Resolution of bleomycin-induced murine pulmonary fibrosis via a splenic lymphocyte subpopulation

Nine-week-old male C57BL/6 mice were purchased from Charles River Laboratories Japan (Yokohama, Japan). We chose this strain because it is well characterized and is susceptible to BLM-induced pulmonary fibrosis. Interleukin (IL)-10 knock-out mice with C57BL genetic background (B6.129P2-Il10tm1Cgn/J) were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). The experimental protocols in this study were approved by the animal care and use committee of Nippon Medical School (Tokyo, Japan, Approval Number; 27–098).

Osmotic pumps (ALZET model 2001; DURECT Corporation, Cupertino, CA, USA) containing 200 μL saline, with or without BLM (100 mg/kg of mouse body weight; Nippon Kayaku Co., Tokyo, Japan), were implanted in recipient C57BL/6 mice on day 0 (Fig. 1a) [13]. Incision wounds were sealed using a surgical suture. BLM was continuously infused via the pumps over 7 days, according to the manufacturer’s instructions.

Spleens were removed from donor C57BL/6 mice without BLM treatment and minced to obtain single-cell suspensions. Recipient C57BL/6 mice treated with BLM received 1 × 10⁵ donor C57BL/6 splenocytes on day 7 or 14 via caudal vein injection. On day 28, mice were sacrificed and the lungs were removed for examination.

Lung samples were fixed in 10% formalin buffer (Wako Pure Chemical Industries, Ltd., Osaka, Japan) for histological examination. Paraffin sections (3.5 μm-thick) were cut from fixed lungs, stained with hematoxylin and eosin (HE) to assess gross morphology and Masson’s trichrome stain to visualize collagen deposition [14], and examined by microscopy. Lung fibrosis was measured using quantitative histology following Ashcroft’s method [15].

To deplete Tregs, we incubated single-cell suspensions of splenocytes (1 × 10⁵) with 1 μL of an anti-CD25 monoclonal antibody (mAb) (clone: PC61; BioLegend, San Diego, CA, USA) dissolved in sterile phosphate-buffered saline (PBS) for 20 min on ice. Then, these cells were adoptively transferred to BLM-treated mice on day 14 via the caudal vein. On day 28, the mice were sacrificed, and the lungs were removed and subjected to HE and Masson’s trichrome staining. The extent of lung fibrosis was measured by quantitative histology according to Ashcroft’s method to determine the effect of CD25 neutralization with the anti-CD25 mAb.

Splenocyte subpopulations were analyzed by evaluating the relative proportions of Tregs, macrophages, and B cells. Single-cell suspensions of splenocytes were stained with anti-mouse CD4 (BioLegend) and CD25 (eBioscience, Waltham, MA, USA) antibodies to analyze Tregs. To analyze macrophages and their polarization, splenocytes were stained with anti-mouse F4/80 (BioLegend), CD80 (eBioscience), and CD206 (BioLegend) antibodies. Splenocytes were stained with an antibody against CD45R (B220; BioLegend) for B cell detection. To eliminate nonspecific staining, isotype- control antibodies, matched to the surface marker antibody’s host species and class, were used. FACS analysis was performed using a BD FACSCanto II and BD FACSVerse flow cytometers (BD Biosciences, San Diego, California, USA). Data collected were analyzed with FlowJo software (Tree Star, Inc., Ashland, Oregon, USA).

After single-cell suspensions were obtained from the spleens of C57BL/6 mice, Tregs were purified using a MiniMacs CD4⁺CD25⁺ Regulatory T-cell Isolation Kit (Miltenyi Biotec, Auburn, CA, USA) according to the manufacturer’s instructions. To achieve the highest purity, we separated positive and negative cell fractions by passing the cells through a second column. BLM-treated recipient C57BL/6 mice received an intravenous tail vein injection of 1 × 10⁶ isolated donor cells on day 14 after initiating BLM treatment. The dose of Tregs was determined according to a study performed by D’Alessio FR and colleagues [16]. Mice were sacrificed on day 28, and the lungs were removed and subjected to histological and biochemical analyses. Blood collected by cardiac punctures and lung homogenates were subjected to cytokine or chemokine analyses.

The total collagen content of the right lung was determined by hydroxyproline assay [17]. After acid hydrolysis of the right lung with 12 N HCl at 100 °C for 20 h in a sealed glass tube (Iwaki, Tokyo, Japan), the hydroxyproline content was determined by high-performance liquid chromatography.

Sections of paraffin-embedded lung lobes were deparaffinized and rehydrated. Antigen retrieval was achieved by boiling at 105 °C for 7 min in 10 mM citrate buffer (pH 6.0), followed by gradual cooling to room temperature. Then, the sections were treated with 3% hydrogen peroxide in methanol for 20 min and blocked with 10% normal goat serum (NICHIREI BIOSCIENCES, INC., Tokyo, Japan) at room temperature for 10 min. Sections were incubated with an anti-FGF9 polyclonal antibody (Abcam, Cambridge, MA, USA) for 30 min at room temperature. For FGF9 staining, tissue sections were incubated with a secondary anti-rabbit antibody (NICHIREI BIOSCIENCES, INC.) for 30 min at room temperature. FGF9 expression was visualized using Histofine Simple Stain Mouse MAX-PO (R) and Histofine Simple Stain AEC Solution (NICHIREI BIOSCIENCES, INC.).

Mouse plasma FGF9 and IL-10 levels were measured using ELISA kits purchased from Cloud-Clone Corp. (Houston, TX, USA) and R&D Systems Inc. (Minneapolis, MN, USA), respectively. Approximately 0.5–1.0 mL of blood was collected from each mouse on day 28 by cardiac puncture, placed in a tube containing EDTA and aprotinin, incubated at 4–8 °C for 15 min, and then centrifuged at 5000 rpm for 2 min at 4 °C to separate the plasma. Plasma samples were stored at − 20 °C until analysis. For chemoattractant chemokine (CC motif) ligand 2 (CCL2) measurement in the lungs, the right lung of each mouse was added to 1.0 mL ice-cold lysis buffer (100 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, and protease inhibitor cocktail [Complete mini; Roche Diagnostics, Basel, Switzerland]), and then homogenates were prepared with a 2-mL tissue grinder (Wheaton Industries, Millville, New Jersey, USA). After centrifuging the homogenate at 10,000×g for 5 min at 4 °C, supernatants were prepared from the lung homogenates, and CCL2 concentration was measured using ELISA kits from Cloud-Clone Corp.

ELISA was performed to determine the concentrations of FGF9, IL-10, and CCL2 following the manufacturer’s protocols. Briefly, supernatants of each sample were diluted in PBS, and 100 μL of the diluted samples was assayed with the kits. The optical density was measured at a wavelength of 450 nm using a microtiter plate reader. All samples were measured in duplicate.

Sections of paraffin-embedded lung lobes were deparaffinized and rehydrated. Antigen retrieval was achieved by boiling at 105 °C for 7 min in 10 mM citrate buffer (pH 6.0) (LSI Medience Corporation, Tokyo, Japan), followed by gradual cooling to room temperature. Then, the sections were blocked with 1% bovine serum albumin (BSA) (Sigma–Aldrich Corporation, St. Louis, MO, USA) for 20 min at room temperature. Tissue sections were incubated with anti-CD45 and anti-Col-I antibodies (Abcam) overnight at 4 °C followed by Alexa Fluor 488-conjugated and Alexa Fluor 568-conjugated antibodies (Abcam) for 1 h at room temperature. VECTASHIELD mounting medium dispersed over the entire section to counterstain the nuclei with 4′,6-diamidino-2-phenylindole (DAPI) (Vector Laboratories, Inc. Burlingame, CA, USA).

Sections of paraffin-embedded lung lobes were deparaffinized and rehydrated. Antigen retrieval was achieved by boiling at 105 °C for 7 min in 10 mM citrate buffer (pH 6.0), followed by gradual cooling to room temperature. Then, the sections were blocked with 1% BSA for 20 min at room temperature. For FGF9 staining, tissue sections were incubated with an anti-FGF9 antibody (Abcam) for 1 h at room temperature followed by an Alexa Fluor 568-conjugated antibody (Abcam) for 30 min at room temperature. Subsequently, they were blocked with 1% BSA for 20 min at room temperature and incubated with an anti-E-cadherin antibody (Merck KGaA, Darmstadt, Germany) at 4 °C overnight followed by an Alexa Fluor 488 antibody for 30 min at room temperature. VECTASHIELD mounting medium dispersed over the entire section to counterstain the nuclei with DAPI.

Either on day 0 or 14 after BLM challenge, the abdominal cavity of mice was opened above the left kidney during isoflurane anesthesia. Then, the spleen vessels were carefully cauterized and the spleen was removed. On day 28, mice were sacrificed and lungs were subjected to HE and Masson’s trichrome staining to quantify the fibrotic score using Ashcroft method.

Animal experiments involved at least 6 mice in each treatment group, unless otherwise stated. Comparisons among multiple groups were analyzed by 1-way analysis of variance with the Tukey–Kramer post-hoc correction to adjust for multiple comparisons. An unpaired 2-tailed Student’s t test was used for single comparisons. Data were analyzed using JMP 9 software, version 9.0.3 (SAS Institute Inc., Cary, NC, USA). Differences were considered statistically significant when P values were less than 0.05. Data are expressed as the mean ± standard deviation.

Because whole splenocytes can ameliorate LPS-induced lung injury in mice [16], as an initial assessment, we examined the potency of the adoptive transfer of splenocytes in ameliorating BLM-induced murine pulmonary fibrosis. BLM (100 mg/kg body weight) was administered to C57BL/6 mice using osmotic pumps (day 0, Fig. 1a). Splenocytes were prepared from the spleens of C57BL/6 mice without BLM treatment and injected (1 × 10⁵ cells/mouse) via the caudal vein on either day 7 or 14 post-BLM challenge. Mice were sacrificed on day 28, and the lungs were removed and subjected to histological examination. Lung histological data obtained on day 28 post-BLM administration showed focal fibroplasias with destruction of the alveolar wall in the group receiving BLM (Fig. 1d and e) but not in the saline group (Fig. 1b and c). Injection of splenocytes on day 14 ameliorated the lesions (Fig. 1h and i); however, no obvious effect of splenocyte injection was observed on day 7 (Fig. 1f and g). To quantify the anti-fibrotic effects of splenocytes in the lungs of BLM-treated mice, we determined the extent of lung fibrosis by quantitative histology according to Ashcroft’s method on day 28 post-treatment. Because BLM administration with osmotic pumps causes lung fibrosis predominantly in the subpleural regions [18, 19], subpleural fibrosis between the groups was compared using a numerical scale. Two blinded observers [KKa and MI] quantified fibrosis in each section. Splenocytes significantly attenuated the fibrosis score when administered to BLM-treated mice on day 14 (Fig. 1j, *P < 0.05); however, administration of splenocytes on day 7 had no effect on pulmonary fibrosis. To characterize the anti-fibrotic effects of splenocytes, we assayed the hydroxyproline content in the lungs on day 28. As shown in Fig. 1k, the adoptive transfer of splenocytes on day 14 post-BLM challenge significantly reduced the hydroxyproline content in the lungs when compared with the BLM treated group (*P < 0.05).

Splenocytes have been reported to be rich in Tregs, and D’Alessio et al. reported that Tregs contributed to the resolution of LPS-induced lung injury in mice [16]. Given that the resolution of BLM-induced murine pulmonary fibrosis was dependent on spleen cells due to their suppressive properties, we hypothesized that specific lymphocyte subsets among splenocytes may be critical in this process. To begin understanding the potential contribution of those cells to BLM responses, we depleted Tregs using an anti-CD25 mAb. Splenocytes were incubated with the anti-CD25 mAb, after which they were injected via the caudal vein on day 14 post-BLM treatment. On day 28, the mice were sacrificed, and their lungs were removed and subjected to histological examination and biochemical analyses (Fig. 2). As shown in Fig. 2e and f, splenocytes depleted of Tregs using an anti-CD25 mAb lost the potential to ameliorate BLM-induced murine pulmonary fibrosis. To quantify this effect, we determined the extent of lung fibrosis by quantitative histology on day 28, according to Ashcroft’s method. Two blinded observers [KKa and MI] quantified the degree of fibrosis in each section. Amelioration of BLM-induced murine pulmonary fibrosis by splenocytes was significantly abrogated by Treg depletion with the anti-CD25 mAb (Fig. 2g, * P < 0.01). These data indicated that CD25-mediated Treg depletion promotes increased lung fibrosis, thereby suggesting that immunomodulatory cells (including Tregs) might mediate the effects of splenocytes on pulmonary fibrosis resolution. A similar trend in the numerical score was observed on the lung hydroxyproline content, although statistical significance was not achieved (Fig. 2h, P = 0.16).

To characterize splenocytes, we examined the number of CD4 and CD25 double-positive cells by immunostaining and FACS analysis of spleen digests from BLM-untreated C57BL/6 mice. As shown in Fig. 2i, Tregs accounted for approximately 11% of the splenocytes. Furthermore, we assayed macrophage subpopulations because diverse activation states of macrophages can result in phenotypic polarization and the formation of pro-inflammatory M1 and pro-healing M2 type macrophages [20]. As shown in Fig. 2j, approximately 10% of splenocytes were macrophages (F4/80), most of which were of the M1 phenotype, in agreement with previous data published elsewhere [21]. B cells constituted approximately 30% of the splenocytes (data not shown).

The data obtained using the anti-CD25 mAb prompted us to examine whether Tregs potently resolve BLM-induced pulmonary fibrosis in mice. To that end, Tregs isolated from spleens of donor C57BL/6 mice were adoptively transferred to recipient mice (1 × 10⁶ cells/mouse) via the caudal vein on day 14 post-BLM challenge (Fig. 3a). Because adoptive transfer of splenocytes on day 14 ameliorated fibrosis, which was abrogated by an anti-CD25 antibody, this time point was selected for the Treg transfers. Thereafter, on day 28, mice were sacrificed, and the lungs were removed and subjected to immunohistochemical and biochemical analyses (Fig. 3). In addition, blood was collected by cardiac punctures to analyze the FGF9 and IL-10 levels, and lung homogenates were prepared to measure CCL2 levels. As shown in Fig. 3h and i, adoptively transferring Tregs on day 14 ameliorated the lesions. To quantify the anti-fibrotic effects of Tregs in the lungs of BLM-treated mice, we determined the extent of lung fibrosis using a quantitative histology according to Ashcroft’s method on day 28. Two blinded observers [KKa and AM] quantified fibrosis in each section. Tregs significantly attenuated the fibrosis score when adoptively transferred to BLM-treated mice on day 14 (Fig. 3j, *P < 0.01). To further characterize the anti-fibrotic effects of Tregs, we quantified the hydroxyproline content in the lungs on day 28. As shown in Fig. 3k, adoptive transfer of Tregs significantly reduced the hydroxyproline content in the lungs when compared with the BLM treated group (*P < 0.05).

FGF9 is a neuroimmune system molecule with manifold effects on embryonic development and mesenchymal proliferation that may be involved in human lung fibrosis [22]. While in vivo data demonstrated that antibody-mediated Treg depletion augmented FGF9 expression in mice overexpressing TGF-β1 [23], the effect of adoptively transferred Tregs on FGF9 expression in BLM-induced lung fibrosis is not yet described. To determine whether Tregs affect FGF9 expression, we performed immunohistochemistry against FGF9. The control and Treg transferred groups had undetectable FGF9 expression (data not shown), which was consistent with a prior report [23]. In contrast, FGF9-positive cells bearing the morphology of alveolar epithelial cells markedly increased in the setting of BLM-induced pulmonary fibrosis (Fig. 4a), whereas FGF9-positive cells decreased following the adoptive transfer of Tregs (Fig. 4b). Composite data represented as the number of FGF9-positive cells revealed that the adoptive transfer of Tregs reduced FGF9-positive cells by 54.2%, as compared to BLM-treated mice without adoptively transferred Tregs (Fig. 4c). To confirm that FGF9-positive cells are epithelial cells, we performed dual staining of these cells using anti-FGF9 and anti-E-cadherin antibodies. As shown in Fig. 4d–f, most of the FGF9-positive cells (red) expressed E-cadherin (green). In addition, we determined the plasma FGF9 levels by ELISA. As shown in Fig. 4g, the plasma FGF9 concentration appeared to increase with BLM treatment, as compared to saline-treated control mice. The increase showed a trend to be reduced to control levels by the adoptive transfer of Tregs, although statistical significance was not achieved.

Tregs are conventionally associated with the production of classic anti-inflammatory cytokines including IL-10, TGF-β, and IL-35, consistent with their anti-inflammatory functions [24]. Among these cytokines, IL-10 has been reported to inhibit BLM-induced lung injury in mice [25, 26]. Therefore, we hypothesized that the Treg-mediated anti-fibrotic effect was due in part to IL-10 production from adoptively transferred Tregs. To test this hypothesis, we collected plasma samples from BLM-treated and Treg-transferred mice, and then assessed IL-10 levels in ELISAs. The data indicated that the plasma IL-10 concentration increased by up to 3.0-fold after BLM treatment (Fig. 5). Unexpectedly, the adoptive transfer of Tregs significantly inhibited IL-10 production in mouse plasma induced by BLM administration.

The above data prompted us to examine the effect of Tregs isolated from IL-10 knock-out mice on pulmonary fibrosis. Tregs isolated from IL-10 knock-out mice spleens were adoptively transferred via the tail vein on day 14 after a BLM-challenge according to the protocol described in Fig. 3a. On day 28, the mice were sacrificed, and the lungs were removed and subjected to immunohistochemical and biochemical analyses (Fig. 6). As shown in Fig. 6e and f, adoptive transfer of Tregs isolated from IL-10 knock-out mice did not ameliorate the fibrosis. We quantified these results using quantitative histology according to Ashcroft’s method on day 28. Two blinded observers [KKa and AM] quantified fibrosis in each section. Although statistical significance was not achieved, a trend was observed indicating that the transfer of Tregs isolated from IL-10 knock-out mice did not attenuate the fibrosis score when compared with transferring Tregs obtained from wild-type C57BL/6 mice (Fig. 6g). We also quantified the hydroxyproline content in the lungs on day 28. As shown in Fig. 6h, a similar trend for the numerical score was observed.

Bone marrow-derived circulating fibrocytes are mesenchymal progenitor cells that express markers compatible with leukocytes, hematopoietic progenitor cells, and fibroblasts [27]. They are chemotactically recruited to sites of tissue injury by chemokines, including CCL2 [28], and promote fibrotic responses [29]. Because Tregs have been found to inhibit fibrocyte recruitment [23], we hypothesized that Treg adoptive transfer affects chemokine production in the lungs. To determine this, we measured CCL2 production by ELISA in BLM-induced murine pulmonary fibrosis lungs that had received Treg adoptive transfer. As shown in Fig. 7a, BLM treatment increased CCL2 production by up to 2.0-fold in the lungs. Adoptive transfer of Tregs significantly reversed this increase to the basal levels, which was accompanied by reduced accumulation of fibrocytes in the lungs (Fig. 7b–g).

Mouse splenocytes can be reduced in number by BLM administration [30], causing us to speculate that spleen is a reservoir of Tregs. Because our observation suggested the participation of spleen cells in the amelioration of pulmonary fibrosis, we were prompted to examine the effect of splenectomy. BLM was administered to C57BL/6 mice using osmotic pumps (day 0, Fig. 8a), after which the spleen vessels were carefully cauterized and spleen was removed either on day 0 or 14. On day 28, the mice were sacrificed and the lungs were subjected to HE and Masson’s trichrome staining to quantify the fibrotic score, using Ashcroft’s method. Two blinded observers [KKa and AM] quantified the degree of fibrosis in each section. As shown in Fig. 8b–f, splenectomy on day 0 significantly ameliorated pulmonary fibrosis, whereas that on day 14 had no effect on fibrosis (data not shown).