β₁ Integrin regulates adult lung alveolar epithelial cell inflammation

Integrins are heterodimeric transmembrane receptors consisting of α and β subunits that bind extracellular matrix (ECM) components; propagate bidirectional signaling (1–5); and regulate critical processes such as adhesion, migration and proliferation that are required for the development of multicellular organisms (6–10). Of the 24 known α-β integrin heterodimers, 12 integrins contain the β₁ subunit. β₁ Integrins are present in epithelial cells, where they mediate cell adhesion to basement membranes and facilitate epithelial tissue organogenesis (9, 11–24). Compared with organ development, the function of integrins in maintaining tissue homeostasis is poorly defined.

Alveoli are complex structures composed of epithelial cells attached to a basement membrane juxtaposed to capillaries and stromal fibroblasts. Epithelial cells are either cuboidal type 2 alveolar epithelial cells (AECs) expressing high levels of surfactant protein C (SP-C) or very thin type 1 AECs in close apposition to capillaries. We previously reported that β₁ integrin regulates branching morphogenesis and alveolarization during lung development (22). Moreover, we showed that genetically deleting β₁ integrin in the developing alveolus results in dilated airspaces, thickened alveolar septa, type 2 AEC hyperplasia, and increased numbers of alveolar macrophages. Macrophage depletion rescued the alveolarization defect in these mice (22). These findings suggest that epithelial β₁ integrin dysfunction has deleterious consequences in lung epithelium through regulation of innate immunity. The mechanisms whereby these epithelial-macrophage interactions occur are uncertain, and, perhaps more importantly, the function of β₁ integrin in the adult lung is not established.

In this study, we deleted β₁ integrin in type 2 AECs after completion of lung development, which occurs by P28. At 2 years of age, the mice developed emphysematous changes in the lung parenchyma, as well as lymphoid aggregates and increased macrophage accumulation, which are characteristic of patients with advanced chronic obstructive pulmonary disease (COPD). This condition was preceded by proliferation of inflamed AECs that exhibited abnormal cell-cell junctions and excessive inflammation. Reduction of monocytes and monocyte-derived macrophages caused rapid onset of emphysema in young mice, suggesting that these cells limit inflammation and injury by clearance of deranged type 2 AECs. Thus, we conclude that under physiological conditions, β₁ integrin plays a critical homeostatic role in lung epithelial cells by suppressing inflammatory signaling.

Conditional β₁ integrin deletion in type 2 AECs results in emphysema and increased inflammation in aged mice. To test the importance of β₁ integrin deletion after development, we crossed integrin β₁^fl/fl (β1^f/f) mice with a doxycycline-inducible (dox-inducible) Cre recombinase under control of the surfactant protein-C (SP-C) promoter (designated as β1^rtTA mice). β₁ Integrin deletion was induced in type 2 AECs by addition of dox to drinking water from day P28, at the completion of lung development, until 2 months of age. Mice were sacrificed at 3 and 24 months of age. Lungs of 24-month-old β1^rtTA mice exhibited emphysema and increased numbers of macrophages (Figure 1, A and B). Lung morphometry quantification by mean linear intercept demonstrated a 60% airspace enlargement in β1^rtTA lungs compared with both β1^f/f and β1^rtTA mice that did not receive dox (Figure 1, C and D). Multiple lobes were sampled to minimize bias introduced by regional differences in alveolar size (25–27). There was evidence of bronchus-associated lymphoid tissue (BALT), which is characteristic of advanced COPD (arrows in Figure 1A, quantified in Figure 1E; and ref. 28). In addition to BALT lesions, histological examination revealed increased macrophages in β1^rtTA lungs, identified by the pan-macrophage marker CD68 (Figure 1F), and an increase in bronchoalveolar lavage fluid (BALF) cell count (1.8 × 10⁵ ± 0.2 × 10⁵ cells/mL from β1^rtTA lungs compared with 0.8 × 10⁵ ± 0.1 × 10⁵ cells/ml from β1^f/f lungs; Supplemental Figure 1A; supplemental material available online with this article; https://doi.org/10.1172/jci.insight.129259DS1).

Figure 1

Deletion of β₁ integrin in type 2 AECs results in emphysema and increased inflammation in aged mice. (A and B) H&E-stained paraffin lung sections show marked airspace enlargement and increased number of BALT lesions in 24-month-old β1^rtTA mice compared with age-matched β1^f/f mice and 24-month-old dox-naive β1^rtTA mice. Arrows indicate BALT lesions. (C) High-power images: Airspace enlargement is visible in β1^rtTA lungs; quantification in D shows increased mean linear intercept in β1^rtTA lungs (10 sections/mouse; n = 11 β1^f/f, n = 9 β1^rtTA + dox mice, n = 5 β1^rtTA – no dox mice). (E) Increased BALT lesions per lung section in β1^rtTA mice. n = 19 β1^f/f, n = 15 β1^rtTA + dox mice; n = 5 β1^rtTA – no dox mice; P = 0.0649 by 1-way ANOVA. (F) β1^rtTA lungs contain increased numbers of CD68⁺ macrophages. Scale bars: 200 μm in A and B, 50 μm in C and F. *P < 0.05 by 1-way ANOVA with Tukey’s test for multiple comparison.

Epithelial dysfunction precedes major morphological changes in β1^rtTA mice. To determine the timing of the structural deficits in β1^rtTA lungs relative to gene deletion, we performed histological examination of 3-month-old mice. We verified the efficiency of β₁ integrin deletion in the lungs of β1^rtTA mice by immunohistochemistry and found it was removed in more than 90% of type 2 AECs (Figure 2, A and B). This finding was confirmed by immunoblotting of primary type 2 AEC lysates from β1^rtTA and β1^f/f mice (Figure 2C). Microscopic examination showed no difference in airspace size in 3-month-old β1^rtTA mice (Figure 3, A and B). By crossing β1^rtTA mice to mice expressing the mTmG reporter (allowing visualization of GFP⁺ progeny derived from cells that had undergone Cre activation), we observed that β1^rtTA; mTmG mice exhibited GFP⁺ type 1 AECs immediately adjacent to β₁-deficient type 2 AECs, suggesting β₁ integrin is not required for type 2–to–type 1 AEC differentiation during homeostasis in the adult lung (Supplemental Figure 1B). β1^rtTA mice did exhibit mild intraseptal edema (arrows in Figure 3C), increased BALF protein (Supplemental Figure 2A), and increased BALF macrophages (Supplemental Figure 2B). Transmission electron microscopy (TEM) revealed intact cell-matrix interactions (arrows in Figure 3D) and defects in tight junctions between type 1 and type 2 AECs. Rather than the normal dark stranded seal demarcating tight junctions at the apical cell-cell junction, β1^rtTA lungs had a deep cleft (Figure 3, D and E, with tight junctions marked by asterisks in E). Consistent with these tight junction abnormalities, β1^rtTA mice had decreased claudin-3 protein levels in primary type 2 AEC lysates (Figure 3F) and decreased mRNA expression of Claudin-4 but not Claudin-18 as measured by quantitative RT-PCR (qPCR) of type 2 AECs (Figure 3G).

Figure 2

β₁ Integrin is deleted in type 2 AECs in β1^rtTA lungs. (A) Immunostaining for pro–SP-C (green) and β₁ integrin (red) demonstrates type 2 AEC–specific deletion of β₁ integrin in 3-month-old β1^rtTA lungs. Arrows indicate the presence/absence of β₁ integrin expression. Scale bar: 5 μm. (B) Type 2 AEC–specific deletion is represented as percentage of pro–SP-C⁺ cells that express β₁ integrin. 100–120 type 2 AECs counted/mouse; n = 3 β1^f/f, n = 4 β1^rtTA mice. (C) Representative Western blot for β₁ integrin on primary type 2 AEC lysate, normalized to GAPDH; representative of 3 separate experiments. *P < 0.05 by 2-tailed Student’s t test.

Figure 3

In the absence of aging, deletion of β₁ integrin in type 2 AECs minimally alters gross alveolar structure but results in epithelial dysfunction. (A and B) H&E-stained paraffin lung sections from 3-month-old β1^f/f and β1^rtTA mice demonstrate equal airspace size. (C) H&E-stained paraffin lung sections show increased intraseptal edema (arrows) in β1^rtTA lungs. (D and insets in E) Transmission electron microscopic images of β1^f/f and β1^rtTA lungs show intact cell-matrix interactions (arrows in D), but clefts at the cell-cell junctions in β1^rtTA lungs (junctions marked by asterisks in E). (F) Representative Western blot for claudin-3 on primary type 2 AEC lysate, with densitometry. n = 6 mice/group, normalized to GAPDH. (G) Gene expression for Claudin-4 and Claudin-18 by qPCR. n = 6 mice/group, normalized to GAPDH. RQ, relative quantitation. Scale bars: 200 μm in A, 25 μm in B, 50 μm in C, 500 nm in D, 250 nm in E. *P < 0.05 by 2-tailed Student’s t test. Images in A–C are representative of 6 mice/group.

We next assessed whether there were abnormalities of type 2 AEC-ECM interactions by visualizing their adherence to the laminin-containing basement membrane. While the basal surface of type 2 AECs appeared to adhere normally to the basement membrane (Figure 4A), we noticed that there were more type 2 AECs in β1^rtTA than β1^f/f mice (Figure 4, B and C). The excess of type 2 AECs, evidenced by pro–SP-C–positive staining, was due to increased cellular proliferation that was identified by Ki-67 immunostaining (Figure 4, D and E). In contrast, no differences in the number of apoptotic type 2 AECs between β1^rtTA and β1^f/f lungs were observed, as demonstrated by dual TUNEL⁺pro–SP-C⁺ cells (Figure 4, F and G). Thus, deletion of β₁ integrin in AECs from 3-month-old adult mice caused subtle structural defects with abnormal tight junctions that likely allowed for paracellular fluid flux leak and type 2 AEC proliferation. Proliferation of type 2 AECs is a well-known feature of inflammatory lung diseases and a recognized consequence of lung injury (29–35); therefore, this finding suggests an ongoing injury-repair cycle.

Figure 4

Loss of type 2 AEC β₁ integrin results in epithelial proliferation. (A) ABCA3⁺ type 2 AECs (green) remain adherent to the basement membrane (laminin in red) in β1^rtTA lungs. (B) Lung sections immunostained for type 2 AEC marker pro–SP-C (red) and laminin γ1 (green) show increased numbers of type 2 AECs in β1^rtTA lungs, as quantified in C. 20 sections/mouse; n = 6 mice/group. (D) Lung sections immunostained for pro–SP-C (red) and the proliferation marker Ki-67 (green) show increased type 2 AEC proliferation in β1^rtTA lungs, as quantified in E. 20 sections/mouse; n = 6 mice/group. (F) No difference in the number of apoptotic type 2 AECs in β1^rtTA and β1^f/f lungs, as identified by TUNEL and pro–SP-C costained type 2 AECs, as quantified in G. n = 6 β1^f/f, n = 5 β1^rtTA mice. Scale bars: 5 μm in A; 50 m in B, D (10 μm for insets), and F. *P < 0.05 by 2-tailed Student’s t test.

β₁ Integrin–deficient type 2 AECs induce increased efferocytosis. We next performed in-depth analysis of the inflammatory status of β1 integrin–deficient mice. When we examined aged β1^rtTA mice, we noted that pro–SP-C staining often colocalized with CD68, suggesting that macrophages phagocytosed AECs in β1^rtTA but not β1^f/f lungs (Figure 5A). This observation is consistent with efferocytosis, a tightly regulated process by which phagocytic cells ingest diseased or dying cells, thereby minimizing inflammation in the microenvironment (36–39). To define the mechanisms whereby this occurred, we cultured primary type 2 AECs and measured secretion of CX3CL1, a “find me” chemokine that attracts phagocytes (40, 41). We also assessed the expression levels of Cd47, whose gene product is an inhibitory “don’t eat me” signal, in freshly isolated primary type 2 AECs (41, 42). We found increased CX3CL1 production and reduced Cd47 mRNA expression by β1^rtTA type 2 AECs relative to type 2 AECs isolated from β1^f/f mice (Figure 5, B and C). These findings support the conclusion that macrophage efferocytosis of β₁ integrin–deficient type 2 AECs is prominent in β1^rtTA lungs.

Figure 5

β1^rtTA mice have increased efferocytosis of type 2 AECs. (A) Aged 24-month-old β1^rtTA lungs exhibit increased colocalization of pro–SP-C (red) and CD68 (green), indicative of increased efferocytosis of type 2 AECs by macrophages. Scale bar: 10 μm. Representative of 3 mice/group. (B) Increased CX3CL1 by ELISA in media collected from primary β1^rtTA type 2 AECs after 24 hours in culture. n = 10 mice/group. (C) Decreased gene expression of CD47 in freshly isolated primary type 2 AECs isolated from β1^rtTA mice. n = 3 mice/group. *P < 0.05 by 2-tailed Student’s t test.

Deleting CCL2-recruited monocytes/macrophages causes severe destruction of alveolar architecture in β1^rtTA mice by decreasing AEC efferocytosis. Our histological examination of both aged and 3-month-old β1^rtTA lungs suggested increased inflammation. Since β1^rtTA lungs had increased macrophages, and β₁-null type 2 AECs exhibited markers of efferocytosis, we tested whether impairment of macrophage recruitment would disrupt homeostasis in 3-month-old β1^rtTA mice. To target these recruited immune cell populations, we crossed β1^f/f and β1^rtTA mice to the Ccr2-null background. CCR2 is the receptor for CCL2, one of the primary monocyte chemokines in the lung. CCR2^–/–;β1^rtTA mice and their CCR2^–/–;β1^f/f littermate controls received dox from P28 until 2 months of age in the same manner as β1^f/f and β1^rtTA mice. In contrast to β1^rtTA mice, 3-month-old CCR2^–/–;β1^rtTA mice exhibited dramatically enhanced lung pathology (Figure 6, A and B), with widespread emphysematous destruction; marked airspace enlargement, quantified by mean linear intercept (Figure 6C); increased inflammatory infiltrates (arrows in Figure 6B); and increased BALF cell counts (Figure 6D). CCR2^–/–;β1^rtTA mice exhibited a large number of CD68⁺ macrophages (Figure 6, E and F) despite loss of CCL2 recruitment due to excessive proliferation of existing resident macrophages as verified by increased Ki-67 staining (Supplemental Figure 3, A–C). A large increase in pro–SP-C⁺ type 2 AECs accompanied the expanded immune cell population (Figure 6E, quantified in Figure 6G). Immunostaining for Ki-67 demonstrated that depletion of CCL2-driven monocytes/macrophages did not change the proliferation rate of AECs compared with β1^rtTA mice (Supplemental Figure 3, D and E). These findings indicate that the increase in epithelial cell numbers in β1^rtTA mice was due to impaired AEC removal rather than increased AEC proliferation. Despite numerous macrophages and an overabundance of type 2 AECs, there was almost no colocalization of CD68 and pro–SP-C in CCR2^–/–;β1^rtTA mice (Figure 6H, quantified in Figure 6I), suggesting minimal efferocytosis in these mice. To directly test whether macrophages from CCR2^–/–;β1^rtTA mice were defective in efferocytosis, we collected macrophages from bronchoalveolar lavage and exposed these cells to fluorescently labeled primary type 2 AECs from β1^rtTA mice (Figure 6J). While macrophages from β1^rtTA and β1^f/f lungs briskly engulfed β₁-deficient AECs, macrophages from CCR2-deficient mice (both CCR2^–/–;β1^rtTA and CCR2^–/–;β1^f/f) ingested far fewer labeled AECs, demonstrating that CCR2-deficient macrophages were less efficient efferocytosis agents. These data strongly suggest that the more severe phenotype in the CCR2^–/–;β1^f/f mice is caused by their inability to remove deranged type 2 AECS and that the efferocytosis function of CCL2-recruited macrophages is to limit inflammation and mitigate lung damage in β1^rtTA mice.

Figure 6

Recruited monocytes/macrophages maintain structural homeostasis in β1^rtTA mice through efferocytosis. (A and B) CCR2^–/–;β1^rtTA lungs show severe remodeling and increased inflammatory infiltrate (arrows) in low-power (A) and high-power (B) images of H&E-stained sections. (C) Increased mean linear intercept in 3-month-old CCR2^–/–;β1^rtTA lungs. 6–10 sections/mouse; n = 6 β1^f/f, n = 6 CCR2^–/–;β1^f/f, n = 6 β1^rtTA, n = 7 CCR2^–/–;β1^rtTA mice/group). (D) Increased BALF cell counts in CCR2^–/–;β1^rtTA mice. n = 17 β1^f/f; n = 6 CCR2^–/–;β1^f/f, n = 21 β1^rtTA, n = 6 CCR2^–/–;β1^rtTA mice/group). (E) Immunostaining for CD68 (green) and pro–SP-C (red) demonstrates increased macrophages and type 2 AECs in CCR2^–/–;β1^rtTA lungs. (F and G) Quantification for numbers of CD68⁺ (F) and pro–SP-C⁺ cells/field (G). 20 sections/mouse; n = 4–6 mice/group. (H and I) Immunostaining demonstrates minimal colocalization of CD68⁺ and pro–SP-C⁺ cells in CCR2^–/–;β1^f/f and CCR2^–/–; β1^rtTA lungs, whereas abundant colocalization was present in β1^rtTA lungs. 20 sections/mouse; n = 4–6 mice/group. (J) Quantification of alveolar macrophage (Alv mac) efferocytosis of fluorescently labeled primary type 2 AECs. n = 8–12 mice/group. Scale bar: 200 μm in A, 50 μm in B and E, 5 μm in H. *P < 0.05 by ordinary 1-way ANOVA with secondary analysis by Tukey’s test for multiple comparisons as indicated.

CD11b⁺CD11c^– monocytes/macrophages efferocytose type 2 AECs in β1^rtTA mice. We next examined the immune cell population in the whole lung by flow cytometry. β1^rtTA lungs contained increased CD45⁺CD11b⁺CD11c^– immune cells, markers consistent with recently recruited monocyte–early macrophages (Figure 7A, gating strategy in Supplemental Figure 4; and refs. 43, 44). We identified this as a mixed population, as cells expressed the monocyte marker Ly6C, the macrophage marker CD64, or both (Figure 7B). Since the CD11b⁺CD11c^– immune cells were differentially enriched in β1^rtTA mice, we collected this population by FACS, cytospinned the cells, and immunostained for pro–SP-C and CD68 to determine whether these cells contributed to the increased efferocytosis seen in β1^rtTA mice. We found that 68% ± 4% of monocytes/macrophages collected from β1^rtTA lungs contained pro–SP-C⁺ material compared with 14% ± 2% of these cells from β1^f/f lungs (Figure 7, C and D). To functionally phenotype these cells in β1^rtTA mice, we collected media from cultured monocytes/macrophages and assayed for cytokine production by cytokine multiplex. The β1^rtTA monocyte-macrophage population secreted only scant amounts of inflammatory cytokines/chemokines, equivalent to expression levels by cells from β1^f/f mice (Supplemental Table 1). Taken together, these data demonstrate that CD11b⁺CD11c^– monocytes/macrophages are critical effector cells for efferocytosis but do not directly contribute to the inflammatory state of β1^rtTA mice.

Figure 7

CD11b⁺CD11c^– monocytes/macrophages efferocytose type 2 AECs in β1^rtTA mice. (A) β1^rtTA lungs contain increased CD45⁺CD11b⁺CD11c^– monocytes/macrophages by flow cytometry. n = 7 β1^f/f, n = 4 CCR2^–/–;β1^f/f, n = 6 β1^rtTA, and n = 7 CCR2^–/–;β1^rtTA mice. (B) CD45⁺CD11b⁺CD11c^– population consists of cells that are both Ly6C⁺ and CD64⁺ in β1^rtTA lungs. n = 6 β1^rtTA mice. (C) Immunostained cytospins from CD45⁺CD11b⁺CD11c^– monocytes/macrophages show increased numbers of CD68⁺pro–SP-C⁺ cells in β1^rtTA lungs, as quantified in D. n = 6 mice/group. Scale bar: 40 μm in C, 10 μm for inset. *P < 0.05 by 2-tailed Student’s t test.

β1 Integrin regulates AEC inflammation. Our data thus far suggest that the β1 integrin–null cells provide an inflammatory stimulus resulting in monocyte-macrophage chemoattraction into the alveolus. These recruited cells function as efferocytotic agents but do not contribute to the inflammatory status of the lungs. Next, we tested whether β₁-deficient AECs drive the inflammatory phenotype in lungs of β1^rtTA mice. Ten of 32 cytokines (31%), including mediators of macrophage chemotaxis and maturation, were significantly increased in the culture media of β1^rtTA AECs compared with that of β1^f/f AECs (Figure 8A and Supplemental Table 2). To define the consequence of increased AEC inflammatory signaling in the whole lung, we performed multiplex analysis on tissue lysates (Figure 8B and Supplemental Table 3). Multiple inflammatory mediators were increased in lungs of β1^rtTA mice compared with β1^f/f controls. Even further increases were seen in CCR2^–/–;β1^rtTA, where inflamed β₁-deficient type 2 AECs remained unchecked by efferocytosis. Since many of the cytokines increased in β1^rtTA and CCR2^–/–;β1^rtTA lungs were recognizable gene products of NF-κB signaling (including KC, IL-6, MIP-2, and G-CSF), we performed immunohistochemistry for activated NF-κB in β₁ integrin–deficient AECs (Figure 8, C and D). Immunofluorescence staining for phospho-p65 (S276), a well-recognized marker of NF-κB activation (45), revealed numerous phospho-p65⁺pro–SP-C⁺ type 2 AECs in lungs from β1^rtTA mice and CCR2^–/–;β1^rtTA mice. Other cell types, in addition to type 2 AECs, exhibited NF-κB activation in β1^rtTA and CCR2^–/–;β1^rtTA lungs. These findings indicate that β₁ integrin deficiency results in a pervasive inflammatory environment in the distal lung with contributions from retained β₁-deficient type 2 AECs.

Figure 8

Loss of β₁ integrin results in widespread lung inflammation. (A) Increased proinflammatory cytokines in media collected from β1^rtTA type 2 AECs compared with β1^f/f type 2 AECs. n = 11–12 mice/group. (B) β1^rtTA and CCR2^–/–;β1^rtTA lungs exhibit increased proinflammatory cytokines in whole lung tissue lysate by cytokine multiplex assay. n = 4–6 mice/group. (C) Immunostaining for phospho-p65 (red) and the type 2 AEC marker ABCA3 (green) demonstrates increased NF-κB activation in type 2 AECs and throughout the tissue in β1^rtTA and CCR2^–/–;β1^rtTA lungs, as quantified in D. Fewer than 10 sections/mouse; n = 4 mice/group. Scale bars: 50 μm in C, 10 μm for insets. *P < 0.05 by 1-way ANOVA with secondary analysis by Tukey’s test for multiple comparisons. a: β1^f/f vs. CCR2^–/–;β1^rtTA, b: CCR2^–/–;β1^f/f vs. CCR2^–/–;β1^rtTA, c: β1^rtTA vs. CCR2^–/–;β1^rtTA.

β₁-Deficient AEC inflammatory mediators are produced as a consequence of ROS generation. Since the generation of ROS has been linked to NF-κB–dependent cytokine expression in epithelial cells and β₁-containing integrins have been shown to modulate ROS signaling (46–50), we measured ROS production in cultured type 2 AECs. We found that β1^rtTA type 2 AECs produced more superoxide (O₂^–) and hydrogen peroxide (H₂O₂) than β1^f/f cells (Figure 9, A and B); however, no differences in mitochondria-derived ROS were detected (Figure 9C). Given the increase in O₂^– generation, we investigated whether the NADPH oxidase (NOX) system was upregulated in cells from β1^rtTA mice. Of the 5 major NADPH isoforms in the lung epithelium, Duox1 expression was markedly increased in freshly isolated β₁–integrin deficient primary type 2 AECs, but there was no significant difference in expression of the NOX isoforms Duox2, Nox1, Nox2, or Nox4 between β₁ integrin–deficient and control cells (Figure 9D).

Figure 9

β₁-Deficient AECs generate excess inflammatory mediators via ROS production. (A and B) Primary β1^rtTA type 2 AECs produce increased superoxide and H₂O₂ by LumiMax assay (A; n = 7 mice/group) and Amplex Red assay (B; n = 5 β1^f/f, n = 7 β1^rtTA mice), respectively. (C) No difference in mitochondrial ROS production in β1^f/f and β1^rtTA type 2 AECs by MitoSOX assay. n = 8–9 mice/group. ex, excitation; em, emission. (D) Increased gene expression of Duox1 in primary type 2 AECs isolated from β1^rtTA mice. n = 6 mice/group. (E) TEMPOL treatment decreases CCL2 secretion by primary type 2 AECs isolated from β1^rtTA mice. 2 mM; n = 4 mice/group). (F) DPI treatment decreases CCL2 secretion by primary type 2 AECs isolated from β1^rtTA mice. 10 μM; n = 7 β1^f/f, n = 8 β1^rtTA mice. (G) CCL2 secretion is not different between β1^f/f and β1^rtTA type 2 AECs after treatment with the NOX1/4 inhibitor GKT137831. 10 μM; n = 5 β1^f/f, n = 5 β1^rtTA mice. (H) Macrophage migration in response to β1^rtTA type 2 AEC media is decreased by treatment with CCL2-neutralizing antibody (NAb) and DPI. n = 4–8 mice/group. *P < 0.05 by 2-tailed Student’s t test. CCR2, recombinant CCR2.

To define whether ROS production stimulated NF-κB–dependent cytokine expression, we measured levels of CCL2, a known downstream cytokine product of NF-κB activation (51–53). We treated type 2 AECs isolated from β1^rtTA and β1^f/f mice with a superoxide dismutase mimetic (TEMPOL) or a pan-NOX inhibitor (DPI) and measured CCL2 concentration in the media by ELISA. Both TEMPOL and DPI treatment decreased CCL2 secretion by β1^rtTA type 2 AECs (Figure 9, E and F). Although a specific Duox1 inhibitor is not available, we narrowed down the NOX subunits potentially regulated by β₁ integrin using the NOX1/4 inhibitor GKT137831 (Figure 9G). In contrast to the pan-NOX inhibitor DPI, treatment with GKT137831 did not reduce CCL2 secretion from β1^rtTA type 2 AECs, implicating NOX2, Duox1, and/or Duox2 as the source of increased ROS in β1^rtTA mice. As Duox1 was the only NOX isoform with increased expression, these data suggest that β₁ integrin regulates ROS production through this isoform in AECs.

To test whether ROS-dependent CCL2 production by β1^rtTA type 2 AECs was in part responsible for increased macrophage infiltration in β1^rtTA mice, we performed chemotaxis assays using WT macrophages collected by bronchoalveolar lavage and conditioned media from cultured type 2 AECs from β1^f/f and β1^rtTA mice. Macrophage migration toward media from β1^rtTA type 2 AECs was greatly enhanced compared with media from control cells, and this increase was completely abrogated by treatment with DPI or neutralizing antibodies to CCL2 (Figure 9H). These findings support the conclusion that β1^rtTA type 2 AECs have persistent ROS production that contributes to CCL2 secretion that induces macrophage migration into the airspaces of β1^rtTA mice.

While numerous studies have defined the critical role of integrins in organ morphogenesis, few have examined their role in tissue homeostasis in adults. In the setting of development, phenotype severity is highly correlated with timing of integrin deletion after conception and is primarily ascribed to defects in cell adhesion and migration. In this study, we defined the role of β₁ integrin in the structurally stable, fully formed alveolus of the lung, where epithelial cells undergo slow turnover and are tightly bound to the basement membrane. We show that deleting β₁ integrin in AECs under these circumstances results in emphysema, a condition characterized by destruction/loss of gas exchange units and chronic inflammation. Surprisingly, there were no adhesion defects in the AECs in our model; however, these cells were highly inflamed, with excessive ROS production that caused increased NF-κB–dependent cytokine production. Thus, β₁ integrin in alveolar epithelial cells has an antiinflammatory role and is required for alveolar homeostasis in the lung.

Our studies provide direct evidence that mice with a targeted deletion of β₁ integrin in type 2 AECs develop aging-related, spontaneous emphysema as quantified by mean linear intercept. This method easily captures one component of the emphysematous phenotype, enlargement of airspaces. Although we did not perform stereological analysis to address alveolar number specifically, we took precautions in our studies to minimize bias in our 2D morphological measurements from sampling (25–27). Loss of β₁ integrin in AECs stimulates ROS production and NF-κB signaling, and subsequently released inflammatory mediators recruit and activate a mixed population of monocytes/macrophages that efferocytose the β₁-deficient AECs. One possible mechanism for the development of emphysema is that macrophages mediate lung destruction via altered protease/antiprotease balance (54–56). These observations are consistent with studies demonstrating a role for excessive ROS and NF-κB activation as initiators of macrophage accumulation and subsequent alveolar injury, resulting in emphysema (57–59). In addition, epithelial apoptosis in combination with ineffective efferocytosis could contribute to the development of emphysema. Both epithelial and endothelial apoptosis can contribute to emphysema independent of inflammation (60–62). Consistent with these potential explanations for development of emphysema in our model, blocking efferocytosis has been shown to potentiate alveolar destruction in murine models of elastase-induced emphysema associated with increased MMP2 and -12 expression (63). It is unclear whether loss of efferocytosis with its antiinflammatory effects or the retention of inflamed β₁-deficient AECs causes emphysema. However, our data indicating that there is no phenotypical difference in the efferocytosing monocytes/macrophages suggest that retained β₁-deficient AECs are the primary driver of emphysema in β1^rtTA mice. Our data also confirmed and extended studies that indirectly implicated β₁-containing integrins in the pathogenesis of emphysema. Mice with impaired fucosylation exhibit an emphysematous lung phenotype, and fucosylation is required for normal α₃β₁ integrin–dependent migration and signaling, suggesting that the phenotype is due to impairedα₃β₁integrin function (64, 65). Similarly, fibulin 5^–/– mice have enlarged airspaces at birth that progressively dilate into adulthood (66). Fibulin 5 is a ligand for α_vβ₃-, α_vβ₅, and α₉β₁ integrins, participates in outside-in integrin signaling, and is crucial for proper assembly of elastic fibers (66, 67).

We show that the inflammatory phenotype of β₁ integrin–deficient AECs, as manifested by increased NF-κB signaling and cytokine production, is mediated at least in part by excessive ROS production. While this phenomenon is well documented in multiple other cell types (68–70), the mechanisms whereby integrins regulate ROS production are poorly understood. Our studies implicate β₁ integrin as a critical negative regulator of the NOX isoform Duox1 in AECs. Previous studies reported that β₁ integrin negatively regulates ROS production through NOX2 in chondrocytes and kidney mesangial cells (46, 47, 71, 72). Thus, integrins play a critical role in regulating ROS production in multiple cell types; however, the mechanisms appear to be cell type specific.

One of the most interesting observations in our study was that genetic depletion of CCR2, which blocks CCL2-mediated recruitment of monocyte-derived macrophages, exacerbates alveolar remodeling in adult β₁–deficient mice. This contrasts with our previous observation that chemical depletion of macrophages using intranasal instillation of clodronate during lung development rescues alveolarization defects (22). These findings expose differential functions of macrophage subtypes and their potentially paradoxical roles in the adult versus developing lung. In development, fetal lung macrophages are essential for normal lung morphogenesis. They actively clear mesenchymal cells through phagocytosis during sacculation, and their response to inflammatory stimuli regulates airway branching through modulation of developmental signals (36–39, 73–75). During homeostasis, macrophages are required for regulation of inflammatory signaling, host defense, and wound healing (76, 77). The majority of efferocytosis activity following injury is accomplished by macrophages, but more recent data suggest that monocytes significantly contribute to efferocytosis and antigen presentation in the presence of apoptotic cells (78). Although monocytes and macrophages efferocytose dying cells during acute injury, their role in chronic inflammation is less well defined (78, 79). In our model, deranged type 2 AECs are efferocytosed by the CD11b⁺CD11c^– monocyte/macrophage population. This is likely a mix of newly recruited monocytes and monocytes transitioning into macrophages. In β1^rtTA mice, homeostatic compensation fails with loss of CCL2-driven monocyte-macrophage recruitment, resulting in an escalation of inflammation associated with diminished efferocytosis. Although determining why CCL2-recruited monocytes/macrophages are necessary for efficient efferocytosis of type 2 AECs will require further study, this finding could have direct implications for human lung diseases, including COPD, in which ineffective efferocytosis has been suggested to be a contributor to pathogenesis (80–86).

Aside from regulating inflammation, β₁ integrin is required for other critical cellular processes in type 2 AECs, including maintenance of tight junctions and control of proliferation. Microscopic examination of β1^rtTA mice revealed abnormal tight junctions associated with decreased Claudin-3 and Claudin-4 expression. Consistent with this observation, proximal kidney tubule cells with β₁ integrin deletion exhibit altered Claudin-2 expression (87). Our epithelial β₁–null mice also share many phenotypic similarities with claudin-deficient mice. Unchallenged claudin-4–deficient mice have normal lung histology but exhibit significantly increased BALF cell counts with hyperoxia exposure and increased CCL2 signaling with mechanical ventilation (88). In addition, claudin-18–null mice develop AEC hyperplasia and macrophage accumulation over time (89–91). Although a causal relationship between β₁ integrin and claudins has not been described, previous in vitro studies demonstrated that ROS can disrupt tight junctions (92–94), suggesting that β₁ integrin–mediated ROS could regulate tight junctions via effects on claudin expression. Another phenotype identified in β1^rtTA mice is AEC proliferation. Developmental deletion of β₁ integrin decreases epithelial proliferation in the kidney, mammary, and submandibular glands (11, 15, 19, 21, 22), whereas increased epithelial proliferation has been reported when β₁ integrin was deleted in the intestine or skin (14, 17). The mechanisms whereby β₁ integrin regulates cell number/density in fully formed organs is unknown; however, this could also be ROS mediated, since multiple investigations have shown that ROS can stimulate cell proliferation.

In conclusion, this study shows that loss of β₁ integrin in type 2 AECs promotes persistent lung inflammation and emphysematous remodeling, which is mitigated by efferocytosis of inflamed AECs by CD11b⁺CD11c^– monocytes/macrophages. Thus, regulation of inflammation is the major function of β₁ integrin in alveolar homeostasis in the lung.

Mice. For timed deletion of β₁ integrin, we crossed transgenic mice with inducible Cre recombinase expression by the dox-inducible reverse tetracycline transactivator under control of the SP-C promoter (SP-C rtTA;Tet-O-Cre) with integrin β₁^fl/fl mice (95, 96). Postdevelopmental type 2 AEC deletion was induced on P28 in these triple transgenic SP-C rtTA;Tet-O-Cre; β1^f/f mice (called β1^rtTA mice) using dox in drinking water (2 g/L × 4 weeks). Control littermate β1^f/f mice received identical dox treatment. To test the role of β₁ integrin in epithelial differentiation during alveolar homeostasis, we crossed β1^rtTA mice to the mTmG Cre recombinase reporter. To test the role of CCL2-recruited monocytes/macrophages in β1 integrin–regulated alveolar homeostasis, we crossed β1^rtTA and β1^f/f mice onto a homozygous null background for Ccr2, the CCL2 receptor. The resulting transgenic CCR2^–/–;SP-C rtTA;Tet-O-Cre;β1^f/f mice (termed CCR2^–/–;β1^rtTA mice) and control littermate CCR2^–/–;β1^f/f mice, received identical dox treatment on P28 to induce β₁ integrin deletion. Integrin β1^f/f mice were a gift from Elaine Fuchs (Howard Hughes Medical Institute, The Rockefeller University, New York, New York, USA). SP-C rtTA, Tet-O-Cre, Ccr2 homozygous null, and mTmG Cre recombinase reporter mice were purchased from the Jackson Laboratory. All mice were on a C57BL/6 background.

Histology and morphological analysis. Lungs from β1^rtTA, β1^f/f, CCR2^–/–;β1^rtTA; and CCR2^–/–;β1^f/f mice were harvested for histological examination. Mice were sacrificed, right ventricle flushed with PBS, and lung inflation fixed at 25 cm with 10% formalin for more than 24 hours prior to paraffin embedding and sectioning. Multiple lobes were sectioned to reduce bias in morphological analysis generated from regional differences in alveolar size (25–27). Mean linear intercept was calculated from images obtained using a Keyence BZ-X710 inverted fluorescence phase contrast microscope with ×40 objective for 6–10 nonoverlapping sections per mouse. For immunohistochemistry stains, paraffin sections were deparaffinized, antigen retrieved, blocked, and incubated with the indicated primary antibody, followed by colorimetric detection by Vector Red (Vector Laboratories). Stained paraffin sections were imaged using a Keyence BZ-X710 inverted fluorescence phase contrast microscope with ×20 objective lens (low-power images) or an Olympus BX41 with ×60 objective lens (high-power images). Immunofluorescence staining was performed on frozen lung sections that were inflation fixed with 2:1 PBS/O.C.T. mixture (Tissue-Tek), embedded, and sectioned at 8-μm thickness. Slides were subsequently fixed with 4% paraformaldehyde, permeabilized with 0.1% Triton-X, blocked with 5% donkey serum, incubated in primary antibody overnight at 4°C, incubated in secondary antibody for 2 hours at room temperature, incubated with DAPI nuclear stain (Vector Laboratories), mounted with ProLong Gold (Thermo Fisher Scientific), and imaged using a Nikon Spinning Disk TiE inverted fluorescence confocal microscope attached to an Andor DU-897 EMCCD camera (×20 or ×60 objective). The following primary antibodies were used: anti–pro–SP-C (Abcam ab90716), rabbit anti-CD68 (Abcam ab125212), rat anti-CD68 (Abcam ab53444), anti–β₁ integrin (Millipore MAB1997), anti–laminin γ1 (Novus NBP2-44751), anti–pan-laminin (Millipore AB2034), anti–Ki-67–FITC (eBioscience 11-5698-80), anti-ABCA3 (Seven Hills WMAB-13H257), and anti–phospho-p65 (S276) (Abcam ab106129). The following secondary antibodies were used: anti-rabbit Alexa Fluor 488 (Life Technologies A21206), anti-rabbit Alexa Fluor 594 (Life Technologies A21207), anti-mouse Alexa Fluor 488 (Life Technologies A11001), and anti-rat Alexa Flour 488 (Life Technologies A21208). TUNEL staining was performed per the manufacturer’s instructions (Roche 11684795910). Quantification of immunostained sections was performed on 20 nonoverlapping images obtained with a ×20 objective.

TEM. Lungs were harvested from 3-month-old β1^rtTA and β1^f/f mice, processed, postfixed with potassium ferrous cyanide, dehydrated with graded acetone, thick sectioned at 1 μm, thin sectioned at 80 nm in the region of interest, and imaged using a Philips FEI T-12 transmission electron microscope in the Vanderbilt Cell Imaging Shared Resource core.

Bronchoalveolar lavage. Sterile saline lavages were performed with 1 mL PBS after sacrifice. Lavage fluid was centrifuged at 270 g at 4°C, and cells were resuspended and counted. The Pierce BCA Protein Assay kit (Thermo Fisher Scientific, 23225) was used to test for BALF protein per the manufacturer’s instructions. For immunofluorescence analysis of immune cells collected by bronchoalveolar lavage, 40,000 cells were spun onto Shandon cytoslides (Thermo Fisher Scientific) at 240 g for 7 minutes, dried, and immunostained per the above protocol.

AEC isolation and collection of conditioned medium. Type 2 AECs were isolated from 3-month-old β1^rtTA and β1^f/f mice as previously described, yielding more than 90% type 2 AECs (22, 97, 98). Briefly, a single-cell suspension was generated with a 40-minute dispase digestion and 100-μm, 40-μm, and-20 μm serial filtration. The suspension was then incubated at 37°C for 2 hours in anti-CD45 (BD 553076) and anti- CD32 (BD 553142) antibody–coated plates for negative selection. The medium containing epithelial cells was collected and spun down, and AECs were plated in 5% bronchial epithelial cell growth medium (BEGM) on Matrigel-coated wells with or without the indicated treatment. Treatment reagents included TEMPOL (Sigma-Aldrich 176141) and DPI (Sigma-Aldrich D2926). Medium was collected at 24 hours for analysis.

Western blotting. Protein (60 μg) collected from type 2 AEC isolations was electrophoresed in a 10% gel and transferred onto nitrocellulose membranes. Membranes were blocked, incubated with primary antibody (anti–β₁ integrin [Millipore MAB1997], anti–claudin-3 [Invitrogen 341700], anti-GAPDH [Invitrogen MA5-15738]), and incubated with Odyssey IRDye 800CW and 680RD secondary antibodies. Signal was detected using a LI-COR Odyssey CLx Near-Infrared Western Blot Detection system.

qPCR. RNA was isolated from freshly isolated primary type 2 AECs using the RNEasy Plus Mini Kit (QIAGEN) and cDNA synthesized using the SuperScript VILO Master Mix kit (Thermo Fisher Scientific). qPCR reactions were performed in triplicate using TaqMan PCR Fast Advanced Master Mix (Applied Biosystems, Thermo Fisher Scientific) on a StepOne Plus PCR System (Applied Biosystems) using the following TaqMan probes (Applied Biosystems, Thermo Fischer Scientific): Claudin-4 Mm00515514_s1, Claudin-18 Mm00517321_m1, CD47 Mm00495011_m1, Duox1 Mm01328685_m1, Duox2 Mm01326247_m1, Nox1 Mm00549170_m1, Nox2 Mm01287743_m1, Nox4 Mm00479246_m1, and GAPDH Mm99999915_g1. Data were normalized to the housekeeping gene GAPDH. Relative quantities were analyzed using β1^f/f values as control.

ELISA and multiplex assay. ELISA for CCL2 and CX3CL1 on AEC conditioned media was performed in triplicate according to the manufacturer’s instructions (R&D Systems, MJE00 and MCX310, respectively). Cytokine/chemokine Magnetic Bead 32-Multiplex Panel (Millipore MCYTMAG-70K-PX32) assay was performed on monocyte/macrophage conditioned media, AEC conditioned media, and whole lung tissue lysates in triplicate per the manufacturer’s instructions. Tissue lysates were generated by sonicating the right upper lobe, centrifuging the tissue mixture, collecting the supernatant, and normalizing to protein. The Multiplex assay was read on the Luminex MAGPIX platform in the Vanderbilt Hormone and Analytical Services Core.

Flow cytometry. We used collagenase XI (Sigma-Aldrich C7657, 0.7 mg/mL) and type IV DNAse (Sigma-Aldrich D5025, 30 μg/ml) digestion and 40-μm filtration to obtain a single-cell whole lung suspension for flow cytometry analysis. Briefly, cells were blocked with anti-CD32 antibody (BD 553142), incubated with conjugated primary antibody, and analyzed using a 5-laser BD LSR II analytical flow cytometer (BD Biosciences) and FlowJo analysis software (Becton, Dickinson, and Co.). Both single antibody and fluorescence-minus-one controls were used for compensation. The following primary conjugated antibodies were used in flow cytometry experiments: CD45-BV650 (BioLegend 103151), CD64-APC (BioLegend 139306), CD11b–PE-Cy7 (BD 561098), CD11c–PE-Cy5 (eBioscience 15-0114-82), and Ly6C-APC-Cy7 (BD 560596).

Efferocytosis assay. Macrophages collected by bronchoalveolar lavage (5 × 10⁴ cells/well) were plated for 4 hours in serum-free media and exposed to fluorescently labeled primary type 2 AECs (Millipore 382065) from β1^rtTA mice (1 × 10⁵ cell/well) for 1 hour. After incubation, nonadherent cells were removed by careful washing, and fluorescence was detected on a Molecular Devices SpectraMax M5 Plate Reader.

Macrophage migration assay. Conditioned medium from primary β1^rtTA and β1^f/f AECs was placed in the bottom chamber of a 5-μm Transwell insert (Corning 3422). WT macrophages (40,000 per insert) were obtained from pooled BALF and placed in the top chamber, incubated at 37°C for 4 hours. Unmigrated macrophages were removed from the top chamber, while migrated macrophages were fixed to the underside of the Transwell membrane and stained using the spHema 3 Manual Staining System (protocol 122-911, Fisher Scientific). Five nonoverlapping images of stained migrated macrophages were taken, and the number of migrated cells/field was quantified.

ROS assays. LumiMax Superoxide Anion Detection Kit (Agilent Technologies 204525) was used to detect superoxide from primary type 2 AECs per the manufacturer’s instructions. Amplex Red assay (Invitrogen, Thermo Fisher Scientific A22188) was used to detect H₂O₂ released from primary type 2 AECs per the manufacturer’s instructions. MitoSOX assay (Thermo Fisher Scientific M36008) was performed on primary type 2 AECs per the manufacturer’s instructions. For inhibitor studies, we used TEMPOL (Sigma-Aldrich 176141, 2 mM) and DPI (Sigma-Aldrich D2926, 10 μM).

Statistics. A 2-tailed Student’s t test was used for comparisons between 2 groups, with results representing mean SEM. For comparisons between 3 or 4 groups, an ordinary 1-way ANOVA was used with secondary analysis by Tukey’s test for multiple comparisons as indicated. For both statistical analyses, P < 0.05 was considered statistically significant.

Study approval. All animal experiments were approved by the Vanderbilt University Medical Center Institutional Animal Care and Use Committee.