Diet-induced obesity in mice impairs host defense against Klebsiella pneumonia in vivo and glucose transport and bactericidal functions in neutrophils in vitro

Abstract

Obesity impairs host defense against Klebsiella pneumoniae, but responsible mechanisms are incompletely understood. To determine the impact of diet-induced obesity on pulmonary host defense against K. pneumoniae, we fed 6-wk-old male C57BL/6j mice a normal diet (ND) or high-fat diet (HFD) (13% vs. 60% fat, respectively) for 16 wk. Mice were intratracheally infected with Klebsiella, assayed at 24 or 48 h for bacterial colony-forming units, lung cytokines, and leukocytes from alveolar spaces, lung parenchyma, and gonadal adipose tissue were assessed using flow cytometry. Neutrophils from uninfected mice were cultured with and without 2-deoxy-d-glucose (2-DG) and assessed for phagocytosis, killing, reactive oxygen intermediates (ROI), transport of 2-DG, and glucose transporter (GLUT1–4) transcripts, and protein expression of GLUT1 and GLUT3. HFD mice had higher lung and splenic bacterial burdens. In HFD mice, baseline lung homogenate concentrations of IL-1β, IL-6, IL-17, IFN-γ, CXCL2, and TNF-α were reduced relative to ND mice, but following infection were greater for IL-6, CCL2, CXCL2, and IL-1β (24 h only). Despite equivalent lung homogenate leukocytes, HFD mice had fewer intraalveolar neutrophils. HFD neutrophils exhibited decreased Klebsiella phagocytosis and killing and reduced ROI to heat-killed Klebsiella in vitro. 2-DG transport was lower in HFD neutrophils, with reduced GLUT1 and GLUT3 transcripts and protein (GLUT3 only). Blocking glycolysis with 2-DG impaired bacterial killing and ROI production in neutrophils from mice fed ND but not HFD. Diet-induced obesity impairs pulmonary Klebsiella clearance and augments blood dissemination by reducing neutrophil killing and ROI due to impaired glucose transport.

INTRODUCTION

The prevalence of obesity has increased dramatically in the United States and other high- and middle-income nations during the past 40 years, contributing to sizeable increases in morbidity and health care costs (1). Recent estimates indicate that 42.4% of adults 20 yr and older in the United States have obesity, based on a body mass index (BMI) ≥30 kg/m² (2). Obesity is a well-known predisposing risk factor for cardiovascular disease, type 2 diabetes, and hypertension, and it was recently shown to have a negative impact on certain viral and bacterial respiratory tract infections (3–5). During the 2009 influenza pandemic, individuals with obesity or metabolic syndrome experienced greater severity of illness and mortality from the H1N1 virus (6). Recently, obesity was associated with a greater severity of illness among patients with COVID-19 (7). Hence, defining the molecular basis of the effects of obesity on host defense against respiratory infections has significant implications for global public health.

Although reports associating obesity with a greater severity of illness from influenza are consistent, the impact of obesity on other human respiratory tract infections is controversial. Obesity was associated with a greater risk of pneumonia in several epidemiologic studies (8–13), but in contrast, others have reported improved outcomes in patients with obesity and pneumonia, relative to nonobese patients (14–18). In humans with obesity, neutrophil antibacterial function was impaired in some (19–21), but not all (22, 23) reports, suggesting that differing prevalence of obesity-associated comorbidities or methodological differences may lead to these divergent results.

Murine models are of considerable potential utility to define mechanistically the effects of obesity on lung host defense. However, despite impairments in antiviral host defense in murine models of obesity following infection with mouse-adapted influenza viruses (24–27), inconsistent results have been reported in models of bacterial pneumonia in obese mice. Leptin-deficient (ob/ob), leptin receptor-deficient (db/db), diet-induced obese, and carboxypeptidase E-deficient (CPE^fat/fat) mice all exhibited impaired pulmonary bacterial clearance of Klebsiella pneumoniae (5, 28) and ob/ob mice exhibited a greater severity of illness from Streptococcus pneumoniae (29). These two organisms are leading causes of hospital-acquired and community-acquired pneumonias, respectively. In contrast, we found no differences in pulmonary host defense between female wild-type control and obese CPE^fat/fat mice infected with S. pneumoniae (30). Reasons for these disparate results besides sex differences include differences in the underlying metabolic defects that produce obesity in genetic versus dietary models; immune suppression associated with the lack of leptin receptor signaling; and pathogen-specific impairments in host defense against bacterial pneumonia. Nevertheless, consistent findings among these reports were impairments in host defense against Klebsiella pneumoniae and reduced neutrophil bactericidal function in obese leptin-deficient mice (5, 28, 29, 31).

Neutrophils play a critical role in pulmonary host defense against Klebsiella pneumonia; impairments in their recruitment to the alveoli and in reactive oxygen intermediate (ROI) production are associated with poor outcomes in human patients and animal models (32–34). The few studies that have evaluated the impact of obesity on neutrophil function in humans have shown inconsistent results in bactericidal endpoints such as phagocytosis, NETosis, reactive oxygen intermediate production, and cytokine production (22, 23, 35, 36). In contrast, impairments in neutrophil recruitment into the alveolar space following infection with K. pneumoniae and reduced phagocytosis of B. burgdorferi were reported in diet-induced obese mice (5, 37). Whether neutrophil-mediated killing of bacteria is reduced in diet-induced obese mice has not, to our knowledge, been determined.

In the present study, we examined the impact of diet-induced obesity on host defense against Klebsiella pneumonia in mice and analyzed mechanisms crucial to control neutrophil bactericidal function.

MATERIALS AND METHODS

Animals

Four-week-old male C57BL/6j mice were purchased from Jackson Laboratories (Cat. No. 000664, Bar Harbor, ME) and were housed in a specific pathogen-free facility with a 12-h light/12-h dark cycle. Starting at 6 wk of age, mice were fed ad libitum either a normal fat diet (ND) composed of 13% fat (5001; Laboratory Diet, St. Louis, MO) or a high-fat diet (HFD) consisting of 60% fat (D12492; Research Diets, New Brunswick, NJ) for 16 wk. Mice were treated according to National Institutes of Health guidelines for the use of experimental animals with the approval of the Institutional Animal Care and Use Committee at the University of Michigan.

Blood Glucose and Body Composition

Blood was collected from the tails of mice in the fed state before euthanasia and glucose was assessed using a Bayer Contour Glucose Meter (Ascensia Diabetes, Parsippany, NJ). Mice were weighed and body composition was determined noninvasively using an EchoMRI 100 (Echo MRI, Houston, TX).

Klebsiella pneumoniae Preparation and Infection

Klebsiella pneumoniae strain 43816, serotype 2, was purchased from the American Type Culture Collection (Manassas, VA) and grown in tryptic soy broth (DIFCO, Detroit, MI) for 18 h at 37°C and 5% CO₂ in air. The suspension of K. pneumoniae was pelleted by centrifugation (20,000 g for 3 min) and resuspended in phosphate-buffered saline (PBS), twice, to wash the bacteria. We determined the concentration of bacteria using a spectrophotometer (A₆₀₀) and a standard curve and obtained the appropriate concentration by serial dilution in PBS.

Following anesthesia of the mice using ketamine and xylazine, a midline incision was made to expose the trachea, a 30-μL inoculum containing 5 × 10³ colony-forming units (CFU) K. pneumoniae was administered via an injection in the trachea using a 26-gauge needle, then the wound was closed using surgical glue (Nexaband, Phoenix, AZ) as previously described (28). To confirm the actual concentration of the inoculum, a sample was grown on soy-based blood agar No. 2 (Cat. No. BD Difco, Detroit, MI) overnight at room temperature.

Determination of Lung and Spleen K. pneumoniae Colony-Forming Units

At 24 and 48 h after K. pneumoniae infection, mice were euthanized by CO₂ asphyxiation, and then lungs and spleen were harvested, homogenized in 0.5 mL of sterile PBS, serially diluted, and plated on soy-based blood agar plates. After 18 h at room temperature, K. pneumoniae colony-forming units (CFUs) were enumerated.

Determination of Cytokines in Lung Homogenates

Using lung homogenates from uninfected mice and the same lung homogenates as for CFU determinations from infected mice, we determined the levels of cytokines [IL-1β (Cat. No. DY401), IL-6 (Cat. No. DY206), IL-17 (Cat. No. DY421), IFN-γ (Cat. No. DY485), CCL1 (Cat. No. DY845), CXCL2 (Cat. No. DY452), and TNF-α (Cat. No. DY220)] using commercially available enzyme-linked immunosorbent assay (ELISA) kits according to the manufacturer’s instructions (R&D Duoset, R&D Systems, Minneapolis, MN).

Recovery and Identification of Lung Leukocytes from Lung Homogenates and Bronchoalveolar Lavage Fluid

In separate groups of mice euthanized at baseline or 24 h after infection, we first depleted intraalveolar leukocytes by bronchoalveolar lavage (BAL) using 2-mL HEPES-buffered saline as previously described (38). Next, for total lung leukocyte counts, lungs were excised and collected in 20 mL of cold Dulbecco’s PBS medium (DPBS; Cat. No. 14190144; Gibco, Thermo Fisher, Carlsbad, CA) with 30 μg/mL DNAase (Cat. No. 10104159001, Sigma-Aldrich, St. Louis, MO). Lung lobes were blended in a Waring blender at low speed for 30 s, and the entire lung homogenate was centrifuged. The resuspended homogenate was filtered through a 40-μm cell strainer (Cat. No. BD 352340, BD Biosciences, San Jose, CA), and recovered cells were washed three times by centrifugation before staining. Leukocytes were enumerated using a hemocytometer.

Leukocyte Isolation from Gonadal White Adipose Tissue

Isolation of leukocytes from the adipose tissue stromal vascular fraction of gonadal white adipose tissue (GWAT) was performed as previously described (39).

Flow Cytometry Protocol

To stain cells recovered from lung homogenates, GWAT, and bronchoalveolar lavage fluid (BALF), leukocytes were suspended in 2-mL DPBS buffer with 2% (vol/vol) fetal bovine serum and incubated with 10 μg/mL of TruStain fcX anti-CD16/32 block (BioLegend, San Diego, CA) for 10 min at 4°C. Next, samples were equally divided, and cocktails of prealiquoted monoclonal antibodies (mAb; 100 μL total volume) were added, using 0.5 μL of mAb per 100 μL of staining buffer. We employed two mAb panels: for myeloid cell types [CD11c-APC-Cy7, CD64-PE, Ly6G-FITC, CD115-APC, Ly6C-PerCPCy5.5, CD45.2-Pacific Blue (eBiosciences), F4/80-PerCP-Cy5.5, CD11c-PE-Cy5, CD11b-AF488 (BioLegend)] and for lymphoid cell types [CD103-PE, B220-PE Texas Red (BD PharMingen, San Jose, CA), CD4-Pacific Blue, CD8-APC-Cy7, CD3e-PerCP-Cy5.5, CD44-AF700, NK1.1-AF647, CD62L-AF488, CD25-PE-Cy7, CD69-PE-Cy5 (BioLegend)]. Macrophages were identified as high autofluorescent CD64⁺ and CD11c⁺ and CD64⁺ and CD11c⁻ cells that were negative for CD3 and Ly6G. Neutrophils were identified as high side scatter cells that were negative for CD11c and abundantly expressing Ly6C. Dendritic cells (DC) were identified as low autofluorescent, CD64⁻, and CD11c⁺. Preparations were incubated for 30 min at 4°C in the dark with constant agitation, and then washed using 2 mL of staining buffer. For data acquisition and analysis, we used a FACScan LSR II flow cytometer (BD Biosciences, San Diego, CA) and FlowJo software, version v10 (Tree Star Inc., Ashland, OR), with instrument configuration as previously described (40). Representative gating of leukocytes in lung homogenates, GWAT, and BALF is shown in Supplemental Figs. S1–S3 (all Supplemental material is available at https://doi.org/10.6084/m9.figshare.16722844.v1).

Recovery of Glycogen-Elicited Peritoneal Neutrophils (PMNs)

Elicited polymorphonuclear neutrophils (PMNs) were obtained from uninfected ND and HFD mice by peritoneal lavage 5 h after an intraperitoneal injection of 1% glycogen solution in saline. Ninety percent of the cells obtained by peritoneal lavage were identified as PMNs by a modified Wright-Giemsa stain (Diff-Quik; American Scientific Products, McGaw Park, IL). Following PMN isolation, the cells were enumerated using a hemocytometer, suspended in RPMI 1640 (Gibco, Grand Island, NY) to a final concentration of 1 × 10⁶ cells/mL.

Recovery of Bone Marrow Neutrophils

Bone marrow cells were isolated from tibia and fibula of mice by flushing with PBS. Bone marrow cells were passed through a syringe with an 18-gauge needle, filtered through a 100-μm mesh, pelleted by centrifugation for 10 min at 300 g at 4°C, and resuspended in red blood cell (RBC) lysis buffer (Cat. No. 00-4333-57, Thermo Fisher Scientific, Waltham, MA). After a second centrifugation, bone marrow cells were resuspended in 45% Percoll (Cat. No. 65455-52-9, Thermo Fisher Scientific) and layered on top of a gradient consisting of 50%, 55%, 62%, and 81% Percoll. After centrifugation at 1,500 g for 30 min, the layer of cells between the 62% and 81% layers of Percoll was removed, and the cells were resuspended in RPMI. The morphology of the cells in suspension was assessed by cytocentrifuging a sample onto glass slides and staining with Diff-Quik. Greater than 90% of the cells were identified as PMNs. The concentration of PMNs was adjusted to 1 × 10⁶ cells/mL in RPMI.

Analysis of Glucose Transporter [(Slc2a1 (GLUT1), Slc2a2 (GLUT2), Slc2a3 (GLUT3), and Slc2a4 (GLUT4)] Gene Expression

RNA was isolated from PMNs using Trizol according to the manufacturer’s protocol (Thermo Fisher Scientific, Inc. (Waltham, MA). Quantitative real-time PCR (RT-qPCR) was performed by synthesizing cDNA from 0.5 to 1 mg of RNA using the High Capacity Reverse Transcription Kit (Cat. No. 4368814, Life Technologies). Primers, cDNA, and Power SYBR Green PCR Master Mix (Cat. No. 4367659, Life Technologies) were combined as instructed by the manufacturer’s guidelines, and quantitative real-time polymerase chain reaction (qPCR) was performed using QuantStudio 5 (Thermo Fisher Scientific) as previously described (41). Messenger RNA (mRNA) expression level was normalized to Gapdh and analyzed using the 2^−ΔΔCT threshold cycle method after evaluation of several reference genes. qPCR primer sequences are listed in Table 1. Data were normalized to the expression level of (Slc2a3) GLUT3, as it was the most abundant transcript for GLUTs in PMNs from mice fed the ND.

Table 1.

Primers used for reverse-transcription qPCR

Gene	Forward Sequence (5′–3′)	Reverse Sequence (5′–3′)
Slc2a1	TGGCGGGAGACGCATAGTTA	CTCCCACAGCCAACATGAGG
Slc2a2	TGTTGGGGCCATCAACATGA	ACATGCCAATCATCCCGGTT
Slc2a3	GAGGAACACTTGCTGCCGAG	CTGGAAAGAGCCGATCGTGG
Slc2a4	GCCCGGACCCTATACCCTAT	GTCACTCGCTGCCGAGG
Gapdh	CACTTGAAGGGTGGAGCCAA	ACCCATCACAAACATGGGGG

Glucose Transport in PMNs

To determine glucose transport into bone marrow PMNs from mice fed ND versus HFD, we assessed uptake of 2-deoxy-d-[¹⁴C (U)] glucose (2-DG). Bone marrow PMNs, isolated as described earlier, were adhered to 24-well polystyrene plates coated with 3.5 μg/cm² of Cell-Tak (Cat. No. CB40240, Corning, Tewksbury, MA) at a concentration of 5 × 10⁵ cells per well and cultured with 2-DG. The cells were washed with PBS twice and incubated with 500-μL Krebs Ringer Bicarbonate Buffer (KRBH; 30 mM HEPES, pH 7.4, 10 mM NaHCO₃, 120 mM NaCl, 4 mM KH₂PO₃, 1 mM MgSO₄, and 1 mM CaCl₂ with 0.5% BSA) for 30 min at 37°C. 2-DG uptake was initiated by the addition of 50 μL of freshly prepared reaction mixture containing 0.1 μCi of 2[¹⁴C (U)]-DG (New England Nuclear, PerkinElmer, Waltham, MA) and 0.05 μL of 200 mM cold 2-DG (Cat. No. D8375, Sigma-Aldrich; final concentration, 20 μM) in KRBH buffer at room temperature. Following incubation for 5 min, the reaction was stopped by adding 50 μL of 200 mM 2‐DG. The wells were washed three times with ice‐cold PBS and scraped following the addition of 0.5 mL of ice cold PBS with 1% SDS. A 10-μL aliquot of each sample was removed to assess protein concentration using a Bradford assay kit (Cat. No. 23200, Thermo Fisher Scientific). Radioactivity of 400 μL of each sample was assessed by scintillation counting in liquid scintillation fluid (PerkinElmer, Waltham, MA). The results were calculated as pM/mg protein/5 min and are expressed as means ± SE of four independent experiments performed in quadruplicate.

Immunoblot Analysis of Glucose Transporters 1 and 3

Neutrophils were obtained from the bone marrow of mice after being fed the ND and HFD for 16 wk as mentioned earlier. Neutrophils were lysed in ice-cold lysis buffer (RIPA buffer; Sigma), and cells were disrupted with sonication (10 bursts at 20% duty/cycle). Two and one-half micrograms of protein, as determined by a Pierce BCA Protein Assay Kit (Pierce Chemical, Rockford, IL), was separated by SDS-PAGE under reducing conditions and transferred to nitrocellulose membranes. Membranes were probed with mouse and rabbit (respectively) primary polyclonal Abs against glucose transporter 1 and 3 (GLUT1; SPM 498; 1:250) and (GLUT3; Ab41525; 1:250; Abcam, Wathman, MA), respectively. Primary Abs were detected using Alexa Fluor 680 goat anti-mouse IgG (H + L; Ref. A21058; titer 1:5,000) and Alexa Fluor 790 goat anti-rabbit IgG (H + L; Ref. A113669; titer 1:10,000; Thermo Fisher Scientific) and visualized with the Li-Cor Odyssey CLx Imaging System (Lincoln, NE). The density values of the GLUT1 and GLUT3 blots were quantified using Image Studio software, version 5.2 (Li-Cor Biosciences) and normalized to Revert Total Protein Stain (Cat. No. 926-11011; Li-Cor Biosciences).

Fluorometric Assay of Neutrophil Phagocytosis

PMN phagocytosis of K. pneumoniae was assessed using a published protocol that employs FITC-labeled heat-killed organisms (42), which were opsonized with 5% normal rat serum in Hanks’ buffered salt solution (Thermo Fisher Scientific) for 15 min as previously described (43). Briefly, 1.25 × 10⁵ PMNs/well were seeded into 384-well tissue culture plates with opaque sides and optically clear bottoms (Cat. No. 142761, Thermo Scientific, Nunc), allowed to adhere for 1 h in RPMI 1640 media. PMNs were infected with opsonized FITC–K. pneumoniae using a multiplicity of infection (MOI) of 150:1 and cultured for 60 min to allow phagocytosis to occur. Trypan blue (Cat. No. 72-57-1, 250 μg/mL; Argos Organics, Thermo Fisher Scientific) was added for 1 min to quench the fluorescence of extracellular bacteria, then fluorescence was determined using a Spectramax Gemini EM fluorometer with 485-nm excitation/535-nm emission (Molecular Devices, Sunnyvale, CA). The phagocytic index was calculated in relative fluorescence units (RFUs) as previously described (42). Four separate experiments were conducted with six replicate wells for every experimental condition.

Bactericidal Assays

We determined the survival of opsonized K. pneumoniae following culture with PMNs as described previously (44). Briefly, 1 × 10⁵ PMNs were adhered to 96-well plates (Corning, Lowell, MA) for 1 h in RPMI and a suspension of K. pneumoniae (1 × 10⁴ CFU) opsonized with 5% normal rat serum was added to each well. The MOI was 0.1 (1:10). In some experiments, PMNs were pretreated with media alone or 10 mM 2-deoxy-d-glucose for 30 min before addition of K. pneumoniae. Next, the plate was centrifuged for 1 min at 10 g and was incubated at 37°C for 90 min. To assess bacterial counts, the supernatant was collected at 45 and 90 min and plated at serial dilutions on blood agar. As a control, wells without PMNs included K. pneumoniae only to permit the observation of bacterial growth during the assay. Results were expressed as percentage of survival of the initial bacterial inoculum.

Reactive Oxygen Intermediate Production

PMNs were adhered to 384-well plates at a concentration of 1.25 × 10⁵ cells/well and cultured with PBS containing 10 μM of the cell-permeant fluorescent probe 2′,7′-dichlorodihydrofluorescein diacetate (H2DCF; Cat. No. D399, Invitrogen, Carlsbad, CA) for 1 h. The medium was then replaced with warmed RPMI, and the cells were cultured with or without 10 mM 2-DG before stimulation with heat-killed K. pneumoniae (HKKP) opsonized with 5% normal rat serum at an MOI of 50:1. ROI production was assessed 90 min later by measuring fluorescence using a Spectramax Gemini XS fluorometer (Molecular Devices) with excitation/emission setting at 493/522 nm.

Statistical Analyses

Statistical analyses were conducted using Prism software (version 7.0, GraphPad Software, La Jolla, CA). Where appropriate, mean values were compared using a paired Student’s t test. Differences were considered significant if P < 0.05. The number of experiments performed for each figure are mentioned in the figure legends. Data are presented as mean values ± SE, unless noted otherwise.

RESULTS

Sixteen Weeks of HFD Increases Body Weight, Body Fat, and Blood Glucose

After 16 wk of ad libitum feeding, animals fed the HFD weighed ∼40% more than those on the ND. Body composition assessment revealed that the significantly increased body weight (Fig. 1A) on HFD was entirely due to increases in body fat content (average ∼20 g difference; Fig. 1B), with no differences in lean body weight (Fig. 1C). In addition, blood glucose levels were higher in mice fed the HFD (Fig. 1D). These results are consistent with previous reports demonstrating that ad libitum access to a HFD for 16 wk results in obesity and diabetes (39).

Figure 1.

Effect of high-fat diet (HFD) intake on body composition and blood glucose. Starting at 6 wk of age, male C57BL/6 mice were fed a normal diet (ND) containing 13% fat (open circles) or HFD containing 60% fat (closed squares) for 16 wk. Body weight (A), lean body mass (B), fat body mass (C), and blood glucose levels (D) were determined as described in materials and methods. Bars represent means ± SE. n = 8 mice per group. *P < 0.05 (compared with ND using a Student’s t test).

Defective Pulmonary Bacterial Clearance in Mice Fed a HFD

Next, we assessed the impact of the HFD feeding on pulmonary host defense against a clinically relevant pathogen in a murine model of Gram-negative pneumonia. At 24 h after infection, the bacterial loads in HFD mice were higher than in ND, both in the primary site of infection in the lungs (2.4-fold) and in the spleens (1.8-fold), indicating hematogenous dissemination (Fig. 2A). Forty-eight hours after infection, the bacterial burdens in HFD mice were 5-log fold and 4-log fold higher in lungs and spleens, respectively (Fig. 2B). These results demonstrate that diet-induced obesity profoundly impairs pulmonary bacterial clearance following K. pneumoniae infection and leads to greater bacterial dissemination to the bloodstream, as reflected in the spleen.

Figure 2.

Greater lung and spleen bacterial loads following K. pneumoniae infection in HFD fed mice. Mice fed ND (open circles) or HFD (closed squares) for 16 wk were intratracheally infected with 5,000 CFUs of K. pneumoniae; lungs and spleens were assessed for bacterial loads 24 h (A) and 48 h (B) later, as described in materials and methods. Bars represent means ± SE of n = 5 mice per group from three independent experiments. *P < 0.05 (compared with ND using a Student’s t test). CFUs, colony-forming units; HFD, high-fat diet; ND, normal diet.

Impact of HFD on Pulmonary Cytokines at Baseline and following K. pneumoniae Infection

Obesity is associated with a chronic state of low-grade systemic inflammation characterized by elevated proinflammatory adipokines and cytokines (45). To determine if the HFD had an impact on pulmonary cytokines at baseline and after infection, we assessed cytokine levels in lung homogenates by ELISA. Unexpectedly, we observed lower levels of IL-1β, IL-6, IL-17, IFN-γ, CXCL2, and TNF-α in lung homogenates recovered at baseline from mice fed the HFD (Fig. 3A). In contrast, mice fed the HFD were found to have higher levels of IL-6, CCL2, and CXCL2 at both 24 and 48 h, and of IL-1β (24 h only), relative to ND-fed mice (Fig. 3, B and C). We did not observe differences post-K. pneumoniae infection between mice fed the two diets in IL-17, IFN-γ, CCL2, or TNF-α, another unexpected finding, as these cytokines have been reported to be elevated in murine models of obesity (46–48).

Figure 3.

Lung homogenates cytokine levels in HFD mice are generally decreased at baseline, but in some cases were elevated during Klebsiella pneumonia. Mice fed a ND (open circles) or HFD (closed squares) for 16 wk were intratracheally infected with 5,000 CFUs of K. pneumoniae. Lung homogenates were assessed for cytokines as described in materials and methods at either baseline (A), 24 h (B), or 48 h (C) following infection. Pooled values from n = 5–6 mice per group from two independent experiments. *P < 0.05 compared with mice fed the ND using the Student’s t test. Bars represent means ± SE. CFUs, colony-forming units; HFD, high-fat diet; ND, normal diet.

Effect of Diet-Induced Obesity on Lung and Gonadal White Adipose Tissue Leukocytes

Because HFD feeding has been shown to increase leukocytes accumulation in white adipose tissue (WAT) (49, 50), we examined the possibility that lung leukocytes might also be impacted by the HFD. After 16 wk of ND or HFD, we detected no differences in total leukocytes, CD64⁺ macrophages, Ly6C⁺ monocytes, neutrophils, or lymphocytes in lung homogenates in mice that were not infected (data not shown).

Twenty-four hours after infection, we observed a trend for more total leukocytes in the lung homogenates of mice fed HFD relative to those fed the ND (3.67 × 10⁷ ± 5.4 × 10⁶ vs. 2.59 × 10⁷ ± 4.78 × 10⁶; P = 0.15; Fig. 4A), a trend (P = 0.064) for higher numbers of CD64⁺CD11c⁺ macrophages, but no significant differences in numbers of CD64⁺CD11c⁻ macrophages, CD64⁻ CD11c⁺ DCs, or neutrophils. At the same time after infection, we found markedly higher numbers of total leukocytes in GWAT (HFD relative to ND; 1.22 × 10⁷ ± 1.01 × 10⁶ vs. 2.3 × 10⁶ ± 2.9 × 10⁵; data not shown), with significantly higher CD64⁺CD11c⁺ and CD64⁺CD11c⁻ macrophages, CD64⁻ CD11c⁺ DCs, Ly6C⁺ monocytes, and neutrophils in mice fed the HFD versus ND (Fig. 4B).

Figure 4.

Leukocyte counts during Klebsiella pneumonia in lung homogenates and gonadal white adipose tissue (GWAT) of mice fed a ND or HFD. Mice fed ND (open symbols) or HFD (solid squares) were infected with 5,000 CFUs of K. pneumonia; 24 h later, CD64⁺ CD11c⁻ macrophages (CD64+Macs) were recovered from lung homogenates (A; pooled values from n = 8 per group from 2 separate experiments) and GWAT (B; n = 5 per group, a representative from 2 separate experiments). *P < 0.05 compared with mice fed the ND using the Student’s t test. Bars indicate mean ± SE. CFUs, colony-forming units; HFD, high-fat diet; ND, normal diet.

Reduced Leukocyte Counts in BALF of Mice Fed the HFD 24 h after Infection

At baseline, there was a nonsignificant trend for lower numbers of total leukocytes recovered from BALF in mice fed HFD relative to ND (229,500 ± 64,899 vs. 434,611 ± 87,183; P = 0.07, n = 9 for each group). In Klebsiella pneumonia, leukocytes are recruited from the peripheral blood in response to chemoattractants produced at the focus of infection within the alveolar space. This recruitment includes a marginated pool of neutrophils within the vascular space. In contrast to the number of leukocytes in lung homogenates (which includes the marginated pool), there were significantly fewer total leukocytes (Fig. 5A) at 24 h after infection in the BALF of mice fed HFD, with lower CD64⁺ macrophages (Fig. 5B), neutrophils (Fig. 5C), CD64⁻ CD11c⁺ DCs, and Ly6C⁺ monocytes (Fig. 5D; significant for all but the last cell type). Thus, in HFD, despite a trend toward greater numbers of multiple types of leukocytes in the lung parenchyma and vasculature relative to ND-fed mice, plus levels of selected cytokines and chemokines that were equivalent or greater, fewer of all those cell types appeared in the air space in response to infection with K. pneumoniae.

Figure 5.

Leukocyte counts during Klebsiella pneumonia in bronchoalveolar lavage fluid (BALF) of mice fed a ND of HFD. Mice fed ND (open symbols) or HFD (solid squares) were infected with 5,000 CFUs of K. pneumonia; 24 h later, cells were recovered from BAL to enumerate total leukocytes (A), CD64⁺ macrophages (CD64+Macs; B), neutrophils (C), and CD64⁻ CDllc⁺ dendritic cells (DCs) and Ly6C⁺ monocytes (D). Pooled values from n = 8–13 mice per group from three separate experiments. *P < 0.05 compared with mice fed the HFD using the Student’s t test. Bars indicate means ± SE. CFUs, colony-forming units; HFD, high-fat diet; ND, normal diet.

Defective Peritoneal-Elicited Neutrophil Phagocytosis, Killing, and ROI Generation in Mice Fed the HFD

We next investigated the effect of HFD on neutrophils, which play an essential role in the defense of the lung against Klebsiella pneumonia. Impairments in neutrophil bactericidal functions are associated with greater bacterial outgrowth, sepsis, and death in animal models of pneumonia (51–53). To obtain neutrophils to evaluate bactericidal function, we lavaged the peritoneal cavities of mice fed the ND or HFD 5 h following glycogen elicitation; the isolated neutrophils were assayed for phagocytosis, killing, and ROI synthesis. Results showed a modest reduction in phagocytosis by neutrophils of K. pneumoniae in vitro (HFD, 81.7 ± 3.3 vs. ND, 100 ± 6.2, P = 0.04 after normalization to the ND). Because neutrophils can kill bacteria via both intracellular and extracellular mechanisms, we assessed the survival of K. pneumoniae cultured with neutrophils over a time course. After 45 min, there were slightly higher numbers of bacteria surviving culture with neutrophils from mice fed HFD (data not shown). At 90 min, there were markedly higher numbers of K. pneumoniae surviving in culture with neutrophils from mice fed the HFD compared with ND-fed mice (87.5% vs. 18.6%). Bacteria that were cultured without neutrophils increased in number to 114.5% of the original inoculum (data not shown). These data demonstrate that the defect in pulmonary bacterial clearance shown in vivo in mice fed a HFD (Fig. 2) was paralleled by a substantial curtailment of neutrophil-mediated killing of K. pneumoniae in vitro (Fig. 6A). Next, we compared neutrophil ROI production, which is crucial for neutrophil-mediated killing of K. pneumoniae (54). Lower ROI synthesis was observed in neutrophils from mice fed the HFD (Fig. 6B), indicating one potential contributing mechanism for defective killing.

Figure 6.

Reduced bactericidal function in neutrophils from mice fed HFD. Peritoneal-elicited neutrophils (PMNs) obtained from ND mice (open squares) or HFD mice (solid circles) were assessed for their ability to kill K. pneumoniae (A) or produce reactive oxygen intermediates (ROI; B) during culture with K. pneumoniae, as described in materials and methods. At the end of the experiment, the wells containing bacteria alone increased in number by 114.5% (data not shown). Results represent means ± SE, for the killing assays from four independent experiments with four replicates per group, for ROI, from four independent experiments with four replicates per group. *P < 0.05 compared with HFD-fed mice using the Student’s t test. HFD, high-fat diet; ND, normal diet; PMNs, polymorphonuclear neutrophils.

HFD Reduces PMN Glucose Uptake and GLUT1 and GLUT3 mRNA and GLUT3 Protein Expression

PMNs are dependent on glucose as the fuel for glycolysis to provide energy required for chemotaxis into the alveolar space, bacterial phagocytosis, and ROS-mediated killing. To determine if the defect in ROI synthesis might be due to impaired glucose uptake, we assessed PMN uptake of C¹⁴-labeled 2-DG. For these experiments, we used bone marrow PMNs, which yielded more consistent cell numbers and measurements of glucose uptake, as compared with glycogen-elicited peritoneal PMNs. Uptake of 2-DG was ∼50% lower in PMNs from mice fed HFD relative to those fed ND (Fig. 7A), suggesting that HFD might lead to reduced PMN glucose transporter expression. Pretreating PMNs with insulin did not change glucose uptake, consistent with the known lack of expression of the insulin responsive transporter GLUT4 by murine PMNs.

Figure 7.

HFD reduces glucose transport and mRNA transcripts for GLUT1 and GLUT3, and GLUT3 protein expression in neutrophils. PMNs were obtained from the bone marrow of ND mice (open bars and open squares) or HFD mice (solid circles). PMNs were adhered to 24-well plates and assessed for the uptake of 2-deoxy-d-[¹⁴C (U)] glucose (2-DG) for 5 min as described in materials and methods. qPCR of GLUT1 (A) and GLUT3 (B) transcripts in PMNs; qPCR 2^ΔΔCT data were normalized to ND GLUT3. None of the samples expressed GLUT2 or GLUT4 mRNA (data not shown). n = 4 samples per group. C and D: immunoblots were performed on PMN lysates to assess the levels of GLUT3 protein as described in materials and methods. C: bar graph represents the density of the blots for GLUT3 normalized for total protein with n = 3 samples per group. D: a representative blot for GLUT3 and total protein in PMN lysates from mice fed the ND or HFD. *P < 0.05 compared with ND-fed mice using the paired Student’s t test. Bars in A–C represent means ± SE. GLUT1–4, glucose transporters 1, 2, 3, and 4; HFD, high-fat diet; ND, normal diet; PMNs, polymorphonuclear neutrophils.

To investigate this reduction in glucose uptake in HFD PMNs, we next assessed the impact of HFD on specific glucose transporter expression. Comparing the levels of GLUT1, GLUT2, GLUT3, and GLUT4 mRNA in PMNs by qPCR, we observed 4-fold and 36-fold lower expression of GLUT1 and GLUT3, respectively, in PMNs obtained from mice fed the HFD as compared with those fed the ND (Fig. 7B). We did not detect mRNA for GLUT2 or GLUT4; we confirmed that our primers were functional using liver and skeletal muscle as controls (data not shown). Immunoblot analyses revealed that GLUT3 protein expression was approximately twofold lower in PMNs from mice fed the HFD as compared with those fed the ND (Fig. 7, C and D). Although we could not detect GLUT1 in PMNs, we did detect the expression of this glucose transporter in alveolar macrophages (data not shown). These data indicate that HFD reduces glucose transport in PMNs by reducing expression of GLUT1 (mRNA only) and GLUT3 (mRNA and protein).

Blocking Glycolysis with 2-DG Reduces Killing and ROI Production by Bone Marrow-Derived Neutrophils Only in ND-Fed Mice

Because the reduction in PMN killing and ROI synthesis in cells from HFD-fed mice was associated with reduced glucose uptake and GLUT3 protein expression, we next investigated whether inhibiting glycolysis with 2-DG would reduce killing and ROI synthesis in bone marrow-derived PMNs. In untreated cells, bacterial killing was reduced by more than 40% in PMNs from mice fed the HFD versus the ND (Fig. 8A). Although 2-DG pretreatment reduced bacterial killing in PMNs from ND mice by more than 40%, it did not decrease killing in PMNs from HFD mice (Fig. 8A). Similarly, 2-DG reduced ROI synthesis in PMNs from ND mice by ∼50% (Fig. 8B). Although ROI synthesis was lower in PMNs from mice fed the HFD versus the ND, 2-DG did not decrease ROI production in PMNs from mice fed the HFD. These results, indicating the dependence on glycolysis for neutrophil bacterial killing and ROI synthesis only in mice fed the ND, support our interpretation that the reduced glycolysis in HFD-fed mice explains their defects in these functions.

Figure 8.

Blocking glycolysis with 2-DG reduces K. pneumoniae killing and ROI production in PMNs from mice fed the ND but not the HFD. Neutrophils were obtained from the bone marrow of ND mice (open bars) or HFD mice (solid bars). PMNs were treated without or with 2-DG (10 mM) for 1 h, then cultured with K. pneumoniae, as described in materials and methods to assess bacterial killing (A; n = 4) or ROI production (B; n = 3). *P < 0.05 compared with ND-fed mice using the Student’s t test. Bars represent means ± SEM. 2-DG, 2-deoxy-d-glucose; BM, bone marrow; HFD, high-fat diet; PMNs, polymorphonuclear neutrophils; ROI, reactive oxygen intermediates.

DISCUSSION

In this report, we demonstrate that HFD-induced obesity impairs pulmonary K. pneumoniae clearance and increases bacterial dissemination in vivo. This impairment was associated with relative increases in the concentrations of multiple inflammatory cytokines in lung homogenates of HFD-fed mice and with a nonsignificant trend for greater numbers of almost every type of inflammatory cell in lung homogenates, which include the marginated intravascular populations. Surprisingly, given their far greater bacterial lung burdens, HFD mice showed lower numbers of all inflammatory cell types assayed in BALF, compatible with a deficit in transmigration into the site of infection in the alveolar space. We also observed a reduction in PMN phagocytosis and killing of K. pneumoniae in vitro that was associated with reduced ROI synthetic capacity. An additional novel and notable finding of this study was a reduction in glucose transport and diminished expression of GLUT1 and GLUT3 transcripts and GLUT3 protein in PMNs recovered from mice fed a HFD. These results suggest that HFD impairs host defense by limiting glucose metabolism, which is crucially important in PMN bactericidal functions. This conclusion was supported by the ability of 2-DG to reduce bacterial killing and ROI synthesis in PMNs from mice fed the ND but not the HFD. These results demonstrate that HFD adversely impacts multiple levels of lung host defense against a key bacterial pathogen associated with nosocomial pneumonias.

Obesity has been characterized as a chronic state of inflammation with elevated systemic proinflammatory cytokine and adipokine levels in serum and adipose tissue (39, 45, 55). However, we found that baseline pulmonary IL-1β, IL-6, IL-17, TNF-α, and CXCL2 concentrations were lower in mice fed the HFD compared with ND. Similar results have been reported in peritoneal macrophages recovered mice fed a HFD versus a low-fat diet (56, 57). After infection, the opposite was found with elevated levels of IL-β, IL-6, CCL2, and CXCL2 in lung homogenates of mice fed the HFD compared with ND at 24 and 48 h after Klebsiella challenge. These differences may be due to the increased bacterial burdens or disparities in cytokine production in resident versus recruited macrophages observed in mice fed the HFD versus ND at both time points after infection. In addition, there was a trend for greater numbers of total lung homogenate leukocytes in mice fed the HFD versus ND, a likely explanation for the elevation in pulmonary cytokines after infection.

Pulmonary clearance of K. pneumoniae strain 43816 is dependent on the robust recruitment of leukocytes to the alveolar focus of infection, as neutralization of chemokines or the selective depletion of either PMNs or monocytes before infection dramatically increases lung bacterial loads (32, 58–61). Relative reductions of nearly all intraalveolar (BALF) leukocytes in mice fed the HFD were associated with higher lung and spleen bacterial burdens. These differences were observed despite elevated levels of pulmonary chemokines CCL2 and CXCL2, lung homogenate leukocytes, and greater GWAT leukocyte counts in mice fed a HFD. Although GWAT leukocytes are known to be elevated at baseline in mice fed a HFD, these differences persist even after infection (62). Interestingly, Ubags et al. (5) reported lower PMN counts in BALF from mice fed a HFD versus a ND following intratracheal LPS instillation, and this difference was associated with impaired PMN chemotaxis in vitro. Collectively, these results suggest that pulmonary bacterial clearance depends crucially on leukocyte recruitment to the alveolar space and that HFD-fed mice likely have reduced PMN chemotaxis following infection.

After recruitment into the alveolar space, PMNs bind, phagocytose, and kill ingested bacteria by releasing proteolytic enzymes, antimicrobial peptides, and reactive oxygen intermediates into the phagolysosome (63). In addition, PMNs can kill bacteria extracellularly by degranulating and by releasing antimicrobial agents and extracellular traps (64). In the current study, the killing of K. pneumoniae following incubation with PMNs from ND mice was ∼70% higher than that of PMNs from HFD mice. This impairment in killing can be attributed to a modest reduction in phagocytosis and a 45% reduction in ROI production, in agreement with previous demonstrations that reduced ROI production by leukocytes or epithelial cells impairs killing of K. pneumoniae (65, 66). It is noteworthy that both phagocytosis and ROI generation were assessed using heat-killed rather than live bacteria, which resist opsonization and would be more physiologically relevant. Similar results were reported by Frydrych et al. (67) who demonstrated that bone marrow-derived PMNs recovered from HFD-induced obese mice exhibited reduced phagocytosis of Escherichia coli and lower ROI relative to nonobese mice. In that study of a murine model of polymicrobial sepsis, diet-induced defective PMN function was associated with greater mortality and higher peritoneal bacterial loads than in nonobese mice (67). Similarly, PMNs from obese human subjects exhibited impairments in killing Staphylococcus aureus (19–21).

Because impairments in PMN recruitment into the alveolar space, phagocytosis, and ROI synthesis suggested a common limitation in cellular energy availability, we evaluated glucose uptake and glucose transporter expression. The energy requirements of PMN bactericidal functions are highly dependent on glucose as a glycolytic substrate (68). We observed a 50% reduction in 2-DG uptake in PMNs from mice fed HFD versus ND, suggesting substantially impaired glucose transport. This difference was associated with markedly lower GLUT1 and GLUT3 mRNA transcripts and GLUT3 protein expression in HFD PMNs. Insulin treatment of PMNs in vitro did not affect glucose transport (data not shown), likely due to the absence in PMNs of GLUT4, the principal insulin-sensitive glucose transporter. Although it is beyond the scope of this report to determine the mechanisms by which HFD reduces GLUT1 and GLUT3 transcripts and protein expression, we speculate that the abundance of lipids may have shifted energy substrate utilization from glucose utilization to fatty acid metabolism. Taken together, these results suggest that the impairments in PMN bactericidal function were due to defects in GLUT1- and GLUT3-mediated glucose uptake.

There were some limitations in our study. First, we exclusively used male mice because in diet-induced obesity, the focus of this study, the obesity phenotype, hyperglycemia, and insulin resistance are more profound in males than females (62). In addition, male mice show increased susceptibility to bacterial pneumonia, a phenomenon also observed in humans (69–71). Future studies will be necessary in both males and females to uncover potential sexual dimorphic responses to the diet-plus-infection model. Another limitation was the use of an HFD as a model of diet-induced obesity, which in humans can also result from excessive calories of any source. Although this approach closely resembles the development of obesity in humans in high-income nations, our results may have been influenced by differences in fat content. Future studies should be conducted using other high-calorie diets or hyperphagic mice fed normal rodent chow. We have used the latter approach to demonstrate that, as compared with WT mice, obese hypothalamic leptin receptor-deficient mice exhibit impaired host defense and greater mortality following influenza infection (72).

In summary, we report a novel mechanism by which diet-induced obesity impairs PMN bactericidal function and host defense against Klebsiella pneumonia in a murine model. Specifically, we have observed that HFD feeding impairs glucose transport and reduces glucose transporter transcripts in PMNs. Future studies are needed to determine the mechanisms by which obesity, alterations in lipid metabolism, or both, affect glucose transport and bactericidal functions in PMNs from transgenic mice and humans with obesity.

SUPPLEMENTAL DATA

Supplemental Figs. S1–S3: https://doi.org/10.6084/m9.figshare.16722844.v1.

GRANTS

This work was funded by a grant from the University of Michigan M-Cubed Program (to P.M., J.L.C., and K.S.); National Institutes of Health (K08 DK101755 and R01 DK115583; to K.S.); DK107535 (to D.B.); and Merit Review Awards I01 BX001389 (to C.M.F.) and I01 CX000911 (to J.L.C.), Department of Veterans Affairs.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

P.M., J.L.C., and K.S. conceived and designed research; P.M., A.M.W., C.A.G., B.B., and C.M.F. performed experiments; P.M., J.L.C., A.M.W., C.M.F., D.B., and K.S. analyzed data; P.M., J.L.C., D.B., and K.S. interpreted results of experiments; P.M. prepared figures; P.M. drafted manuscript; P.M., J.L.C., A.M.W., C.A.G., B.B., C.M.F., D.B., and K.S. edited and revised manuscript; P.M., J.L.C., A.M.W., C.A.G., B.B., C.M.F., D.B., and K.S. approved final version of manuscript.