Non-classical monocytes sense Hypoxia, regulate Pulmonary Vascular Remodeling, and Promote Pulmonary Hypertension

Abstract

An increasing body of evidence suggests that bone marrow derived myeloid cells play a critical role in the pathophysiology of pulmonary hypertension (PH). However, the true requirement for myeloid cells in PH development has not been demonstrated and a specific disease-promoting myeloid cell population has not been identified. Utilizing bone marrow chimeras, lineage labelling, and proliferation studies, we determined that, in murine hypoxia-induced PH, Ly6C^lo non-classical monocytes are recruited to small pulmonary arteries and differentiate into pulmonary interstitial macrophages (IMØ). Accumulation of these non-classical monocyte-derived pulmonary IMØ around pulmonary vasculature is associated with increased muscularization of small pulmonary arteries and disease severity. To determine if the sensing of hypoxia by non-classical monocytes contributes to the development of PH, mice lacking expression of Hif1α in the Ly6C^lo monocyte lineage were exposed to hypoxia. In these mice, vascular remodeling and PH severity were significantly reduced. Transcriptome analyses suggest that the Ly6C^lo monocyte lineage regulates PH through complement, phagocytosis, antigen presentation, and chemokine/cytokine pathways. Consistent with these murine findings, relative to controls, lungs from PAH patients displayed a significant increase in the frequency of non-classical monocytes. Taken together, these findings show that, in response to hypoxia, non-classical monocytes in the lung sense hypoxia, infiltrate small pulmonary arteries, and promote vascular remodeling and the development of PH. Our results demonstrate that myeloid cells, specifically cells of the non-classical monocyte lineage play a direct role in the pathogenesis of PH.

Introduction

Pulmonary arterial hypertension (PAH) is a heterogeneous group of diseases characterized by abnormal remodeling and, ultimately, ablation of pulmonary intralobar arteries. It is thought that in pre-disposed individuals, environmental and genetic factors promote endothelial cell dysfunction or damage and the recruitment and proliferation of vascular smooth muscle cells and pericytes, which leads to intimal, medial, and adventitial hypertrophy, muscularization of small arteries, and an eventual increase in pulmonary vascular resistance and pressure. The exact cause of this pulmonary vascular remodeling remains unknown, but it has traditionally been viewed as a disease of endothelial and vascular smooth muscle cells and the signaling pathways that regulate these cell types (1). However, more recently, a body of evidence has emerged suggesting that bone marrow derived myeloid cells play an important role in PAH pathophysiology (2-4).

It has long been recognized that the development of pulmonary hypertension (PH) is associated with the infiltration of myeloid cells into the pulmonary vasculature, although the functional significance of this has not been clear (5-7). A causative role for myeloid cells in PH has been suggested by studies in which manipulations that alter myeloid cell numbers also regulate the severity of PH (8-13). It has even been suggested that PAH involves a primary bone marrow defect, as patients with PAH and their unaffected family members display abnormalities in the myeloid lineage (3, 14). This may be related to mutations in bone morphogenetic protein receptor type 2 (BMPR2), which are present in 70% of familial PAH cases and 10–25% of idiopathic PAH cases (15). In humans with PAH, BMPR2 mutations are associated with increased production of the monocyte chemoattractant granulocyte-macrophage colony-stimulating factor (GM-CSF) and increased infiltration of GM-CSF receptor expressing macrophages in pulmonary arteries (16). In mouse models, GM-CSF infusion increased and GM-CSF blockade decreased vascular myeloid cell in filtration, small pulmonary artery muscularization, and the development of PH (16). Moreover, bone marrow transplantation with BMPR2 mutant cells caused PH in mice, while wild-type cells are protective in BMPR2 mutant mice (17). Together, these studies strongly suggest that myeloid cells play a causative role in the development of PH; however, a direct role for a specific disease-inciting myeloid cell type has never been demonstrated.

Macrophages display significant diversity and plasticity, and the functionality of specific macrophage subtypes is determined by both their origins and location (18-20). Tissue macrophages may arise from one of three sources: fetal macrophages, CX3CR1^loLy6C^hi classical monocytes, and CX3CR1^hiLy6C^lo non-classical monocytes (21, 22). Fetal macrophages arise in the yolk sac or fetal liver and seed tissues during development. In the lung, these become resident pulmonary macrophages, which, under homeostatic conditions, are maintained via proliferation through the entire lifespan of the animal (21, 23). Classical monocytes are bone marrow-derived, and are rapidly recruited to sites of inflammation via the activity of the chemokine receptor CCR2, and mediate acute inflammatory and fibrotic responses (24). Non-classical monocytes are also bone marrow-derived, patrol the microvasculature by crawling on the endothelium, infiltrate tissues in response to inflammatory stimuli, seemingly act as inflammation-initiating sentinel cells, and regulate the resolution of inflammation, endothelial health, and tissue repair (25-27) Previously, we demonstrated that PH is increased in CCR2-deficient mice, strongly suggesting that classical monocytes do not directly contribute to the development of PH (28, 29). In this study, using chronic hypoxia models of PH, we identify specific myeloid cell types that infiltrate small pulmonary arteries, and by sensing of hypoxia, contribute to the development of PH.

Material and Methods

Animal

Male CD45.2 C57BL/6 mice at 6 weeks of age were purchased from Charles River Laboratories (Morrisville, NC). CD45.1 C57BL/6, Cx3cr1^gfp/gfp, and Hif-1α^flox/flox were purchased from Jackson Laboratories (Bar Harbor, Maine). Cx3cr1-cre transgenic animals were developed and characterized in the laboratory of Michael Gunn (30). All mice were housed in a barrier and specific pathogen-free facility at Duke University School of Medicine (Durham, NC). All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at Duke University. For hypoxia exposure, animals were placed at 18,000 feet altitude in an environmentally controlled hypobaric chamber. The duration of exposure is specified in each experiment. For hypoxia plus SU5416 (Sigma Aldrich, St. Louis, MO) exposures, animals were treated with a weekly dose of subcutaneous 20mg/mg SU5416 dissolved in 5% ethanol, 5% dimethyl sulfoxide in carboxymethyl cellulose (CMC).

Tissue Samples

Human lung samples were obtained from two sources. Control human lung tissues were obtained from organs declined for transplantation. All control lungs were from subjects without a history of smoking or chronic lung disease obtained from the University of North Carolina Marisco Lung Institute/Cystic Fibrosis Center Tissue Procurement and Cell Culture Core (under an approved protocol at University of North Carolina at Chapel Hill). Lung explants from patients with Group I pulmonary arterial hypertension (PAH) were obtained at the time of lung transplant at Duke University under a Duke University Institutional Review Board approved protocol. Fresh lung tissues were processed and digested into single cell suspensions and prepared for multiparameter flow cytometry as described (31).

Cardiac Function Evaluation

Cardiac function evaluations were performed in mice as described (28). Briefly, animals were anesthetized, intubated, and ventilated. The right jugular vein was cannulated with a fluid-filled PE-10 tube. RV pressure, heart rate, and respiratory rate were recorded using PowerLab (ADInstruments, Colorado Springs, CO; RRID:SCR_001620).

Bone Marrow Chimera Generation

CD45.2 male animals were treated intra-peritoneally with 25mg/kg of Busulfan (Sigma Aldrich, St. Louis, MO) dissolved in a 30% DMSO/PBS solution at 72 and 48 hours prior to transferring bone marrow cells. Bone marrow cells (1 × 10⁷) derived from 6-8 week old CD45.1 male animals were injected via the retro-orbital sinus into CD45.2 male recipients. Animals were maintained on sulfamethoxazole/trimethoprim supplemented water for 14 days. To assess chimerism, blood samples were collected 3 weeks after cell transfer and analyzed by flow cytometry as described (32).

Flow Cytometry

Phenotyping of murine and human pulmonary immune repertoires were performed using multiparameter flow cytometry and a panel of antibodies previously described (32). Briefly, tissues were dissociated into single cell suspensions and stained with a panel of antibodies. Data were acquired using an LSRII flow cytometer (BD Bioscience, San Jose, CA; RRID:SCR_002159). Data were analyzed using Flowjo X (BD Bioscience, San Jose, CA; RRID:SCR_008520). Absolute numbers of each immune cell type were calculated by multiplying the total cell number recovered from each lung digest by the proportion of a particular cell type as percent of live single cells.

Immunofluorescence

Murine lung tissues were fixed in 4% paraformaldehyde (Sigma Aldrich, St. Louise, MO), and washed with phosphate-buffered-saline (PBS) solution. Fixed tissues were embedded in Optimal Cutting Temperature compound (OCT). Frozen tissue sections of 6-8μm were prepared. Immunofluorescence staining was performed using rat anti-mouse CD64 (AT152-9; Bio-Rad, Hercules, CA; RRID:AB_2687456), chicken anti-green fluorescent protein (GFP) (GFP-1020; Aves Labs, Tigard, OR), rat anti-mouse CD169 (3D6; Biolegend, San Diego, CA; RRID:AB_10915134). 4’,6’-diamidino-2-phenylindole (DAPI) was used for nuclear staining. CD64 and CD169 staining was performed using tyramide amplification (PerkinElmer Tyramide Plus, Waltham, MA). Small pulmonary arteries of similar sizes were selected blindly across treatment or animal groups for imaging. Confocal images were obtained with a Zeiss 710 inverted confocal microscope using 20x objective (Zeiss, Cambridge, UK). Images were converted to JPEG file format using ImageJ (National Institute of Health; RRID:SCR_003070).

Quantitative Real-Time RT-PCR

Analyses of mRNA levels were performed using real-time RT-PCR. Total RNA was isolated from the specified immune cell subpopulations purified from the lung using a BD Aria II sorter. Total RNA was isolated using RNAeasy Plus Micro Kit (Qiagen, Germantown, MD). Subsequently, cDNA was synthesized using QuantiTect Reverse-Trasncription kit (Qiagen, Germantown, MD). Amplification was performed using PowerUp SYBR Green Master Mix (Thermo Fisher, Scientific, Carlsbad, CA) on a QuantStudio Real-Time PCR system (Thermo Fisher Scientific, Carlsbad, CA). Primers for Hif1α are the following: forward primer (TCATCAGTTGCCACTTCCCCAC) and reverse primer (CCGTCATCTGTTAGCACCATCAC). Gene expression was normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH). Using GAPDH forward primer (AGGTCGGTGTGAACGGATTTG) and reverse primer (TGTAGACCATGTAGTTGAGGTCA). Changes in gene expression between Hif1α-sufficient and Hif1α-deficient cells were quantified using delta-delta-Ct calculation.

PKH26PCL labelling and Proliferation in vivo

7-8 week old C57BL/6 males were injected with 5μM of PHK26PCL in Diluent B (Sigma Aldrich, St. Luis, MO) via the retro-orbital sinus. Animals were also treated with 50mg/kg of Edu (5-ethynyl-2’-deoxyuridine) weekly for 3 weeks by intraperitoneal injection. Lungs were harvested and prepared as single cell suspensions as previously described (32). Cellular proliferation was detected using Click-iT Plus EdU Proliferation Kit for flow cytometry (Thermo Fisher Scientific, Carlsbad, CA). Data were collected using a BD LSRII (BD Bioscience, San Jose, CA; RRID:SCR_002159) and analyzed using Flowjo X (BD Bioscience, San Jose, CA; RRID:SCR_008520).

RNA sequencing

Animals were exposed to 18,000 feet altitude plus 20mg/kg SU5416 weekly for 3 weeks. Lungs were harvested and prepared for sorting as described above. For RNAseq, 250 non-classical monocytes or pulmonary interstitial macrophages (IMØ) were directly sorted into 1X lysis buffer containing RNase inhibitor (Takara Bio Inc., U.S.A.). Libraries were prepared using SMARTer@ Stranded Total RNA-seq Kit - Pico Input Mammalian library Prep kit per the manufacturer’s protocol. Library qualities and quantities were verified using Agilent Bioanalyzer system (Agilent, Santa Clara, CA) and Qubit fluorometric quantification (Thermo Fisher Scientific, Carlsbad, CA). Sequencing was performed using an Illumina Hiseq 2500 (Illumina, San Diego, CA), with 126 pair-ended runs, and approximately depth of 50 million reads per library. The sequencing reads were trimmed to remove the Illumina adapters and any low-quality base at the ends (the Phred quality score <30) by using Cutadapt (v1.12) (33). Subsequently, concordant pair end reads for each sample were successfully aligned to the mouse reference transcriptomes, GRCm38, using Tophat (v2.1.1) (34). Finally, the read-count-per-gene measurements for each sample were performed by htseq-count (HTSeq v0.6.0) (35). The read counts were then filtered and normalized by estimated size factors by using the R package “DESeq2” (36). The gene expression differences across the treatment groups were then evaluated using the default generalized linear model in DESeq2. Genes passing the threshold, an FDR of <5%, were considered as significantly differentially expressed.

Quantification of small pulmonary arterial muscularization and remodeling

Murine lung tissues were perfused with PBS, followed by fixation in 4% paraformaldehyde. Harvested tissues were washed extensively with PBS. All lobes of the lungs were sliced in cross-section (perpendicular to main pulmonary arteries), at approximately 0.5 cm thickness. All lung sections were embedded as single block in paraffin. Tissue sections of 5μm were obtained and stained with anti-human von Willebrand factor (vWF) and alkaline phosphatase conjugated mouse anti-human α-smooth muscle cell actin (SMA) (1A4; Sigma-Aldrich, St. Luis, MO) and developed with VectorRed Alkaline Phosphatase (VectorLabs, Burlingame, CA). Tissues were counterstained with Hematoxylin QS (VectorLabs, Burlingame, CA). The entire slide was imaged and stitched with Zeiss Axio Imager Wide field Fluorescence Microscope (Zeiss, Cambridge, UK). In a blinded fashion, all vessels under 50μm on the entire slides were categorized as no muscularization (absence of SMA staining around the vessel), partially muscularized (SMA staining in parts of the vessel), and fully muscularized (SMA staining encircles the entire circumference of the vessel). Vessel muscularization was expressed as the percentage of total vessels enumerated.

Statistical Analyses

All data are expressed as mean (±SEM) across experimental repeats, as stated. Group comparisons were performed in Graphpad PRISM 6 (La Jolla, CA; RRID:SCR_002798) using Student’s t test, one-way ANOVA, or two-way ANOVA. P values of <0.05 were considered statistically significant. All experiments were repeated at least three times.

Results

Accumulation of interstitial macrophages in hypoxic small pulmonary arteries.

To identify the myeloid cell types that accumulate in lungs during hypoxia, we used flow cytometric analysis to quantify myeloid populations in the lungs of hypoxic and normoxic mice (Fig S1) (32, 37). Based on our prior finding that myeloid cell infiltration peaked at 3 weeks of hypobaric hypoxia exposure, C57BL/6 animals were exposed to normoxia or hypoxia, with or without Su5416, for 3 weeks (28). Su5416 is a tyrosine kinases inhibitor that is thought to induce endothelial cell dysfunction, leading to development of severe pulmonary hypertension in mice (28, 38). Relative to normoxic controls, mice exposed to either hypoxia or hypoxia plus Su5146 exhibit a 2-3 fold increase in the number of CD11b⁺CD64⁺ pulmonary interstitial macrophages (IMØ) (Fig 1A). No significant changes in the numbers CD11b⁻CD64⁺ alveolar macrophages (AMØ) or CD11c⁺MHCII⁺CD24⁺CD64⁻ dendritic cells (DC) were observed (Fig 1A). Consistent with our previous report, hypoxia exposed animals displayed a significant reduction in the numbers of both classical (Ly6C^hi) and non-classical (Ly6C^lo) monocytes in the lungs (Fig 1A) (28). However, in hypoxia plus Su5416 treated animals, compared to normoxia plus Su5416 treated controls, monocyte reduction was only observed in the non-classical (Ly6C^lo), but not classical (Ly6C^hi), subset (Fig 1A). No significant changes in granulocyte populations, including neutrophils and eosinophils, were observed (data not shown). Overall, these findings demonstrate that IMØ are the only myeloid cell type to increase in the lungs of mice exposed to hypoxia.

Figure 1.

Perivascular accumulation of pulmonary IMØ. A) Absolute numbers of IMØ, AMØ, dendritic cell, and monocyte subsets in animals exposed to normoxia vs. hypoxia, with or without Su5416 for 3 weeks (n=3 per group). Data is representative of > 3 experiments. Statistical analyses: two way ANOVA. Values displayed as Mean ± SEM. *p<0.05; **p<0.01; and ***p<0.001. B) Immunofluorescence staining of lung tissues derived from animals exposed to normoxia vs. hypoxia, with or without Su5416 for 3 weeks. Lung tissues were stained with anti-αSMA (red); anti-CD64 (magenta); anti-GFP (green); and DAPI. Merged images show CD64⁺ GFP⁺ IMØ (white), around remodeled small pulmonary arteries (red).

To determine if IMØ are also the cells that infiltrate small pulmonary arteries in response to hypoxia, we exposed Cx3cr^wt/gfp mice to normoxia or 3 weeks of hypoxia in the presence or absence of Su5416 and examined the localization of IMØ in lung sections by confocal microscopy. Compared to hypoxia alone, addition of Su5416 treatment induces increased pulmonary vascular remodeling and severity of PH (38). In Cx3cr^wt/gfp mice, all pulmonary macrophages express CD64, but only IMØ express green fluorescent protein (GFP)⁺, leaving AMØ GFP⁻ (32). In both models of hypoxia-induced PH, lungs of normoxic mice displayed no muscularization of small pulmonary arteries, as demonstrated by a lack of α-smooth muscle actin (SMA) staining and no accumulation of GFP⁺ CD64⁺ IMØ around the small pulmonary arteries (Fig 1B, columns 1 and 2). In contrast, the lungs of hypoxic mice displayed robust muscularization of small pulmonary arteries (SMA⁺) and infiltration of these remodeled small pulmonary arteries by GFP⁺ CD64⁺ IMØ (Fig 1B, columns 3 and 4). To confirm that the small pulmonary artery infiltrating cells were macrophages, we also examined expression of CD169, a marker expressed on IMØ and AMØ but not monocytes or DC (32). The lungs of hypoxic mice displayed infiltration of SMA⁺ small pulmonary arteries by GFP⁺CD169⁺ IMØ (Fig S2A). Moreover, the infiltration of small pulmonary arteries by IMØ appeared to occur to a greater extent in smaller vessels (<50μm) (Fig S2B). These findings demonstrate that IMØ are the myeloid cell type that infiltrate small pulmonary arteries during the development of hypoxic PH.

Hypoxia-induced interstitial macrophages are bone marrow derived.

Because IMØ may arise from either resident fetal derived macrophages or circulating bone marrow derived monocytes, we determined the origin of the IMØ that accumulate in lungs in response to hypoxia. We transplanted congenic CD45.1⁺ bone marrow into CD45.2⁺ recipients after myeloid cell ablation with busulfan. The recipients were exposed to normoxia or hypoxia, and the origin of IMØ was examined (Fig 2A). Analyses of peripheral blood 3 weeks after transplant demonstrated effective chimerism in which >85% of circulating monocytes, neutrophils, and eosinophils were of donor (CD45.1⁺) origin (Fig 2B). Consistent with prior reports, busulfan treatment generally preserves the CD45.2⁺ recipient origin of lung resident tissue macrophages but allows the replacement of bone marrow-derived circulating monocytes with those of CD45.1⁺ donor origin (Fig 2C-D) (39). As with Cx3cr^wt/gfp mice, recipients exposed to hypoxia displayed a 2-3-fold increase in the frequency of lung IMØ (Fig. 2E-F). In normoxia, ~25% of IMØ were CD45.1⁺ (Fig 2G-H). In contrast, in the lungs of hypoxic mice, over 55% of IMØ were CD45.1⁺ donor origin (Fig 2G-H). This enrichment of CD45.1⁺ donor cells in IMØ with hypoxia exposure demonstrates that the vast majority of newly arriving IMØ are bone marrow derived.

Figure 2.

Infiltrating pulmonary IMØ in PH are derived from circulating bone marrow cells. A) Schema of CD45.1:CD45.2 bone marrow chimera generation and hypoxia exposure. B) Flow cytometry assessment of chimeric efficiency. Graph depicts CD45.1 expression (donor origin) on blood leukocytes in chimeric animals. C) Histograms depicting preservation of CD45.2⁺ recipient-derived resident IMØ and AMØ, but replacement of monocytes by CD45.1⁺ donor cells in chimeric animals. D) Bar graph showing percent of IMØ, AMØ, or monocytes that express CD45.2 and, thus, are derived from recipient origin. (n=4/group). E) Flow cytometric analyses showing accumulation of pulmonary IMØ in hypoxic vs. normoxic chimeric animals. F) Bar graph quantifying IMØ changes in normoxic vs. hypoxic chimeras. Graph depicts IMØ as percent of total leukocytes (CD45+) in normoxia- vs. hypoxia-exposed animals (n=5/group). G) Histogram showing percent of CD45.1⁺ IMØ in the lung in normoxic vs. hypoxic animals. H) Bar graph depicts percent of IMØ that express CD45.1 in normoxic vs. hypoxic animals (n=5/group). Data shown are representative of three experiments. Statistical analyses were done using unpaired t tests. Bar graph measurements are displayed as Mean ± SEM.

Hypoxia-induced interstitial macrophages arise from non-classical monocytes

Bone marrow derived IMØ may arise from either classical or non-classical monocytes. Classical monocytes (Ly6c^hi) are dependent on CCR2 signaling to traffic into the tissue (40, 41). Since macrophage numbers increased to the same extent in hypoxic wild-type and CCR2^−/− mice, this strongly suggested that hypoxia-induced IMØ do not arise from classical monocytes (28). To determine if hypoxia-induced IMØ arise from non-classical monocytes, we initially considered tracking the fate of these cells after adoptive transfer; however, mice do not contain sufficient numbers of non-classical monocytes in blood, spleen or bone marrow to allow effective adoptive transfer. We therefore performed in vivo cell labeling using PKH26PCL, a dye that is retained by tissue macrophages and non-classical (Ly6C^lo) monocytes (Fig 3A) (42). To optimize and validate cell type labelling by PKH26PCL, we examined dye uptake and retention by immune cells in the lung and blood. At 24 hours after PKH26PCL administration, all circulating and tissue immune cells were labelled with PKH26PCL (data not shown). However, four days after PKH26PCL administration, as expected, there was selective dye retention by tissue IMØ and AMØ (Fig 3C). In addition to tissue macrophages, in both blood and lung, PKH26PCL was retained in non-classical (Ly6C^lo) monocytes, but not classical (Ly6C^hi) monocytes (Fig 3B-C). Other immune cells, including dendritic cells and neutrophils did not retain the PKH26PCL dye (Fig 3C). To determine cellular origin of infiltrating IMØ in response to hypoxia, mice were injected intravenously with PKH26PCL. Four days after administration of PKH26PCL, animals were exposed to normoxic vs. hypoxic condition, with or without Su5416, for 21 days (Fig 3A). Additionally, to determine the extent that expanded IMØ population was due to the resident MØ proliferation, mice were also injected with 5-Ethynyl-2’-deoxyuridine (Edu) at weekly intervals (Fig 3A, D-E). Lung tissues were then harvested, and IMØ labelling by PKH26PCL and Edu were examined (Fig 3D-G). Similar to findings in Figs 1 and 2, in animals exposed to hypoxia alone or hypoxia plus Su5416, compared to controls, pulmonary IMØ were increased 2-3 fold (Data not shown). With or without Su5416 exposure, IMØ derived from the lungs of normoxic mice demonstrated a low level (~7%) of proliferation (Fig 3D-E). In mice exposed to 21 days of hypoxia, IMØ proliferation, as measured by Edu incorporation, remained low and similar to that of normoxic animals (~7%) (Fig 3D-E). Consistent with above finding that the majority of expanding IMØ derive from circulating bone marrow cells, this Edu staining further demonstrates that expansion of IMØ was not due to proliferation of PHK26PCL⁺ resident IMØ. The staining also demonstrated that, similar to normoxic animals, nearly all IMØ from hypoxic animals are PKH26PCL⁺ (Fig 3F-G). Thus, the expanding IMØ population in hypoxic animal must arise from the circulating PKH26PCL⁺ cell type: non-classical monocytes. Taken together, these findings demonstrate that circulating non-classical monocytes serve as the major precursors to hypoxia-induced pulmonary IMØ.

Figure 3.

Ly6C^lo non-classical monocytes are the progenitors of accumulating pulmonary IMØ. A) Schema depicting non-classical monocyte labelling, Edu treatment, and timing of hypoxia exposure. B) Histograms showing that on the dayof hypoxia exposure (day 0), circulating blood Ly6C^lo non-classical monocytes, but not Ly6C^hi classical monocytes, are labelled by PKH26PCL. Dark gray filled histogram represent cells derived from diluent only animals. Solid black lines represent cells derived from PKH26PCL treated animals. C) Histograms showing, on day 0 of hypoxia exposure, PKH26PCL staining of myeloid cell types in the lung. D) Contour plots depicting percent of Edu+ IMØ, as a measure of cellular proliferation, in controls vs. hypoxia or hypoxia + Su5416 treatment for 21 days. E) Bar graph quantifying percent of Edu+ IMØ in controls vs. hypoxia or hypoxia + Su5416 exposed animal (n=3/group). F) Histograms showing PKH26PCL labelling in pulmonary IMØ in controls vs. hypoxia or hypoxia + Su5416 treated animals. G) Bar graph quantifying PKH26PCL mean fluorescent intensity (MFI) in IMØ (n=3/group). Data shown are representative of three experiments. Unpaired t tests were performed for statistical analyses. Bar graph measurements are displayed as Mean ± SEM.

Non-classical monocytes sense hypoxia to promote hypoxia-induced PH.

To determine if IMØ derived from non-classical monocytes play a causative role in the development of hypoxia-induced PH, we examined mice in which the alpha subunit of hypoxia-inducible factor 1 (HIF-1α) was selectively eliminated in the non-classical monocyte lineage using Hif1α^flox/flox mice. HIFs are the critical oxygen-sensing molecules in mammalian cells and animals heterozygous for Hif-1α are protected from hypoxia-induced PH (43, 44). Since CX3CR1 is expressed most highly in Ly6C^lo non-classical monocyte, we used Cx3cr1-cre mice to target the non-classical monocyte lineage (30, 32). The specificity of Cre expression in these mice was examined by crossing them to Rosa26-fGFP reporter mice. Cx3cr1cre:Rosa26-fgfp mice displayed expression of farnesylated GFP (fGFP) on Ly6C^lo, but not Ly6C^hi monocytes, in both blood and lung (Fig S3A). GFP expression was also detected in ~50% of lung macrophages and ~60% of dendritic cells (Fig S3A). There was no GFP expression in CD45⁻ non-immune cells in the lung, demonstrating that there was no off-target gene deletion in non-immune cells (Data not shown). When crossed to Rosa26-dtr mice, Ly6^lo, but not Ly6C^hi monocytes were depleted when the resulting Cx3cr1cre:Rosa26-dtr mice were treated with diphtheria toxin (Fig S3B).

Cx3cr1cre;Hif1α^Δ/Δ mice displayed a selective deletion of Hif1α in non-classical monocytes. By semi-quantitative PCR, there was a ~90% reduction of Hif1α expression in Ly6C^lo monocytes, but normal expression levels in Ly6C^hi monocytes (Fig 4A). Interestingly, while fGFP expression in Cx3cr1cre:Rosa26-fgfp mice was detected in subpopulations of CD64⁺ macrophages and IA/IE⁺CD11c⁺CD64⁻CD24⁺ DC, by semi-quantitative PCR, Hif1α was not effectively deleted in these cell types (Fig 4B-C, and S3A). Although Hif1α was not deleted in macrophages under steady state conditions (Fig 4A), Hif1α deletion was observed in IMØ derived from hypoxic animals (Fig S3C). Hif1α deletion had no significant effect on pulmonary immune cell composition (Fig S3D), baseline body weight, heart rate, or hemoglobin levels (Table S1). Relative to Hif1α^flox/flox controls, normoxic Cx3cr1cre;Hif1α^Δ/Δ mice displayed no abnormalities in right ventricular systolic pressures (RVSP), right ventricular hypertrophy, as assessed by Fulton’s index [RV/(LV+S)], or pulmonary vessel muscularization in either the absence or presence of SU5416 (Fig 4D-F). However, after 4 weeks of hypobaric hypoxia, Cx3cr1cre;Hif1α^Δ/Δ mice displayed a significant reduction in RVSP compared to Hif1α^Δ/Δ littermates (Fig 4D). Similarly, when exposed to hypoxia plus SU5416, Cx3cr1cre;Hif1α^Δ/Δ mice displayed significant reductions in RVSP, right ventricular hypertrophy, and pulmonary vessel muscularization compared to Hif1α^Δ/Δ littermates (Fig 4B-E). Associated with the decreased muscularization of small pulmonary arteries in hypoxia-exposed Cx3cr1cre;Hif1α^Δ/Δ mice, there was a decreased accumulation of CD64⁺ pulmonary IMØ around small pulmonary arteries (Fig 4G). These findings demonstrate that non-classical monocytes are the myeloid cell type that senses hypoxia, infiltrates small pulmonary arteries, differentiates into IMØ, and directly contributes to the development and/or progression of PH.

Figure 4.

Ly6C^lo monocytes sense hypoxia and modulate hypoxia-induced PH. A-C) Quantitative PCR depicting Hif1α expression in subsets of monocytes (A), total MØ (B), DC (B), and MØ subsets (C) derived from untreated Cx3cr1cre;Hif1α^Δ/Δ vs. Hif1α^f/f animals. Representative of three experiments. D) Bar graph depicts RVSP measurements in animals exposed to normoxia (N), hypoxia (H); normoxia plus Su5416 (N+Su5416); and hypoxia plus Su5416 (H+Su5416) (n=6-12/group). E) Bar graph depicts Fulton index [RV/(LV+S)] measurements in animals exposed to two models of hypoxia-induced PH (n= 8-21/group). F) Bar graph depicting percent of muscularized small pulmonary arteries in animals exposed to N+Su5416 vs. H+Su5416 (n=6/group). All lobes of the lungs were examined in cross-sections of approximately 0.5 cm thickness. All lung sections were embedded as single block in paraffin. Tissue sections of 5μm were obtained and stained with anti-human von-Willebrand factor (vWF) and anti-human α-smooth muscle cell actin (SMA). Entire slide was imaged and stitched into single image. In a blinded fashion, vessels under 50μm on the entire slides were categorized as no muscularization (absence of SMA staining around the vessel), partially muscularized (SMA staining in parts of the vessel), and fully muscularized (SMA staining encircles the entire circumference of the vessel). Muscularized vessels were expressed as percent of total vessels enumerated. N = no muscularization; P = partial muscularization; and F = complete muscularization. G) Immunofluorescence images of lung tissues derived from Hif1α^flox/flox vs. Cx3cr1cre;Hif1α^Δ/Δ exposed to two hypoxia-induced PH models. Red = anti-αSMA; Cyan = anti-CD64; Blue = DAPI; and White = co-localization of CD64+ IMØ (Cyan) around muscularized small pulmonary arteries (Red). Statistical analyses were performed with two way ANOVA. Measurements displayed as Mean ± SEM. *p<0.5; **p<0.01; ***p<0.001; and ****p<0.0001.

Impaired maturation of Hif1α-deficient non-classical monocytes into mature disease-promoting MØ.

The above results raise questions concerning the mechanisms by which hypoxia-stimulated non-classical monocytes and IMØ may stimulate the development of PH. To identify functional pathways in these cell types that may contribute to PH pathogenesis, we performed RNA sequencing (RNAseq) of non-classical monocytes and IMØ purified from the lungs of Cx3cr1cre;Hif1α^Δ/Δ and control Hif1α^flox/flox mice after 3 weeks of exposure to hypoxia plus SU5416 (Fig 5A). Approximately 20 ~ 30 million 126 base pair paired-end reads were generated for each sample. Subsequently, 75% (15 - 23 million) quality concordant pair end reads for each sample were successfully aligned to the mouse reference transcriptomes, GRCm38. Finally, the read-count-per-gene measurements for each sample were performed to convert the mapped reads to read counts for a total of 26,608 genes. The read counts were then filtered and normalized by estimated size factors by using the R package “DESeq2” (Love et al. 2014). The gene expression differences across the treatment groups were then evaluated using the default generalized linear model in DESeq2. Genes passing the threshold, an FDR of <5%, were considered to be significantly differentially expressed. Comparing Hif1α-deficient to Hif1α-sufficient non-classical monocytes exposed hypoxia plus Su5416, 314 differentially expressed genes were identified, with 260 down-regulated and 54 up-regulated genes. For Hif1α-deficient IMØ, 1577 differentially expressed genes were identified (1003 down-regulated and 574 up-regulated).

Figure 5.

Transcriptomic analyses of pulmonary IMØ derived from hypoxic Cx3cr1cre;Hif1α^Δ/Δ vs. Hif1α^flox/flox animals revealed macrophage functions that may contribute to PH pathogenesis. A) Principal component analyses clustering of Ly6C^lo monocytes and IMØ sorted from H+Su5416 exposed Hif1α^flox/flox vs. Cx3cr1cre;Hif1α^Δ/Δ animals (n=3/group). B) Heat map showing decreased relative expression of macrophage maturation markers between IMØ derived from hypoxic Cx3cr1cre;Hif1α^Δ/Δ vs. Hif1α^flox/flox. C) Heat maps depicting IMØ derived from Cx3cr1cre;Hif1α^Δ/Δ animals have decreased expression of genes associated with phagocytosis-initiating pathways, including FcɣR-mediated phagocytosis and complement pathways. D) Heat maps showing IMØ derived from Cx3cr1cre;Hif1α^Δ/Δ animals have decreased expression of genes associated with many pathways involved in protein processing and antigen presentation. These include phagosome, lysosome, protein processing in endoplasmic reticulum, and presentation of antigen onto cell surface. Heat maps are expressed in logCPM (log counts per million).

Relative to controls, Cx3cr1cre;Hif1α^Δ/Δ IMØ expressed lower levels of macrophage maturation markers including CD64 (Fcɣr1), F4/80 (Emr1), CD88 (C5ar1), and LysM (Lyz2) (Fig 5B). In addition to these conventional markers for macrophage identification, Cx3cr1cre;Hif1α^Δ/Δ IMØ also down-regulated pathways associated with classical macrophage functions, including phagocytosis and complement activation (Fig 5C). These macrophages also down-regulated pathways associated with antigen presentation, which is essential for subsequent activation of adaptive immune response. In Cx3cr1cre;Hif1α^Δ/Δ IMØ, all processes required for efficient antigen presentation including phagosomes, lysosomes, protein processing in the endoplasmic reticulum (ER), and antigen presentation onto cell surface were reduced (Fig 5D). Overall, these findings suggest an impaired maturation of Cx3cr1cre;Hif1α^Δ/Δ non-classical monocytes into disease promoting IMØ, with a reduced capacity to mediate phagocytosis, complement activation, antigen presentation, and subsequent activation of innate and adaptive immune responses.

Both IMØ and non-classical monocytes from Cx3cr1cre;Hif1α^Δ/Δ animals expressed significantly lower levels of cytokines that have been associated with PH, including interleukin-1 (IL-1), interleukin-6 (IL-6), tumor necrosis factor (TNF) and transforming growth factor beta (TGFβ) (Fig 6). These findings suggest that non-classical monocytes and non-classical monocyte-derived infiltrating IMØ sense hypoxia and promote PH.

Figure 6.

Decreased expression of chemokines and cytokines associated with PH progression in Ly6C^lo monocytes and IMØ derived from hypoxic Cx3cr1cre;Hif1α^Δ/Δ animals. Heat map of differential IMØ and Ly6C^lo monocyte chemokine, cytokine, and receptor expressions between Hif1α^flox/flox vs. Cx3cr1cre;Hif1α^Δ/Δ. Heat map are expressed in logCPM (log counts per million).

Non-classical monocytes accumulate in the lungs of PAH patient.

The above findings suggest that, in hypoxic mice, the activity of non-classical monocytes in the lungs plays a key role in the development of PH. To determine if cells of the non-classical monocyte lineage accumulate in human PAH lungs, we examined monocyte populations in peripheral lung tissues from control and PAH patients using flow cytometry and the gating strategy shown in Figure 7C (31). Control lungs were derived from human donors who were declined at the time of transplant. PAH lung tissues were obtained at the time of transplant. In control lungs, the vast majority of monocytes were CD14^hiCD16^lo, the human equivalent of murine classical monocytes (Fig 7A and B). In comparison, PAH lungs displayed a 5-6 fold increase in CD14^loCD16^hi non-classical monocytes, the human equivalent of Ly6C^lo non-classical monocytes (Fig 7A and B). These findings demonstrate that non-classical monocytes specifically accumulate in the lungs of human PAH patients, suggesting that these cells contribute to the pathogenesis of PAH in humans.

Figure 7.

Flow cytometric analyses of human PAH lung explants revealed increased non-classical monocyte accumulations in diseased lungs. A) Bar graphs depict increased frequency of CD14^loCD16⁺ non-classical monocytes in human PH lungs, compared to controls (n=3 per group). Statistical analyses performed using unpaired t tests. Bar graphed as Mean ± SEM. *p<0.05. B) Representative dot plots showing monocyte subpopulations in control and PAH lung tissues. C) Gating strategy of human lung tissues for identification of monocyte subpopulations. R1 represents single cells. R2 identifies CD45⁺ leukocytes. Live CD45+ cells are included in R3. Granulocytes, including neutrophils and eosinophils, were excluded (R4). CD206⁻CD14⁺ (R5) cells are monocytes. Monocyte subpopulations were identified based on the expression of CD14 and CD16 (CD14⁺ classical monocytes, CD14⁺CD16⁺ monocytes, and CD14^loCD16⁺ non-classical monocytes).

Discussion

Myeloid cell infiltration of the pulmonary vasculature is a common feature of PH in humans and all animal models; however, the specific infiltrating cell type has not been identified and the causal relationship between this inflammatory response and disease pathogenesis has remained unclear. Here, using mouse models of hypoxic PH, we can draw three novel conclusions about the myeloid cells that infiltrate the pulmonary vasculature during the development of PH. First, we show that the vasculature-infiltrating cells are IMØ. Second, that these cells arise from circulating non-classical monocytes. Third, that these non-classical monocyte-derived vascular infiltrating macrophages sense hypoxia and directly contribute to PH pathogenesis.

It has been previously thought that DC are the main accumulating myeloid cell type in PH (6, 8). In contrast, we found that IMØ are the only myleoid cell type that accumulates in lungs in hypoxia-induced PH. Our conclusion is based on the finding that the only cells that accumulate in PH lungs are CD45⁺, CD11c^int/hi, CD11b^hi, CD64^hi, CX3CR1⁺ and CD169⁺, while being Ly6G⁻, Ly6C⁻, CD24⁻, and Siglec F⁻, and that the vasculature-infiltrating cells are CD64⁺, CX3CR1⁺ and CD169⁺. This is the phenotype of pulmonary IMØ as we and others have described (32, 37). The finding of expanded pulmonary IMØ in PH is consistent with our preliminary findings and other recent reports (45-47). Previous reports of DC accumulation likely stem from the fact that pulmonary IMØ also express CD11c and major histocompatibility complex II (MHCII), which were commonly used to identify DCs, but are now known to be expressed by macrophages. Thus, the myeloid cells that accumulate in PH are pulmonary IMØ.

Our second conclusion is that vasculature-infiltrating IMØ arise primarily from circulating bone marrow-derived non-classical monocytes. This conclusion is based on three findings: 1) that the cells that accumulate in hypoxic lungs are of donor origin after bone marrow transplantation; 2) that after labeling circulating bone marrow-derived non-classical but not classical monocytes with PKH26PCL, the vast majority of IMØ in hypoxic lungs are are PKH26PCL⁺; and 3) IMØ in hypoxic lungs display only minimal proliferation. This conclusion is consistent with our prior finding that classical monocytes, which depend on CCR2 for trafficking into tissue, are not major precursors to the accumulating pulmonary IMØ (28). Also, the decrease in Ly6C^lo non-classical monocyte numbers in the lungs of animals exposed to hypoxia or hypoxia+Su5416 likely reflects maturation of monocytes into monocyte-derived macrophages (28). Mature macrophages are speicalized cells with enhanced phagocytic capaity. PKH26PCL was designed to label highly phagocytic cells and has been used to differentiate infiltrating Ly6c^hi classical monocyte-derived macrophages from resident macrophages. Labeling of monocyte subsets for in vivo imaging and tracking has been challenging (25). In this study, we demonstrate that PKH26PCL administration and timing can be optimized to label monocyte subsets differentially. Prior studies suggest that Ly6C^lo non-classical monocytes represent a group of more mature and specialized monocytes that patrol the vasculature (25). The enhanced ability of Ly6C^lo non-classical monocytes to take up and retain PKH26PCL, compared to Ly6C^hi classical monocytes, is consistent with the view that Ly6C^lo non-classical monocytes represent a more mature population along the monocyte-macrophage continuum (25). Addtionally, our findings that cells of Ly6C^lo non-classical monocyte lineage accumulate in small pulmonary arteries in PH further support a specialized role for these cells in regulating vassel homeostasis and disease (27, 48).

The accumulation of myeloid cells has been described extensively in humans and animal models of PH. The extent to which these cells play a direct causal role in PH pathogenesis has remained contraversial due to the fact that evidence for such a role for these cells has been limited. Our third and key conclusion is that cells of the non-classical monocyte lineage play a direct causal role in PH development. We find that the sensing of hypoxia by this lineage promotes vascular remodeling and contributes significantly to the development of hypoxia-induced PH. Using Cx3cr1cre to target Hif1α deletion in Ly6C^lo non-classical lineage, we show that Hif1α-deficient animals have a reduced severity of PH. Based on the expression of Cx3cr1cre:fGFP, we had expected that Cx3cr1cre would be expressed and reduce Hif1α transcripts in subpopulations of pulmonary macrophages and dendritic cells (Fig 4B-C, and Fig S3). However, by semi-quantitative PCR, Hif1α transcripts remained intact in pulmonary macrophages and dendritic cells. This finding demonstrates that specific gene deletions are affected by factors beyond presence or absence of cre recombinase. Potential determinants include timing and the expression level of cre recombinase in relation to targeted gene locus accessbility (49, 50). Monocytes are drived from a distinct precursor lineage, compared to majority of resident pulmonary macrophages and classical dendritic cells (19). This distinct cell differentiation and maturation program may affect the efficiency of Hif1α deletion. Thus, the efficiency and specificity of targeted gene deletion should not be inferred based on reporter expression, but should be confirmed empirically (51). Using Cx3cr1cre:Hif1α, we are not able to examine the contribution of resident macrophages to PH pathogenesis. However, the finding of Hif1a gene deletion in IMØ derived from hypoxic but not normoxic animals is consistent with the view that, in response to hypoxia, non-classical monocytes serve as progenitors for infiltrating IMØ (Fig 4A and Fig S3C). Thus, our findings suggest a direct causal role of infiltrating Ly6C^lo non-classical monocytes to PH development. This conclusion is supported by previous studies in which interventions that increased cell numbers that would include non-classical monocytes, their precursors, or their derivatives increased the severity of PH. On the other hand. interventions that decreased the number of such cells also decreased PH severity (4, 8, 10, 11, 16). Our findings suggest a model in which hypoxia, working via HIF activation, stimulates non-classical monocytes to infiltrate small pulmonary arteries, differentiate into IMØ, and produce factors that stimulate pulmonary vascular remodelling.

At present, the specific factors that stimulate pulmonary vascular remodelling remain to be identified. In our transcriptome analyses, by PCA scatter, there was not complete separation between IMØ derived from Hif1α-sufficient and Hif1α-deficient animals. This may be because our sorted IMØ population would have included Hif1α-sufficient resident IMØ (Fig 5A). Consistent with the mixture of Hif1α-sufficient resident IMØ and Hif1α-deficient IMØ in our RNA sequencing analyese, and the increased absolute number of pulmonary IMØ (~2 fold) in hypoxic animals, our transcriptomic analyses displayed a ~50% reduction in Hif1α expression between IMØ derived from Hif1α-deficient vs. Hif1α-sufficient hypoxic animals (Fig 1A, and Fig S3C). Even with this mixture of infiltrating and resident IMØ population, we identified significant alterations in IMØ derived from non-classical monocyte-specific Hif1α-deficient animals. Hif1α-deficient IMØ express lower levels of MØ-specific transcripts (Fcɣr1/CD64, Emr1/F4-80, CD68, C5ar1/CD88, and Lyz2), suggesting these MØs are immature. Consistent with this immature phenotype, many pathways associated with quientecential MØ functions are decreased, including phagocytosis, complement activation, and processes leading to antigen presentation. Antigen presentation by MØs are important for regulating both T can B cell functions. Thus, Hif1α-deficient IMØ may have a reduced capacity to activate or perpetuate subsequent innate and adaptive immune response. In addition, non-classical monocytes and IMØ derived from targeted Hif1α-deleted animals display reduced transcripts for both M1 and M2 cytokines and chemokines that have been implicated in PH development and progression, including Il-6, TNF, and TGFβ (11, 15, 52). Thus, rather than skewing MØ M1 vs. M2 profiles, deletion of Hif1α in non-classical monocyctes prohibits maturation of these monocytes into disease-promoting MØs. Thus, non-classical monocyte-derived cells orchestrate a vascular microenvironment, through immune activation and cytokine production, that promotes vascular remodeling.

With respect to human PH, our finding that cells of the non-classical monocyte lineage are uniquely increased in lung explants from patients with Group I pulmonary arterial hypertension (PAH) supports the view that cells of this lineage may play a role in human PH. One significant difference between our murine and human results was that in the murine studies we demonstrated an accumulation of pulmonary IMØ while, in Group I PAH lung explants, we found an accumulation of non-classical monocytes. This difference may be explained by the disparate disease stages. Mouse models of PH generally produce relatively mild and reversible pathology that is similar to early stages of disease. Human PAH lung explants are derived from patient with non-reversible, end-stage disease. It is likely that cellular differentiation kinetics differ between disease stages and species. Despite this limitation, our findings support a role for the non-classical monocyte lineage in PH pathogenesis. This view is consistent with studies showing that humans with mutations in BMPR2 display increased production of GM-CSF, a potent monocyte chemoattractant, in small pulmonary arteries and infiltration of vessels by GM-CSFRa⁺CD68⁺ cells, the phenotype of monocyte-macrophages (16). Our results support the view that BMPR2 mutations promote PAH, at least in part, by stimulating the accumulation of IMØ lineage cells in small pulmonary arteries. This would represent one “hit” in the “multiple hit” model of PAH pathogenesis (53).

It is known that multiple distinct cell types contribute to the development of PH through the sensing of hypoxia. The specific deletion of Hif2α in endothelial cells reduces the severity of PH in hypoxic models (54, 55). Pulmonary smooth muscle cell-specific HIF1α has also been shown drive to disease development (56, 57). Until now, direct evidence that immune cells also contribute to PH through the sensing of hypoxia has been lacking (58). Here, we demonstrated that non-classical monocyte-derived pulmonary IMØ sense hypoxia and directly contribute to PH pathogenesis. This suggests that in multiple types of PH, infiltration of IMØs into small pulmonary arteries may be stimulated by hypoxia itself in the context of alveolar hypoventilation, pulmonary emboli, systemic inflammation, or local infection (59, 60). If such IMØ infiltration is a common phenomenon, it would predispose those affected to the development of PH. As such, specific factors produced by IMØ that stimulate pulmonary vascular remodeling may represent attractive therapeutic targets for multiple types of PH in patients.

Key Points.

• Ly6C^lo non-classical monocytes infiltrate the pulmonary vasculature and promote PH.

• Hif1α regulates the maturation of non-classical monocytes into infiltrating IMØ.

• Infiltrating IMØ promote PH by modulating local immune and cytokine responses.

Abbreviations

AMØ

Alveolar macrophages

BMPR2

Bone morphogenic protein receptor 2

DC

Dendritic Cells

fGFP

Farnesylated GFP

GM-CSF

Granulocyte-macrophage colony-stimulating factor

Hif1α

Hypoxia-inducible factor-1α

H

Hypoxia

IMØ

Interstitial macrophages

N

Normoxia

PAH

Pulmonary arterial hypertension

PH

Pulmonary hypertension

SMA

α-Smooth muscle cell actin