albicans or curdlan stimulation (Fig 7A). We also observed increased phosphorylation of Glut1 within 30 minutes of fungal or curdlan stimulation (Fig 7B). Moreover, human neutrophils exhibited increased in vitro glucose uptake (measured by 2-NBDG incorporation) during C. albicans stimulation. A Glut1-specific inhibitor (WZB117) blocked both glucose uptake and fungicidal activity in neutrophils (Fig 7C, D). Collectively, these data show an essential role for Glut1 in ensuring the ‘metabolic fitness’ of human neutrophils that enable them to mediate immunity to C. albicans infections.

Fig 7: Glut1 is critical for fungicidal function of human neutrophils.

Healthy donor neutrophils (n=10) were stimulated with C. albicans (C. a) (MOI=0.2) for 3 h. (A) Gene expression of SLC2A1 was measured by qPCR. (B) Neutrophils isolated from healthy volunteers were stimulated with C. a (MOI=1) or curdlan (100 μg/ml) for 30 min. Glut1 phosphorylation was assessed by western blot. Representative image and ImageJ quantification of band intensities are shown. Healthy donor neutrophils were stimulated with C. a (MOI=1) in the presence or absence of Glut1 inhibitor (WZB117: 10 μM) for 1.5 h. (C) Glucose uptake by neutrophils was detected by flow cytometry, and (D) candidacidal activity of neutrophils was assessed by in vitro fungal killing assay. Statistical analysis by Student’s T test (A, D) and One-way ANOVA (B, C). Data are represented as mean ± SEM (A, B). See also Table S1.

Discussion

The regulation of cellular metabolic pathways is a critical determinant of immune function, a field that has gained renewed appreciation in recent years. To date, most studies of immunometabolism focus on how T cells and macrophages are impacted, with surprisingly little emphasis on understanding metabolic regulation of neutrophil function. Neutrophils are short lived, terminally differentiated cells and execute rapid antimicrobial function in diverse tissues and have unique metabolic requirements (Burn et al., 2021). How neutrophils dynamically regulate metabolism to fuel antifungal activity is poorly understood. We show that neutrophils upregulate Glut1 expression and glucose transport across the cell membrane in candidiasis. Recognition of fungal β-glucan via CLRs induced rapid phosphorylation, surface localization and transcription of Glut1 in a dectin-1 dependent manner. Mice with a neutrophil-specific deletion of Glut1 showed increased susceptibility to candidiasis. We also demonstrated that β-glucan-mediated upregulation of Glut1 is essential for antifungal functions in human neutrophils. Overall, these results reveal that fungal sensing ensures ‘metabolic fitness’ of neutrophils, which is required for host defense against candidiasis.

We demonstrate that Glut1 transcript and protein expression were upregulated in neutrophils in disseminated candidiasis. Although baseline Glut3 mRNA was expressed at a higher level than Glut1, Glut3 mostly remained unchanged during fungal stimulation, indicating that Glut1 and Glut3 are differentially regulated in neutrophils. The transcription factor RelA binds to the Glut1 promoter (Wang et al., 2019), thus confirming the role of NF-κB in the selective regulation of Glut1 in neutrophils. In contrast, Glut3 is upregulated in primary human monocytes following C. albicans stimulation (Wu et al., 2021). In a recently published report, mice with a neutrophil-specific deletion of Glut3 showed diminished number of neutrophils in the blood, indicating that Glut3 is indispensable for neutrophil development and survival (Ancey et al., 2021). Deletion of Glut1 partially inhibited neutrophil-mediated glucose transport, suggesting that Glut3 may contribute to increased glucose uptake during fungal stimulation. In-depth studies utilizing tamoxifen inducible Cre mice to ablate Glut3 are required to interrogate the role of Glut3 in host defense against candidiasis.

The recognition of fungal PAMPs via CLRs is crucial for innate immunity and clearance of fungal pathogens (Netea et al., 2006; Taylor et al., 2007). Here, we show that fungal β-glucan recognition via dectin-1 is also required for metabolic reprogramming of neutrophils, which is essential for the antifungal activities. Intriguingly, the lack of dectin-1 partially inhibited the upregulation of Glut1, indicating the contribution of additional PRRs. Accordingly, selective inhibitors of Syk and PKCδ completely abrogated Glut1 upregulation in neutrophils. We observed that mannan was unable to induce Glut1 expression, which may be due to low dectin-2 expression on mouse neutrophils (Taylor et al., 2014). The impact of fungal PAMPs on alternative metabolic pathways including fatty acid oxidation and glutaminolysis, known to regulate neutrophil function during limited glucose availability in tissues, warrants careful investigation (Kumar and Dikshit, 2019).

While most antifungal activities were diminished in the absence of Glut1, chemotaxis was unaffected in neutrophils. This observation is in line with our prior study showing that uremic neutrophils with compromised Glut1-mediated glucose uptake exhibited normal migration during disseminated candidiasis (Jawale et al., 2020). The neutrophils respond to chemotactic signals instantly and make extensive cytoskeleton reorganization, which is an energy dependent process. However, there are several possibilities that can explain this result. Numerous reports suggested that mitochondria and TCA cycle regulate chemotaxis in neutrophils (Bao et al., 2015; Chen et al., 2006; Zhou et al., 2018). Accordingly, carbonyl cyanide 4-(trifluoromethoxy) phenylhydrazone treatment, a potent uncoupler of mitochondrial oxidative phosphorylation, suppressed neutrophil migration (Fossati et al., 2003). Alternatively, energy derived from break down of stored glycogen by glycogenolysis may compensate for the loss of glucose uptake during chemotaxis.

We show a dynamic and kinetically distinct regulation of Glut1-mediated glucose uptake in neutrophils during candidiasis. Following fungal recognition via CLRs, the early stage enables rapid glucose transport and is mediated by PKCδ-driven phosphorylation of pre-formed Glut1 in cytoplasmic vesicle. These early events are followed by increased Glut1 gene transcription and Glut1 accumulation to replenish Glut1 level inside the cell and sustain antifungal activities by neutrophils, dependent on the β-glucan/dectin-1/CARD9-axis. The physiological significance of early glucose uptake is underscored by the fact that PKCδ, but not CARD9, promotes neutrophil ROS production and C. albicans killing within 3 h (Li et al., 2016). Interestingly, dectin-1/Akt/mTOR/HIF-1α and the noncanonical Raf-1 pathways have been implicated in increased glucose consumption and a shift towards glycolysis dependent metabolic pathway in ‘trained’ human monocytes with trained immunity (Cheng et al., 2014). Although Raf-1 pathway is dispensable for Glut1-mediated glucose uptake in neutrophils, we showed that mTORC1 inhibition suppressed glucose incorporation in neutrophils in fungal infection (Jawale et al., 2020). Moreover, only Syk inhibition but not PKCδ or NK-κB inhibitors showed a greater reduction in Slc2a1 mRNA expression than Card9^−/− neutrophils, suggesting a Syk dependent but Card9 independent signaling mechanisms in Slc2a1 gene regulation in neutrophils. Although the small molecule inhibitors used in the present study showed no effect on neutrophil viability, potential off-target effects of these inhibitors on Glut1 regulation cannot be completely ruled out.

The neutrophils are exposed to a variety of tissue micro-environments during infection, which poses metabolic stress on these innate cells. The neutrophils respond to these changing environments by enhancing glucose uptake to execute antimicrobial activities (Weerasinghe and Traven, 2020). Moreover, recent studies have indicated that neutrophils exhibit preferences for utilizing alternative metabolic pathways depending on the availability of nutrients in tissues (Jeon et al., 2020; Kumar and Dikshit, 2019). Hence, we tested the ability of PMN^ΔGLUT1 animals to control fungal infection in OPC, where neutrophils play an important role in fungal clearance in the oral cavity. In contrast to kidney fungal burden in DC, PMN^ΔGLUT1 mice exhibited moderate impairment in fungal clearance in the oral cavity during OPC, thus phenocopying Card9^−/− mice (Bishu et al., 2014). In this context, it is important to note that MRP8 promoter was originally described to drive Cre-mediated deletion in 80–90% of neutrophils, with minor leakage (<20%) into some monocyte/macrophage populations (Abram et al., 2014).

A comprehensive understanding of the metabolic regulator(s) in neutrophils could be helpful in unraveling the fundamental mechanisms of antifungal immunity, with broader implications for other infections and autoinflammatory diseases. Additionally, this knowledge may allow timely and selective targeting of metabolic pathways to augment antifungal effector mechanisms of neutrophils without causing bystander damage to host tissues.

STAR METHODS

Resource Availability

Lead Contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Partha S. Biswas (psb13@pitt.edu).

Materials Availability

This study did not generate new unique reagents.

Data and Code Availability

• The study did not create large data sets that required deposition in a public data base. Microscopy data reported in this paper will be shared by the lead contact upon request.

• The study did not create original code.

• Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

Experimental models and subject details

Mice

C57BL/6 (WT) mice were purchased from Taconic Biosciences Inc. MRP8-Cre⁺, Card9^−/−, Clec7a^−/− and Myd88^−/− mice were purchased from the Jackson laboratory. Slc2a1^fl/fl mice are under Material Transfer Agreement (MTA) from University of Colorado, USA. To generate mice with conditional deletion of Glut1 in neutrophils, we crossed the Slc2a1^fl/fl mice with MRP8-Cre-IRES/GFP mice. All mice were housed in specific pathogen free conditions under the supervision of Division of Laboratory Animal Resources, University of Pittsburgh. All animal experiments were conducted following the NIH guidelines under protocols approved by the University of Pittsburgh Institutional Animal Care and Use Committee.

Mouse model of disseminated candidiasis and oropharyngeal candidiasis (OPC)

C. albicans (Strain SC5314) was grown in yeast extract-peptone-dextrose (YPD) broth at 30°C for 16 to 20 hours in a shaker incubator. Eight to ten weeks old male mice were infected with 10⁵ CFU of C. albicans per mouse via the lateral tail vein and daily monitored for signs of illness, weight loss, and survival. Mice showing >20% weight loss or signs of severe pain or distress were sacrificed for humane purpose. For fungal burden evaluation, mice were sacrificed at day 3 p.i. and kidneys were weighed and homogenized using a GentleMACS. Serial dilutions of homogenates were plated on YPD agar, and fungal burden was represented as CFU per gram of kidney. Formalin-fixed tissue sections were stained with Periodic Acid-Schiff (PAS) stain for visualization of C. albicans.

OPC was induced as previously described (Solis and Filler, 2012). Briefly, mice were sublingually inoculated with a cotton ball saturated in C. albicans (Strain SC5314) for 75 min under anesthesia. Then the mice were daily monitored for signs of illness and weight loss. On day 5 p.i., mice tongue was weighed and homogenized. Fungal burdens were determined by plating serial dilutions of homogenates on YPD agar, and fungal burden was represented as CFU per gram of tongue.

Human subjects

All human subjects’ studies were approved by the University of Pittsburgh Institutional Review Board. The biological samples were obtained after providing written informed consent. The healthy donor blood samples (n=10) were obtained as part of a study entitled “Banking of Biological Samples and Collection of Clinical Data for Connective Tissue Disease” at the Division of Rheumatology and Clinical Immunology, University of Pittsburgh. All demographic information including race, gender and age of the study subjects are provided as Supplementary Table 1.

Neutrophil Culture

Isolated mouse or human neutrophils were used immediately for in vitro experiments. Neutrophils were cultured in RPMI 1640 Complete Medium at 37°C with 5% CO₂. The cells were seeded in 24-well flat bottom plates with a concentration of 1.5 × 10⁶/well and stimulated for 30 min to 6 hours depending on the experiment.

Method details

Mouse and human neutrophil isolation

Mouse femur and tibia BM-derived neutrophils were isolated using Neutrophil Isolation Kit, following manufacturer instructions. Neutrophil purity (>90%) was checked by FACS (liveLy6G⁺CD11b⁺). Human peripheral blood neutrophils were isolated from healthy donors using MACSxpress Human Neutrophil Isolation Kit, following manufacturer instructions, and checked for purity (>95%) by FACS.

Glucose uptake measurement

BM neutrophils were seeded in 96-well flat bottom plates and incubated at 37°C for 30 min. Cells were stimulated with C. albicans (MOI=1) or curdlan (100 μg/ml) for 1.5 hours or 6 hours. The cells were washed with PBS and 100 μl 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino]-2-deoxy-D-glucose (2-NBDG, 100 μM in PBS) was added. Cells were incubated at 37 °C for another 30 min. The glucose uptake by neutrophils was detected by FACS.

For glucose uptake measurement in kidney-infiltrating neutrophils, mice were infected with 10⁵ CFU of C. albicans via the lateral tail vein. After 24 hours, mice were injected i.v. with 200 μL 2-NBDG (2.5 mM in PBS) or glucoseCy5 (37 μM in PBS). Thirty min later, kidney infiltrating neutrophils were analyzed for glucose uptake by flow cytometry (FACS).

Glut1 expression

BM neutrophils were seeded in 24-well flat bottom plates and incubated at 37°C for 30 min. Cells were stimulated with C. albicans (MOI=1) or curdlan (100 μg/ml) for 3 hours or 6 hours. For FACS analysis, the cells were fixed and anti-Glut1 antibody or Rabbit IgG isotype control antibody were used for intracellular staining. For Western blot detection of Glut1, neutrophils were lysed and total protein was stained for Glut1 using anti-Glut1 antibody.

For Glut1 protein level measurement in kidney-infiltrating neutrophils, mice were systemically infected with 10⁵ CFU of C. albicans. After 24 hours, kidney cell suspensions were prepared and Glut1 protein expression in neutrophils was measured by FACS.

Imagestream analysis

BM neutrophils were seeded in 24-well flat bottom plates and incubated at 37°C for 30 min. Cells were stimulated with C. albicans (MOI=1) or curdlan (100 μg/ml) for 1.5 hours. For Imagestream analysis, the cells were stained with anti-Ly6G antibody followed by intracellular staining with anti-Glut1 antibody. The co-localization of Ly6G and Glut1 fluorescence was detected by Imagestream analysis and expressed as percentage of surface expressed (Ly6G co-lacalized) Glut1 out of total Glut1 protein.

For Glut1 localization in kidney-infiltrating neutrophils, mice were infected with 10⁵ CFU of C. albicans. After 24 hours, kidney cell suspensions were subjected to Imagestream analysis to detect Glut1 localization in neutrophils.

Inhibitors for in vitro studies

Inhibitors used for the in vitro experiments include 2 μM BAY61–3606 (Syk inhibitor), 10–50 μM Rottlerin (PKCδ inhibitor), 1 and 10 μM GW5074 (Raf-1 inhibitor), 1 μM BAY11–7085 (NFκB inhibitor), and 10 μM WZB117 (Glut1 inhibitor). All the inhibitors were incubated with neutrophils 30 min before stimulation. The stock solution of BAY61–3606 (10 mM), Rottlerin (20 mM), GW5074 (10 mM), BAY11–7085 (10 mM) and WZB117 (10 mM) was made by dissolving each inhibitor in DMSO. The stock solutions were further diluted using media to make working concentrations.

Flow cytometry

Single cell suspensions from tissues were prepared as previously described (Jawale et al., 2020). Briefly, spleens were subjected to mechanical dissociation followed by red blood cell lysis. For kidneys, mice were perfused with PBS containing 10 mM EDTA before the harvest. Kidneys were digested with collagenase IV for 20 min and homogenates were passed through 70 μm cell strainers.

Staining of mouse cells was performed with surface antibodies (CD45, CD11b, Ly6G, Ly6C, F4/80, CD34, CD3, CD19, B220 and C-kit) and intracellular antibody (Glut1). Cells were fixed and permeabilized before intracellular staining. Dead cells were excluded using the Live/Dead Ghost Dye Violet 510 dye. To quantify the number of cells, cell counting beads were added before the run. The samples were acquired on BD LSR Fortessa cytometer and analyzed with FlowJo software.

In vivo neutrophil phagocytosis and Candida killing assay

In vivo neutrophil phagocytosis and Candida killing assay was performed as previously described (Jawale et al., 2020). Briefly, dTomato⁺ C. albicans were incubated with 0.5 mg/mL Biotin-XX, sulfo-SE for 2 h and then 0.02 mg/mL streptavidin conjugated to Alexa Fluor 633 for another 30 min. Mice were infected with 5 × 10⁶ CFU of AF633⁺tdTomato⁺ C. albicans via lateral tail vein. Mice were sacrificed at 2 h p.i., and phagocytosis/intracellular killing by kidney-infiltrating neutrophils were measured by FACS. The streptavidin-AF633⁺dTomato⁺ C. albicans emit two fluorescence signals: dTomato⁺AF633⁺ is emitted by live C. albicans, while dead C. albicans emits only AF633 fluorescence signal (dTomato⁻AF633⁺).

In vitro C. albicans killing assay

BM neutrophils (5 × 10⁴/well) were seeded in a flat bottom 96-well plate. After 30 min, 10⁴ CFU of C. albicans were added to the wells. The number of live C. albicans was assessed by plating serial dilutions on YPD agar at 3 h post-incubation. The percent killing was expressed as [1 − (CFU of C. albicans in the presence of neutrophils/CFU of C. albicans cells in the absence of neutrophils)] × 100%. Fold change C. albicans killing was expressed as percent killing in experimental group/percent killing in control group.

NETosis assay

BM neutrophils were incubated with C. albicans for 3 h. The cells were fixed (4% PFA) and permeabilized and NET formation was measured by staining with Sytox Orange and an anti-neutrophil elastase polyclonal antibody. The secondary antibody used was goat anti-rabbit Cy5. The staining was visualized using an EVOS FL Auto microscope (Life Technologies).

Giemsa Staining of blood smear

Peripheral blood smears of PMN^WT and PMN^ΔGLUT1 mice were stained with Giemsa stain and microscopically evaluated for the morphology of neutrophils. The staining was visualized using an EVOS FL Auto microscope.

Transwell migration assay

BM neutrophils were subjected to transwell migration assay (inserts with 3 μm pore size) in the presence or absence of recombinant murine CXCL2 (100 ng/ml). Ninety min post-incubation, the numbers of cells in the lower and upper chambers were quantified by flow cytometry analysis and expressed as Chemotactic Index (Number of cells in the lower chamber/Number of cells in the upper chamber).

Granulopoiesis measurement

BM cells from femur and tibia were stained with anti-Ly6G, anti-CD34, anti-CD3, anti-CD19, anti-B220 and anti-C-kit antibodies and analyzed by flow cytometry analysis.

Apoptosis detection

Mice were infected with 10⁵ CFU of C. albicans via the lateral tail vein. After 24 hours, kidney cell suspensions were prepared and early and late apoptotic neutrophils were detected using Annexin V Apoptosis Detection Kit, according to manufacturer instructions, followed by flow cytometry analysis.

ROS measurement

Mice were infected with 10⁵ CFU of C. albicans via the lateral tail vein. After 24 hours, kidney cell suspensions were prepared and resuspended with RPMI complete medium. CellROX^® Deep Red reagent was added to the cell suspensions and incubated at 37°C for 30 min. The ROS production was measured by flow cytometry analysis.

Western blot analysis

Neutrophils were lysed by 1×NP-40 lysis buffer supplemented with protease inhibitor cocktail. Lysates were separated by SDS-PAGE and transferred to polyvinylidene difluoride membranes. After incubation with primary and secondary antibodies, protein bands were detected using an enhanced chemiluminescence detection system and developed with a FluorChem E imager (ProteinSimple).

RNA extraction and qPCR

RNA was extracted from tissues or neutrophils using RNeasy kits. Complementary DNA was synthesized by SuperScript III First Strand Kits. Quantitative real-time PCR (qPCR) was performed with the PerfeCTa SYBR Green FastMix and analyzed on an ABI 7300 real-time instrument. Primers were obtained from QuantiTect Primer Assays. The expression of each gene was normalized to that of Gapdh.

Seahorse metabolic assay

BM neutrophils from PMN^WT and PMN^ΔGLUT1 mice were isolated and plated on Cell-Tak coated Seahorse culture plates (5 × 10⁵ cells/well) in DMEM without glucose. After stimulated with/without curdlan (Cur) (10 μg/ml) at 37°C for 3 h, the cells were analyzed using a Seahorse XFe96 Analyzer (Agilent). EACR (Basal extracellular acidification rate) was detected in the presence of glucose (25mM), oligomycin (2 mM), 2-deoxyglucose (100 mM) to obtain maximal and control EACR values.

Untargeted high-resolution LC-HRMS

Neutrophils from PMN^WT and PMN^△GLUT1 mice were either stimulated with curdlan (10 μg/ml) for 3 h (8 replications per condition) or left unstimulated. Metabolic quenching and polar metabolite pool extraction from neutrophils was performed by adding 500μL 80% methanol/ 0.1% formic acid spiked with deuterated (D3)-creatinine and (D3)-alanine, (D4)-taurine and (D3)-lactate as an internal standard for a final concentration of 10μM. After 5 min of vertexing, the supernatant was cleared of protein by centrifugation at 16,000×g. 2μL of cleared supernatant was subjected to online LC-MS analysis.

Analyses were performed by untargeted LC-HRMS. Briefly, samples were injected via a Thermo Vanquish UHPLC and separated over a reversed phase Thermo HyperCarb porous graphite column (2.1×100mm, 3μm particle size) maintained at 55°C. For the 20 min LC gradient, the mobile phase consisted of the following: solvent A (water / 0.1% FA) and solvent B (ACN / 0.1% FA). The gradient was the following: 0–1min 1% B, increase to 15%B over 5 minutes, continue increasing to 98%B over 5 min, hold at 98%B for 5 min, re-equilibrate at 1%B for 5 min. The Thermo IDX tribrid mass spectrometer was operated in both positive and ion mode, scanning in ddMS2 mode (2 μ scans) from 70 to 800 m/z at 120,000 resolutions with an AGC target of 2×10⁵ for full scan, 2×10⁴ for ms2 scans using HCD fragmentation at stepped 15, 35, 50 collision energies. Source ionization setting was 3.0 and 2.4kV spray voltage, respectively for positive and negative mode. Source gas parameters were 35 sheath gas, 12 auxiliary gas at 320°C, and 8 sweep gas. Calibration was performed prior to analysis using the PierceTM FlexMix Ion Calibration Solutions. Integrated peak areas were then extracted manually using Quan Browser (Xcalibur ver. 2.7) and normalized to the peak areas of the spiked internal standards to control for both sample and instrument variation. Untargeted differential comparisons were performed using Compound Discoverer 3.0 to generate a ranked list of significant compounds with tentative identifications from BioCyc, KEGG, and internal compound databases.

Quantification and statistical analysis

All data are expressed as means ± SEM. Statistical analyses were performed using the analysis of variance (ANOVA), log-rank, Mann-Whitney, or Student’s t test through GraphPad Prism 8 program. *P < 0.05; **P < 0.01; ***P < 0.001.

KEY RESOURCES TABLE.

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
PerCP-Cy™5.5 Rat Anti-Mouse CD45 (Clone 30-F11 (RUO))	BD Biosciences	Cat # 561869; RRID: AB_10895563
Anti-phospho GLUT-1 Antibody (Ser226)	Millipore	Cat # ABN991
APC anti-mouse CD45 Antibody (Clone 30-F11)	Biolegend	Cat # 103112; RRID: AB_312977
CD45 Monoclonal Antibody, eFluor 450, eBioscience™ (Clone 30-F11)	Invitrogen	Cat # 48-0451-82; RRID: AB_1518806
Alexa Fluor® 700 anti-mouse/human CD11b Antibody (Clone M1/70)	Biolegend	Cat # 101222; RRID:AB_493705
Ly-6G Monoclonal Antibody, FITC, eBioscience™ (Clone 1A8-Ly6g)	Invitrogen	Cat# 11-9668-82; RRID: AB_2572532
Ly-6G Monoclonal Antibody (1A8-Ly6g)	Invitrogen	Cat# 48-9668-82; RRID:AB_2637124
Ly-6G Monoclonal Antibody, PE, eBioscience™ (Clone 1A8-Ly6g)	Invitrogen	Cat# 12-9668-82; RRID: AB_2572720
Ly-6C Monoclonal Antibody, PerCP-Cyanine5.5, eBioscience™ (Clone HK1.4)	Invitrogen	Cat# 45-5932-80; RRID: AB_2723342
Brilliant Violet 605™ anti-mouse F4/80 antibody, BioLegend (Clone BM8)	Biolegend	Cat# 123133; RRID:AB_2562305
F4/80 Monoclonal Antibody, APC, eBioscience™ (Clone BM8)	Invitrogen	Cat# 17-4801-82; RRID: AB_469452
Rabbit IgG Isotype Control (Alexa Fluor® 488 Conjugate)	Cell Signaling Technology	Cat# 4340S; RRID:AB_561545
Rabbit (DA1E) mAb IgG XP® Isotype Control	Cell Signaling Technology	Cat# 3900; RRID:AB_1550038
Alexa Fluor 647-AffiniPure Goat Anti-Rabbit IgG (H+L) antibody,	Jackson Immuno	Cat# 111-605-003; RRID: AB_2338072
PE Rat Anti-Mouse c-KIT (Clone 2B8)	BD Biosciences	Cat# 553355; RRID: AB_394806
Alexa Fluor® 700 Rat Anti-Mouse CD34 (Clone RAM34)	BD Biosciences	Cat# 560518; RRID: AB_1727471
PerCP anti-mouse/human CD45R/B220 Antibody (Clone RA3-6B2)	Biolegend	Cat# 103233; RRID: AB_893355
PerCP/Cy5.5 anti-mouse CD19 Antibody (Clone 1D3/CD19)	Biolegend	Cat# 152405; RRID: AB_2629814
PerCP Hamster Anti-Mouse CD3e (Clone 145-2C11)	BD Biosciences	Cat# 553067; RRID: AB_394599
Pacific Blue™ anti-mouse Lineage Cocktail	Biolegend	Cat# 133310; RRID:AB_11150779
CD16/CD32 Monoclonal Antibody (93), PerCP-Cyanine5.5,	eBioscience™	Cat # 45-0161-80; RRID:AB_996661
APC anti-mouse Ly-6A/E (Sca-1) Antibody	Biolegend	Cat # 108111; RRID:AB_313348
APC anti-mouse CX3CR1 Antibody	Biolegend	Cat # 149007; RRID:AB_2564491
Anti-Glucose Transporter GLUT1-FITC antibody	Abcam	Cat # ab195359; RRID:AB_2714026
Phospho-PKCdelta (Tyr311) Antibody	Cell Signaling Technology	Cat # 2055; RRID:AB_330876
PKCδ (D10E2) Rabbit mAb	Cell Signaling Technology	Cat # 9616; RRID:AB_10949973
IκB-α Antibody (C-21): sc-371	Santa Cruz	Cat # SC-371; RRID:AB_2235952
Anti-GLUT-1 Antibody, CT	Millipore	Cat # 07-1401; RRID:AB_1587074
Anti-Glucose Transporter GLUT1 antibody [EPR3915]	Abcam	Cat# ab115730; RRID:AB_10903230
Anti-Glucose Transporter GLUT3 antibody [EPR10508(N)1 - N-terminal	Abcam	Cat# ab191071; RRID: AB_2736916
Anti-beta Actin antibody [AC-15] (HRP)	Abcam	Cat# ab49900; AB_867494
Anti-Neutrophil Elastase antibody	Abcam	Cat# ab21595; RRID:AB_446409
Goat anti-rabbit IgG (HPL) HRP conjugated	Invitrogen	Cat# G21234; RRID:AB_2536530
Bacterial and virus strains
Biological Samples
Healthy human peripheral blood samples	Division of Rheumatology and Clinical Immunology, University of Pittsburgh.	See Table S1
Chemicals, Peptides, and Recombinant Proteins
WZB117	Sigma	Cat# SML0621; CAS: 1223397-11-2
mannan	sigma	Cat# M7504; CAS: 903688-8
curdlan	sigma	Cat# C7821; CAS: 54724-00-4
Zymosan	sigma	Cat# Z4250; CAS: 58856-93-2
Rottlerin	Tocris	Cat# 1610; CAS: 82-08-6
Bay-61-3606	sigma-Calbiochem	Cat# 574714; CAS: 732938-37-8
Bay 11-7085	sigma-Calbiochem	Cat# B5681; CAS: 196309-76-9
GW5074	Sigma	Cat# G6416; CAS: 220904-83-6
CellROX® Deep Red reagent	Invitrogen	Cat#C10422
SYTOX™ Orange Nucleic Acid Stain	Sigma	Cat#S11368
Phorbol-12-myristate-13-acetate	Sigma-Calbiochem	Cat# 524400; CAS: 16561-29-8
Recombinant Murine MIP-2 (CXCL2)	Peprotech	Cat# 250-15
Ghost Dye™ Violet 510	TONBO biosciences	Cat# 13-0870-T100
CountBright™ Absolute Counting Beads	Invitrogen	Cat# C36950
GlucoseCy5	Dr. Greg M. Delgoffe, University of Pittsburgh	Watson et al., 2021
2-NBDG	Cayman Chemical Company	Cat#11046; CAS: 186689-07-6
Biotin-XX, sulfo-SE	Invitrogen	Cat# B6352
Collagenase IV	ThermoFisher Scientific	Cat# 17104019
Gibco™ ACK Lysing Buffer	ThermoFisher Scientific	Cat# A1049201
Corning™ EDTA, 0.5M, pH8.0	Fisher Scientific	Cat# MT-46034CI
DNAsel	MP Biomedicals	Cat# 190062
Streptavidin, Alexa Fluor 633 conjugate	Life Technologies Corporation	Cat# S21375
Giemsa Stain (Powder/Certified Biological Stain)	Fisher Chemical™	Cat# G146-5
Protease inhibitor cocktail	Roche	Cat# 11697498001
Critical Commercial Assays
PE Annexin V Apoptosis Detection Kit I	BD Pharmingen	Cat# 559763
Neutrophil Isolation kit mouse	Miltenyi Biotec	Cat# 130-097-658
SuperScript™ III First-Strand Synthesis kit	Invitrogen	Cat# 18080051
RNeasy Mini Kit	Qiagen	Cat# 74106
MACSxpress® Neutrophil Isolation kit, Human	Miltenyi Biotec	Cat# 130-104-434
Deposited Data
Experimental Models: Cell Lines
Experimental Models: Organisms/Strains
Mouse: C57BL/6NTac	Taconic Biosciences	Model# B6-M
Mouse: Card9−/−: B6.129-Card9tm1Xlin/J	The Jackson Laboratory	Cat# 028652
Mouse: Dectin-1−/−: B6.129S6-Clec7atm1Gdb/J	The Jackson Laboratory	Cat# 012337
Mouse: MRP8-Cre+: B6.Cg-Tg(S100A8-cre,-EGFP)1Ilw/J	The Jackson Laboratory	Cat # 021614
Mouse: Myd88−/−: B6.129P2(SJL)-Myd88tm1.1Defr/J	The Jackson Laboratory	Cat# 009088
Mouse: Slc2a1fl/fl	University of Colorado; MTA	N/A
Mouse: Act1−/−	Dr. Sarah L. Gaffen University of Pittsburgh	N/A
C. albicans: SC5314	Dr. Sarah Gaffen, University of Pittsburgh	ATCC: MYA-2876
dTomato+ C. albicans reporter strain	Dr. Michail Lionakis, NIH	N/A
Oligonucleotides
Primers	QIAGEN	See Table S2
Recombinant DNA
Software and Algorithms
Image J	National Institute of Health	https://imagej.nih.gov/ij/
FlowJo v10	Tree Star	https://www.flowjo.com/
Alpha View SA v3.4.0	ProteinSimple	https://www.proteinsimple.com/
GraphPad Prism 8	GraphPad	https://www.graphpad.com/
IDEAS applications 6.1	Luminex	https://www.luminexcorp.com/flow-cytometry-and-imaging/
EndNote ×9	EndNote	https://endnote.com/
Compound Discoverer 3.0	Thermo Fisher	https://assets.thermofisher.com/
Other

Highlights.

• Neutrophils upregulate Glut1 and glucose uptake in disseminated candidiasis

• The β-glucan/dectin-1-axis transcriptionally and post-translationally modulates Glut1

• Glut1 deficiency in neutrophils exacerbates susceptibility to disseminated candidiasis

• Glut1 is indispensable for antifungal activity in mice and human neutrophils