Bone marrow transplantation rescues monocyte recruitment defect and improves cystic fibrosis in mice

Abstract

Cystic fibrosis (CF) is an inherited life-threatening disease accompanied by repeated lung infections and multi-organ inflammation that affects tens of thousands of people worldwide. The causative gene, cystic fibrosis transmembrane conductance regulator (CFTR), is mutated in CF patients. CFTR functions in epithelial cells have traditionally been thought to cause the disease symptoms. Recent work has shown an additional defect: monocytes from CF patients show a deficiency in integrin activation and adhesion. Since monocytes play critical roles in controlling infections, defective monocyte function may contribute to CF progression. In this study, we demonstrate that monocytes from CFTR^ΔF508 mice (CF mice) show defective adhesion under flow. Transplanting CF mice with wild-type (WT) bone marrow after sublethal irradiation replaced most (60–80%) CF monocytes with WT monocytes, significantly improved survival, and reduced inflammation. WT/CF mixed bone marrow chimeras directly demonstrated defective CF monocyte recruitment to the bronchoalveolar lavage and the intestinal lamina propria in vivo. WT mice reconstituted with CF bone marrow also show lethality, suggesting that the CF defect in monocytes is not only necessary but also sufficient to cause disease. We also showed that monocyte-specific knockout of CFTR retards weight gains and exacerbates DSS-induced colitis. Our findings show that providing WT monocytes by bone marrow transfer rescues mortality in CF mice, suggesting that similar approaches may mitigate disease in CF patients.

Introduction

Cystic fibrosis (CF) is one of the most common monogenic diseases (more than 70,000 cases worldwide, ~1,000 new cases each year) (1), which is caused by variations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. CFTR is an anion-conducting transmembrane channel that is very important in mucus formation and the ion homeostasis of cells (2). More than 2000 gene variants have been identified, which are grouped into six types of mutations (2). In humans, CF disease is dominated by lung and pancreas pathologies. The dominant pathology in the lung is inflammation caused by failure to clear microorganisms and the generation of a toxic pro-inflammatory microenvironment (2–4). CFTR dysfunction in the pancreas results in pancreatic insufficiency in most CF patients (2).

CFTR mutations result in mucus dysfunction (5) and impaired mucociliary clearance in sino-pulmonary tissue. The resulting viscous mucus becomes infected with pathogens, such as Pseudomonas aeruginosa (6–9), Staphylococcus aureus (10, 11), Haemophilus influenza (12), and Aspergillus species (13). These recurrent infections greatly exacerbate the symptoms of CF (4, 5, 14) and impact the patients’ quality of life. Current clinical management of CF includes prophylactic (flucloxacillin) and therapeutic antibiotics (nebulized tobramycin, colistin, and aztreonam), DNAse (Dornase alfa), CFTR potentiators and correctors (Ivacaftor, Lumacaftor), and mucus thinners (Hypertonic saline) (15). Although progress has been made (16), there is currently no cure for CF.

There are several animal models of CF, some of which mimic human pathology. In several mouse models of CF (17, 18), homologues of common human CFTR mutations have been knocked into the mouse CFTR locus. For the present mechanistic study, we use the CFTR^ΔF508 mouse (19), a widely used and well-accepted mouse model of CF (20, 21), although it is dominated by intestinal pathology, mucus accumulation in the crypts of Lieberkühn, goblet cell hypertrophy, hyperplasia, and eosinophilic concretion in the crypts, ultimately resulting in constipation, rectal prolapse, ileus and death (22).

A recent study reported that mutations of CFTR commonly found in CF patients cause a profound monocyte adhesion defect (23, 24). The authors demonstrated expression of CFTR in human monocytes and found a significant defect of their integrin activation and static adhesion. A similar defect of static adhesion was reported in CFTR^ΔF508 mouse monocytes, resulting in a ~60% (human) or ~40% (mouse) loss of monocyte static adhesion. Adhesion under flow was not studied. Activation of the three main monocyte integrins (25), β₂ (α_Lβ₂, α_Mβ₂) and α₄ (α₄β₁), was found to be defective. Remarkably, this defect is restricted to monocytes, with no defect in neutrophils and a minor defect (~20–30%) in lymphocytes. These features and the molecular mechanism make it different from the known leukocyte adhesion deficiencies (LAD) I, II, and III (26), suggesting that CF is associated with a new leukocyte adhesion deficiency, LAD IV (24).

Monocytes are important guardians of epithelial surfaces. They serve by directly secreting cytokines (27) and killing pathogens (28, 29). Monocytes can differentiate into macrophages and inflammatory dendritic cells (30). Macrophages clear apoptotic debris (efferocytosis), secrete cytokines for tissue homeostasis and survey the environment (31). When macrophages detect pathogens or danger signals, they directly phagocytose and kill bacteria (29, 32). They also produce many inflammatory cytokines and chemokines that orchestrate the recruitment of other immune and inflammatory cells (27), which are involved in the inflammatory antimicrobial response, wound healing, and fibrosis (33). Monocyte-derived dendritic cells migrate to the draining lymph nodes, where they present antigens to CD4 and CD8 T cells and thus control the adaptive immune response.

Given the expression of CFTR in monocytes (23), and given the monocyte adhesion defect, we reasoned that monocyte recruitment to mucosal sites might be defective in CFTR^ΔF508 mice (CF mice), resulting in weakened host defense and increased pathogen burden. Here, we report that the monocyte recruitment is indeed defective. We reasoned that correcting CFTR in monocytes might improve CF disease burden and clinical outcomes. To test this, we performed bone marrow transplantation (BMT) to generate mice that were defective in CFTR in hematopoietic cells, non-hematopoietic cells, both, or neither. We observed improvement of CF symptoms and longer survival in mice that received wild-type (WT) bone marrow (BM). Mechanistically, we show that CF monocytes have a severe recruitment defect to the intestinal lamina propria and the bronchoalveolar space. Since it is now possible to correct genetic defects in hematopoietic stem cells (HSCs) (34, 35), these data suggest that autologous transfer of engineered HSCs or allogeneic BMT should be attempted in patients with CF.

Materials and Methods

Reagents

Recombinant mouse P-selectin-Fc, ICAM-1-Fc, and CCL2 were purchased from R&D Systems. Casein blocking buffer, Penicillin-Streptomycin solution, pHrodo™ Red Staphylococcus aureus bioparticles, and dextran sulfate sodium (DSS) were purchased from Thermo Fisher Scientific. Roswell Park Memorial Institute (RPMI) medium 1640 without phenol red, phosphate-buffered saline (PBS) without Ca²⁺ and Mg²⁺, Hanks’ balanced salt solution (HBSS) with phenol red without Ca²⁺ and Mg²⁺ were purchased from Gibco. Fetal bovine serum (FBS) and human serum albumin (HSA) were purchased from Gemini Bio Products. Phorbol 12-myristate 13-acetate (PMA), Type VIII collagenase, Dnase, and N-Acetyl-L-cysteine I were purchased from Sigma-Aldrich. Phycoerythrin (PE) or PE-Cyanine 7 (Cy7)-conjugated anti-CD115 antibody (clone AFS98), Alexa Fluor 647 (AF647), AF700, or Brilliant Violet 650 (BV650)-conjugated anti-Ly6G antibody (clone 1A8), BV785-conjugated anti-CD11b antibody (clone M1/70), BV605-conjugated anti-CD11c antibody (clone N418), BV570-conjugated anti-Ly6C antibody (clone HK1.4), BV711-conjugated anti-TCRβ antibody (clone H57–597), Allophycocyanin (APC)-conjugated anti-CD117 antibody (clone 2B8), APC-Cy7-conjugated anti-CD19 antibody (clone 6D5), PE-conjugated anti-Ly6A/E antibody (clone D7), BV650-conjugated anti-NK1.1 antibody (clone PK136), PE-conjugated anti-F4/80 antibody (clone BM8), FITC-conjugated anti-CCR2 antibody (clone SA203G11), unconjugated anti-CD11b blocking antibody (clone M1/70), and unconjugated CD18 blocking antibody (clone M18/2) were purchased from Biolegend. Ghost Dye™ Violet 510 was purchased from Tonbo biosciences. Fluorescein‎ (FITC)-conjugated anti-CD45.1 antibody (clone A20) was purchased from BD Biosciences. Peridinin Chlorophyll Protein Complex (PerCP)-Cy5.5-conjugated anti-CD45.2 antibody (clone 104), PE-eFluor610-conjugated anti-CD4 antibody (clone GK1.5), eFluor450-conjugated anti-CD8a antibody (clone 53–6.7) were purchased from eBioscience. Red blood cell (RBC) lysis buffer was purchased from Invitrogen. iC3b was purchased from Complement Technology. EasySep™ Mouse Monocyte Isolation Kit was purchased from STEMCELL Technologies.

Mice

C57BL/6J WT mice (000664; JAX), DsRed mice (006051; JAX; WT C57BL/6J background), CD45.1 mice (002014; JAX; WT C57BL/6J background), CFTR^ΔF508 or CF mice (002515; JAX; C57BL/6J background), and CSFR1-cre mice (029206; JAX; C57BL/6J background) were obtained from the Jackson Laboratory. CFTR^flox/flox (Cftr^tm1Cwr; C57BL/6J background) mice were developed by and obtained from the Cystic Fibrosis Mouse Models Core at Case Western Reserve University. CD45.1/CD45.2 mice were bred by C57BL/6J WT and CD45.1 mice. Mice were fed a standard rodent chow diet and were housed in microisolator cages in a pathogen-free facility in the La Jolla Institute for Immunology or UConn Health. Mice were euthanized by CO₂ inhalation. All experiments followed guidelines of the La Jolla Institute for Immunology and UConn Health Animal Care and Use Committee, and approval for the use of rodents was obtained from the La Jolla Institute for Immunology and UConn Health according to criteria outlined in the Guide for the Care and Use of Laboratory Animals from the National Institutes of Health.

BMT

WT or CF Recipients (C57BL/6J background littermates, male or female, one-month-old) were irradiated in two doses of 550 or 350 rads each (for a total of 1,100 or 700rads) four hours apart (RS-2000 X-Ray irradiator, Rad Source). Bone marrow cells from both femurs and tibias of donor mice (WT or CF mice, sex-matched, one-month-old) were collected under sterile conditions. Bones were centrifuged for the collection of marrow, and unfractionated bone marrow cells were washed, resuspended in PBS, injected retro-orbitally into the lethally irradiated mice (one donor to five recipients). Recipient mice were housed in a barrier facility under pathogen-free conditions before and after bone marrow transplantation. After bone marrow transplantation, mice were provided autoclaved acidified water with antibiotics (trimethoprim-sulfamethoxazole) and were fed autoclaved food. The survival of mice was monitored. Mice were used for further experiments eight weeks after bone marrow reconstitution. In some experiments, DsRed WT mice were used as recipients or donors to test the reconstructive rate of BMT by flow cytometry.

In the mixed chimeric BMT, CD45.1/CD45.2 or WT C57BL/6J mice (eight weeks old males) were irradiated in two doses of 550 rads each (for a total of 1,100 rads) four hours apart (RS-2000 X-Ray irradiator, Rad Source). Bone marrow cells from both femurs and tibias of donor mice (CF male and CD45.1 WT male, eleven weeks old) were collected under sterile conditions. Bones were centrifuged for the collection of marrow, and unfractionated bone marrow cells were washed, resuspended in PBS, mixed at a ratio of 1:1 or 1:2, confirmed by flow cytometry, and injected retro-orbitally into the lethally irradiated mice (one donor to five recipients). Recipient mice were housed in a barrier facility under pathogen-free conditions before and after bone marrow transplantation. After bone marrow transplantation, mice were provided autoclaved acidified water with antibiotics (trimethoprim-sulfamethoxazole) and were fed autoclaved food. Mice were used for further experiments eight weeks after bone marrow reconstitution.

Bodyweight and hematology

The bodyweight of aged CF and WT littermates (40–72 weeks) and young CFTR^flox/flox Csfr1-cre^+/− and CFTR^flox/flox Csfr1-cre^−/− littermates (age from 3–6 weeks) were recorded. Blood counts of WT or CF littermates (age from 10–25 weeks) were taken via retro-orbital bleeding and analyzed by an automatic analyzer (Hemavet 950FS, DREW Scientific). Mice were anesthetized by the inhalation of the isoflurane/oxygen gas mixture during the bleeding.

Flow cytometry

Blood was obtained by cardiac puncture with an ethylenediaminetetraacetic acid (EDTA)-coated syringe. Red blood cells were lysed in RBC Lysis Buffer according to the manufacturer’s protocol. Lungs were lavaged with PBS containing 2mM EDTA, dissociated by gentleMACS™ Dissociator (Miltenyi), and filtered through a 70 μm strainer. Ten centimeters of small intestines from the end of the stomach, which contains the duodenum and a part of the jejunum, were collected. For the preparation of lamina propria (LP) cells, fat and connective tissue were removed from the intestines. Intestines were opened longitudinally, cut into 1-cm pieces, and washed 3× 10 minutes in HBSS containing 5% FBS, 2 mM EDTA, 100 mM N-acetyl cysteine, and 10 mM HEPES to remove mucus and epithelial cells. Tissue was digested with 1 mg/mL collagenase VIII (Sigma) and 20 μg/mL DNase I (Sigma) for 20 minutes at 37°C. Digested material was dissociated by gentleMACS™ Dissociator (Miltenyi) and filtered through a 70 μm strainer. Bone marrow cells were collected from both femurs and tibias. In some experiments, mice were challenged intranasally with mouse CCL2 100ng/mouse 12 hours prior to the harvest. Blood was collected via retro-orbital bleeding. Bronchoalveolar lavage fluid (BAL) was collected by intratracheal lavage with PBS before the collection of lungs.

All samples were collected in PBS with 2 mM EDTA to prevent cation-dependent cell-cell adhesion and were stored on ice during transportation, staining, and analysis. Cells were resuspended in 100 μl flow staining buffer (1% BSA and 0.1% sodium azide in PBS). Cells were stained with Ghost Dye™ Violet 510 for analysis of viability. Fcγ receptors were blocked for 15 min, and surface antigens on cells were stained for 30 min at 4°C with directly conjugated fluorescent antibodies (anti-CD45.1-FITC, anti-CD45.2-PerCP-Cy5.5, anti-CD11b-BV785, anti-CD11c-BV605, anti-CD115-PE-Cy7, anti-Ly6C-BV570, anti-Ly6G-AF700, anti-TCRβ-BV711, anti-CD4-PE-eFluor610, anti-CD8a-eFluor450, anti-CD19-APC-Cy7, anti-CD117-APC, anti-Ly6A/E-PE, and anti-NK1.1-BV650 for blood and bone marrow cells; anti-CD45.1-FITC, anti-CD45.2-PerCP-Cy5.5, anti-CD11b-BV785, anti-CD11c-BV605, anti-CD115-PE-Cy7, anti-Ly6C-BV570, anti-Ly6G-AF700, anti-TCRβ-BV711, anti-CD4-PE-eFluor610, anti-CD8a-eFluor450, anti-CD19-APC-Cy7, anti-F4/80-PE, and anti-NK1.1-BV650 for BAL, lung, and LP cells). Forward and side-scatter parameters were used for the exclusion of doublets from the analysis. Cell fluorescence was assessed with an LSRII (BD Biosciences) and was analyzed with FlowJo (BD, version 10.4). Lung and LP monocytes or macrophages were normalized by blood monocytes to eliminate the reconstructive rate difference of WT and CF BM after BMT.

In CCR2 expression experiment, blood was obtained by retro-orbital bleeding. Red blood cells were lysed in RBC Lysis Buffer according to the manufacturer’s protocol. Fcγ receptors were blocked for 15 min, and surface antigens on cells were stained for 30 min at 4°C with directly conjugated fluorescent antibodies (anti-Ly6G-BV650, anti-CD115-PE, and anti-CCR2-FITC). Forward and side-scatter parameters were used for exclusion of doublets from the analysis. Cell fluorescence was assessed with an LSRII (BD Biosciences) and was analyzed with FlowJo (BD, version 10.4). Monocytes were gated as CD115⁺Ly6G⁻ cells, and the median fluorescence intensities (MFI) of FITC were used to quantify CCR2 expression.

In the phagocytosis assay, pHrodo™ Red Staphylococcus aureus bioparticles (1.33 mg/mL) were immobilized by iC3b (0.33 mg/mL) at 4ºC overnight and washed twice with PBS before use. Bone marrow cells were collected from both femurs and tibias of WT and CF mice. Monocytes were purified using EasySep™ Mouse Monocyte Isolation Kit according to the manufacturer’s protocol. Purified monocytes were resuspended in 1.5×10⁶/mL, incubated with 10 μM PMA and 20 μg/mL CD11b, CD18 blocking antibodies, or isotype controls at room temperature (RT), 300 rpm (plate shaker) for 15 minutes, and incubated with iC3b-immobilized pHrodo™ Red Staphylococcus aureus bioparticles (66.6 μg/mL) at 37 ºC, 100 rpm (shaking incubator) for one hour. After two washes with 1% PFA and one wash with PBS with 2 mM EDTA, cell fluorescence was assessed with an LSRII (BD Biosciences) and was analyzed with FlowJo (BD, version 10.4). The MFI of pHrodo™ Red Staphylococcus aureus bioparticles in monocytes was used to quantify phagocytosis. The MFI was normalized to the CD18 blockade sample of each mouse to eliminate the background of β2 integrin-independent phagocytosis.

Histology

Intestines were rinsed, opened on the anti-mesenteric side, cut into three strips, and placed in parallel on a biopsy pad in a cassette. Samples were then fixed in zinc formalin, embedded in paraffin, and cut into 3- to 5-μm sections. The tissues were stained with hematoxylin and eosin (H&E) and periodic acid-Schiff (PAS) for histological assessment by a single investigator who was blinded to the experimental design. Slides were digitized on AxioScan Z1 slide scanner using 40× 0.95NA objective (Zeiss). The height of intestinal villi and crypt and the percentage of intestinal crypts that contain mucus were quantified in ZEN Blue software (Zeiss).

Microfluidic perfusion assay

The assembly of the microfluidic devices used in this study and the coating of coverslips with recombinant mouse P-selectin-Fc and ICAM-1-Fc have been described previously (36–39). Briefly, cleaned coverslips were coated with P-selectin-Fc (2 μg ml⁻¹) and ICAM-1-Fc (10 μg ml⁻¹) for 2 hours and then blocked for 1 hour with casein (1%) at RT. After coating, coverslips were sealed to polydimethylsiloxane chips by magnetic clamps to create flow chamber channels ~29 μm high and ~300 μm across. By modulating the pressure between the inlet well and the outlet reservoir, 6 dyn cm⁻² wall shear stress was applied in all experiments.

For the in vitro adhesion assay of monocytes, isolated mouse BM cells (10⁷ cells ml⁻¹) from WT or CF mice were incubated with PE-conjugated anti-CD115 antibody and AF647-conjugated anti-Ly6G antibody for 10 minutes at RT and perfused through the microfluidic device over a substrate of recombinant mouse P-selectin-Fc and recombinant mouse ICAM-1-Fc at a wall shear stress of 6 dyn cm⁻². After cells were rolling on the substrate, 100 ng ml⁻¹ mouse CCL2 was perfused. The processes of rolling and arrest were recorded by epifluorescence microscopy using an Olympus IX71 inverted microscope equipped with a 40×0.95NA objective.

DSS-induced colitis

Colitis was induced by adding 2% DSS in the drinking water for five days, followed by regular water. Mice (males, 11 weeks old) were monitored and weighed every day for two weeks and then sacrificed. Bodyweight data is normalized by the initial body weight to determine the weight lost.

Statistics

Statistical analysis was performed with Prism 6 (GraphPad). Data are presented as survival curve (Fig. 1A,2A,3A), mean ± SD plus individual data points (Fig. 1B–F, I–L,2C,D,F,3C–E,4B–G,5, Supplemental Figure 3,2B,C,E), and individual data points with linear regressions (Supplemental Figure 1). Survival curves were compared using the Gehan-Breslow-Wilcoxon test. The means for the data sets were compared using the Student t-tests with equal variances. Linear regressions were compared using the F-test. P values less than 0.05 were considered significant.

FIGURE 1.

Leukocytosis and monocyte adhesion and phagocytosis deficiency in CFTR^ΔF508 (CF) mice. (A-B) the survival curve (A, n=21 WT and 11 CF) and weight (B, n=12) of CF and WT mice. (C-F) Hematology for total leukocytes (C), monocytes (D), neutrophils (E), and lymphocytes (F) in CF mice (n=17) compared to WT controls (n=18). (G-H) Monocytes and neutrophils in bone marrow (BM) cells are distinguished by staining of CD115 and Ly6G staining, respectively, in flow cytometry (G) and by fluorescence microscopy in a microfluidic flow chamber (H). Both monocytes (red) and neutrophils (cyan) adhered on the substrate of P-selectin/ICAM-1 at a shear stress of 6 dyn·cm⁻² (H). The scale bar is 10 μm. (I-J) Number of rolling (white) and arrested monocytes at rest and after infusion of CCL2 (100 ng·mL⁻¹) (I). The percentage of arrested as a fraction of all monocytes in a field of view (J) for WT and CF monocytes with or without the stimulation of CCL2. Mean±SD of n=9 fields-of-view. (K) CCR expression of WT and CF blood monocytes. Mean±SD of n=9 mice. (L) The MFI of phagocytized pHrodo™ Red Staphylococcus aureus bioparticles in PMA-stimulated monocytes purified from WT or CF BM in presents with isotype control (open boxes), CD11b blocking antibody (light grey boxes), or CD18 blocking antibody (dark grey boxes). Mean±SD of monocytes from n=9 mice. n.s. p>0.05, *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001 by Gehan-Breslow-Wilcoxon test (A) and student T-test (B-F,I-L).

FIGURE 2.

WT Bone marrow transplantation partially rescues the survival and intestine inflammation in CFTR^ΔF508 mice. (A) the survival curve of CFTR^ΔF508 mice (700 rads) transplanted with WT (green curve, n=9;) or CFTR^ΔF508 (orange curve, n=5) BM. (B) Typical H&E staining images of intestines from CFTR^ΔF508 mice transplanted with CFTR^ΔF508 or WT BM. (C-D) Quantification of crypts (C, Mean±SD of n=9) and villi (D, Mean±SD of n=27) height of intestines from CFTR^ΔF508 mice transplanted with CFTR^ΔF508 or WT BM. n=9. (E) A typical PAS staining image of intestine from a CFTR^ΔF508 mouse transplanted with WT BM. White arrows indicate mucus in crypts. (F) Percentage of intestine crypts that contain mucus in it. CFTR^ΔF508 mice are transplanted with WT or CFTR^ΔF508 BM. Mean±SD of n=27. n.s. p>0.05, *p<0.05, **p<0.01in the Gehan-Breslow-Wilcoxon test (A) and student t-test (C-D,F). Scale bars are 100 μm.

FIGURE 3.

CFTR^ΔF508 Bone marrow transplantation induces the disease in WT mice. (A) the survival curve of WT mice (1100 rads) transplanted with WT (blue curve, n=21) or CFTR^ΔF508 (red curve, n=6) BM. (B) Typical H&E staining images of intestines from WT mice transplanted with CFTR^ΔF508 or WT BM. (C-D) Quantification of crypts (C, Mean±SD of n=9) and villi (D, Mean±SD of n=27) height of intestines from WT mice transplanted with CFTR^ΔF508 or WT BM. n=9. (E) Percentage of intestine crypts that contain mucus in it. WT mice are transplanted with WT or CFTR^ΔF508 BM. Mean±SD of n=27. n.s. p>0.05, *p<0.05, ***p<0.001 in the Gehan-Breslow-Wilcoxon test (A) and student t-test (C-E). Scale bars are 100 μm.

FIGURE 4.

Monocyte recruitment in mixed bone marrow chimeras. (A) Schematic showing the establishment of CFTR^ΔF508/WT mixed bone marrow chimeras. (B-C) CFTR^ΔF508 and WT monocytes (CD115⁺CD11b⁺CD11c⁻F4/80⁻, B) and macrophages (F4/80⁺CD11b⁺, C) in the lungs of unchallenged mixed chimeric mice as assessed by flow cytometry. (D-E) Mixed chimeric mice were challenged with CCL2 (intranasal 100 ng/mice, 12 hours before the harvest) and the BAL/lung monocyte ratio (D) or Ly6C^hi BAL/lung monocyte ratio (E) was calculated for CFTR^ΔF508 and WT monocytes. (F-G) The small intestinal lamina propria leukocytes (LPL) of mixed chimeric mice were harvested assessed for monocytes (CD11b⁺CD11c⁻F4/80⁻Ly6G⁻TCR⁻Cd19⁻NK1.1⁻, F) and macrophages (F4/80⁺, G) by flow cytometry. Mean±SD of n=10 mice for B-C, F-G and 14 mice for D-E. n.s. p>0.05, *p<0.05, **p<0.01 in the student T-test.

FIGURE 5.

Monocyte-specific CFTR deficiency retards weight gains and exacerbates DSS-induced colitis in mice. (A) Body weights of Cftr^flox/floxCsfr1-cre^+/− and Cftr^flox/floxCsfr1-cre^−/− littermates at indicated ages. Mean±SD of n=24 Cftr^flox/floxCsfr1-cre^+/− and 25 Cftr^flox/floxCsfr1-cre^−/− mice. (B) Body weights of Cftr^flox/floxCsfr1-cre^+/− and Cftr^flox/floxCsfr1-cre^−/− littermates at indicated days after DSS-induced colitis. Mean±SD of n=7 mice each. *p<0.05, **p<0.01, ****p<0.0001 in the student T-test. ####p<0.0001 in the 2-way ANOVA test.

Results

As expected, CF mice died spontaneously under specific-pathogen-free conditions. The first mice succumbed at 23 days of age, with half of all deaths occurring by 40 days and ~91% CF mice succumbing by 59 days (Fig. 1A). Like WT (CFTR^WT/WT) mice, heterozygous mice (CFTR^ΔF508/WT) survived for at least 110 days. At the same age (paired comparison of littermates), CF mice had significantly lower body weights (Fig. 1B, Supplemental Figure 1) and significantly elevated blood leukocyte counts (Fig. 1C). Monocytes, lymphocytes, and neutrophils were all elevated (Fig. 1D–F).

Adhesion defect under flow and phagocytosis defect of CF monocytes

Previous studies (23) showed that CF monocytes have an adhesion defect under static conditions. However, in vivo, monocytes must adhere to endothelial cells under flow conditions. To test monocyte adhesion under flow, we used a microfluidic device (36, 40). The best monocyte-specific marker is CD115 (Colony-stimulating factor 1 receptor, CSF1R), and the best marker for neutrophils is Ly6G (Fig. 1G). Using monoclonal antibodies to CD115 (red) and Ly6G (cyan), we visualized monocytes and neutrophils in flow chambers coated with P-selectin (to support rolling (36, 41, 42)) and ICAM-1 (to bind β2 integrins and support arrest (36, 42)) (Fig. 1H). In these experiments, arrest was triggered by infusing CCL2 (Chemokine C-C motif ligand 2, also knowns as monocyte chemoattractant protein-1, MCP-1) at a concentration of 100 ng·mL⁻¹. This is known to trigger rapid integrin activation and arrest of monocytes (43, 44). The same number of WT or CF BM cells (5×10⁷ mL⁻¹) were perfused in the flow chambers. Rolling and arrest were monitored by epifluorescence microscopy (Supplemental Movie 1). The total number of cells (monocytes plus neutrophils) per field-of-view was the same in flow chambers perfused with WT or CF BM cells. The number of arrested WT, but not CF monocytes, doubled in response to CCL2 (Fig. 1I, J). We also assessed expression of CCR2, the CCL2 receptor, on blood monocytes from WT and CF mice and found no significant difference (Fig. 1K). Thus, we conclude that CCL2-induced arrest under flow is completely abolished in CF monocytes.

Besides adhesion, complement-mediated phagocytosis of monocyte also depends on β2 integrin activation (45). Complement-mediated phagocytosis of C3b-coated beads and Pseudomonas aeruginosa has been reported to be deficient in blood monocytes from CF patients (46). We tested whether purified monocytes from CF mice have a similar defect in phagocytizing Staphylococcus aureus bioparticles (Fig. 1L). We found that PMA pre-stimulated monocytes from CF mice phagocytized significantly fewer iC3b-immobilized Staphylococcus aureus bioparticles compared to monocytes from WT mice on a per-cell basis. This phagocytosis defect in CF monocytes is Mac-1 (integrin α_Mβ₂, CD11b/CD18) dependent because CD11b or CD18 antibody blockade reduced the phagocytosis of WT monocytes, but not the phagocytosis of CF monocytes.

Transplanting WT BM relieves CF disease in mice

Histological assessment of the small intestine revealed the previously described increased crypt depth and villus height in CF mice (Supplemental Figure 2A). For both parameters, the differences between CF and WT mice were significant (Supplemental Figure 2B, C). CF mice also showed more mucus accumulation in the crypts of Lieberkühn than WT mice (Supplemental Figure 2D), resulting in a significantly larger percentage of crypts containing mucus (Supplemental Figure 2E).

Next, we asked whether reconstituting CFTR in hematopoietic stem cells (and therefore monocytes) was sufficient to improve the health of CF mice. To this end, we conducted BMT by injecting sterile WT or CF (control) BM cells into sub-lethally (700 rads) irradiated CF recipient mice. Reconstituting CF mice with WT BM resulted in 60–80% WT monocytes (Supplemental Table 1) and significantly prolonged their survival (Fig. 2A). The median survival time increased from 34 to 99 days, and 33% of the mice survived for up to 225 days. Interestingly, all mice that survived to 110 days (80 days after BMT) remained alive for the entire experiment. Pathologic evaluation of the bone marrow-transplanted mice showed that reconstituting CF mice with WT BM restored the crypt height defect (Fig. 2B, C), but not the villus height defect (Fig. 2D). This treatment also reduced the mucus percentage as assessed by histology (Fig. 2E, F).

Transplanting CF BM induces disease in WT mice

In a reverse approach, we tested whether the CFTR-dependent monocyte adhesion defect was sufficient to confer disease to WT mice. We reconstituted lethally (1100 rads) irradiated WT mice with CF or WT (control) BM (Fig. 3A). This is a model of complete bone marrow replacement, and almost all hematopoietic cells are donor-derived. The reconstitution efficiency of BM in mice upon the 1100 rads lethally irradiation was >90% (Supplemental Table 1). The epithelial cells are host-derived and thus retain intact, functional CFTR. All WT to WT bone marrow transplanted control mice survived for the entire duration of the experiment (232 days), but half of the mice that were reconstituted with CF BM succumbed by 41 days.

Reconstituting WT mice with CF BM induced the crypt height defect (Fig. 3B, C), but did not change villus height or mucus percentage (Fig. 3D, E). Thus, we conclude that CFTR^ΔF508 in hematopoietic cells (by BMT of CFTR^ΔF508 cells) might induce CF disease in WT mice. Taken together, these data show that the CFTR defect in bone marrow cells may be both necessary and sufficient to drive CF disease in mice. CFTR-dependent monocyte adhesion defect may contribute to CF pathology. Moreover, we observed an increased number of CCR2+ Ly6C^hi inflammatory monocytes in the bone marrow of CF mice compared to controls (Supplemental Figure 3). This pro-inflammatory phenotype of monocytes in CF mice may also contribute to CF pathology.

Recruitment defect of CF monocytes in vivo

Having shown the monocyte arrest defect under flow, we next asked whether this arrest defect translated into an in vivo monocyte recruitment defect of CF monocytes. To stringently investigate the recruitment of monocytes to the intestine and lung, we used mixed BM chimeras (36). In these experiments, lethally irradiated WT mice were reconstituted with both WT and CF BM (Fig. 4A) at an approximately 1:1 ratio. WT cells were CD45.1 and CF cells were CD45.2, two functionally equivalent alleles of CD45 that are distinguishable by flow cytometry. Both monocytes and macrophages were identified by flow cytometry. In unchallenged mice, there was no difference in the number of WT and CF monocytes in the lung (Fig. 4B), but the lungs contained significantly more WT than CF macrophages (Fig. 4C). This finding suggests that the monocyte-to-macrophage differentiation, a process known to require integrin signaling (47, 48), may be defective in CF monocytes. To directly test monocyte recruitment in vivo, we challenged mixed CF/WT bone marrow chimeric mice with intranasal CCL2. Significantly more WT than CF monocytes appeared in the BAL, demonstrating defective CF monocyte recruitment in vivo (Fig. 4D). This was true for all monocytes and specifically the Ly6C-hi subset (Fig. 4E). In the small intestinal lamina propria, the findings were similar: we found no difference between WT and CF monocytes in unchallenged mice (Fig. 4F), but more WT than CF macrophages in lamina propria leukocytes (LPL, Fig. 4G). Taken together, these findings indicate that the CF monocyte defect manifests in defective elicited monocyte recruitment and defective monocyte to macrophage differentiation.

Monocyte specific CFTR defect leads to severe gut inflammation

To investigate the monocyte-specific contribution in CF gut inflammation, we investigated CFTR^flox/flox Csfr1-cre^+/− mice. Of 27 CFTR^flox/flox Csfr1-cre^+/− mice, three died on days 21, 26, 30, respectively. Of 29 CFTR^flox/flox Csfr1-cre^−/− littermates, one died on day 31. We observed a slight but significant decrease in body weight gain of CFTR^flox/flox Csfr1-cre^+/− mice compared to CFTR^flox/flox Csfr1-cre^−/− littermates (Fig. 5A).

Next, we tested the contribution of monocyte-specific CFTR defect in a DSS-induced colitis model. We found that CFTR^flox/flox Csfr1-cre^+/− mice lose significantly more bodyweight after DSS administration compared to CFTR^flox/flox Csfr1-cre^−/− littermates (Fig. 5B). These results suggested that CFTR deficiency in monocytes contributes to the pathogenesis of gut inflammation.

Discussion

This study demonstrates that BMT rescues the CF phenotype in the CFTR^ΔF508 mouse model. Although this model does not exactly recapitulate human CF disease, this fundamental discovery suggests that BMT mitigated some aspects of CF disease by restoring monocyte/macrophage function and mucosal host defense. Survival and inflammation of CF mice were both improved by the hematological reconstitution using WT BM. We show that CF monocytes fail to adhere under flow, show reduced recruitment in response to CCL2, and contribute less to macrophages in the lung and intestinal lamina propria than WT monocytes. We show that the CFTR^ΔF508 in monocytes might be both necessary (BMT with WT bone marrow into CF mice improves the disease) and sufficient (BMT with CF bone marrow into WT mice induces the disease) for full manifestation of CF disease.

Besides the monocyte recruitment defect, CF monocytes may have other functional defects when challenged by inflammation and infections. Upon LPS stimulation, monocytes from CF patients have been reported to show reduced inflammatory cytokine expression (lower TNFα, lower IL-6, lower IL-12, lower IL-23) and increased production of anti-inflammatory IL-10 compared to WT controls (49). Taken together with our findings, this suggests complex defects encompassing monocyte arrest, recruitment, cytokine production, and differentiation into macrophages. The common denominator of these defects may be defective β₂ integrin activation in CF monocytes because β₂ integrins are known to be involved in many myeloid cell functions (50). Exactly why CF monocytes have a defect in differentiating to macrophages is beyond the scope of the present work and will be addressed in future studies.

Whether or not CF monocytes show defective phagocytosis remains controversial. Complement-mediated phagocytosis of C3b-coated beads and Pseudomonas aeruginosa has been reported to be deficient in blood monocytes from CF patients (46). In another study, the phagocytosis of Escherichia coli was increased in CF monocytes (49). These findings suggest that some but not all phagocytosis mechanisms are affected by dysfunctional CFTR. Antigen presentation to CD4 T cells by MHC-II is decreased in CF monocytes (49). A recent study showed that WT CFTR associates with PTEN, triggering the anti-inflammatory PI3K/Akt pathway in response to Toll-like receptor 4 (TLR4). This helps enhance Pseudomonas aeruginosa clearance (7), suggesting a mechanism that may fail in CF patients.

CFTR is expressed in the monocyte but not the neutrophil plasma membrane (46). Consequently, CFTR deficiency did not affect the adhesion of human neutrophils (23), which are the most abundant leukocytes in human blood and play essential roles in immune defense (36). However, CFTR mRNA is expressed in human neutrophils (51). CFTR protein is found expressed in the phagolysosome membrane of neutrophils, which affects bacterial killing, degranulation, and disease outcome (51–56). Thus, correcting CFTR by BMT may also restore the function of neutrophils and improve CF by promoting bacterial killing.

Our study is not the first CF study to use BMT, but it is the first to focus on monocyte defects. Previous investigators hypothesized that stem cells may exist among lung epithelial cells (57–59) or gut epithelial cells (59, 60). These studies provided evidence that hematopoietic stem cells and multipotent mesenchymal stromal cells can home to the lung or intestine. Whether they can differentiate into functional epithelial cells is less clear. A mild but significant rescue (~10%) of epithelial electrophysiology was shown in distal colons of CF mice, which have a decrease of forskolin-induced transepithelial current compared to WT controls (59, 60). However, the epithelial reconstitution was incomplete, even under optimized conditions (57). Optimized WT to CF BMT improved bacterial clearance and survival after infection with Pseudomonas aeruginosa (57). However, it is not clear whether the above clinical improvements may be due to an effect of BMT on putative epithelial stem cells. Although it has been shown that transferring nonhematopoietic BM cells can be differentiated to lung epithelial cells (61), there is no evidence showing that WT nonhematopoietic BM cells can rescue lung infections in CF mice. Importantly, at the time of these studies (47–50), the monocyte recruitment defect was not known. In light of the present findings, it seems more likely that the benefit observed in (57) was due to the functional rescue of myeloid cell function rather than epithelial cell function. A recent study supported our idea that, in a Pseudomonas aeruginos lung infection model, transferring CF BM to WT mice induced more death, and transferring WT BM to CF mice rescued the survival (62). Results in Pseudomonas aeruginosa-infected Cftr KO and KI mice showed that Cftr anti-inflammation/infection effect is myeloid-specific (62). These are consistent with our results.

Other studies have shown that CF macrophages exhibited pro-inflammatory cytokine expression, upregulated TLR4 signaling, and decreased anti-bacterial response after LPS stimulation compared to WT macrophages (63–65). In these studies, it is not clear whether the observed differences are a cause or a consequence of the ongoing CF disease process with infections. After transplantation of WT BM cells, the LPS-induced pro-inflammatory cytokine production in CF mice was reduced (63). However, monocyte recruitment was not studied. Myeloid cell recruitment to the lung is a multi-step process, where the myeloid cells first leave the blood compartment to appear in the lung interstitium and then must negotiate the epithelial layer to appear in the bronchoalveolar lavage (66). Our present findings suggest that the monocyte recruitment defect is localized to the transepithelial and not the transendothelial migration. The recruitment defect is not detected under homeostatic conditions but appears when mice are challenged with CCL2. CF patients likely are in a permanently challenged condition because of the bacterial load in the bronchoalveolar mucus.

Taken together, we show that restoration of WT monocytes by partial WT BMT is sufficient to improve health and survival in CF mice. Many CF patients are diagnosed at a young age, and BMT is more successful in children than in adults. Thus, partial BMT could be an appealing strategy for the management of CF disease (67). It appears that an antibody-based approach for BMT (without irradiation) may be safe and effective (68). Thus, this might be suitable to be tested for CF treatment. Unlike in malignant diseases, lethal irradiation is not required and partial restoration with 60–80% WT monocytes is sufficient. Moreover, it is now possible to correct gene defects in HSCs (34, 35) by gene therapy approaches. Thus, transfer of engineered autologous HSCs may become a practical treatment for patients with CF.

Key Points.

1. Monocytes from CFTR^ΔF508 mice have a defect in β2 integrin-dependent adhesion.

2. Transfering WT bone marrow rescues CF disease.

3. Monocyte-specific CFTR KO retards weight gains and exacerbates DSS-induced colitis

Abbreviations used in this article:

CF

cystic fibrosis

CFTR

cystic fibrosis transmembrane conductance regulator

BMT

bone marrow transplantation

WT

wild-type

LAD

leukocyte adhesion deficiencies

HSCs

hematopoietic stem cells

EDTA

ethylenediaminetetraacetic acid

LPL

lamina propria leukocytes

CCL2

Chemokine C-C motif ligand 2

MCP-1

monocyte chemoattractant protein-1

BAL

bronchoalveolar lavage fluid

DSS

dextran sulfate sodium

PMA

Phorbol 12-myristate 13-acetate