Loss of immune tolerance to self-antigens can promote chronic inflammation and disrupt the normal function of multiple organs, including the lungs. Degradation of elastin, a highly insoluble protein and a significant component of the lung structural matrix, generates proinflammatory molecules. Elastin fragments (EFs) have been detected in the serum of smokers with emphysema, and elastin-specific T cells have also been detected in the peripheral blood of smokers with emphysema. However, an animal model that could recapitulate T cell–specific autoimmune responses by initiating and sustaining inflammation in the lungs is lacking. In this study, we report an animal model of autoimmune emphysema mediated by the loss of tolerance to elastin. Mice immunized with a combination of human EFs plus rat EFs but not mouse EFs showed increased infiltration of innate and adaptive immune cells to the lungs and developed emphysema. We cloned and expanded mouse elastin-specific CD4+ T cells from the lung and spleen of immunized mice. Finally, we identified TCR sequences from the autoreactive T cell clones, suggesting possible pathogenic TCRs that can cause loss of immune tolerance against elastin. This new autoimmune model of emphysema provides a useful tool to examine the immunological factors that promote loss of immune tolerance to self.

Elastin is a matrix protein that composes over 90% of assembled elastic fibers in the extracellular space and provides the required tissue strength and elasticity necessary for multiple organs (1). Specifically, proper function of the lungs, vascular structures, and skin depends on their flexibility; as such, they contain a much higher amount of elastin per dry weight than other organs (2). Under steady-state, biogenesis of matrix molecules includes regular reorganization; however, extracellular elastin matrix assembly, known as elastogenesis, primarily occurs during organ development and remains highly stable throughout life (3). Thus, elastin degradation due to abnormal exposure to elastolytic enzymes expressed by innate immune cells can result in organ dysfunction and life-threatening diseases of the lung (48) and vasculature (912).

Cigarette smoking causes a distinct pattern of lung parenchyma destruction characterized by loss of tissue elasticity and generation of elastin fragments (EFs) found in the serum (13, 14). We and others have shown that chronic exposure to cigarette smoke recruits innate and adaptive immune cells into the lung (5, 1518). Activated innate immune cells (e.g., macrophages and neutrophils) release several elastin-degrading enzymes, including neutrophil elastase, matrix metalloproteinase (MMP) 9, and MMP12, which can either directly cleave elastin or block alpha-1 anti-trypsin, the absence of which is associated with severe emphysema (8, 19, 20). In addition to innate immune cells, activated adaptive immune cells (T and B lymphocytes) are recruited to the lungs of smokers, and adoptive transfer of CD8+ T cells has been shown to induce lung inflammation and emphysema (2124).

We and others have shown that smokers who develop emphysema harbor activated Th1 and Th17 cells expressing IFN-γ and IL-17A, respectively, in the lungs when compared with control subjects (2527). Consistently, CD4+ T cells isolated from the peripheral blood of smokers with emphysema show increased IFN-γ and IL-17A expression in response to EFs, which can be inhibited in the presence of MHC class II blocking Abs (28, 29). The significance of adaptive immunity against elastin was shown in a longitudinal study in which the magnitude of autoreactive immune responses to EFs correlated with the severity of physiological decline over 3 years (30). Moreover, we have shown that autoreactive T cell responses significantly correlate with emphysema severity and lung function decline (28, 29). Collectively, human studies suggest that elastin-specific autoreactive T cells persist in smokers with emphysema despite smoking cessation, which may contribute to progressive inflammation and result in the destruction of several elastin-rich organs.

Despite recent advances and a better understanding of the pathophysiological effects of chronic cigarette smoke–induced lung inflammation, little is known about the loss of immune tolerance to elastin. In this paper, we provide the methods that we used to develop a novel mouse model of emphysema that reproduces autoimmune inflammation against elastin that is found in smokers. Repeated immunization using non-self EFs (human and rat), but not mouse elastin, successfully broke tolerance against elastin in mice; the model recapitulated cigarette smoke–induced emphysema characterized by airspace enlargement and inflammatory cell infiltration in elastin-rich organs. The precise contribution of EF reactive T cells to tissue damage is not fully known; however, we cloned autoreactive T cells and identified several potential pathogenic TCRs against mouse elastin. In this study, we describe the in vivo method for induction of EF-specific T cell responses and isolation of both mouse and human TCR sequences for in vitro analysis of EF-specific TCR function. This novel experimental mouse model provides a valuable tool that could be used to determine the underlying mechanisms involved in the loss of immune tolerance to EFs in elastin-rich organs. Furthermore, we anticipate that the techniques used to develop this model could be easily adapted to establish novel mouse models for other autoimmune diseases.

C57BL/6 mice (7–8-wk-old females) were purchased from The Jackson Laboratory (Bar Harbor, ME). All experimental protocols were approved by the Institutional Animal Care and Use Committee of Baylor College of Medicine and followed the National Research Council Guide for the Care and Use of Laboratory Animals.

PBMCs were isolated from smokers with emphysema, and T cells reactive to human lung EFs (hEFs) were isolated and cloned as previously described (29). Studies were approved by the Institutional Review Board at Baylor College of Medicine, and informed consent was obtained from all patients.

TC-378-4, a human elastin-specific CD4+ T cell clone isolated from PBMCs from a smoker with emphysema (29), was used to generate human EF-specific TCRs expressing transfectant thymoma cell line.

Total RNA from the TC-378-4 clone was extracted with TRIzol (no. 15596026; Thermo Fisher Scientific, Waltham, MA) following the manufacturer’s instructions. cDNA was synthesized with TCR-specific reverse transcription (RT) primers to increase specificity using the High-Capacity cDNA Reverse Transcription Kit (catalog no. 4368814; Applied Biosystems, Forster City, CA), 10× buffer, 25× dNTP mix, 50 U RT enzyme, 40 U RNase inhibitor, and 325 nM final concentration of each TCR-specific RT-PCR primer in a total 20-μl volume. Primer sequences were as follows: 5′-AGCTGGACCACAGCCG-3′ (TRAC cDNA), 5′-GAAATCCTTTCTCTTGACCATG-3′ (TRBC1 cDNA), and 5′-GCCTCTGGAATCCTTTCTCT-3′ (TRBC2 cDNA). The reaction mix was incubated at 25°C for 10 min, 45°C for 120 min, and 85°C for 5 min.

Following RT, a multiplex nested PCR was carried out in two rounds of PCR for TCR α- and TCR β-chain amplification separately with reverse-transcribed cDNA (50–200 ng). In the multiplex PCR (first-round PCR), the V region of TCR α- or TCR β-chain was amplified using a pool of multiplex V region primers (Vα-pool or Vβ-pool) along with an external constant region primer (TRACext or TRBCext). Details related to the design of TCR primer sequences have been published (31). Briefly, Vα-pool and Vβ-pool were generated with 44 and 40 primers to detect each V region of TCRα, and TCRβ, respectively. A total 25-μl PCR was prepared with 5× GoTaq buffer, 80 μM dNTPS, 3% DMSO, 2 μM primer pool (final concentration of each primer was 46 nM), 200 nM constant primer, 1 U GoTaq DNA polymerase (PAM8298; VWR), and 2.5 μl of RT reaction mixture. The PCR cycles were 2 min at 94°C, followed by 34 cycles of 20 s at 94°C, 30 s at 54°C, and 1 min at 72°C, followed by 7 min at 72°C.

For the nested PCR (second-round PCR), 2.5 μl of the first-round PCR was used as a template. Similarly, two separate PCRs were prepared for TCRα and TCRβ amplification in a final volume of 25 μl. Reaction mix contained 5× GoTaq buffer containing loading dye, 80 μM dNTPS, 3% DMSO, 200 nM primer adaptor primer (TRAVada or TRBVada), 200 nM internal constant primer (TRACint or TRBCint), and 1 U GoTaq DNA polymerase (PAM8298; VWR). The PCR cycles were 2 min at 94°C, followed by 34 cycles of 20 s at 94°C, 30 s at 56°C, and 1 min at 72°C, followed by 7 min at 72°C. Then, 5 μl of the final reaction was run on 1% agarose gel to confirm successful PCR, and the 20-μl remainder was purified for Sanger sequencing using a PCR purification kit (no. 11-305C; Zymo Research).

For Sanger sequencing, the TRAVada and TRBVada primers were used for TCRα and TCRβ, respectively. TCR sequences were analyzed with the ImMunoGeneTics (IMGT)/V-Quest tool (http://www.imgt.org/).

The primers used in the multiplex nested PCRs contain restriction sites for subsequent subcloning into the template retroviral vector pMSCVII-Ametrine (pMIA) to generate TCR expressing transfectant thymoma cell lines. The vector was modified to incorporate restriction enzyme cut sites for TCR cloning (32, 33).

Subcloning of the purified TCRα or TCRβ PCR products into pCR-Blunt II-TOPO vector (K280020; Thermo Fisher Scientific) were performed using the nested reaction products. To generate blunt ends, the PCR products were treated with DNA polymerase I Large (Klenow) (M220C; Promega) for 30 min at 37°C, followed by heat inactivation for 5 min at 70°C. Klenow-treated reaction products were ligated with pCR-Blunt II-TOPO vector at room temperature for 15 min as recommended by the manufacturer. TOP10 competent cells (K280020; Thermo Fisher Scientific) were used for transformations. The insertion of each TCR α- and TCR β-chain was confirmed by test digestion with BstBI (R0519; New England Biolabs [NEB]) and MfeI (R3589; NEB) for TCRα and SnaBI (R0130; NEB) and SacII (R0157; NEB) for TCRβ. Following confirmation of the correct insertion sites, Sanger sequencing was performed.

TCRβ containing pCR-Blunt II-TOPO vector was digested with BstBI and MfeI, and DNA was purified with a gel DNA purification kit (no. 11-300; Zymo Research). In parallel, the template retroviral vector pMIA was digested with the same restriction enzymes and gel-purified using a gel DNA purification kit. Before ligation, the digested template vector was treated with 10 U CIP enzyme (M0290; NEB) to prevent self-ligation for 1 h at 37°C, followed by heat inactivation for 10 min at 75°C, and then was purified with a PCR purification kit (no. 11-305C; Zymo Research). The ligation reaction was performed with the insert at 6 M excess to the vector and ∼150 ng of total DNA using DNA ligase for 30 min at room temperature. For transformations, DH5α competent cells (no. 18265017; Invitrogen, Carlsbad, CA) were used. The successful insertion of the TCR β-chain was confirmed by test digestion with the restriction enzymes and Sanger sequencing. For TCRα insertion into TCRβ-containing retroviral vector, a similar process was carried out with different restriction enzymes, SnaBI and SacII.

A retroviral vector was generated using 4 μg of human TCR-pMIA (vector generated described above), 4 μg of pEQ-Pam3(-E) (packaging vector), and 2 μg of pVSVg (envelope vector) as provided in Transit 293T transfection reagent (MIR2700; Mirus) for HEK293T cells. After 24 h, the media was replaced with 5% FCS complete DMEM, and 24 h later, supernatant from hTCR-transfected HEK293T cells was harvested and filtered through 0.45-μm filters. Mouse 4G4 thymoma cells expressing human CD4 (1 × 106) were suspended in 4 ml of supernatant from hTCR-transfected HEK293T cells mixed with polybrene at 6 μg/ml, placed in a six-well plate, and spun at 1800 rpm for 90 min at 20°C. The second round of transduction was repeated the next day as described above, and after an additional 24 h in culture, cells were placed in complete RPMI 1640 (C-RPMI) supplemented with 10% heat-inactivated FBS. C-RPMI was prepared by adding 2-ME (5 × 10−5 M), l-glutamine (100×), sodium pyruvate (100×), MEM nonessential amino acids (100×), MEM vitamin (100×), penicillin/streptomycin (100×), and gentamicin (10 μg/ml). The hTCR-transduced 4G4 cells were sorted by FACS for mouse CD3–Brilliant Violet (BV) 510+, hTCR–Ametrine+, and human CD4–GFPhigh cells representing stable and functional hTCR:mouse CD3 complexes (m:hTCR) on the cell surface. The m:hTCR-sorted cells were expanded for 3–5 d in a six-well plate, and cell purity (>95%) was confirmed using flow cytometry.

To identify HLA restriction, genomic DNA was isolated from 5 × 105 TC-378-4 clone using a DNA isolation kit (no. 69504; Qiagen) and used for high-resolution next-generation sequencing to identify HLA typing (Table I). Based on the HLA typing, we used the B-Lymphoblastoid cell line (no. 09420, purchased from International Histocompatibility Working Group, Seattle, WA) as APCs. m:hTCR-transduced 4G4 cells (1 × 104) were cultured with APCs (1 × 105) in the presence of 30 μg/ml hEFs (QP45; Elastin Products, Owensville, MO) for 4 d. For HLA blocking experiments, APCs were incubated with anti-DQ, anti-DR, or isotype control Abs (2.5 μg/ml) for 1 h at 37°C before coculture with m:hTCR cells. Mouse IL-2 production was measured in the supernatant using ELISA according to the manufacturer’s instructions (no. 555148; BD Biosciences, San Jose, CA).

C57/BL6 mice (7–8-wk-old females) purchased from The Jackson Laboratory were exposed to active smoke from commercial cigarettes (Marlboro 100 long; Marlboro) (15). In the cigarette smoke chamber exposure model, cigarettes are burned at a rate of 4–5 min/cigarette, which mimics the exposure to heated smoke. Smoke during each cycle is forced using 4 l of air/min into the exposure chamber intermittently; each smoke cycle is designed to mimic the puffing of actual human smokers and to prevent asphyxiation of the mice. Smoke cycles provide 5 s of heated cigarette smoke, which is interrupted by 25 s of 4 l of air/min using a timer-controlled two-way valve system (Humphrey, Kalamazoo, MI). The concentration of particulate matter and carbon monoxide during 30-min smoke cycles of one cigarette in the mouse smoke chamber resulted in mean total particulate concentration of 550 ± 50 mg/m3 (±SD), and the average CO concentration in parts per million (v/v) during for one 30-min smoke cycle experiment was 757 parts per million (34). Mice were exposed to four cigarettes 5 d/wk for a total of 6 mo. We used the number of cigarettes that approximated moderate to heavy smoking habits (e.g., >20 pack year smoke exposure) in humans.

C57BL/6 female mice (7–8 wk old) were purchased from The Jackson Laboratory. hEFs (no. QP45), rat lung alpha EFs (rEFs; product ID RA50), and mouse lung EFs (mEFs; no. MLP54) were purchased from Elastin Products. On day 1, mice were immunized using s.c. injections with a mixture of 25 μg of hEFs and 25 μg of rEFs or with 50 μg of mEFs in CFA (no. 263910; BD Biosciences) with 250 μg of Mycobacterium tuberculosis (no. 231141; BD Biosciences). After that, mice were immunized once a week by s.c. injection with a mixture of 25 μg of hEFs and 25 μg of rEFs or with 50 μg of mEFs in IFA for a total of 7 wk (see schema in Fig. 3).

For analysis of experimental emphysema, immunized mice were euthanized, and bronchoalveolar lavage (BAL) fluid, lungs, spleen, and aorta were collected. BAL fluid was collected by instilling and withdrawing 0.8 ml of sterile PBS twice through the trachea. Total and differential cell counts in the BAL fluid were determined with the standard hemocytometer and HEMA3 staining (Biochemical Sciences, Swedesboro, NJ) using 150 μl of BAL fluid prepared with cytospin slides. Dissected lung tissues, spleen, and aorta were used to prepare single-cell suspensions and/or histopathological studies using H&E staining as described previously (35).

To determine the severity of lung parenchymal destruction (emphysema), we used microcomputed tomography (microCT) and mean linear intercept (MLI) as previously described (34). The Animal Phenotyping Core at Baylor College of Medicine performed microCT studies on live, anesthetized mice. Briefly, mice were placed in microCT (Gamma Medica, Salem, NH), and completed images of the chest were used to quantitate emphysema using Amira 3.1.1. software (FEI, Hillsboro, OR) (35).

Quantification of emphysema using MLI was done on blinded samples using 10 fields that were randomly selected from the left lobe of lung. Parallel lines were placed on the selected field, and the number of intercepts was measured. MLI was calculated by dividing the length and the number of lines per field, multiplied by the number of intercepts.

Single-cell suspensions from thoracic aorta were prepared by enzymatic digestion as previously described, with minor modifications (36, 37). Briefly, thoracic aorta segments were dissected following vasculature perfusion with 2 mM EDTA buffer (5 ml), PBS (10 ml), and FACS buffer (10 ml), respectively. Thoracic aorta, the segment above the diaphragm, was cut into small pieces and digested in 3 ml of enzyme mixture containing 400 U/ml collagenase type I (no. C0130; Millipore Sigma), 120 U/ml collagenase type XI (no. C7657; Millipore Sigma), 60 U/ml hyaluronidase type I (no. H3506; Millipore Sigma), and 60 U/ml DNase I (no. 11284932001; Millipore Sigma) in RPMI supplemented with 10% heat-inactivated FBS (hiFBS) for 50 min at 37°C. Digested products in suspensions were filtered through a 40-μm cell strainer (BD Falcon, San Jose, CA), followed by RBC lysis (ACK lysis buffer; Millipore Sigma) for 3 min at room temperature.

Single-cell suspensions from mouse lung, draining lymph nodes, or spleen were prepared by mechanically mincing collected organs through a 40-μm strainer, followed by RBC lysis for 3 min at room temperature as described (38).

Cloning of EF-specific CD4+ T cells was accomplished by a serial dilution method as previously described, with minor modification (39). Live CD3+CD4+CD25 T cells were sorted using flow cytometry from single-cell suspensions of the mouse lung, lung draining lymph nodes, or spleen. The sorted cells were cocultured with gamma-irradiated (30 Gy [the international system unit of absorbed dose of ionizing radiation]) CD3 splenocytes isolated from CD45.1 congenic mice as APCs in a 1:2 ratio (T cells:APCs) in C-RPMI supplemented with 0.5% heat-inactivated mouse serum (SM-0100HI; Equitech-Bio) in the presence of mEFs (30 μg/ml). After 3 d, the culture medium was replaced with C-RPMI supplemented with 10% hiFBS. After 2 d, the cells were placed in C-RPMI supplemented with 5% hiFBS. Ficoll Pague Plus (GE Healthcare) gradient was used to remove dead cells by centrifugation at 500 × g for 20 min at room temperature. The collected live cells were placed in C-RPMI supplemented with 10% hiFBS and 10% T cell culture supplement (TCCS) without conA (no. 354116; BD Biosciences). After 3 d, the culture medium was replaced with C-RPMI supplemented with 10% TCCS and incubated for 2 d. All steps from coculture with APCs in the presence of mEFs were repeated for two additional rounds. After the third round of stimulation with mEFs, T cells were diluted in C-RPMI supplemented with 10% hiFBS and 10% TCCS at serial concentrations in a 96-well plate at concentrations of one cell/100 μl and 10 cells/100 μl. APCs (7 × 105 cells/100 μl) were added to each well in the presence of mEFs (final concentration 30 μg/ml). The medium was replaced every 3 d for 2 wk, and at the same time, T cells were restimulated by adding 7 × 105 cells/100 μl of APCs in the presence of mEFs (30 μg/ml) until the growth of the cloned T cells becomes visible.

Mouse T cell clones reactive to mEFs were used to identify mouse elastin-specific TCRs. RNA from T cell clone was extracted with TRIzol (no. 15596026; Thermo Fisher Scientific) following the manufacturer’s instructions. RNA from each T cell clone was reverse-transcribed using a High-Capacity cDNA Reverse Transcription Kit (catalog no. 4368814; Thermo Fisher Scientific) and mouse TCR-specific RT primers to increase specificity, 10× buffer, 25× dNTP mix, 50 U RT enzyme, and 20 U RNase inhibitor, and 26 μM was used as the final concentration of each TCR-specific RT-PCR primer in a 20-μl volume. Primer sequences were as follows: 5′-CTCAGCGTCATGAGCAGG-3′ (TRAC cDNA), 5′-CCATAGCCATCACCACCAG -3′ (TRBC1 cDNA), and 5′-CCATGGCCATCAGCACTAG -3′ (TRBC2 cDNA). The reaction mix was incubated at 25°C for 10 min, 45°C for 45 min, and 85°C for 5 min.

Following RT, we used the multiplex nested PCR method designed with minor modifications (40) in two rounds of amplification. The first round of PCR was performed with oligonucleotide mixture of 23 TRAV (each 363 nM final concentration), 19 TRBV forward (each 363 nM final concentration), and single TRAC (3.3 μM final concentration) and TRBC (3.3 μM final concentration) reverse primers in total volume of 25 μl containing 5× GoTaq buffer, 80 μM dNTPS, 0.5 U GoTaq DNA polymerase (PAM8298; VWR), and 2.5 μl of RT reaction product. The PCR cycles were 5 min at 94°C, followed by 36 cycles of 30 s at 94°C, 30 s at 55°C, and 1 min at 72°C, followed by 7 min at 72°C.

For the second round of amplification, 2.5 μl of the first PCR product was used as a template for amplification of TCRα and TCRβ in two separate reactions using a corresponding internal sequence primer mix. The reaction components were similar to the first-round PCR, using a mixture of 23 TRAV forward primers and single TRAC reverse for TCRα and 19 TRBV forward and single TRBC reverse for TCRβ. The PCR cycles were 5 min at 94°C, followed by 36 cycles of 30 s at 94°C, 30 s at 55°C, and 1 min at 72°C, followed by 7 min at 72°C.

The PCR products were visualized on a 1% agarose gel and purified using a PCR purification kit (K220001; Thermo Fisher Scientific). The purified products were sequenced using TRAC or TRBC internal reverse primers for α and β PCR products, respectively. TCR sequences were analyzed with the IMGT/V-Quest tool (http://www.imgt.org/).

CD4+ T cells from the lung or spleen of air- or cigarette smoke–exposed mice were isolated using mouse anti-CD4–conjugated MicroBeads (no. 130-117-043; Miltenyi Biotec), followed by AutoMACS (Miltenyi Biotec) positive selection. Single cells from the negative selection were used to isolate CD11c+ cells for APCs by positive selection with anti-CD11c–conjugated MicroBeads (no. 130-108-338; Miltenyi Biotec). Isolated CD4+ T cells (0.5 × 106) were cocultured with gamma-irradiated APCs (0.5 × 105) in a 10:1 ratio for 3 d with or without mEFs (30 μg/ml). After 3 d of coculture, supernatants were collected and used to measure concentrations of cytokines IFN-γ and IL-17A by Luminex.

T cell clone autoreactivity was measured using coculture with gamma-irradiated CD3 splenocyte APCs isolated using MicroBeads (no. 130-094-973; Miltenyi Biotec) with or without mEFs (30 μg/ml). T cell responses to mEFs were recorded as fold change (cytokines detected in the presence of mEFs divided by cytokines measured in the absence of mEFs) (29).

Flow cytometry was performed using a BD LSR II (BD Biosciences), and data were analyzed with FlowJo software (Tree Star, Ashland, OR). UV live/dead fixable dead cell stain (Thermo Fisher Scientific) was used to exclude dead cells. For intracellular cytokine staining of IFN-γ and IL-17A, cells were stimulated in 10% FBS containing RPMI with PMA (10 ng/ml; Sigma-Aldrich, St.Louis, MO), ionomycin (1 μg/ml; Sigma-Aldrich), and brefeldin A (10 μg/ml; Sigma-Aldrich) overnight. Cells were stained with following anti-mouse Abs: Pacific blue–conjugated anti-CD3e (500A2), PE–Cy5–conjugated anti-CD4 (RM4-5), and allophycocyanin–Cy7–conjugated anti-CD8a (53-6.7) purchased from eBioscience (San Diego, CA). Cells were then fixed with FACS lysing solution (BD Biosciences), permeabilized, and stained with PE-conjugated anti–IL-17A (TC11-18H10; Thermo Fisher Scientific) and allophycocyanin-conjugated anti–IFN-γ (XMG1.2; Thermo Fisher Scientific). For transcription factor analysis, cells were stained with PE-RORγt (BD2; Thermo Fisher Scientific), PE–Cy7–T-bet (4B10; Thermo Fisher Scientific), Alexa Fluor 647–Foxp3 (FJK-16s; Thermo Fisher Scientific), and BV421–Gata3 (16E10A23; BioLegend) according to the manufacturer’s protocol (Thermo Fisher Scientific). For neutrophil and conventional dendritic cell analyses, the following Abs were used; eFluro450-B220 (RA3-6B2; Thermo Fisher Scientific), PE-CD11b (M1/70; BioLegend), allophycocyanin–CD11c (N418; Thermo Fisher Scientific), BUV395-CD45 (BD30-F11; BD Biosciences,), or BV510–Ly6G (1A8; BD Biosciences).

Total RNA from thoracic aorta tissue treated with RNAlater RNA stabilization reagent (no. 1018087; Qiagen) was extracted using RNeasy fibrous tissue isolation kit (no. 74704; Qiagen) following the manufacturer’s instructions. Total RNA from mouse BAL cells was extracted with TRIzol (Invitrogen) following the manufacturer’s instructions. cDNA was synthesized with the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Forster City, CA). Gene probes, MMP9 (Mm00600164_g1), and MMP12 (Mm00500554_m1) were purchased from Thermo Fisher Scientific. All data were normalized to 18S rRNA (Hs99999901_s1) expression.

Statistical analyses were performed using GraphPad Prism version 8 for Mac OS X (GraphPad Software, La Jolla, CA). All data shown in the figures are the mean ± SEM. We used one-way ANOVA with a Bonferroni correction for multiple comparisons or the Student t test using two-tailed parameters.

We have previously identified and cloned several elastin-specific CD4+ T cells from PBMCs isolated from smokers with emphysema (29). Elastin-specific T cell clones and bulk T cells isolated from PBMCs in smokers showed specificity for hEFs, as determined by increased cytokine secretion and a requirement for MHC class II recognition (2729). To determine the optimal conditions that could be used to develop a novel elastin-induced model of emphysema, we sequenced TCR α- and TCR β-chains of one the human T cell clones, TC-378-4, using multiplex nested PCR and designed primers complementary to all human TCRα and TCRβ V regions as previously described (31) (Fig. 1A–C). We next subcloned the purified human TCRα and TCRβ PCR products to a retroviral vector to generate human elastin-specific TCR by transducing murine surface TCR–deficient 4G4 thymoma cells with retroviral vector expressing elastin-specific hTCR, which we labeled as m:hTCR (Fig. 1D, Table I).

FIGURE 1.

Workflow schematic for human EF-specific TCR identification and the results. (A) Reverse-transcribe mRNA from human EF-specific T cell clone (TC-378-4) isolated from emphysema patient using TCR-specific primers. (B) Detail of the multiplex nested PCR (two rounds of PCR) to amplify hTCR α- and β-chains (left). The amplified PCR products on 1% agarose gel (right). (C) Sequencing result. (D) Construct map after hTCR α- and β-chain serial insertion into a retroviral vector to generate hTCR-expressing 4G4 cells (m:hTCR cells; left). Representative flow cytometry showing successful expression of the hTCR on the cell surface of a transduced 4G4 thymoma cell line gated on Ametrine (right). (E) HLA restriction assay with HLA blocking Abs (DQ/DR) or isotype control Ab (2.5 μg/ml). HLA-matched APCs were pretreated with blocking Ab for 1 h at 37°C and cultured with m:hTCR cells for 4 d in the presence of hEFs (30 μg/ml). **p < 0.01, ***p < 0.001, as determined by one-way ANOVA test with Bonferroni correction for multiple comparisons.

FIGURE 1.

Workflow schematic for human EF-specific TCR identification and the results. (A) Reverse-transcribe mRNA from human EF-specific T cell clone (TC-378-4) isolated from emphysema patient using TCR-specific primers. (B) Detail of the multiplex nested PCR (two rounds of PCR) to amplify hTCR α- and β-chains (left). The amplified PCR products on 1% agarose gel (right). (C) Sequencing result. (D) Construct map after hTCR α- and β-chain serial insertion into a retroviral vector to generate hTCR-expressing 4G4 cells (m:hTCR cells; left). Representative flow cytometry showing successful expression of the hTCR on the cell surface of a transduced 4G4 thymoma cell line gated on Ametrine (right). (E) HLA restriction assay with HLA blocking Abs (DQ/DR) or isotype control Ab (2.5 μg/ml). HLA-matched APCs were pretreated with blocking Ab for 1 h at 37°C and cultured with m:hTCR cells for 4 d in the presence of hEFs (30 μg/ml). **p < 0.01, ***p < 0.001, as determined by one-way ANOVA test with Bonferroni correction for multiple comparisons.

Close modal
Table I.
HLA typing result
DRB1
DRB3DRB4DQB1
DQA1
07:01 13:03 01:01 01:03 02:02 03:01 02:01 05:05 
DRB1
DRB3DRB4DQB1
DQA1
07:01 13:03 01:01 01:03 02:02 03:01 02:01 05:05 

Next, we deep-sequenced the TC-378-4 clone to identify the endogenous MHC class II molecules required for activation of m:hTCR in functional assays (Table II). Coculture of m:hTCR cells with endogenous MHC class II APCs resulted in increased IL-2 production when compared with control (Fig. 1E). Furthermore, blocking HLA-DQ molecule with the anti-DQ Ab, but not anti-DR or isotype control, attenuated IL-2 production in m:hTCR transfectant cells stimulated with hEFs. These findings suggested that I-A molecule expressed in C57BL/6J strain, a mouse counterpart of human DQ (41), is required to present EFs by APCs in mice (Fig. 1E). Therefore, we used the C57BL/6J strain to develop a novel Ag-induced model of emphysema.

Table II.
Identified TCRs of mouse elastin-specific T cell clones
OrganClone No.TCRαTCRβ
TRAVTRAJCDR3αTRBVTRBJTRBDCDR3β
Lung A5 5-4 or 5D-4 52 CAASDTNTGANTGKLTF 19 2-3 CASSRGQGHAETLYF 
C5 14D-1 43 CAASWDNNAPRF 15 2-4 CASSVDWGNTLYF 
E1 17 58 CALEGQGTGXKLSF 13-2 2-5 CASGATGGADDTQ 
G1 12-2 or 12D-1 23 CALSDQNYNQGKLIF 2-5 CASSPDWDGDTQYF 
G2 14D-1 TVFWVKTQVVGQLTF 1-2 CASSQEMDSDYTF 
H4 10 47 CAARSKDYANKMIF 31 2-3 CAISSAETLYF 
Spleen A6 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
B3 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
B4 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
D5 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
E1 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
F3 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
H4 6-5 or 6D-5 16 CAXSEPSSGQKLVF 13-2 2-1 CASGDTGGYAEQFF 
OrganClone No.TCRαTCRβ
TRAVTRAJCDR3αTRBVTRBJTRBDCDR3β
Lung A5 5-4 or 5D-4 52 CAASDTNTGANTGKLTF 19 2-3 CASSRGQGHAETLYF 
C5 14D-1 43 CAASWDNNAPRF 15 2-4 CASSVDWGNTLYF 
E1 17 58 CALEGQGTGXKLSF 13-2 2-5 CASGATGGADDTQ 
G1 12-2 or 12D-1 23 CALSDQNYNQGKLIF 2-5 CASSPDWDGDTQYF 
G2 14D-1 TVFWVKTQVVGQLTF 1-2 CASSQEMDSDYTF 
H4 10 47 CAARSKDYANKMIF 31 2-3 CAISSAETLYF 
Spleen A6 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
B3 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
B4 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
D5 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
E1 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
F3 6-6 or 6D-6 34 CALLSSNTNKVVF 31 2-5 CAWSLGGAHQDTQYF 
H4 6-5 or 6D-5 16 CAXSEPSSGQKLVF 13-2 2-1 CASGDTGGYAEQFF 

TCR V, D, and J region nomenclature follows that of IMGT.

We have previously shown that CD4+ T cells isolated from the PBMCs of smokers with emphysema differentiate to Th1 and Th17 cells in response to hEFs, indicating a strong association between anti-elastin autoimmunity and emphysema in smokers (27). Although the cigarette smoke–induced model of emphysema induces robust Th1- and Th17-mediated lung inflammation (15, 34), anti-elastin autoimmunity in this model had not been examined. To determine whether cigarette smoke induces elastin-specific T cells, we examined mEF-specific autoreactive T cells in C57BL/6J mice exposed to 6 mo of cigarette smoke. Compared with air-exposed control mice, CD4+ T cells from emphysematous lungs exposed to cigarette smoke reacted to mEFs, as determined by increased expression of IFN-γ (Fig. 2A), indicating similar autoreactive immune responses to those detected in the lungs of smokers with emphysema (28, 29). However, mEFs failed to induce IL-17A in CD4+ T cells under the same conditions (Fig. 2B).

FIGURE 2.

mEF-specific autoreactive CD4+ T cells from cigarette smoke–exposed mice. Lung CD4+ T cells of air- or cigarette smoke–exposed mice were cultured with gamma-irradiated CD11c+ cells with or without mouse EFs (30 μg/ml) for 3 d. After stimulation, the supernatants were harvested and used for the cytokine measurement. The concentrations of IFN-γ (A) and IL-17A (B) were plotted as fold change over nil stimulation. Each dot represents a data point from an individual mouse. Data are from two independent experiments. *p < 0.05, as determined by the Student t test.

FIGURE 2.

mEF-specific autoreactive CD4+ T cells from cigarette smoke–exposed mice. Lung CD4+ T cells of air- or cigarette smoke–exposed mice were cultured with gamma-irradiated CD11c+ cells with or without mouse EFs (30 μg/ml) for 3 d. After stimulation, the supernatants were harvested and used for the cytokine measurement. The concentrations of IFN-γ (A) and IL-17A (B) were plotted as fold change over nil stimulation. Each dot represents a data point from an individual mouse. Data are from two independent experiments. *p < 0.05, as determined by the Student t test.

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Our findings so far indicate that cigarette smoke exposure induces Th1-specific loss of tolerance against elastin in C57BL/6J mice but also highlight differences between animal models and human emphysema, with the former occurring in the absence of respiratory infection, whereas the latter is often complicated with repeated bouts of microbial infections (4244).

Therefore, to address the role of autoimmunity in the pathogenesis of emphysema, we immunized mice using EFs. To accomplish this goal, we first chose C57BL/6J mice to immunize against mEFs isolated from the lungs of congenic mice. We found that repeated immunization with mEFs isolated from the lungs of C57BL/6J mice did not result in the induction of elastin-specific T cells, lung inflammation, or emphysema (Supplemental Fig. 1). We reasoned that immunization with a mixture of hEFs and rEFs (h+rEFs), which have 72.6 and 91% homology with mEFs, respectively (45), could induce autoimmunity in mice. Therefore, we next immunized with h+rEFs using the same protocol (Fig. 3A). We found that immunization with h+rEFs resulted in emphysema, as determined by increased total lung volume quantified by microCT (Fig. 3B, 3C) and enlarged alveolar spaces detected by H&E staining of lung sections (Fig. 3D). Unbiased lung morphometry measurement (MLI) (Fig. 3E) also showed emphysema in h+rEF-immunized mice when compared with control (PBS-immunized) mice. Examination of lung inflammatory cells showed significantly increased numbers of macrophages, lymphocytes, and neutrophils (Fig. 3F–H) that were present in the BAL fluid compared with control mice. Increased lung inflammation and emphysema was consistent with increased expression of MMP9 and MMP12 mRNA expression (Fig. 3I, 3J).

FIGURE 3.

Non-self EF immunization–induced emphysema mouse model. (A) Schematic diagram of EF immunization. Mice were immunized with 25 μg hEFs and 25 μg rEFs in CFA with 250 μg M. tuberculosis by s.c. injection on day 1. Mice were then repeatedly immunized once a week (a total of seven times) with 25 μg hEFs and 25 μg rEFs (h+rEFs) in IFA. Mice were analyzed 1 wk after the last injection. (B) Representative three-dimensional images and (C) microCT quantification of lung volume from PBS- or mixture of h+rEFs–immunized mice. Average lung volume is shown below (B). (D) Representative images of H&E-stained lung sections; insets represent original magnification ×200. Scale bars, 100 μm. (E) MLI measurement from the indicated groups of mice. BAL fluid analyses from the same group of mice showing total (F), macrophages (Mac) (F), lymphocytes (lymph) (G), and neutrophils (neut) (H). Expression of MMP9 (I) and MMP12 (J) in BAL cells isolated from PBS- or mixture of h+rEFs–immunized mice. Data were combined with two different experiments. Results are represented as mean ± SEM from three independent experiments with four to five mice in each group. *p < 0.05, **p < 0.01, as determined by the Student t test.

FIGURE 3.

Non-self EF immunization–induced emphysema mouse model. (A) Schematic diagram of EF immunization. Mice were immunized with 25 μg hEFs and 25 μg rEFs in CFA with 250 μg M. tuberculosis by s.c. injection on day 1. Mice were then repeatedly immunized once a week (a total of seven times) with 25 μg hEFs and 25 μg rEFs (h+rEFs) in IFA. Mice were analyzed 1 wk after the last injection. (B) Representative three-dimensional images and (C) microCT quantification of lung volume from PBS- or mixture of h+rEFs–immunized mice. Average lung volume is shown below (B). (D) Representative images of H&E-stained lung sections; insets represent original magnification ×200. Scale bars, 100 μm. (E) MLI measurement from the indicated groups of mice. BAL fluid analyses from the same group of mice showing total (F), macrophages (Mac) (F), lymphocytes (lymph) (G), and neutrophils (neut) (H). Expression of MMP9 (I) and MMP12 (J) in BAL cells isolated from PBS- or mixture of h+rEFs–immunized mice. Data were combined with two different experiments. Results are represented as mean ± SEM from three independent experiments with four to five mice in each group. *p < 0.05, **p < 0.01, as determined by the Student t test.

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We next determined whether h+rEF immunization induces innate and adaptive inflammation in the lung parenchyma. Single-cell examination of lungs from h+rEF-immunized mice showed increased relative abundance of lung CD11c+CD11bHigh conventional dendritic cells (Fig. 4A, 4B) and Ly6G+CD11b+ neutrophils (Fig. 4C, 4D) when compared with control (PBS-immunized) mice. We next examined whether, in the same immunized mice, there is an induction of adaptive immune cells. Intracellular cytokine staining of CD4+ cells showed increased expression of IL-17A (Th17)– and IFN-γ (Th1)–producing cells in the lungs of h+rEF-immunized mice compared with controls (Fig. 4E–G). We also found an increased relative abundance of CD8+ CTLs expressing IL-17 and CD8+ CTLs expressing IFN-γ T cells (Fig. 4H, 4I) in the same group. Notably, mice immunized with hEFs alone failed to develop lung inflammation or emphysema (Supplemental Fig. 2). Although immunization with rEFs alone resulted in a mild increase in BAL cell numbers, enlarged alveolar space, and increased Th17 cells in the lung (Supplemental Fig. 3), the changes were less robust when compared with the combined h+rEF immunization protocol (Figs. 3, 4).

FIGURE 4.

Non-self EF immunization induces inflammation in the lung. Representative (A) and cumulative (B) flow cytometry analyses of B220CD11c+CD11bHigh mDCs in the lung of PBS- or mixture of h+rEFs–immunized mice. Representative (C) and cumulative (D) flow cytometry analyses of CD45+Ly6G+CD11b+ neutrophils (Neut) in the lungs of the same group of mice. Representative intracellular staining (E) and cumulative analysis of IL-17A (Th17)– (F) and IFN-γ (Th1)–expressing (G) CD4+ T cell subsets in the lung. Cumulative intracellular cytokine staining of IL-17A (Tc17)– (H) and IFN-γ (Tc1)–expressing (I) CD8+ T cell subsets in the lung. Results are represented as mean ± SEM from three independent experiments with five mice in each group. *p < 0.05, **p < 0.01, as determined by the Student t test.

FIGURE 4.

Non-self EF immunization induces inflammation in the lung. Representative (A) and cumulative (B) flow cytometry analyses of B220CD11c+CD11bHigh mDCs in the lung of PBS- or mixture of h+rEFs–immunized mice. Representative (C) and cumulative (D) flow cytometry analyses of CD45+Ly6G+CD11b+ neutrophils (Neut) in the lungs of the same group of mice. Representative intracellular staining (E) and cumulative analysis of IL-17A (Th17)– (F) and IFN-γ (Th1)–expressing (G) CD4+ T cell subsets in the lung. Cumulative intracellular cytokine staining of IL-17A (Tc17)– (H) and IFN-γ (Tc1)–expressing (I) CD8+ T cell subsets in the lung. Results are represented as mean ± SEM from three independent experiments with five mice in each group. *p < 0.05, **p < 0.01, as determined by the Student t test.

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We next explored whether h+rEF immunization induced inflammation in other elastin-rich organs (e.g., aorta). Consistent with Th1 and Th17 lung inflammation, we found increased relative abundance of RORγt+ T (Th17)– and T-bet+ T (Th1)–expressing T cells in the thoracic aorta (Fig. 5A, 5B). Furthermore, we found increased expression of Mmp9 and Mmp12 in the same tissue, indicating organ-specific effects of inflammation on the affected organ (Fig. 5C, 5D). h+rEF-immunized mice also showed increased Th1 and Th17 cells in the spleen, indicating the systemic impact of immunization in this model (data not shown). Together, these findings strongly support that immunization with non-self EFs (e.g., h+rEFs), but not mEFs, induces systemic and organ-specific inflammation in mice.

FIGURE 5.

Non-self EF immunization induces inflammation in the thoracic aorta. (A) Representative staining of transcription factors (Foxp3 and RORγt) and cumulative analysis of RORγt-expressing CD4+ T cells (Th17) in the thoracic aorta from PBS- or mixture of h+rEFs–immunized mice. (B) Representative staining of transcription factors (T-bet and GATA3) and cumulative analysis of T-bet–expressing CD4+ T cells (Th1) in the thoracic aorta from the same group of mice. Expression of MMP9 (C) and MMP12 (D) in the thoracic aorta isolated from the same group of mice. Results are represented as mean ± SEM from two independent experiments with five mice in each group. *p < 0.05, as determined by the Student t test.

FIGURE 5.

Non-self EF immunization induces inflammation in the thoracic aorta. (A) Representative staining of transcription factors (Foxp3 and RORγt) and cumulative analysis of RORγt-expressing CD4+ T cells (Th17) in the thoracic aorta from PBS- or mixture of h+rEFs–immunized mice. (B) Representative staining of transcription factors (T-bet and GATA3) and cumulative analysis of T-bet–expressing CD4+ T cells (Th1) in the thoracic aorta from the same group of mice. Expression of MMP9 (C) and MMP12 (D) in the thoracic aorta isolated from the same group of mice. Results are represented as mean ± SEM from two independent experiments with five mice in each group. *p < 0.05, as determined by the Student t test.

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Our findings thus far provided evidence that immunization with r+hEFs but not mEFs induces systemic innate and adaptive immune responses in mice. We next isolated CD4+ T cells from the lung and spleen of h+rEF-immunized mice and cocultured them with irradiated CD11c+ APCs isolated from the spleen of congenic mice (Supplemental Fig. 4A). After initial stimulation with mEFs, T cell cultures were subjected to three additional rounds of stimulus to enrich for mEF-specific T cell clones. Before each series of stimulation, dead cells were removed, and live T cells were quantified. As expected, T cells expanded in response to mEFs, suggesting that h+rEF immunization results in the induction of autoreactive T cells in mice (Fig. 6).

FIGURE 6.

Proliferation of autoreactive T cells in response to mEFs. Expansion of mEF-specific CD4+ T cells from the lung (A) and spleen (B) of control (PBS) or h+rEF-immunized mice was monitored using a hemocytometer prior to serial dilution and clonal expansion. Cells were stimulated with mEFs (30 μg/ml) on days 0, 14, and 28. Dead cells were removed on days 5, 22, and 34. Serial dilution and clonal expansion were done on day 34.

FIGURE 6.

Proliferation of autoreactive T cells in response to mEFs. Expansion of mEF-specific CD4+ T cells from the lung (A) and spleen (B) of control (PBS) or h+rEF-immunized mice was monitored using a hemocytometer prior to serial dilution and clonal expansion. Cells were stimulated with mEFs (30 μg/ml) on days 0, 14, and 28. Dead cells were removed on days 5, 22, and 34. Serial dilution and clonal expansion were done on day 34.

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Next, the Ag-enriched T cells were used for T cell cloning by serial dilution, which resulted in multiple clones (Supplemental Fig. 4B). To identify the mEF-specific TCR, each T cell clone was expanded and subjected to multiplex nested PCR and TCR sequencing (Supplemental Fig. 4C, 4D). We identified two unique TCRs from a total of seven T cell clones generated from splenocytes of immunized mice and six unique TCRs from a total of six T cell clones isolated from the lung cells (Table II). Finally, we validated whether T cell clones were reactive against mEFs. Compared with CD4+ T cells isolated from control mice, T cells cloned from spleen and lung of r+hEF mice stimulated with mEFs exhibited increased IFN-γ secretion (Fig. 7A, 7C).

FIGURE 7.

Responses of T cell clones from the lung and the spleen against mEFs. Each mEF-specific T cell clone generated from (A and B) the lung (total of six different T cell clones) and (C and D) the spleen (a total of seven different T cell clones) was cultured with gamma-irradiated CD3 splenocytes with or without mEFs (30 μg/ml) for 3 d. CD4+ T cells isolated from splenocytes were used as control (Ctrl; n = 4). After coculture, the supernatants were harvested and used for the measurement of cytokine concentrations. The concentrations of IFN-γ (A and C) and IL-17A (B and D) were plotted as fold change over nil stimulation. Each dot represents a data point from an individual T cell clone. Results are mean ± SEM. *p < 0.05, as determined by the Student t test.

FIGURE 7.

Responses of T cell clones from the lung and the spleen against mEFs. Each mEF-specific T cell clone generated from (A and B) the lung (total of six different T cell clones) and (C and D) the spleen (a total of seven different T cell clones) was cultured with gamma-irradiated CD3 splenocytes with or without mEFs (30 μg/ml) for 3 d. CD4+ T cells isolated from splenocytes were used as control (Ctrl; n = 4). After coculture, the supernatants were harvested and used for the measurement of cytokine concentrations. The concentrations of IFN-γ (A and C) and IL-17A (B and D) were plotted as fold change over nil stimulation. Each dot represents a data point from an individual T cell clone. Results are mean ± SEM. *p < 0.05, as determined by the Student t test.

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In this study, we provide an Ag-specific model of emphysema as an efficient tool that could be used to determine the pathophysiological role of adaptive immunity in cigarette smoke–induced lung disease. This model recapitulates smoke-induced lung disease in humans because we showed loss of immune tolerance to elastin, induction of autoreactive T cells from systemic and elastin-rich organs, and pathophysiological changes in the lungs consistent with emphysema. Although instillation of elastase in the lungs has been used as a non-smoke model of emphysema, to our knowledge, immunization with EFs has not been shown to induce emphysema in mice.

Autoreactive T cells against elastin have been cloned from peripheral blood of smokers with emphysema and shown to secrete several proinflammatory cytokines in response to their cognate Ags (46). Although human association studies have provided strong support for cigarette smoke–mediated induction of autoimmunity as a potential mechanism for emphysema, whether loss of tolerance to elastin, independent of cigarette smoke, could cause emphysema has not been previously shown. This report provides the methods employed that successfully broke tolerance to EFs and resulted in the generation of autoimmune inflammation in multiple elastin-rich organs in C57BL/6J mice. We show in this study that immunization with mouse EFs failed to break tolerance to elastin; however, a combination of human and rat EFs was sufficient to induce autoimmunity in mice.

The elastin gene, ELN, contains a higher number of repeated elements than the rest of the genome, whereas an alignment report of elastin DNA sequences in vertebrates shows a little over 64% identity at the nucleotide level and 72% similarity at the amino acid level between human and mouse (45). Interestingly, the similarity between rat and mouse at the amino acid level is reported at 91%, indicating substantial similarities between rodents, which diverge from other mammals (45). Nonetheless, immunization of mice with mEFs failed to induce lung inflammation and emphysema. Prior studies have shown that immunization with unaltered (endogenous) proteins promotes Ag-specific regulatory T cells (Tregs) (47, 48); similarly, autoantigen-specific T cells against apolipoprotein B detected in healthy cohorts were shown to be phenotypically biased toward Tregs (49). However, whether such regulatory cells are present in elastin-mediated autoimmunity remains unknown and would be of importance to examine in the future studies.

Other autoimmune models have used human protein (e.g., myelin oligodendrocyte glycoprotein) to induce T cell–mediated autoimmunity in experimental autoimmune encephalomyelitis in mice, a model for neuronal degeneration that mimics encephalopathy seen in patients with multiple sclerosis (50). The mechanism by which combined h+rEFs can promote stronger autoimmune responses could be related to the strength and diversity of homologous Ags presented to T cells. Prior studies have established that sequence homology between self- and xeno-derived peptides could be the underlying mechanism resulting in cross-activation of autoreactive T cells (51). Specifically, the similarity between foreign and self-elastin peptides (human 71% and rat 91% homology with mouse elastin, respectively) can expose T cells to multiple cryptic epitopes to promote epitope spreading and/or induce molecular mimicry to expand autoreactive T cells. Notably, immunization with xenomolecules (human collagen peptides, etc.) can promote tissue-specific autoimmunity in mice (52), but to our knowledge, development of immunity to mouse elastin induced by h+rEF molecules has not been previously reported. The introduction of this breakthrough model opens a new paradigm and allows investigations into specific roles of TCR-specific pathogenic T and B lymphocytes that promote loss of immune tolerance in emphysema. A caveat of the current study is the time course whereby the persistence of autoreactive T cells remains to be clarified. However, several well-established mouse models of autoimmune diseases (e.g., experimental autoimmune encephalomyelitis, arthritis, colitis, etc.) result in a transient loss of immune tolerance (5355); despite their transient nature, these models have provided invaluable new information to our understanding of the pathophysiology of autoimmunity. Future studies should include an exact time course to examine whether and how autoimmunity can persist in this new model of emphysema.

Induction of autoimmune diseases requires multiple known and elusive etiologic factors that include environmental triggers, genetic predisposition, and threshold variability for autoimmunity in different organs (52). Consistently, development of emphysema in smokers is highly variable, with some estimates approximating 25% of all smokers (5658), indicating that a strong genetic susceptibility factor contributes to this disease. These findings highlight the need for an animal model that could be used to determine the stability of TCR/HLA/peptide trimolecular complex to understand the pathobiology of emphysema.

Cigarette smoke represents one of the most critical environmental triggers associated with the induction of autoimmune inflammation in humans, including rheumatoid arthritis, multiple sclerosis, and, as we have shown, emphysema (15, 5961). Multiple animal models of chronic exposure to cigarette smoke have been developed to study the acute and chronic inflammatory recruitment of immune cells in the lung (62, 63). However, a cigarette smoke–induced emphysema mouse model has some limitations to study anti-elastin autoimmunity because cigarette smoke contains over 5000 chemicals, which can strongly activate innate immune cells producing elastolytic enzymes, resulting in parenchymal destruction like emphysema, even in the absence of adaptive immune cells (64). As such, the new model could provide the necessary tool to determine factors that allow the loss of immune tolerance to self.

A second critical factor in the development of autoimmunity in humans is the contribution of the HLA polymorphism. For instance, expression of DQB1*03:01 in rheumatoid arthritis patients has been found to be significantly associated with an increased incidence of chronic obstructive pulmonary disease and emphysema (65, 66). Using EF-specific autoreactive T cell clones generated from emphysema patients, we found that HLA-DQ (DQA1*02:01 and 05:05, DQB1*02:02 and *03:01) corresponding to I-A molecule of MHC class II is required to present EFs in mice (Fig. 1, Table I).

The s.c. immunization of human and rat EFs induced organ-specific inflammation (e.g., lungs and aorta) resembling smoke-induced inflammatory disease found in susceptible smokers (27, 28). Specifically, immunized mice developed inflammatory responses in the lung, characterized by increased innate immune cells (mDCs and neutrophils) and infiltration of adaptive immune cells (T cells) as well as lung parenchymal destruction. Moreover, for the first time, to our knowledge, we generated EF-specific autoreactive T cell clones from proliferating cells in response to mEFs, supporting the loss of tolerance against self-antigen.

Using this model, we have cloned multiple elastin-specific T cells with potential pathogenic autoimmune responses. TCR plays an essential role in the regulation of immunological tolerance. Specifically, TCR affinity induces the differentiation of T cell lineages such as Tregs and removes autoreactive T cells during thymic selection. The highly variable CDR3 region of TCR provides antigenic specificities (67), a tool that could be used to characterize the dynamics of pathogenic T cell responses in autoimmunity in this disease model. In this study, we identified several CDR3 regions of TCR using autoreactive T cell clones against mEFs. Future studies will characterize pathogenic TCRs that could play a critical role in the induction of inflammation in the lung and emphysema.

In conclusion, we provide the methods for development of the first (to our knowledge) mouse model of anti-elastin autoimmunity that results in emphysema. This model could be used to examine the critical genetic and environmental factors that reduce susceptibility to loss of immune tolerance to self-proteins. Furthermore, the roadmap to generate m:hTCR-expressing cell lines could be used to develop retrogenic models to identify pathogenic autoreactive T cells in future studies. Similarly, future studies could determine the tolerogenic epitopes that could induce and expand Ag-specific Tregs to maintain the balance between inflammation and tolerance for immune homeostasis.

This work was supported by the expert assistance of Joel M. Sederstrom.

This work was supported by National Institutes of Health R01 AI135803-01 (to F.K. and D.B.C.), VA Merit CX000104 and R01 ES029442-01 (to F.K.), HL140398 (to D.B.C.), and National Institutes of Health R01 AI125301 (to M.B.), and by the Cytometry and Cell Sorting Core at Baylor College of Medicine with funding from the National Institutes of Health (AI036211, CA125123, and RR024574).

The online version of this article contains supplemental material.

Abbreviations used in this article:

     
  • BAL

    bronchoalveolar lavage

  •  
  • BV

    Brilliant Violet

  •  
  • C-RPMI

    complete RPMI

  •  
  • EF

    elastin fragment

  •  
  • hEF

    human lung EF

  •  
  • hiFBS

    heat-inactivated FBS

  •  
  • h+rEF

    hEF and rEF

  •  
  • IMGT

    ImMunoGeneTics

  •  
  • mEF

    mouse lung EF

  •  
  • m:hTCR

    hTCR:mouse CD3 complex

  •  
  • microCT

    microcomputed tomography

  •  
  • MLI

    mean linear intercept

  •  
  • MMP

    matrix metalloproteinase

  •  
  • NEB

    New England Biolabs

  •  
  • pMIA

    pMSCVII-Ametrine

  •  
  • rEF

    rat lung alpha EF

  •  
  • RT

    reverse transcription

  •  
  • TCCS

    T cell culture supplement

  •  
  • Treg

    regulatory T cell.

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The authors have no financial conflicts of interest.

Supplementary data