Bone Marrow Transplantation Rescues Monocyte Recruitment Defect and Improves Cystic Fibrosis in Mice

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 proinflammatory 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 sinopulmonary tissue. The resulting viscous mucus becomes infected with pathogens, such as Pseudomonas aeruginosa (6–9), Staphylococcus aureus (10, 11), Haemophilus influenzae (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), homologs 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 an ∼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 (LADs) I, II, and III (26), suggesting that CF is associated with a new LAD, that is, 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 Ags 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. In this study, 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 (BM) 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) BM. Mechanistically, we show that CF monocytes have a severe recruitment defect to the intestinal lamina propria (LP) and the bronchoalveolar space. Because 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.

Recombinant mouse P-selectin-Fc, ICAM-1-Fc, and CCL2 were purchased from R&D Systems. Casein blocking buffer, penicillin-streptomycin solution, pHrodo red S. aureus bioparticles, and dextran sulfate sodium (DSS) were purchased from Thermo Fisher Scientific. RPMI 1640 medium without phenol red, PBS without Ca²⁺ and Mg²⁺, and HBSS with phenol red without Ca²⁺ and Mg²⁺ were purchased from Life Technologies. FBS and human serum albumin were purchased from Gemini Bio-Products. PMA, type VIII collagenase, DNase, and N-acetyl-l-cysteine I were purchased from Sigma-Aldrich. PE or PE-Cy7–conjugated anti-CD115 Ab (clone AFS98), Alexa Fluor (AF)647, AF700, or Brilliant Violet (BV)650–conjugated anti-Ly6G Ab (clone 1A8), BV785-conjugated anti-CD11b Ab (clone M1/70), BV605-conjugated anti-CD11c Ab (clone N418), BV570-conjugated anti-Ly6C Ab (clone HK1.4), BV711-conjugated anti-TCRβ Ab (clone H57-597), allophycocyanin-conjugated anti-CD117 Ab (clone 2B8), allophycocyanin-Cy7–conjugated anti-CD19 Ab (clone 6D5), PE-conjugated anti-Ly6A/E Ab (clone D7), BV650-conjugated anti-NK1.1 Ab (clone PK136), PE-conjugated anti-F4/80 Ab (clone BM8), FITC-conjugated anti-CCR2 Ab (clone SA203G11), unconjugated anti-CD11b blocking Ab (clone M1/70), and unconjugated CD18 blocking Ab (clone M18/2) were purchased from BioLegend. Ghost Dye Violet 510 was purchased from Tonbo Biosciences. Fluorescein (FITC)-conjugated anti-CD45.1 Ab (clone A20) was purchased from BD Biosciences. PerCP complex-Cy5.5–conjugated anti-CD45.2 Ab (clone 104), PE-eFluor 610–conjugated anti-CD4 Ab (clone GK1.5), and eFluor 450–conjugated anti-CD8a Ab (clone 53-6.7) were purchased from eBioscience. RBC lysis buffer was purchased from Invitrogen. iC3b was purchased from Complement Technology. An EasySep mouse monocyte isolation kit was purchased from STEMCELL Technologies.

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.

WT or CF recipients (C57BL/6J background littermates, male or female, 1 mo old) were irradiated in two doses of 550 or 350 rad each (for a total of 1100 or 700 rad) 4 h apart (RS-2000 x-ray irradiator, Rad Source). BM cells from both femurs and tibias of donor mice (WT or CF mice, sex matched, 1 mo old) were collected under sterile conditions. Bones were centrifuged for the collection of marrow, and unfractionated BM cells were washed, resuspended in PBS, 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 BMT. After BMT, 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 8 wk after BM 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 WT C57BL/6J mice (8-wk-old males) were irradiated in two doses of 550 rad each (for a total of 1100 rad) 4 h apart (RS-2000 x-ray irradiator, Rad Source). BM cells from both femurs and tibias of donor mice (CF male and CD45.1 WT male, 11 wk old) were collected under sterile conditions. Bones were centrifuged for the collection of marrow, and unfractionated BM 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 BMT. After BMT, mice were provided autoclaved acidified water with antibiotics (trimethoprim/sulfamethoxazole) and were fed autoclaved food. Mice were used for further experiments 8 wk after BM reconstitution.

The body weight of aged CF and WT littermates (40–72 wk) and young CFTR^flox/floxCsfr1-cre^+/− and CFTR^flox/floxCsfr1-cre^−/− littermates (age from 3 to 6 wk) were recorded. Blood counts of WT or CF littermates (age from 10 to 25 wk) were taken via retro-orbital bleeding and analyzed by an automatic analyzer (Hemavet 950FS, DREW Scientific). Mice were anesthetized by inhalation of an isoflurane/oxygen gas mixture during the bleeding.

Blood was obtained by cardiac puncture with an EDTA-coated syringe. RBCs were lysed in RBC lysis buffer according to the manufacturer’s protocol. Lungs were lavaged with PBS containing 2 mM EDTA, dissociated by a gentleMACS dissociator (Miltenyi Biotec), 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, was collected. For the preparation of LP cells, fat and connective tissue were removed from the intestines. Intestines were opened longitudinally, cut into 1-cm pieces, and washed three times for 10 min 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 min at 37°C. Digested material was dissociated by a gentleMACS dissociator (Miltenyi) and filtered through a 70-μm strainer. BM cells were collected from both femurs and tibias. In some experiments, mice were challenged intranasally with mouse CCL2 (100 ng/mouse) 12 h prior to the harvest. Blood was collected via retro-orbital bleeding. Bronchoalveolar lavage fluid (BALF) 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 of 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 Ags on cells were stained for 30 min at 4°C with directly conjugated fluorescent Abs (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-eFluor 610, anti–CD8a-eFluor 450, anti–CD19-allophycocyanin-Cy7, anti–CD117-allophycocyanin, anti–Ly6A/E-PE, and anti–NK1.1-BV650 for blood and BM cells; and 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-eFluor 610, anti–CD8a-eFluor 450, anti–CD19-allophycocyanin-Cy7, anti–F4/80-PE, and anti–NK1.1-BV650 for BALF, 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 LSR II (BD Biosciences) and was analyzed with FlowJo (BD Biosciences, 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 experiments, blood was obtained by retro-orbital bleeding. RBCs were lysed in RBC lysis buffer according to the manufacturer’s protocol. Fcγ receptors were blocked for 15 min, and surface Ags on cells were stained for 30 min at 4°C with directly conjugated fluorescent Abs (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 LSR II (BD Biosciences) and was analyzed with FlowJo (BD Biosciences, version 10.4). Monocytes were gated as CD115⁺Ly6G⁻ cells, and the mean fluorescence intensities (MFI) of FITC were used to quantify CCR2 expression.

In the phagocytosis assay, pHrodo red S. aureus bioparticles (1.33 mg/ml) were immobilized by iC3b (0.33 mg/ml) at 4°C overnight and washed twice with PBS before use. BM cells were collected from both femurs and tibias of WT and CF mice. Monocytes were purified using an 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 Abs, or isotype controls at room temperature (RT), spun at 300 rpm (plate shaker) for 15 min, and incubated with iC3b-immobilized pHrodo red S. aureus bioparticles (66.6 µg/ml) at 37°C, 100 rpm (shaking incubator) for 1 h. After two washes with 1% paraformaldehyde and one wash with PBS with 2 mM EDTA, cell fluorescence was assessed with an LSR II (BD Biosciences) and was analyzed with FlowJo (BD Biosciences, version 10.4). The MFI of pHrodo red S. 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.

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 H&E and periodic acid–Schiff for histological assessment by a single investigator who was blinded to the experimental design. Slides were digitized on an AxioScan Z1 slide scanner using a ×40/0.95 numerical aperture 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).

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) and ICAM-1-Fc (10 μg/ml) for 2 h and then blocked for 1 h 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 wide. 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) from WT or CF mice were incubated with PE-conjugated anti-CD115 Ab and AF647-conjugated anti-Ly6G Ab for 10 min 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 mouse CCL2 was perfused. The processes of rolling and arrest were recorded by epifluorescence microscopy using an Olympus IX71 inverted microscope equipped with a ×40/0.95 numerical aperture objective.

Colitis was induced by adding 2% DSS in the drinking water for 5 d, followed by regular water. Mice (males, 11 wk old) were monitored and weighed every day for 2 wk and then sacrificed. Body weight data were normalized by the initial body weight to determine the weight lost.

Statistical analysis was performed with Prism 6 (GraphPad Software). Data are presented as survival curve (Figs. 1A, 2A, 3A), mean ± SD plus individual data points (Figs. 1B–F, 1I–L, 2C, 2D, 2F, 3C–E, 4B–G, 5, Supplemental Figs. 2B, 2C, 2E, 3), and individual data points with linear regressions (Supplemental Fig. 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. A p value <0.05 was considered significant.

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

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 (CSF1 receptor), and the best marker for neutrophils is Ly6G (Fig. 1G). Using mAbs to CD115 (red) and Ly6G (cyan), we visualized monocytes and neutrophils in flow chambers coated with P-selectin [to support rolling (36, 40, 41)] and ICAM-1 [to bind β₂ integrins and support arrest (36, 40)] (Fig. 1H). In these experiments, arrest was triggered by infusing CCL2 (also known as MCP-1) at a concentration of 100 ng/ml. This is known to trigger rapid integrin activation and arrest of monocytes (42, 43). The same number of WT or CF BM cells (5 × 10⁷ ml) were perfused in the flow chambers. Rolling and arrest were monitored by epifluorescence microscopy (Supplemental Video 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, 1J). 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 monocytes also depends on β₂ integrin activation (44). Complement-mediated phagocytosis of C3b-coated beads and P. aeruginosa has been reported to be deficient in blood monocytes from CF patients (45). We tested whether purified monocytes from CF mice have a similar defect in phagocytizing S. aureus bioparticles (Fig. 1L). We found that PMA-prestimulated monocytes from CF mice phagocytized significantly fewer iC3b-immobilized S. aureus bioparticles compared with 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 Ab blockade reduced the phagocytosis of WT monocytes, but not the phagocytosis of CF monocytes.

Histological assessment of the small intestine revealed the previously described increased crypt depth and villus height in CF mice (Supplemental Fig. 2A). For both parameters, the differences between CF and WT mice were significant (Supplemental Fig. 2B, 2C). CF mice also showed more mucus accumulation in the crypts of Lieberkühn than WT mice (Supplemental Fig. 2D), resulting in a significantly larger percentage of crypts containing mucus (Supplemental Fig. 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 sublethally (700 rad) irradiated CF recipient mice. Reconstituting CF mice with WT BM resulted in 60–80% WT monocytes (Supplemental Table I) and significantly prolonged their survival (Fig. 2A). The median survival time increased from 34 to 99 d, and 33% of the mice survived for up to 225 d. Interestingly, all mice that survived to 110 d (80 d after BMT) remained alive for the entire experiment. Pathologic evaluation of the BM-transplanted mice showed that reconstituting CF mice with WT BM restored the crypt height defect (Fig. 2B, 2C), but not the villus height defect (Fig. 2D). This treatment also reduced the mucus percentage as assessed by histology (Fig. 2E, 2F).

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 rad) irradiated WT mice with CF or WT (control) BM (Fig. 3A). This is a model of complete BM replacement, and almost all hematopoietic cells are donor derived. The reconstitution efficiency of BM in mice upon the 1100 rad lethal irradiation was >90% (Supplemental Table I). The epithelial cells are host derived and thus retain intact, functional CFTR. All WT-to-WT BM transplanted control mice survived for the entire duration of the experiment (232 d), but half of the mice that were reconstituted with CF BM succumbed by 41 d.

Reconstituting WT mice with CF BM induced the crypt height defect (Fig. 3B, 3C), but it did not change villus height or mucus percentage (Fig. 3D, 3E). 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 BM cells may be both necessary and sufficient to drive CF disease in mice. The CFTR-dependent monocyte adhesion defect may contribute to CF pathology. Moreover, we observed more CCR2⁺Ly6C^hi inflammatory monocytes in the BM of CF mice compared with controls (Supplemental Fig. 3). This proinflammatory phenotype of monocytes in CF mice may also contribute to CF pathology.

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 ∼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 (46, 47), may be defective in CF monocytes. To directly test monocyte recruitment in vivo, we challenged mixed CF/WT BM chimeric mice with intranasal CCL2. Significantly more WT than CF monocytes appeared in the BALF, 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 LP, 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 LP leukocytes (Fig. 4G). Taken together, these findings indicate that the CF monocyte defect manifests in defective elicited monocyte recruitment and defective monocyte-to-macrophage differentiation.

To investigate the monocyte-specific contribution in CF gut inflammation, we investigated CFTR^flox/floxCsfr1-cre^+/− mice. Of 27 CFTR^flox/floxCsfr1-cre^+/− mice, three died on days 21, 26, and 30, respectively. Of 29 CFTR^flox/floxCsfr1-cre^−/− littermates, one died on day 31. We observed a slight but significant decrease in body weight gain of CFTR^flox/floxCsfr1-cre^+/− mice compared with CFTR^flox/floxCsfr1-cre^−/− littermates (Fig. 5A).

Next, we tested the contribution of the monocyte-specific CFTR defect in a DSS-induced colitis model. We found that CFTR^flox/floxCsfr1-cre^+/− mice lose significantly more body weight after DSS administration compared with CFTR^flox/floxCsfr1-cre^−/− littermates (Fig. 5B). These results suggested that CFTR deficiency in monocytes contributes to the pathogenesis of gut inflammation.

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 LP than do WT monocytes. We show that the CFTR^ΔF508 in monocytes might be both necessary (BMT with WT BM into CF mice improves the disease) and sufficient (BMT with CF BM 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 TNF-α, lower IL-6, lower IL-12, lower IL-23) and increased production of anti-inflammatory IL-10 compared with WT controls (48). 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 (49). 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 CF monocytes show defective phagocytosis remains controversial. Complement-mediated phagocytosis of C3b-coated beads and P. aeruginosa has been reported to be deficient in blood monocytes from CF patients (45). In another study, the phagocytosis of Escherichia coli was increased in CF monocytes (48). These findings suggest that some but not all phagocytosis mechanisms are affected by dysfunctional CFTR. Ag presentation to CD4 T cells by MHC class II is decreased in CF monocytes (48). A recent study showed that WT CFTR associates with PTEN, triggering the anti-inflammatory PI3K/Akt pathway in response to TLR4. This helps enhance P. aeruginosa clearance (7), suggesting a mechanism that may fail in CF patients.

CFTR is expressed in the monocyte but not the neutrophil plasma membrane (45). 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 (50). CFTR protein is found expressed in the phagolysosome membrane of neutrophils, which affects bacterial killing, degranulation, and disease outcome (50–55). 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, to our knowledge, it is the first to focus on monocyte defects. Previous investigators hypothesized that stem cells may exist among lung epithelial cells (56–58) or gut epithelial cells (58, 59). 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 with WT controls (58, 59). However, the epithelial reconstitution was incomplete, even under optimized conditions (56). Optimized WT-to-CF BMT improved bacterial clearance and survival postinfection with P. aeruginosa (56). 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 (60), there is no evidence showing that WT nonhematopoietic BM cells can rescue lung infections in CF mice. Importantly, at the time of these studies (45–49), the monocyte recruitment defect was not known. In light of the present findings, it seems more likely that the benefit observed in Duchesneau et al. (56) was due to the functional rescue of myeloid cell function rather than epithelial cell function. A recent study supported our idea that, in a P. aeruginosa lung infection model, transferring CF BM to WT mice induced more death, and transferring WT BM to CF mice rescued the survival (61). Results in P. aeruginosa–infected Cftr knockout and knockin mice showed that the Cftr anti-inflammation/infection effect is myeloid specific (61). These findings are consistent with our results.

Other studies have shown that CF macrophages exhibited proinflammatory cytokine expression, upregulated TLR4 signaling, and decreased the anti-bacterial response after LPS stimulation compared with WT macrophages (62–64). 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 proinflammatory cytokine production in CF mice was reduced (62). However, monocyte recruitment was not studied. Myeloid cell recruitment to the lung is a multistep 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 (65). 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 (66). It appears that an Ab-based approach for BMT (without irradiation) may be safe and effective (67). 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.

We thank A. Denn from the Microscopy and Histology Core Facility at the La Jolla Institute for Immunology for help in obtaining the scientific data presented in this paper. We thank Dr. P. Quinton from the Department of Pediatrics in the School of Medicine at the University of California San Diego for help in editing the manuscript. We thank Dr. J. Rivera-Nieves from the Inflammatory Bowel Disease Center in the Division of Gastroenterology at the University of California San Diego for advice on histology.

The authors have no financial conflicts of interest.