Open access peer-reviewed chapter

Animal Models of Chronic Obstructive Pulmonary Disease

Written By

Lillian Chow, David Smith, Khushboo Chokshi, Wendy Ezegbunam, Prangthip Charoenpong, Kimberly Foley, Adrian Cargill and Patrick Geraghty

Submitted: 01 March 2017 Reviewed: 30 June 2017 Published: 20 December 2017

DOI: 10.5772/intechopen.70262

From the Edited Volume

COPD - An Update in Pathogenesis and Clinical Management

Edited by Cormac McCarthy

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Chronic obstructive pulmonary disease (COPD) is the third leading cause of death in the USA and currently there are minimal therapies specific for the treatment of COPD. To advance our knowledge on COPD pathogenesis and develop new therapeutics, animal models are needed that represent key clinical and pathologic features of the human disease. The primary animal models utilized to study COPD rely on several factors associated with disease progression, i.e. genetic and epigenetic changes, environmental exposures and the microbial flora of the lungs. Here, a systematic approach was taken to summarize and evaluate the current animal models employed to study COPD pathogenesis, comorbidities and exacerbations. The strengths and limitations of these disease models are also delineated. The rodent COPD models have been extensively utilized but several studies have highlighted the potential of larger animals as an additional approach. Due to the inherent heterogeneity of COPD, the usefulness of certain animal models may be limiting but still represent helpful means to explore gene functional studies, testing new therapeutics and the exploring the significance of microbial floral changes. Therefore, interpreting the findings from animal models for the study of COPD represents a critical approach in deciding possible future human therapeutics.


  • disease models
  • COPD
  • animal
  • cigarette smoke
  • lung remodeling
  • comorbidities
  • exacerbation

1. Introduction

Chronic obstructive pulmonary disease (COPD) is one of the major cause of morbidity and mortality worldwide and a significant economic and social burden worldwide [1, 2]. COPD is defined as a disease state characterized by airflow limitation that is not fully reversible, associated with an abnormal inflammatory response of the lungs to noxious particles or gases [3]. Airflow is limited by airway inflammation [4], loss of lung elasticity [5], lung tissue destruction [6] and the narrowing of small airways [7, 8]. Most pulmonologists believe that parenchymal destruction (emphysema) and small-airway obstruction (chronic bronchitis) are the most significant phenotypes of COPD, as they both contribute to airflow limitation [9]. Therefore, when designing models of disease these two phenotypes are crucial when interoperating data to clinical significance. Emphysema is further characterized by augmented inflammation, irreversible destruction of alveolar walls leading to airspace enlargement, loss of elastic recoil and hyperinflation [10]. Most COPD models investigate emphysema characteristics.

Cigarette smoke (CS) inhalation is the primary etiologic risk factor associated with COPD. The age-adjusted mortality for COPD has increased over the past three decades [11] highlighting the need for better therapies [12, 13], increased COPD research and enhanced utilization of research models. In the US, the CS prevalence is estimated to be 15–19% of the US population (age 18 or older) [14] and smoking will remain a major public health issue in the coming years. It is estimated that 16% of all 8th graders have tried smoking and 17% of high school students continue smoking beyond graduation [15]. The smoking prevalence globally remains high, with CS rates at 28% in China [16], 27% in Germany, and 36% in Russia [17, 18]. These statistics will ensure that this disease continues to be a major public health issue for the foreseeable future and CS-induced COPD will likely continue to rise over the next decades. Projects predict COPD to be the third leading cause of death globally by 2030 [19]. Beyond smoking cessation, very limited options currently exist for therapeutic intervention to halt COPD progression. The mechanistic basis underlying the pathogenesis of both emphysema and chronic bronchitis is very complex, involving a combination of recurrent inflammation, enhanced autophagy, oxidative stress, protease/antiprotease imbalance, tissue injury, repair and cell death [20]. These changes are all modulated by environmental exposures and host genetics [21]. Clinical studies demonstrate that all smokers experience a range of pulmonary inflammation but only 15–20% of smokers develop severe progressive emphysema [22]. This disparity underscores the importance of susceptibility factors, a subset of which are almost certainly controlled by host genetics in addition to environmental exposures. Chronic bronchitis is almost three times more frequently diagnosed compared to emphysema in the COPD population in the US but emphysema is associated with a greater frequency of patient deaths [23]. To this extent, new research approaches are required to advance our understanding of the key players involved in COPD development and to identify new therapeutics for future treatment of both emphysema and chronic bronchitis.

To better understand this complex disease, multiple research model approaches are need. Much of our current understanding of the normal functioning of the lung and mechanisms of lung disease comes from these studies utilizing animals. A model is defined as a simplified system that is accessible and easily manipulated. The natural course of COPD in humans can take decades. However, the ideal models of COPD would involve inexpensive species which could grow quickly and in which disease induction occurs over a short period of time while still mimicking the human condition. There are other factors that must be discussed as well when considering a model of COPD, such as sex, age, animal strain/background, exposure dosing and frequency which can all significantly impact on data interpretation. In this chapter, we will outline several animal models that have been used to study the biological processes believed to play major roles in the pathogenesis of COPD. Currently, the majority of COPD research is conducted in rodents. We will also discuss whether data obtained from rodents can shed light on COPD in humans.


2. Models of COPD

While cigarette consumption is the main risk factor for COPD, up to 10–15% of COPD cases are not related to CS exposure. Therefore, other factors can contribute to COPD initiation and progression, and multiple models are required to decipher the key mechanisms driving the disease. At this time, the majority of animal models for COPD research focus on emphysematous characteristics. Three major experimental approaches are commonly employed for the induction of emphysema-like symptoms in animals: elastase lung instillation, CS exposure and genetic manipulation. These three approaches are employed because CS is the primary etiologic risk factor associated with COPD and its comorbidities, an imbalance between elastase and anti-elastase activity results in enzymatic degradation of elastin and emphysema formation [24], and predisposing genetic factors are associated with COPD initiation and progression [25, 26, 27]. It is also known that CS has a major impact on elastase imbalance [28] and can also modulate gene expression [29]. Therefore, CS inhalation is the preferred approach and the following sections will outline these exposure models. Several other models of chronic bronchitis will also be discussed in addition to the emphysema models. The primary approaches to establish a model of COPD in animals is summarized in Figure 1.

Figure 1.

Typical mouse models for COPD. (A) Mice are administered a single or multiple intratracheal doses of PPE to induce an emphysema phenotype. One weeks onwards, there is significant evidence for airspace enlargements. Images show a typical profile following a single 1.2 Unit intranasal dose of PPE or saline and examination of the airways 3-weeks later by H&E staining. (B) A nose or head only (NHO) cigarette smoke exposure system requires animals to be restrained while they inhale the cigarette smoke. The whole-body system streams CS into a larger exposure system chamber and allows animals access to food and water without restraint.

2.1. Elastase model

In 1963, Laurell and Eriksson reported that patients deficient for α1-antitrypsin developed emphysema at an earlier time in life due to an abundance of elastase [30]. This discovery is the primary clinical observation that influenced the development of the elastase model of emphysema. In 1965, Gross et al. [31] developed the first animal model of protease-induced induction of emphysema in rats by instillation of the plant protease, papain, into the lung. Subsequently, numerous other proteases have been utilized to induce emphysema [32, 33, 34, 35], including cathepsin B and proteinase 3. The most common protease utilized currently in animal models is elastase, primarily isolated from porcine. Porcine pancreatic elastase (PPE) is inexpensive and administration of PPE has been shown to induce features that resemble panacinar emphysema and lung damage throughout the organ. This approach has been utilized in many species of animals for several decades. The most frequently documented species, administration routes of PPE and exposure outcomes are summarized in Table 1, and will be discuss here in depth.

Animal Reference Strain Administration Outcomes
Mouse [41, 44, 45, 46, 47, 48, 49] C57BL/6 and C57BL/6J Intratracheal PPE Enhanced airspace enlargements, lung volume and capacity, lung dendritic cells, MMPs, fibronectin degradation, epithelial cell apoptosis, alveolar epithelial damage, macrophages, neutrophils, lymphocytes, eosinophils, NK cells, CD8+ cells, A2M, HMOX1, MMP2, TIMP1, TNF-α, IL-1β, IL-4, IL-5, IL-6, IFN-γ and TGF-β. Reduced α1-antitrypsin, VEGF and DFCO
[39, 41, 50] BALB/c and BALB/cJ Intratracheal PPE Enhanced airspace enlargements, lung volume and capacity, compliance, hyperinflation, elastic and collagen fiber content, BALF hemoglobin content, M1 macrophages, neutrophils, lymphocytes, eosinophils, NK and NKT cells, CD4+ and CD8+ cells, NRF2, A2M, HMOX1, Arg1, Fizz1, MMP12, MMP2, TIMP1, SLPI, PrxI, HO-1, GST-Yc, NQO1, TNF-α, IL-1β, IL-4, IL-5, IL-6, IL-18, iNOS, IL-17A, IFN-γ and TGF-β. Reduced α1-antitrypsin, VEGF and DFCO
[51, 52] A/J Endotracheal PPE Enhanced airspace enlargements and lesions in the lung parenchyma distal to the terminal bronchioles
[53, 54] Swiss Intratracheal PPE Enhanced airspace enlargements, hyperinflation of the alveoli, alveolar collapse, MMP9 and TNF-α. Reduced VEGF and lung elastance
[55, 56] FVB/J Intratracheal PPE Enhanced airspace enlargements, lung volume, BALF hemoglobin content and alveolar surface area. Increased TGF-β, FGF-2 and GAG in BALF
Rat [57, 58, 59] Wistar Intratracheal PPE Enhanced airspace enlargements, NO release and leukocyte infiltration
[60] Sprague-Dawley Intratracheal PPE Enhanced airspace enlargements and increased dynamic compliance
Hamster [61, 62] Intratracheal PPE Increased secretory cell metaplasia
Guinea pig [63, 64, 65] Intratracheal PPE Evidence of interstitial edema, degradation of fibrous tissue, elastin degradation and enhanced alveolar enlargements. Loss of epithelial cilia and detachment of epithelial cells from the basement membrane. Increase tracheal hyperresponses to histamine, diaphragm fatigue, PMNs in tracheal submucosa blood vessels and infiltration of macrophages into the parenchyma
Rabbit [62, 66, 67] Intratracheal PPE Enhanced airspace enlargements, apoptosis and 8-OHdG in lung tissues and reduced DLCO
[68, 69] New Zealand White Endotracheal aerosolized PPE Enhanced airspace enlargements, static compliance increase and decrease in expiratory flow and conductance
Pig [70] Yorkshire PPE instilled into left lower lobe bronchus Evidence of early edema, panlobular, centrilobular and paraseptal emphysema
Dog [33, 71] Endotracheal PPE Enhanced airspace and septal destruction and enhanced airway enlargements
[72] Beagle PPE injected through bronchi Emphysema determined by chest computed tomography and histology
Sheep [73, 74, 75] Endotracheal and intrabronchial PPE Enhanced lung resistance, bronchoconstriction and BALF levels of tissue kallikrein. Decreased tracheal mucus velocity

Table 1.

Animal models of COPD induced by elastase.

Abbreviations: GAG, glycosaminoglycan; NO, nitric oxide; PMNs, polymorphonuclear leukocytes; 8-OHdG, 8-hydroxy-deoxyguanosine; MMP, matrix metalloproteinases; NK, natural killer; IL, interleukin; TIMP, tissue inhibitor of metalloproteinases; TNF, tumor necrosis factor; IFN, interferon; TGF, transforming growth factor; NO, nitric oxide; SLPI, secretory leukocyte peptidase inhibitor; iNOS, nitric oxide synthases; VEGF, vascular endothelial growth factor; DLCO, diffusing capacity of the lungs for carbon monoxide; FGF, fibroblast growth factors; GAG, glycosaminoglycans.

As mentioned already, well-established genetic rodent background strains that utilize PPE-induced emphysema have been widely used as a model of COPD. Employing these strains/backgrounds, data from multiple studies are more reproducible and can be compared between groups. The animal genetic background is also critical for an appropriate control. Each strain has unique background alleles that may interact with and modify the expression of other genes upon establishment of a disease model or upon a stimulus, such as CS or elastase. Mice with different genetic backgrounds show disparate susceptibility to the development of emphysema, when utilizing the elastase model of emphysema [36]. Globally, C56BL/6 mice are the primary background strain used to generate many types of genetically engineered mice, but they are one of many strains. Other strains, including BALB/c mice, are documented to be more sensitive to dose and time dependent to PPE injury, as demonstrated by significantly greater mortality, weight loss, decline in lung function, immune cell infiltration and loss of alveolar tissue [36]. This may be due to several genetic differences between strains. BALB/c and C57BL/6J mice have differing type 1 and type 2 cytokine-mediated inflammation responses that could play key roles in determining the resistance or susceptibility to many diseases. Altered allergy responses in subgroups of patients is frequently observed in COPD, frequently coinciding with eosinophilia and enhanced allergy responses or independent of allergies [37, 38]. Therefore, it is critical to select the appropriate mouse background when utilizing mice for COPD studies and the correct controls.

Most PPE-induced emphysema studies highlight the proteolytic activity of elastase on lung structural changes, such as higher airspace enlargements (measured by mean linear intercept) both in mice and in rats (Table 1). Furthermore, several studies report changes in extracellular matrix (ECM) composition, such as disorganized elastin, degradation of proteoglycans, and abnormal collagen remodeling [39]. However, these effects are dependent on several factors, including animal strain, PPE dose at each instillation, and number of PPE challenges. Mice subjected to repeat PPE administrations within 1-week developed a more severe phenotype, with enhanced alveolar destruction, weight loss, diaphragmatic dysfunction, exercise intolerance, and pulmonary arterial hypertension observed [40]. The pulmonary changes observed in PPE-induced emphysema in mice persisted for 6 months after injury induction [40] and therefore make this exposure model a useful tool in studying lung repair. Limjunyawong and colleagues [41] performed the most extensive study on PPE-induced emphysema in mice, utilizing two mouse strains, multiple doses of PPE over several weeks and extensive pulmonary function analysis. This study found strain, time and dose dependence on PPE-induced mortality, body weight loss, decline in lung function, lung inflammation and loss of alveolar tissue [41]. They also found heterogeneity in signaling associated with emphysema in each mouse strain.

Over the past few decades, other rodents have been utilized in models of PPE-induced COPD, such as rat, guinea pig, and hamster. These and other animal models are utilized as the mouse lung differs to human lungs in total lung capacity, left lung lobe number, lung pleural thickness, parenchyma percentage of the lung, alveoli size, blood-gas barrier thickness, trachea cartilage structure, number of respiratory bronchioles and airway generations, epithelium thickness and airway lumen size [42]. These other rodent models have several phenotypes that more closely reflect the human lung than the mouse. The lung volume and alveolar size have a closer resemblance to human lungs [42]. The airway of most rodents (except guinea pig) do not respond to leukotrienes, important mediators that cause bronchoconstriction in humans [43]. Depending on the species utilized, several confounding factors may not be functional in the model. Larger animal models of PPE-induced emphysema, utilizing rabbits, dogs, pigs and sheep, have observed enhanced airspace and septal destruction, and enhanced airway enlargements (see Table 1). Despite the many advantages of these larger species as an animal model of COPD, these species have not been widely used, probably due to limitations in terms of cost and reagent availability. Larger animal models do offer better understanding of lung structure than smaller animal models but most PPE-induced studies in these species were performed over 40 years ago, and the imaging and pulmonary function equipment was limited to address many parameters. Revisiting this approach in these species may be beneficial.

Overall, the intrapulmonary administration of tissue degrading enzymes, such as elastase, represents a useful approach to studying emphysema, especially when focusing on mechanisms to repair. PPE-based models are an attractive approach, since it is a simple exposure protocol with a single (or multiple) lung administration leading to significant and rapid changes. However, comparing this rapid method to the lifetime development of human COPD is very difficult since this method bypasses a large number of biological mediators. Equally, this method may reflect one subtype of emphysema and may not represent other sub groups of the disease. Therefore, the protease based models encompass several important features of human COPD but other methods, such as inhalation exposures, may also be required to better represent the phenotype of the human disease. The elastase model may be suited to tissue repair research.

2.2. Smoke exposure models

Like PPE-induced emphysema, a variety of animal species have been exposed to CS to mimic the human disease. Rodents (mice, rats, guinea pigs and hamsters) are the most commonly utilized species in CS exposure systems, but rabbits, ferrets, pigs, sheep and dogs have also been used (see Table 2 for a summary of multiple studies). In parallel to PPE models, the severity of emphysema induced by the CS exposure model can be influenced by a variety of factors, such as differences in animal strains, smoke concentration, and duration of exposure, and the sex of the animal. The background strain of a mouse is an important factor in CS exposure with differing strains having varying susceptibilities or resistances to the development of CS-induced emphysema [76]. Smoke concentration measured in total particulate matter (TPM) per liter can also affect CS-induced emphysema development. Mice exposed to NHO CS inhalation, at concentrations of 75, 250, and 600 mg of TPM/L, show only a 13% increase in airspace enlargements (mean linear intercept) in animals exposed to 600 mg TPM/L after 28 weeks of exposure [77]. The TPM of mainstream CS is comprised of 4–9% of the total weight of cigarette smoke and is made up of many components, including polycyclic aromatic hydrocarbons, nitrosamines, phenol and nicotine. The TPM measurement excludes the gas and vapors in smoke. Therefore, regulated smoke concentration and the duration of CS exposure are critical steps in establishing a CS exposure model in animals. To prevent contrasting results when doing CS studies, Kentucky reference cigarettes (from the Center for Tobacco Reference Products) are used by many research groups throughout the world to aid in the standardization of the CS exposure model. Other species are very susceptible to CS-induced COPD, such as guinea pigs [78] and ferrets [79]. They develop COPD-like lesions and emphysema-like airspace enlargement within a few months of active CS exposure. By contrast, rat strains seem to be more resistant to the induction of emphysema-like lesions [80, 81, 82, 83]. Equally, animal sex has been reported as a confounding factor in COPD susceptibility. One example are female A/J mice which develop emphysema 6 weeks earlier than their male counterparts [84].

Animal Reference Strain Exposure Outcomes
Mouse [87, 89, 94] C57BL/6 NHO Enhanced lung hyperinflation, total lung capacity, compliance, alveolar enlargements, systolic blood pressure, circulating platelets and erythrocyte numbers, attenuate alveolar macrophage responses to inflammation, production of reactive oxygen species in heart and kidney and lipid peroxidation in heart, liver and kidneys. Decreased total nitric oxide plasma concentration
[29, 109, 110, 111, 112, 113] C57BL/6 Whole-body Enhanced airspace enlargements, lung capacity, compliance, lung inflammation, serum TNF-α, vascular and myocyte dysfunction, TLR9 signaling, protease activity, lung cell apoptosis, and cytokine production. Reduced weight gain, muscle mass and type I and IIA oxidative fibers, lung tissue elastance, lung PTP1B activity, EPAS1 lung expression
[90, 91] BALB/c NHO Increase in oxidative stress in lung and heart, mitochondrial respiratory dysfunction, chronic inflammation, mucus hypersecretion, airway remodeling and emphysema. Reduced lung function
[114, 115] BALB/c Whole-body Induced lung neutrophilia, IL1α, IL1β and CXCL1
[92, 93] A/J NHO Enhanced susceptibility to tumorigenesis, hyperplasia, metaplasia, and inflammation of the nose and larynx and proliferative lesions of the lungs. Delayed cutaneous wound healing
[29, 116, 117, 118] A/J Whole-body Enhanced airspace enlargement, right ventricle heart hypertrophy, lung cell apoptosis, infiltration of macrophages, neutrophils, lymphocytes, kinase activity, cytokine and protease expression. Reduced FEF50%/FVC, EPAS1 lung expression
[115] Swiss Whole-body Enhanced mast cell recruitment to cutaneous wound
[119] FVB/NJ Whole-body Enhanced airspace enlargements, immune cell recruitment to the lungs, RSV lung infection, lung cell apoptosis, BALF protein concentration, expression of S100A9 and MCP-1. Reduced PTP1B activity
[120] FVB CSE IP injection Enhanced airspace enlargements, prothymosin α expression, NFκB activity, MMP2 and MMP9
[76, 121] AKR/J Whole-body Increased inflammation, Th1 responses, macrophages, neutrophils, and T cells, oxidative stress levels in diaphragm and gastrocnemius. Reduced weight gain
Rat [80, 81, 82, 83] Wistar Whole-body Increased gastric ghrelin, plasma TNF-α. Reduced food intake, weight gain, abdominal fat, plasma levels of leptin, insulin-like growth factor-1
[122, 123] Sprague-Dawley Whole-body Areas of blebbing and microvillus-like projections from the luminal surface, and micro-thrombi proximal to intercostal branches. Enhanced lung parenchymal destruction, pulmonary hypertension and pulmonary inflammation
Hamster [5, 99] Whole-body Induced elevation of right ventricular systolic pressures, medial hypertrophy of pulmonary arterioles, lung chymase activity, Ang II levels and enhanced TGF-β1/Smad signaling. Reduces production of lysyl oxidase and the resynthesis of cross-linked elastin
Guinea pig [97, 98, 99] Hartley Whole-body Increased pulmonary artery pressure, right ventricle hypertrophy, raised respiratory resistance, airspace enlargement, intrapulmonary vessel remodeling, immune cell infiltration, CatK and CHOP expression, ERK and JNK phosphorylation. Decreased elastin and the loss of type III collagen in the alveolar walls
Ferret [79] Mustela putorius furo NHO Increased early-morning spontaneous coughs, sporadic infectious exacerbations, airway obstruction, goblet cell metaplasia/hyperplasia and mucus expression in small airways
Rabbit [101, 102, 103, 104, 105] New Zealand White Whole-body Decreased alveolar count, IRAK degradation. Increased ductal/destructive fraction, lung destruction, apoptosis, airspace enlargements, immune cell infiltration, intraparenchymal vascular congestion and thrombosis, intraparenchymal hemorrhage, respiratory epithelial proliferation, alveolar destruction, emphysematous changes and bronchoalveolar hemorrhage, lung and aorta expression of MMP1 and lung expression of TLR4
Pig [124] Lungs only Enhanced vasodilator response in the bronchial circulation, bronchodilatation and bronchial vasodilatation
Dog [125] Greyhounds Whole-body Increased parenchymal damage and inflammation
[126, 127] Beagle Through a tracheostoma Histological evidence of emphysema
Sheep [128] Unilateral inhalation Increased gas exchange impairment and metabolic activity

Table 2.

Animal models of COPD induced by cigarette smoke.

Abbreviations: ERK, extracellular signal-regulated kinase, JNK, c-Jun N-terminal kinase; TLR, toll-like receptor; EPAS1, endothelial PAS domain protein 1; CXCL, chemokine (C-X-C motif) ligand; NFκB, nuclear factor kappa-light-chain-enhancer of activated B cells; FEF, forced expiratory flow; FVC, forced vital capacity; MCP1, monocyte chemoattractant protein-1; Th1, type 1 T helper; Smad, mothers against decapentaplegic homolog; Cat, cathepsin; CHOP, C/-EBP homologous protein.

The experimental details of CS smoke and tobacco usage in humans are so varied that, currently, no single experimental CS exposure system can replicate the diversity of human smoking patterns. Therefore, CS models probably reflects only facets of COPD but they still represent the closest model of early human disease. In the following sections, we will outline the COPD models that utilize CS exposure. There are advantages and disadvantages with both types of CS exposure approaches, such as animal number, exact quantifiable exposure, exposure duration and end points of study. Additionally, a detailed breakdown of every published CS exposure publication up to 2013 is summarized by Leberl and colleagues [85].

2.2.1. Nose or head only exposure

The nose or head only (NHO) exposure model (sometimes referred to as mainstream CS exposure) was developed for the induction of COPD in animals that allows quantifiable concentrations administered to animals. Several species, including mice, are obligate nose breathers, which allows direct delivery of CS or aerosolized solutions to be delivered to the lungs. The NHO system requires restraining the animal so that their nose is inserted into a cone where they inhale the cigarette smoke [86]. During this exposure time animals are deprived of food and water. This method generates a uniform exposure that produces the desired emphysematous changes. However, the prolonged periods of restraint are stressful for the mice and the machine can usually accommodate only a limited number of mice (e.g. the Jaeger system has eighteen ports which is depicted in Figure 1B). NHO exposures are usually for short exposure times (30–60 min) and CS exposures are repeated several times daily [87]. The level of exposure can range from 75 to 600 mg TPM/m3 [77, 87]. In animal research, TPM concentrations is typically a defining factor for characterizing whether a study is a passive or secondhand exposure model, in addition to a NHO or whole-body method. A recent study examining gene expression profiles in CS exposure models in mice compared to human cohorts demonstrated that low TPM induces genes mainly related to xenobiotic/detoxification responses, while higher TPM activated immune/inflammatory and xenobiotic/detoxification responses [88]. In the same study, one human cohort clustered closer to low TPM but another cohort clustered closer to a high TPM [88]. Therefore, certain biological features of the CS exposure model are dose dependent and may represent a critical factor in establishing a model.

There are multiple companies that manufacture NHO exposure systems, e.g. Scireq, CH Technologies, Buxco Research Systems, In-Tox Products and Promech Lab AB. NHO exposure machines are typically automated for unattended operation (delivery, lighting, positioning, and ejection of cigarettes). They control the CS puff volumes and rates, are adjustable for direct smoke or indirect CS exposures, and contain a sealed chamber that prevents smoke leakage. The machines are relatively compact in size. Typically, an animal is placed in an individual exposure tube, with a rubber seal around the animal's neck to minimize exposure to the head. The tube is mounted onto the exposure tower (see Figure 1B) and CS is applied to the system. Some exposure models are equipped with a pneumotachograph, to measure respiratory flow from movement of the animal's chest wall. Alternatively, these models can be modified to test potential new aerosolized therapies.

Similar to PPE-induced emphysema models, mice are the primary species utilized in CS exposure models. C57BL/6 mice are frequently utilized in NHO exposures [87, 89], but others have utilized BALB/c [90, 91] and A/J mice [92, 93]. NHO exposure to CS has been documented to result in increased lung hyperinflation, total lung capacity, compliance, alveolar enlargements, systolic blood pressure, circulating platelets and erythrocyte numbers. This exposure model attenuates alveolar macrophage responses to inflammation, production of reactive oxygen species in heart and kidney and lipid peroxidation in heart, liver and kidneys and reduced total nitric oxide plasma concentrations in C57BL/6 mice [87, 89, 94].

Gaschler and colleagues [89] report that NHO and whole body CS exposure systems attenuate innate immune responses in a comparable manner. However, some outcomes appear to be different between both exposures, with NHO smoke exposures failing to produce soleus muscle weight reduction [87]. TPM concentrations [88] and exposure duration may also be confounding factors leading to differences in these models. Several larger animal species are more susceptible to CS exposure and NHO exposure resulting in phenotype changes similar to the human disease. Ferrets have several similarities to human airway physiology and submucosal gland distribution. In a recent NHO chronic CS exposure model, ferrets demonstrated clinical features close to human COPD, such as early-morning spontaneous coughs, sporadic infectious exacerbations, airway obstruction, goblet cell metaplasia/hyperplasia and increased mucus expression in small airways [79]. Despite larger animals displaying a lung morphology closer to human, the mouse model is the favored model of NHO CS exposure induced COPD. However, new models like the ferret may be a better model to evaluate and characterize further in future studies.

2.2.2. Whole-body exposure

Whole-body systems (sometimes referred to as side-stream CS exposure) can expose animals to a mixture of both passive and mainstream smoke, released from the burning cigarette and puffed through the cigarette [95]. The passive and mainstream smoke streams are mixed and then propelled by a fan to a chamber containing the mice that are housed within their cages. The advantage of this system is that the mice freely move about and have access to food and water (Figure 1B). Thus, mice in this system can be exposed for longer periods of time daily. In addition, the whole-body exposure system allows for the exposure of large groups of mice. Some systems allow the exposure of greater than 120 mice simultaneously, enabling researchers to use large numbers of mice and to perform multiple experiments without stressing the animals beyond the CS exposure. There are several companies that manufacture whole-body exposure systems, e.g. Scireq and Teague Enterprises. These systems are also automated to help with ease of use.

Similar to the other COPD models already discussed, rodents are the most common species used in whole-body CS exposure models due to the wide variety of applicable gene expression manipulations. Several recent advances in pulmonary function testing in mice [96] will further enhance the usage of mice in CS exposure models. Other animal models include the guinea pig model, which has been shown to be susceptible to whole-body CS-exposures. CS exposure in guinea pigs induces several pulmonary and cardiovascular changes, such as pulmonary artery pressure, right ventricle hypertrophy, raised respiratory resistance, airspace enlargement, intrapulmonary vessel remodeling, loss of elastin and type III collagen in the alveolar walls, immune cell infiltration, protease and kinase activity [97, 98, 99]. Measurable emphysematous changes are also detected in rats following 2-months of whole body CS exposure [100]. Very few larger animals have been utilized in the whole-body exposure system, perhaps due to insufficient access to chambers large enough to contain these animals. Several studies have successfully exposed rabbits to whole body CS [101, 102, 103, 104, 105]. Mice do not produce MMP1, the major collagenase associated with COPD [106], but rabbits express MMP1 and its induction is CS triggered [103]. For the study of proteins not expressed in smaller animals or expressed at a different frequency to human, several larger animals may represent a better model option. Rabbits are relatively easy to handle and several transgenic animals are available [107], making them a good model for investigating lung-related diseases. Rabbits are also large enough to allow non-lethal monitoring of physiological changes during the course of the CS exposure but rabbits still have multiple differences in lung physiology to human [108]. Importantly, when deciding on a CS exposure model of emphysema, it must be noted that both NHO and whole-body exposure methods result in emphysema and other COPD symptoms in animals, and both represent good but not perfect models of COPD.

2.3. Genetic models of COPD

Prior to genome-wide association studies (GWAS) on COPD samples, several genes have been investigated in animals for susceptibility to COPD development and pathogenesis (Table 3 is a summary from multiple mouse studies). GWAS and other genomic based studies have identified many further genes that are associated with COPD [26, 29] but functional studies are required to prove their importance in lung function and disease development. Currently, mice represent the most favored laboratory animal species to manipulate gene expression. The discovery of new gene manipulation approaches may alter this view, such as CRISPR-Cas9. CRISPR-Cas9 approaches in larger animals is an exciting new method for future COPD research. Several existing models will be briefly outlined here that alter susceptibility to environmental exposure induced COPD or genetic alterations that result in spontaneous COPD development. There are two major approaches, i.e. gain-of function or loss-of-function. These gene manipulations can be targeted within the whole body or within specific tissue or cell types. Gain-of-function is achieved by gene overexpression in transgenic mice or expressing a human gene or variant of that gene. Alternatively, loss of function is achieved by specifically targeting loss of expression of a gene, by direct or chemical mutagenesis. The majority of studies performed in COPD utilize whole body knockout animals, where the gene was genetically manipulated in the embryo prior to development. Several genes have been linked with COPD since several knockout animals develop spontaneous COPD as they age, such as Itgb6−/− or klotho−/− mice [129, 130]. Equally, introducing a human gene into a mouse can result in COPD development without environmental exposures, such as lung expression of human MMP1 in mice [131]. Most genetic manipulations require a “second-hit” to observe lung changes causing disease. Inducing a COPD phenotype by CS inhalation or PPE instillation in animals deficient for Nrf2 [132], Ptp1b [119], Gpx1 [133, 134] or p53 [135] exaggerates COPD-like changes. However, most genetic manipulations have been associated with protecting against CS or PPE induced COPD, e.g. Mmp12−/− [136], Cav1−/− [137], iNOS−/− [138], Tnfr−/− [139] and Mrp1−/−Mdr1a−/−Mdr1b−/− [140] mice. Since COPD is an extremely complex heterogenous disease, many of these genes and others may all contribute to disease initiation and progression. The GWAS studies in humans have identified several other possible targets for future investigation, e.g. HHIP, CHRNA5, HTR4, FAM13A, RIN3, TGFB2, GSTCD-NPNT, CYP2A6, IL27-CCDC101, ADGRG6-GPR126, THSD4, ADAM19, TET2, CFDP1, AGER, ARMC2, RARB, EEFSEC, DSP, MTCL1, SFTPD, IREB2, HHIP, and FAM13A [141, 142]. The mouse model still remains the best species to manipulate gene expression in the lungs but new techniques will aid in larger animals being utilized in the coming years.

Reference Mouse Exposure COPD susceptibility Outcomes
[129] Itgb6−/− Increased Spontaneous development of age-related lung emphysema due to lack of ITGB6-TGF-β1 regulation of the MMP12 expression
[130] Klotho−/− Increased Increased aging, enlargement of the air spaces, destruction of the alveolar walls
[131] Human MMP1 expression Increased Increased disruption of the alveolar walls and coalescence of the alveolar spaces
[136] Mmp12−/− CS Resistant Reduced smoke induced alveolar macrophage infiltration and emphysema
[139] Tnfr−/− CS Resistant Reduced airspace enlargements and neutrophil infiltration
[138] iNOS−/− CS Resistant Mice protected against emphysema and pulmonary hypertension
[143] Cd8−/− CS Resistant Blunted inflammatory response and did not develop emphysema
[111] Tlr9−/− CS Resistant Protected from smoke-induced airspace enlargements, loss of lung function, inflammation, protease activity, apoptosis, and inflammation
[144] SOD1 overexpression CS or elastase Resistant SOD1 prevented smoke or elastase induced airspace enlargements, neutrophil infiltration and lipid peroxidation product accumulation
[137] Cav1−/− CS Resistant Reduced senescence of lung fibroblasts and pulmonary emphysema
[140] Mrp1−/−Mdr1a−/−Mdr1b−/− CS Resistant Reduced inflammatory and emphysema
[145] Il-18Ra−/− CS Partially protected Decreased inflammation and emphysema
[144] Il-1R−/− CS Partially protected No induction of inflammatory cell infiltration, small airway remodeling or matrix breakdown
[146] Adiponectin−/− CS Partially protected No induction of inflammatory cell infiltration, airspace enlargements, tissue elastance or TNFα
[147] SOD3 overexpression CS or elastase Partially protected SOD3 prevented smoke or elastase induced airspace enlargement, impairment of lung function and exercise capacity
[148, 149] Il-17a−/− CS Partially protected Exacerbated macrophage and γδ T cells frequency, which trigger emphysema
[119] Ptp1b−/− CS Increased Enhanced airspace enlargements, immune cell recruitment, apoptosis, and inflammation
[132] Nrf2−/− CS Increased More inflammation, oxidative stress, apoptosis and reduced antioxidants
[133, 134] Gpx1−/− CS Increased Enhanced airspace enlargements, BALF neutrophils, macrophages, proteases, IL-17A, MIP1α, NFκB and AP1 activation. Reduced PP2A and PTP1B activity
[135] p53−/− Elastase Increased Increased emphysema severity, macrophages, neutrophils, BALF CCL2, BALF TNF-α and lung oxidative stress

Table 3.

Gene modulated models of COPD in mice.

Abbreviations: ITGB6, Integrin Subunit Beta 6; SOD, superoxide dismutase; Cav1, caveolin 1; Mrp1, multi-drug resistance associated protein 1; Mdr1, multi-drug resistance gene 1; AP1, activator protein 1, CCL, chemokine (C-C motif) ligand.

2.4. Models of bronchitis

As mentioned previously, the second phenotype in COPD is an airway model for chronic bronchitis. Several studies have focused on induction of bronchitis phenotypes in animals (see Table 4 for a summary of several studies). Bronchitis models utilize noxious inhalants such as sulfur dioxide (SO2), nitrogen dioxide (NO2), or ozone. SO2 is a gaseous irritant which can be used to induce COPD-like lesions in animal models. Exposure to high concentrations of SO2 daily results in chronic injury and repair of epithelial cells in rats [150, 151]. The exposure to high-levels of this gas ranging from 200 to 700 ppm for 4 to 8 weeks has been demonstrated to lead to neutrophilic inflammation, morphological signs of mucus production and mucus cell metaplasia and damage of ciliated epithelial in rats [150, 151] and ferrets [152]. Exposure of SO2 to capsaicin-treated rats results in increased airway smooth muscle mass and increased airway responsiveness observed in these animals [153]. NO2 is another gas that induces COPD-like lesions in a concentration, duration of exposure, and species susceptibility dependent manner [154]. Exposure to NO2 (50–150 ppm) can reduce animal survival due to extensive pulmonary injury, including pulmonary edema, hemorrhage and pleural effusion. Sub lethal levels of NO2 causes damage to cilia, hypertrophy of the bronchiolar epithelium and a type II pneumocyte hyperplasia in rats and hamsters [155, 156]. Mice exhibit similar lung responses to NO2 exposure [154]. The administration of ozone [157] also causes significant lung injury with several features associated with human COPD. All three of these exposure models were utilized in several species of animals but are rarely utilized in current studies of COPD. A recent NHO chronic CS exposure model in ferrets demonstrated clinical features close to human bronchitis [79] and suggests that the animal species may be a deciding factor in bronchitis research. Overall, new approaches are needed to access bronchitis in animal models of COPD.

Animal Reference Strain Exposure Outcomes
Mouse [158] BALB/c SO2 Enhanced neutrophilic inflammation, epithelial sloughing, ET-1 and TGFβ expression
[154] C57BL/6 NO2 Increased airway (neutrophils and macrophages), goblet cell hyperplasia, collagen deposition in the lung parenchyma and airspace enlargements
Rat [150, 151] Sprague Dawley SO2 Increased pulmonary resistance, airway responsiveness to methacholine, immune cell infiltration (notably neutrophils), immune dysregulation, oxidative stress, mucin, accumulation of surfactant in lamellar bodies of alveolar type II cells, BALF levels of lactate dehydrogenase and N-acetyl glucosaminidase activity. Reduced dynamic compliance
[159, 160] Sprague Dawley NO2 Increased dopamine D(2) receptor expression
Ferret [152] Mustela putorius furo SO2 Observation of lesions with changes in ciliated cells, edema and cell infiltration. Increased coughing, nasal discharge and dried mucus
[161] Mustela putorius furo NO2 Reduced thoracic clearance function
Hamster [160] DSN NO2 Moderate/severe bronchiolitis and alveolitis development. Increased mitosis in Club cells and BALF surfactant
Guinea pig [162] SO2 Increased hyperresponsiveness to intravenously administered serotonin, degeneration, desquamation of epithelium, and edema of the lamina propria of the trachea and bronchi
[163] Hartley NO2 Increased BALF eosinophils and neutrophils, microvascular leakage in the trachea and main bronchi.
Rabbit [164] SO2 Increase in sputum viscosity. Decreased respiratory rate and pO2 blood level
[165] NO2 Enhanced destructive of walls and abnormal enlargement of the distal air spaces
Dog [166] SO2 Significant ciliated cell damage
[167] NO2 Changes in edema, congestion, interstitial irritation, bronchiolitis, and interstitial fibrosis

Table 4.

Animal models of bronchitis.

Abbreviation: ET-1, endothelin-1.

2.5. New models of COPD

Several novel models of COPD have been utilized recently, including investigation of biomass fuel and individual components of CS, i.e. nicotine. Several of these models will briefly be discussed in this section. One research group has developed a rapid COPD model in mice by daily injecting CS extract (CSE) into the abdominal peritoneal cavity [120, 168]. Airway enlargements and injury of cardiac and skeletal muscles are reported within 6 weeks of initiation [120, 168] but comparative studies to whole-body or NHO systems are required. Recent genetic analyses have identified α3 and α5 nicotine acetylcholine receptors (nAchR) as susceptibility loci for COPD [169]. The airway epithelium expresses α3, α4, α5, α7, α9, β2 and β4 subunits for nAchRs [170], and their expression are highest on the apical cell surface, where exposure to inhaled nicotine occurs [171]. Several animal studies have already utilized nicotine in models of COPD, airway lung injury and tumorigenesis. CS from high-dose nicotine cigarettes induces more emphysematous changes than low-dose nicotine cigarettes in PPE pre-treated rats [172]. Nicotine enhances airway hyperreactivity in lipopolysaccharides (LPS)-challenged mice and inflammation in lung epithelial cells [173]. Nicotine also suppresses apoptosis in lung tumors [174].

These observations are important considering several new approaches for nicotine replacement therapy now being marketed, such as e-cigarettes. E-cigarettes are a relatively new product that has grown in popularity over recent years. E-cigarettes are devices that effectively deliver vaporized liquid nicotine to the lungs. The user can choose the nicotine concentration and devices can deliver a range of volumes. However, several additional studies have suggested that these products require further testing in animal models and human studies to evaluate short and long term effects. Recently, the US Surgeon General concluded that e-cigarette use among the younger population is now a significant public health concern [175]. The use of e-cigarettes enhances oxidative stress and inflammation in mice [176] and impairs immune defense against bacterial and viral infection [177]. Nicotine in e-cigarettes has also been implicated as the driving force of these changes in mice, with long-term inhalation of nicotine-containing e-cigarettes shown to increase airway hyperreactivity, distal airspace enlargement, mucin production, cytokine and protease expression [178]. There are some direct human disease correlations, with normal human bronchial epithelial (NHBE) cells grown at the air liquid interface exposed to nicotine-containing e-cigarette vapor showing impaired ciliary beat frequency, airway surface liquid volume, conductance of ion channels, decreased expression of genes associated with ciliogenesis and increased cytokine production [178]. The animal models used for these studies use either the e-cigarette device connected to a whole-body CS exposure model, outlined in section b.ii, or a whole-body nebulizer. Many companies have modified their NHO or whole-body exposure systems for e-cigarette research, which will expand on new animal models for lung research. Therefore, these models and others should be utilized in future studies of COPD and nicotine replacement therapy.

Models of air pollution have been used to examine several phenotypes of COPD, with exposures to diesel exhaust particulates (DEP) and other biomass fuels under scrutiny. Animals exposed to DEP daily via intratracheal instillation for 3-months have decreased IFN-γ levels in bronchoalveolar lavage (BAL) but elevated CD3+ T and CD8+ T cells increased in the lung parenchyma and airway enlargement [179]. Equally, exposure of mice to smoke or particles generated from dung [180] and wood biomass [181] have been associated with several parameters of COPD. These models used modified CS whole-body exposure systems. Studies like these are important as it is estimated that air pollutants and biomass smoke could contribute to over 4 million deaths annually in the developing world [182] and contribute to the 10–15% of COPD cases not related to CS exposure. Therefore, new models or modification of the current group of COPD models are required to identify new biological targets of COPD and potential new therapies for treatment.


3. Exacerbation models

Exacerbations are one of the most important outcomes for COPD patients, as they contribute to decreasing quality of life, loss of lung function, and increasing mortality and healthcare cost [183]. Modeling acute exacerbations of COPD (AECOPD) in animals is challenging due to the complex clinical manifestations. Most AECOPD are triggered by infection, usually viral [184] or bacterial [185]. This association of the presence of a microbial pathogen at the onset of AECOPD is the primary basis for the majority of AECOPD animal models. These models typically entail exposure of an animal, usually a mouse, to CS followed by infection with a virus or bacteria. Live pathogens can be difficult to work with and often require special containment facilities. Therefore, some groups have used purified components of the pathogen, such as lipopolysaccharides (LPS) [186, 187] or Staphylococcus aureus enterotoxin B (SEB) [188], or a stimulus that will mimic the immune response upon infection, such as polyinosinic:polycytidylic acid (poly(i:c)) [189, 190]. Several of these approaches and outcomes are summarized in Table 5. The most frequently used live pathogens utilized in AECOPD models are influenza [188, 191], rhinovirus [192], respiratory syncytial virus (RSV) [112, 119], Streptococcus pneumoniae [193] and non-typeable Haemophilus influenzae (NTHi) [193]. Most models utilize the CS exposure system, but several have favored the PPE method [192, 193]. The general consensus appears to be that the COPD status enhances the lungs susceptibility to infection, inflammation and worsens lung function in a similar manner to the human disease. A study by Foronjy and colleagues [112] infected mice multiple times with RSV during a chronic CS exposure to mimic repeat viral AECOPD observed in the human disease. Repeated infection heightened airspace enlargements, inflammation, cytokines, chemokines, protease activity, apoptosis and fibrotic areas, while reducing lung phosphatase activity [112]. The same group recently infected mice systemically with chimeric HIV-1 virus that is capable of establishing chronic infection in immunocompetent mice [194] and exposed these mice to CS [118]. This altered HIV-1 infection enhanced CS-induced COPD features in the lungs of these mice and this new murine model can be utilized to study HIV-related COPD, which results in a heightened form of the disease in the HIV+ population [195, 196, 197, 198]. Therefore, several animal models exist that can aid in our understanding of AECOPD.

Animal Reference Pathogen Exposure Outcomes
Mouse [188, 191] Influenza CS Enhanced airway damage, inflammation and alters IL33 responses
[192] Rhinovirus PPE Increased airway inflammation, obstruction, goblet cell metaplasia, total lung volume and alveolar chord length. Reduced lung elastance
[112, 119] RSV CS Increased airspace enlargements, inflammation, cytokines, chemokines, protease activity, apoptosis and fibrotic areas. Reduced phosphatase activity
[118] Modified HIV CS Reduced FEF50%/FVC and enhanced distal airspace enlargement, inflammation, apoptosis and protease activity
[189, 190] Poly(i:c) CS Enhanced pulmonary inflammation, airway hyper responsiveness, corticosteroid resistance, remodeling, apoptosis, fibrosis, TLR3 signaling, type I IFNs, IL-18, IL-12/IL-23 p40, IFN-γ, PKR activation and eIF2α
[193] Streptococcus pneumoniae PPE Increased mortality, inflammatory cells in BALF, and MMP-12, as well as enhanced emphysema progression
[199, 200] NTHi CS Increased pulmonary inflammation (MCP-1, -3, and -5, IP-10, and MIP-1γ)
[186] LPS CS Enhanced susceptibility to airway lung injury
[188] SEB CS Increased BALF lymphocytes and neutrophils, CD8+ T lymphocytes and granulocytes in lung tissue, IL-13, CXCL13, CCL19 and goblet cell hyperplasia
Rat [201] NTHi CS Increased airway bacterial load, aggravated mucus hypersecretion and delayed mucociliary clearance
Guinea pig [187] LPS CS Enhanced airway resistance, lung volume, neutrophils, epithelial hyperplasia and emphysema

Table 5.

Animal models of COPD exacerbations.

Abbreviations: eIF2α, eukaryotic initiation factor-2alpha; PKR, protein kinase; SEB, Staphylococcus aureus enterotoxin B; NTHi, non-typeable Haemophilus influenzae.


4. Modeling COPD comorbidities in animals

COPD is frequently observed with comorbidities that directly affect patient’s quality of life, increase the risk for exacerbations and increase rate of mortality [202]. There are many manifestations of COPD comorbidities, such as lung cancer, skeletal muscle wasting (cachexia), diaphragm muscle dysfunction, cardiovascular disease, osteoporosis and diabetes [202]. Several studies have suggested that a large proportion of COPD-associated deaths are attributed to comorbidities [203, 204]. Given that comorbidities have a significant impact on COPD patient survival and quality of life, researchers have utilized animal models to investigate systemic comorbidities associated with COPD. CS exposure models show extrapulmonary manifestations, similar to those frequently observed in COPD. Mice and rats exposed to CS have reduced body weight, fat mass, hind-limb skeletal muscles mass, grip strength and aerobic endurance [82, 205]. CS exposure also enhances pulmonary artery wall thickening, increased contractility and endothelial dysfunction in guinea pigs [99]. Chronic CS exposure was recently shown to alter several cardiovascular parameters, such as enlarging ventricular end systolic and diastolic diameters, reducing myocardial and cardiomyocyte contractile function, and disrupting intracellular Ca2+ homoeostasis, regulating fibrosis, apoptosis and mitochondrial damage [206]. Mice and guinea pig exposed to CS, displaying lung inflammation and emphysema, have pulmonary hypertension and significant impairments to right ventricular diastolic and systolic function and contractility [99, 207]. CS in mice also causes hypertension, endothelial dysfunction and cardiac remodeling [208]. Chronic CS exposure have also been documented to increase systolic blood pressure, circulating platelets and erythrocyte numbers, attenuate alveolar macrophage responses to inflammation, production of reactive oxygen species in heart and kidney and lipid peroxidation in heart, liver and kidneys in mice [94]. Multiple elastase instillations in mice leads to the development of pulmonary arterial hypertension [209]. Diaphragm muscle dysfunction can contribute to COPD comorbidity. Diaphragm fatigue and ROS production are observed in guinea pig PPE-induced emphysema [65] and mouse smoke-induced emphysema [121], which could contribute to this comorbidity. Enhanced susceptibility to tumorigenesis is also demonstrated in mice exposed to smoke [93]. The hamster inhalation model is the only model in which tumor induction by CS has been reproducibly achieved [210]. These combined changes would be predicted to contribute to multiple comorbidities in COPD patients. Research focusing on models of COPD comorbidities will greatly improve our understanding of key factors in loss of quality of life and mortality in the COPD patient population, and additional novel animal models investigating the link between COPD and comorbidities are needed.


5. Animal model limitations and what is missing in these models

Models of disease try to mimic the disease as closely as possible. Despite no disease model being perfect, many models are helpful tools in deciphering the pathogenesis of the disease. Regarding COPD and the models outlined here, all models yield several phenotypes of the disease. Each researcher must evaluate the outcomes they desire to observe and choose the appropriate model that best suits their needs. For example, most COPD models cannot reproduce the features of severe emphysema as observed in humans, i.e. GOLD stages 3 or 4. Most models, especially the mouse smoke model, only represent early COPD (i.e. similar to human GOLD stages 1 of COPD) regardless of exposure time. It is estimated that obstruction of 75% of the small airways is required before changes can be detected by routine pulmonary function tests (e.g. forced expiratory volume in 1 s (FEV1)) in humans [211]. The majority of morbidity and mortality occurs in COPD patients with severe disease [3]. This is a major challenge to undertake when developing a reliable model of COPD as this requires a considerable amount of time and enhances the frequency of animal death prior to testing the conditions of the model. If a severe stage of COPD is required for investigation, the PPE model may be the best model currently available to examine COPD. However, if researchers are examining disease initiation or progression the PPE model would not be ideal. Another challenge in animal research is in the measurement of lung function in very small mammals, such as mice. The use of the enhanced pause (Penh) in conscious mice as an indicator of airflow obstruction is not ideal and invasive methods remain more reliable. Several new technical advances have aided in enhancing the pulmonary function readouts in animals but further work is needed to correlate animal data to the human disease.

The many differences in lung morphology and physiology between humans and the animals utilized in research will definitely impact on data interpretation, such as mice being obligate nose breathers, having lower numbers of cilia, fewer Club cells, differences in airway mucus production and reduced submucosal glands in the trachea. However, the experimental approach must also undergo scrutiny. Studies do not define whether a new therapeutic treatment is started following disease initiation or in parallel with the disease triggering exposure (i.e. CS or PPE exposure). Many treatments may be useful in preventing disease initiation but will have little impact on the lungs following disease establishment. Several recent studies are now addressing this experimental problem but scientific reviewers need to assert this difference in studies. In mice, lung lesions and inflammation induced by CS inhalation do not progress after cessation of CS exposure but airway enlargements persist [212]. This may have a large impact on the methodological design of experiments in mice. Whether other species recover following CS exposure is unknown. Equally, new genetic targeted approaches should be employed after disease establishment. Genes associated with disease progression can be selectively knocked down or enhanced after disease initiation, rather than utilizing whole body knockout animals that may have lung disease phenotypes due to embryo development problems. Modifying current methodology in these ways will further aid in advancing current animal model data and interpretations. Future animal model COPD studies should assess the histopathological patterns of COPD and examine functional parameters of human COPD using imaging, airflow limitation, mucus hypersecretion, corticosteroid resistance, β-adrenergic bronchodilator responses, chronic cough and exacerbations.

Overall, several major factors limit the interpretation of data from animal models to the human disease. Many investigators expect the animal model to exactly mirror the human condition but COPD is a complex and heterogenous disease and the human disease requires further sub-characterization prior to designing animal models or interpreting current data from animal models. We must also remember that without disease models, the burden on patients would be vast to participate in clinical trials and donate samples. Equally, not utilizing animal models would only allow clinicians to undertake COPD research. Deciphering the key players of a disease requires multiple approaches from many research fields and animal models may have a significant impact on COPD treatment in a similar manner to oncology and cardiovascular research. There are many benefits in development and use of animal models of COPD, especially the understanding of the fundamentals of immune and inflammatory mechanisms. Clinicians will benefit from the input of animal researchers, immunologists, microbiologists, bioinformaticians and statisticians.


6. Conclusion

Current animal model technology allows researchers to investigate the mechanisms of airways dysfunction, the influence of genetics and the immunological factors that define physiological and signaling changes that drive COPD. There are numerous well-established exposure models of COPD that exhibit several characteristics of the human disease, but no model captures every phenotype of the human condition. Mice are the most common species utilized for COPD research as they are small in size, they have a fast gestation period, and cost of feeding and housing are far less than larger animals. Also, the mouse genome is extensively characterized and many genetically modifiable mice are available, as well as equipment and biological reagents designed specifically for mouse anatomy. It is far more cost effective to generate genetically manipulated mice than it is to do so in other species. Another advantage to the use of mouse models is that exposure to CS for 1 year represents approximately 50% of the animal’s lifetime, thereby allowing a better representation of lifetime CS exposure. While there are many physiological differences between the mouse and human pulmonary systems, the mouse models of COPD represent good tools to further our understanding of the human disease.



Patrick Geraghty’s research is currently supported by grants from the Flight Attendant Medical Research Institute and the Alpha-1 Foundation. The authors would like to also thank the Division of Pulmonary and Critical Care Medicine and Departments of Medicine and Cell Biology of SUNY Downstate Medical Center for their support, especially Dr. Robert Foronjy and Dr. Moro Salifu.


  1. 1. Murphy BS, Xu J, Kochanek KD. Deaths: Preliminary Data for 2010. N.V.S. Reports, Editor. Hyattsville, MD: National Center for Health Statistics; 2012
  2. 2. Raherison C, Girodet PO. Epidemiology of COPD. European Respiratory Review. 2009;18(114):213-221
  3. 3. Rabe KF, et al. Global strategy for the diagnosis, management, and prevention of chronic obstructive pulmonary disease: GOLD executive summary. American Journal of Respiratory and Critical Care Medicine. 2007;176(6):532-555
  4. 4. Miller M, et al. Persistent airway inflammation and emphysema progression on CT scan in ex-smokers observed for 4 years. Chest. 2011;139(6):1380-1387
  5. 5. Osman M, et al. Cigarette smoke impairs elastin resynthesis in lungs of hamsters with elastase-induced emphysema. The American Review of Respiratory Disease. 1985;132(3):640-643
  6. 6. Coxson HO, et al. A quantification of the lung surface area in emphysema using computed tomography. American Journal of Respiratory and Critical Care Medicine. 1999;159(3):851-856
  7. 7. Stewart JI, Criner GJ. The small airways in chronic obstructive pulmonary disease: Pathology and effects on disease progression and survival. Current Opinion in Pulmonary Medicine. 2013;19(2):109-115
  8. 8. Hogg JC, et al. The nature of small-airway obstruction in chronic obstructive pulmonary disease. The New England Journal of Medicine. 2004;350(26):2645-2653
  9. 9. McDonough JE, et al. Small-airway obstruction and emphysema in chronic obstructive pulmonary disease. The New England Journal of Medicine. 2011;365(17):1567-1575
  10. 10. Snider GL. Chronic obstructive pulmonary disease: A definition and implications of structural determinants of airflow obstruction for epidemiology. The American Review of Respiratory Disease. 1989;140(3 Pt 2):S3-S8
  11. 11. Miller N, et al. Trends in chronic obstructive pulmonary disease mortality. Missouri Medicine. 2000;97(3):87-90
  12. 12. Hogg JC. A brief review of chronic obstructive pulmonary disease. Canadian Respiratory Journal. 2012;19(6):381-384
  13. 13. Kim V, Criner GJ. Chronic bronchitis and chronic obstructive pulmonary disease. American Journal of Respiratory and Critical Care Medicine. 2013;187(3):228-237
  14. 14. Schroeder SA, Koh HK. Tobacco control 50 years after the 1964 surgeon general's report. JAMA. 2014;311(2):141-143
  15. 15. Arrazola RA, Dube SR, King BA. Tobacco product use among middle and high school students—United States, 2011 and 2012. MMWR. Morbidity and Mortality Weekly Report. 2013;62(45):893-897
  16. 16. Li Q, Hsia J, Yang G. Prevalence of smoking in China in 2010. The New England Journal of Medicine. 2011;364(25):2469-2470
  17. 17. WHO. Report on the Global Tobacco Epidemic, 2009: Implementing Smoke-Free Environments, in Tobacco Free Inititative. World Health Organization, WHO Press; 2009. pp. 1-136
  18. 18. McCartney G, et al. Contribution of smoking-related and alcohol-related deaths to the gender gap in mortality: Evidence from 30 European countries. Tobacco Control. 2011;20(2):166-168
  19. 19. Mathers CD, Loncar D. Projections of global mortality and burden of disease from 2002 to 2030. PLoS Medicine. 2006;3(11):e442
  20. 20. Fischer BM, Pavlisko E, Voynow JA. Pathogenic triad in COPD: Oxidative stress, protease-antiprotease imbalance, and inflammation. International Journal of Chronic Obstructive Pulmonary Disease. 2011;6:413-421
  21. 21. Yoshida T, Tuder RM. Pathobiology of cigarette smoke-induced chronic obstructive pulmonary disease. Physiological Reviews. 2007;87(3):1047-1082
  22. 22. Kohansal R, et al. The natural history of chronic airflow obstruction revisited: An analysis of the Framingham offspring cohort. American Journal of Respiratory and Critical Care Medicine. 2009;180(1):3-10
  23. 23. Statistics NCFH. Chronic Obstructive Pulmonary Disease (COPD) Includes: Chronic Bronchitis and Emphysema. Centers for Disease Control and Prevention. 2015.
  24. 24. Snider GL. Experimental studies on emphysema and chronic bronchial injury. European Journal of Respiratory Diseases. Supplement. 1986;146:17-35
  25. 25. Wilk JB. et al. Framingham Heart Study genome-wide association: Results for pulmonary function measures. BMC Medical Genomics. 2007;8(Suppl 1):S8
  26. 26. Pillai SG, et al. A genome-wide association study in chronic obstructive pulmonary disease (COPD): Identification of two major susceptibility loci. PLoS Genetics. 2009;5(3):e1000421
  27. 27. Wilk JB, et al. A genome-wide association study of pulmonary function measures in the Framingham Heart Study. PLoS Genetics. 2009;5(3):e1000429
  28. 28. MacNee W. Pathogenesis of chronic obstructive pulmonary disease. Proceedings of the American Thoracic Society. 2005;2(4):258-266; discussion 290-1
  29. 29. Yoo S, et al. Integrative analysis of DNA methylation and gene expression data identifies EPAS1 as a key regulator of COPD. PLoS Genetics. 2015;11(1):e1004898
  30. 30. Laurell CB, Eriksson S. The electrophoretic a1-globulin pattern of serum in a1-antitrypsin deficiency. Scandinavian Journal of Clinical and Laboratory Investigation. 1963;15:132-140
  31. 31. Gross P, et al. Experimental emphysema: Its production with papain in normal and silicotic rats. Archives of Environmental Health. 1965;11:50-58
  32. 32. Janoff A. Elastases and emphysema. Current assessment of the protease-antiprotease hypothesis. The American Review of Respiratory Disease. 1985;132(2):417-433
  33. 33. Janoff A. et al. Lung injury induced by leukocytic proteases. The American Journal of Pathology. 1979;97(1):111-136
  34. 34. Lesser M, Padilla ML, Cardozo C. Induction of emphysema in hamsters by intratracheal instillation of cathepsin B. The American Review of Respiratory Disease. 1992;145(3):661-668
  35. 35. Kao RC, et al. Proteinase 3. A distinct human polymorphonuclear leukocyte proteinase that produces emphysema in hamsters. The Journal of Clinical Investigation. 1988;82(6):1963-1973
  36. 36. Affairs, D.o.V. 2011 Survey of Veteran Enrollees' Health and Reliance Upon VA: With Selected Comparison to the 1999-2010 Surveys. Department of Veterans Affairs and Veterans Health Administration. 2012.
  37. 37. Singh B, Arora S, Khanna V. Association of severity of COPD with IgE and interleukin-1 beta. Monaldi Archives for Chest Disease. 2010;73(2):86-87
  38. 38. Singh D, et al. Eosinophilic inflammation in COPD: Prevalence and clinical characteristics. The European Respiratory Journal. 2014;44(6):1697-1700
  39. 39. Padilha GA, et al. Therapeutic effects of LASSBio-596 in an elastase-induced mouse model of emphysema. Frontiers in Physiology. 2015;6:267
  40. 40. Luthje L, et al. Exercise intolerance and systemic manifestations of pulmonary emphysema in a mouse model. Respiratory Research. 2009;10:7
  41. 41. Limjunyawong N, et al. Experimental progressive emphysema in BALB/cJ mice as a model for chronic alveolar destruction in humans. American Journal of Physiology- Lung Cellular and Molecular Physiology. 2015;309(7):L662-L676
  42. 42. Irvin CG, Bates JH. Measuring the lung function in the mouse: The challenge of size. Respiratory Research. 2003;4:4
  43. 43. Drazen JM, et al. Comparative airway and vascular activities of leukotrienes C-1 and D in vivo and in vitro. Proceedings of the National Academy of Sciences of the United States of America. 1980;77(7):4354-4358
  44. 44. Takahashi S, et al. Reversal of elastase-induced pulmonary emphysema and promotion of alveolar epithelial cell proliferation by simvastatin in mice. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2008;294(5):L882-L890
  45. 45. Harada H, et al. Upregulation of lung dendritic cell functions in elastase-induced emphysema. International Archives of Allergy and Immunology. 2009;149(Suppl 1):25-30
  46. 46. Moreno JA, et al. High-density lipoproteins potentiate alpha1-antitrypsin therapy in elastase-induced pulmonary emphysema. American Journal of Respiratory Cell and Molecular Biology. 2014;51(4):536-549
  47. 47. Hou HH, et al. Elastase induced lung epithelial cell apoptosis and emphysema through placenta growth factor. Cell Death & Disease. 2013;4:e793
  48. 48. Ueno M, et al. Alendronate inhalation ameliorates elastase-induced pulmonary emphysema in mice by induction of apoptosis of alveolar macrophages. Nature Communications. 2015;6:6332
  49. 49. Cloonan SM, et al. Mitochondrial iron chelation ameliorates cigarette smoke-induced bronchitis and emphysema in mice. Nature Medicine. 2016;22(2):163-174
  50. 50. Ishii Y, et al. Transcription factor Nrf2 plays a pivotal role in protection against elastase-induced pulmonary inflammation and emphysema. The Journal of Immunology. 2005;175(10):6968-6975
  51. 51. Valentine R, et al. Morphological and biochemical features of elastase-induced emphysema in strain A/J mice. Toxicology and Applied Pharmacology. 1983;68(3):451-461
  52. 52. Munoz-Barrutia A, et al. Quantification of lung damage in an elastase-induced mouse model of emphysema. International Journal of Biomedical Imaging. 2012;2012:734734
  53. 53. Cheng SL, et al. Prevention of elastase-induced emphysema in placenta growth factor knock-out mice. Respiratory Research. 2009;10:115
  54. 54. Santos LM, et al. Bronchial responsiveness in an elastase-induced mouse model of emphysema. Respiratory Physiology & Neurobiology. 2014;194:9-14
  55. 55. Lucey EC, et al. Retinoic acid does not affect alveolar septation in adult FVB mice with elastase-induced emphysema. Respiration. 2003;70(2):200-205
  56. 56. Buczek-Thomas JA, et al. Elastase mediates the release of growth factors from lung in vivo. American Journal of Respiratory Cell and Molecular Biology. 2004;31(3):344-350
  57. 57. Furuya N, et al. Cell therapy with adipose tissue-derived stem/stromal cells for elastase-induced pulmonary emphysema in rats. Regenerative Medicine. 2012;7(4):503-512
  58. 58. Bianchi A, et al. Three-dimensional accurate detection of lung emphysema in rats using ultra-short and zero echo time MRI. NMR in Biomedicine. 2015;28(11):1471-1479
  59. 59. Boiati RF, et al. Evaluation of the anti-inflammatory activity of atorvastatin and its effect on alveolar diameter in a model of elastase-induced emphysema in rats. Drug Research (Stuttg). 2015;65(10):540-544
  60. 60. Onclinx C, et al. Elastase-induced pulmonary emphysema in rats: Comparison of computed density and microscopic morphometry. Radiology. 2006;241(3):763-770
  61. 61. Breuer R, et al. Proteolytic activity of human neutrophil elastase and porcine pancreatic trypsin causes bronchial secretory cell metaplasia in hamsters. Experimental Lung Research. 1985;9(1-2):167-175
  62. 62. Lungarella G, et al. Effect of the novel synthetic protease inhibitor furoyl saccharin on elastase-induced emphysema in rabbits and hamsters. Experimental Lung Research. 1986;11(1):35-47
  63. 63. Miyazaki N, et al. Ultrastructural studies of elastase-induced experimental emphysema. Japanese Journal of Experimental Medicine. 1979;49(4):241-250
  64. 64. Willet KE, Mitchell HW. Effect of elastase instilled into the trachea on airways mechanics in guinea pigs. Pulmonary Pharmacology. 1988;1(3):139-152
  65. 65. Tamaoki J. Effects of elastase-induced emphysema on histochemical properties of guinea pig diaphragm. Respiration. 1988;54(1):16-23
  66. 66. Brenner M, et al. Effects of a novel implantable elastomer device for lung volume reduction surgery in a rabbit model of elastase-induced emphysema. Chest. 2002;121(1):201-209
  67. 67. Baila B, et al. Tetomilast attenuates elastase-induced pulmonary emphysema through inhibition of oxidative stress in rabbits. Biological and Pharmaceutical Bulletin. 2012;35(4):494-502
  68. 68. Chen JC, et al. An animal model for lung volume reduction therapy of pulmonary emphysema. Journal of Investigative Surgery. 1998;11(2):129-137
  69. 69. Nishi Y, et al. Elastase-induced changes in lung function: Relationship to morphometry and effect of drugs. Pulmonary Pharmacology & Therapeutics. 2003;16(4):221-229
  70. 70. Noma S, et al. Sequential morphologic changes of elastase-induced pulmonary emphysema in pig lungs. Evaluation by high-resolution computed tomography. Investigative Radiology. 1991;26(5):446-453
  71. 71. Woods JC, et al. Magnetization tagging decay to measure long-range (3)He diffusion in healthy and emphysematous canine lungs. Magnetic Resonance in Medicine. 2004;51(5):1002-1008
  72. 72. Gotoh M, et al. Development of a canine model of pulmonary emphysema and imaging of the emphysematous lung with infrared thoracoscopy. Journal of Thoracic and Cardiovascular Surgery. 2003;126(6):1916-1921
  73. 73. Scuri M, et al. Inhaled porcine pancreatic elastase causes bronchoconstriction via a bradykinin-mediated mechanism. Journal of Applied Physiology (Bethesda, Md.: 1985). 2000;89(4):1397-1402
  74. 74. Susskind H. et al. Assessment of elastase-induced structural and functional changes in sheep lungs. Bulletin Européen de Physiopathologie Respiratoire. 1985;21(2):149-157
  75. 75. Scuri M, Sabater JR, Abraham WM. Hyaluronan blocks porcine pancreatic elastase-induced mucociliary dysfunction in allergic sheep. Journal of Applied Physiology (Bethesda, Md.: 1985). 2007;102(6):2324-2331
  76. 76. Guerassimov A. et al. The development of emphysema in cigarette smoke-exposed mice is strain dependent. American Journal of Respiratory and Critical Care Medicine. 2004;170(9):974-980
  77. 77. Hodge-Bell KC, et al. Pulmonary inflammation in mice exposed to mainstream cigarette smoke. Inhalation Toxicology. 2007;19(4):361-376
  78. 78. Wright JL, Churg A. Cigarette smoke causes physiologic and morphologic changes of emphysema in the guinea pig. The American Review of Respiratory Disease. 1990;142(6 Pt 1):1422-1428
  79. 79. Raju SV, et al. A ferret model of COPD-related chronic bronchitis. JCI Insight. 2016;1(15):e87536
  80. 80. Tomoda K, et al. Changes of ghrelin and leptin levels in plasma by cigarette smoke in rats. Journal of Toxicological Sciences. 2012;37(1):131-138
  81. 81. Esquivel AL, et al. The effect of obesity and tobacco smoke exposure on inflammatory mediators and matrix metalloproteinases in rat model. Toxicology Mechanisms and Methods. 2014;24(9):633-643
  82. 82. Kamiide Y, et al. Chronic exposure to cigarette smoke causes extrapulmonary abnormalities in rats. Environmental Toxicology and Pharmacology. 2015;39(2):864-870
  83. 83. Gomita Y, et al. Effects of standard cigarette and nicotine-less cigarette smoke inhalings on nicorandil plasma levels in rats. Pharmacology. 1990;40(6):312-317
  84. 84. March TH, et al. Modulators of cigarette smoke-induced pulmonary emphysema in A/J mice. Toxicological Sciences. 2006;92(2):545-559
  85. 85. Leberl M, Kratzer A, Taraseviciene-Stewart L. Tobacco smoke induced COPD/emphysema in the animal model-are we all on the same page? Frontiers in Physiology. 2013;4:91
  86. 86. Nemmar A, et al. Evaluation of the pulmonary effects of short-term nose-only cigarette smoke exposure in mice. Experimental Biology and Medicine (Maywood, NJ). 2012;237(12):1449-1456
  87. 87. Rinaldi M, et al. Long-term nose-only cigarette smoke exposure induces emphysema and mild skeletal muscle dysfunction in mice. Disease Models & Mechanisms. 2012;5(3):333-341
  88. 88. Dvorkin-Gheva A, et al. Total particulate matter concentration skews cigarette smoke's gene expression profile. ERJ Open Research. eCollection 2016 Oct;2(4): pii: 00029-2016
  89. 89. Gaschler GJ, et al. Cigarette smoke exposure attenuates cytokine production by mouse alveolar macrophages. American Journal of Respiratory Cell and Molecular Biology. 2008;38(2):218-226
  90. 90. Raza H, John A, Nemmar A. Short-term effects of nose-only cigarette smoke exposure on glutathione redox homeostasis, cytochrome P450 1A1/2 and respiratory enzyme activities in mice tissues. Cellular Physiology and Biochemistry. 2013;31(4-5):683-692
  91. 91. Beckett EL, et al. A new short-term mouse model of chronic obstructive pulmonary disease identifies a role for mast cell tryptase in pathogenesis. Journal of Allergy and Clinical Immunology. 2013;131(3):752-762
  92. 92. Brown BG, et al. The effect of a 2-h exposure to cigarette smoke on the metabolic activation of the tobacco-specific nitrosamine 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanone in A/J mice. Chemico-Biological Interactions. 2001;138(2):125-135
  93. 93. Hamm JT, et al. Histological alterations in male A/J mice following nose-only exposure to tobacco smoke. Inhalation Toxicology. 2007;19(5):405-418
  94. 94. Nemmar A, et al. Short-term systemic effects of nose-only cigarette smoke exposure in mice: Role of oxidative stress. Cellular Physiology and Biochemistry. 2013;31(1):15-24
  95. 95. Foronjy RF, et al. Structural emphysema does not correlate with lung compliance: Lessons from the mouse smoking model. Experimental Lung Research. 2005;31(6):547-562
  96. 96. Shalaby KH, et al. Combined forced oscillation and forced expiration measurements in mice for the assessment of airway hyperresponsiveness. Respiratory Research. 2010;11:82
  97. 97. Golovatch P, et al. Role for cathepsin K in emphysema in smoke-exposed guinea pigs. Experimental Lung Research. 2009;35(8):631-645
  98. 98. Geraghty P, Wallace A, D'Armiento JM. Induction of the unfolded protein response by cigarette smoke is primarily an activating transcription factor 4-C/EBP homologous protein mediated process. International Journal of Chronic Obstructive Pulmonary Disease. 2011;6:309-319
  99. 99. Dominguez-Fandos D, et al. Sildenafil in a cigarette smoke-induced model of COPD in the guinea-pig. The European Respiratory Journal. 2015;46(2):346-354
  100. 100. Kratzer A, et al. Role of IL-18 in second-hand smoke-induced emphysema. American Journal of Respiratory Cell and Molecular Biology. 2013;48(6):725-732
  101. 101. Xiao R, Goldklang MP, D'Armiento JM. Parenchymal airspace profiling: Sensitive quantification and characterization of lung structure evaluating parenchymal destruction. American Journal of Respiratory Cell and Molecular Biology. 2016;55(5):708-715
  102. 102. Goldklang MP, et al. Single-photon emission computed tomography/computed tomography imaging in a rabbit model of emphysema reveals ongoing apoptosis in vivo. American Journal of Respiratory Cell and Molecular Biology. 2016;55(6):848-857
  103. 103. Geraghty P, Dabo AJ, D'Armiento J, TLR4 protein contributes to cigarette smoke-induced matrix metalloproteinase-1 (MMP-1) expression in chronic obstructive pulmonary disease. Journal of Biological Chemistry. 2011;286(34):30211-30218
  104. 104. Lemaitre V, Dabo AJ, D'Armiento J. Cigarette smoke components induce matrix metalloproteinase-1 in aortic endothelial cells through inhibition of mTOR signaling. Toxicological Sciences. 2011;123(2):542-549
  105. 105. Sezer M, et al. Effects of caffeic acid phenethyl ester on the histopathological changes in the lungs of cigarette smoke-exposed rabbits. Basic and Clinical Pharmacology and Toxicology. 2007;101(3):187-191
  106. 106. Imai K, et al. Human collagenase (matrix metalloproteinase-1) expression in the lungs of patients with emphysema. American Journal of Respiratory and Critical Care Medicine. 2001;163(3 Pt 1):786-791
  107. 107. Zabetian M, Tahmoorespur M, Hosseini K. The applications of transgenic rabbits in agriculture and biomedicine. Journal of Animal and Veterinary Advances. 2011;10(6):780-790
  108. 108. Kamaruzaman NA, et al. The rabbit as a model for studying lung disease and stem cell therapy. BioMed Research International. 2013;2013:691830
  109. 109. Gosker HR, et al. Extrapulmonary manifestations of chronic obstructive pulmonary disease in a mouse model of chronic cigarette smoke exposure. American Journal of Respiratory Cell and Molecular Biology. 2009;40(6):710-716
  110. 110. Tang K, Wagner PD, Breen EC. TNF-alpha-mediated reduction in PGC-1alpha may impair skeletal muscle function after cigarette smoke exposure. Journal of Cellular Physiology. 2010;222(2):320-327
  111. 111. Foronjy R, et al. TLR9 expression is required for the development of cigarette smoke-induced emphysema in mice. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2016;311(1):L154-L166. DOI: 10.1152/ajplung.00073.2016
  112. 112. Foronjy RF, et al. Respiratory syncytial virus infections enhance cigarette smoke induced COPD in mice. PLoS One. 2014;9(2):e90567
  113. 113. Phillips B, et al. A 7-month cigarette smoke inhalation study in C57BL/6 mice demonstrates reduced lung inflammation and emphysema following smoking cessation or aerosol exposure from a prototypic modified risk tobacco product. Food and Chemical Toxicology. 2015;80:328-345
  114. 114. Botelho FM, et al. IL-1alpha/IL-1R1 expression in chronic obstructive pulmonary disease and mechanistic relevance to smoke-induced neutrophilia in mice. PLoS One. 2011;6(12):e28457
  115. 115. Cardoso JF, et al. Effects of cigarette smoke in mice wound healing is strain dependent. Toxicologic Pathology. 2007;35(7):890-896
  116. 116. Braber S, et al. Inflammatory changes in the airways of mice caused by cigarette smoke exposure are only partially reversed after smoking cessation. Respiratory Research. 2010;11:99
  117. 117. Geraghty P, Hardigan A, Foronjy RF. Cigarette smoke activates the proto-oncogene c-src to promote airway inflammation and lung tissue destruction. American Journal of Respiratory Cell and Molecular Biology. 2014;50(3):559-570
  118. 118. Geraghty P, et al. HIV infection model of chronic obstructive pulmonary disease in mice. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2017;312(4):L500-L509. DOI: 10.1152/ajplung.00431.2016
  119. 119. Foronjy RF, et al. Protein tyrosine phosphatase 1B negatively regulates S100A9-mediated lung damage during respiratory syncytial virus exacerbations. Mucosal Immunology. 2016;9(5):1317-1329
  120. 120. Su BH, et al. Prothymosin alpha overexpression contributes to the development of pulmonary emphysema. Nature Communications. 2013;4:1906
  121. 121. Barreiro E, et al. Cigarette smoke-induced oxidative stress in skeletal muscles of mice. Respiratory Physiology & Neurobiology. 2012;182(1):9-17
  122. 122. Pittilo RM, et al. Effects of cigarette smoking on the ultrastructure of rat thoracic aorta and its ability to produce prostacyclin. Thrombosis and Haemostasis. 1982;48(2):173-176
  123. 123. Lee JH, et al. Simvastatin inhibits cigarette smoking-induced emphysema and pulmonary hypertension in rat lungs. American Journal of Respiratory and Critical Care Medicine. 2005;172(8):987-993
  124. 124. Alving K, Fornhem C, Lundberg JM. Pulmonary effects of endogenous and exogenous nitric oxide in the pig: Relation to cigarette smoke inhalation. British Journal of Pharmacology. 1993;110(2):739-746
  125. 125. Hernandez JA, et al. Pulmonary parenchymal defects in dogs following prolonged cigarette smoke exposure. The American Review of Respiratory Disease. 1966;93(1):78-83
  126. 126. Auerbach O, et al. Emphysema produced in dogs by cigarette smoking. JAMA. 1967;199(4):241-246
  127. 127. Cahan WG, Kirman D. An effective system and procedure for cigarette smoking by dogs. Journal of Surgical Research. 1968;8(12):567-575
  128. 128. Schroeder T, et al. PET imaging of regional 18F-FDG uptake and lung function after cigarette smoke inhalation. Journal of Nuclear Medicine. 2007;48(3):413-419
  129. 129. Morris DG, et al. Loss of integrin alpha(v)beta6-mediated TGF-beta activation causes Mmp12-dependent emphysema. Nature. 2003;422(6928):169-173
  130. 130. Suga T, et al. Disruption of the klotho gene causes pulmonary emphysema in mice. Defect in maintenance of pulmonary integrity during postnatal life. American Journal of Respiratory Cell and Molecular Biology. 2000;22(1):26-33
  131. 131. D'Armiento J, et al. Collagenase expression in the lungs of transgenic mice causes pulmonary emphysema. Cell. 1992;71(6):955-961
  132. 132. Rangasamy T, et al. Genetic ablation of Nrf2 enhances susceptibility to cigarette smoke-induced emphysema in mice. The Journal of Clinical Investigation. 2004;114(9):1248-1259
  133. 133. Geraghty P, et al. The glutathione peroxidase 1-protein tyrosine phosphatase 1B-protein phosphatase 2A axis. A key determinant of airway inflammation and alveolar destruction. American Journal of Respiratory Cell and Molecular Biology. 2013;49(5):721-730
  134. 134. Duong C, et al. Glutathione peroxidase-1 protects against cigarette smoke-induced lung inflammation in mice. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2010;299(3):L425-L433
  135. 135. Chrusciel S, et al. Lack of transcription factor p53 exacerbates elastase-induced emphysema in mice. American Journal of Respiratory Cell and Molecular Biology. 2016;54(2):188-199
  136. 136. Hautamaki RD, et al. Requirement for macrophage elastase for cigarette smoke-induced emphysema in mice. Science. 1997;277(5334):2002-2004
  137. 137. Volonte D, et al. Caveolin-1 expression is required for the development of pulmonary emphysema through activation of the ATM-p53-p21 pathway. Journal of Biological Chemistry. 2009;284(9):5462-5466
  138. 138. Seimetz M, et al. Inducible NOS inhibition reverses tobacco-smoke-induced emphysema and pulmonary hypertension in mice. Cell. 2011;147(2):293-305
  139. 139. Churg A, et al. Tumor necrosis factor-alpha drives 70% of cigarette smoke-induced emphysema in the mouse. American Journal of Respiratory and Critical Care Medicine. 2004;170(5):492-498
  140. 140. van der Deen M, et al. Reduced inflammatory response in cigarette smoke exposed Mrp1/Mdr1a/1b deficient mice. Respiratory Research. 2007;8:49
  141. 141. Morrow JD, et al. Functional interactors of three genome-wide association study genes are differentially expressed in severe chronic obstructive pulmonary disease lung tissue. Scientific Reports. 2017;7:44232
  142. 142. Hobbs BD, et al. Genetic loci associated with chronic obstructive pulmonary disease overlap with loci for lung function and pulmonary fibrosis. Nature Genetics. 2017;49(3):426-432
  143. 143. Maeno T, et al. CD8+ T Cells are required for inflammation and destruction in cigarette smoke-induced emphysema in mice. The Journal of Immunology. 2007;178(12):8090-8096
  144. 144. Foronjy RF, et al. Superoxide dismutase expression attenuates cigarette smoke- or elastase-generated emphysema in mice. American Journal of Respiratory and Critical Care Medicine. 2006;173(6):623-631
  145. 145. Kang MJ, et al. IL-18 is induced and IL-18 receptor alpha plays a critical role in the pathogenesis of cigarette smoke-induced pulmonary emphysema and inflammation. The Journal of Immunology. 2007;178(3):1948-1959
  146. 146. Miller M, et al. Adiponectin-deficient mice are protected against tobacco-induced inflammation and increased emphysema. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2010;299(6):L834-L842
  147. 147. Yao H, et al. Extracellular superoxide dismutase protects against pulmonary emphysema by attenuating oxidative fragmentation of ECM. Proceedings of the National Academy of Sciences of the United States of America. 2010;107(35):15571-15576
  148. 148. Shan M, et al. Cigarette smoke induction of osteopontin (SPP1) mediates T(H)17 inflammation in human and experimental emphysema. Science Translational Medicine. 2012;4(117):117ra9
  149. 149. Bozinovski S, et al. Innate cellular sources of interleukin-17A regulate macrophage accumulation in cigarette- smoke-induced lung inflammation in mice. Clinical Science (London). 2015;129(9):785-796
  150. 150. Shore S, et al. Increased airway responsiveness to inhaled methacholine in a rat model of chronic bronchitis. American Journal of Respiratory and Critical Care Medicine. 1995;151(6):1931-1938
  151. 151. Kodavanti UP, et al. The spontaneously hypertensive rat: An experimental model of sulfur dioxide-induced airways disease. Toxicological Sciences. 2006;94(1):193-205
  152. 152. Miller ML, et al. Effect of exposure to 500 ppm sulfur dioxide on the lungs of the ferret. Respiration. 1985;48(4):346-354
  153. 153. Long NC, et al. Airway hyperresponsiveness in a rat model of chronic bronchitis: Role of C fibers. American Journal of Respiratory and Critical Care Medicine. 1997;155(4):1222-1229
  154. 154. Wegmann M, et al. NO2-induced airway inflammation is associated with progressive airflow limitation and development of emphysema-like lesions in C57bl/6 mice. Experimental and Toxicological Pathology. 2005;56(6):341-350
  155. 155. Rombout PJ, et al. Influence of exposure regimen on nitrogen dioxide-induced morphological changes in the rat lung. Environmental Research. 1986;41(2):466-480
  156. 156. Parkinson DR, Stephens RJ. Morphological surface changes in the terminal bronchiolar region of NO2-exposed rat lung. Environmental Research. 1973;6(1):37-51
  157. 157. Mudway IS, Kelly FJ. Ozone and the lung: A sensitive issue. Molecular Aspects of Medicine. 2000;21(1-2):1-48
  158. 158. Cai C, et al. Prior SO2 exposure promotes airway inflammation and subepithelial fibrosis following repeated ovalbumin challenge. Clinical and Experimental Allergy. 2008;38(10):1680-1687
  159. 159. Peiser C, et al. Dopamine D2 receptor mRNA expression is increased in the jugular-nodose ganglia of rats with nitrogen dioxide-induced chronic bronchitis. Neuroscience Letters. 2009;465(2):143-146
  160. 160. Foster JR, et al. A comparative study of the pulmonary effects of NO2 in the rat and hamster. British Journal of Experimental Pathology. 1985;66(2):193-204
  161. 161. Rasmussen RE, et al. Effects of nitrogen dioxide on respiratory tract clearance in the ferret. Journal of Toxicology and Environment Health. 1994;41(1):109-120
  162. 162. Ito M, et al. Effect of clenbuterol on sulfur dioxide-induced acute bronchitis in guinea pigs. Research Communications in Molecular Pathology and Pharmacology. 1995;87(2):199-209
  163. 163. Papi A, et al. Bronchopulmonary inflammation and airway smooth muscle hyperresponsiveness induced by nitrogen dioxide in guinea pigs. European Journal of Pharmacology. 1999;374(2):241-247
  164. 164. Miyata T, et al. Effect of N-acetylneuraminic acid on respiratory tract secretion and inflammation in the bronchitic rabbit. Archives Internationales de Pharmacodynamie et de Thérapie. 1990;304:277-289
  165. 165. Haydon GB, et al. Nitrogen dioxide-induced emphysema in rabbits. The American Review of Respiratory Disease. 1967;95(5):797-805
  166. 166. Man SF, et al. Effects of SO2 exposure on canine pulmonary epithelial functions. Experimental Lung Research. 1989;15(2):181-198
  167. 167. Hine CH, et al. Pulmonary changes in animals exposed to nitrogen dioxide. Proceedings of the Western Pharmacology Society. 1964;7:19-22
  168. 168. Zhang Y, et al. Intraperitoneal injection of cigarette smoke extract induced emphysema, and injury of cardiac and skeletal muscles in BALB/C mice. Experimental Lung Research. 2013;39(1):18-31
  169. 169. Pillai SG, et al. Loci identified by genome-wide association studies influence different disease-related phenotypes in chronic obstructive pulmonary disease. American Journal of Respiratory and Critical Care Medicine. 2010;182(12):1498-1505
  170. 170. Zia S, et al. Nicotine enhances expression of the alpha 3, alpha 4, alpha 5, and alpha 7 nicotinic receptors modulating calcium metabolism and regulating adhesion and motility of respiratory epithelial cells. Research Communications in Molecular Pathology and Pharmacology. 1997;97(3):243-262
  171. 171. Maouche K, et al. {alpha}7 nicotinic acetylcholine receptor regulates airway epithelium differentiation by controlling basal cell proliferation. The American Journal of Pathology. 2009;175(5):1868-1882
  172. 172. Diamond L, et al. Augmentation of elastase-induced emphysema by cigarette smoke. Effects of reduced nicotine content. The American Review of Respiratory Disease. 1988;138(5):1201-1206
  173. 173. Xu Y, Zhang Y, Cardell LO. Nicotine exaggerates LPS-induced airway hyperreactivity via JNK-mediated up-regulation of toll-like receptor 4. American Journal of Respiratory Cell and Molecular Biology. 2014;51(3):370-379
  174. 174. Paleari L, et al. Role of alpha7-nicotinic acetylcholine receptor in human non-small cell lung cancer proliferation. Cell Proliferation. 2008;41(6):936-959
  175. 175. Services, U.S.D.o.H.a.H.. E-Cigarette Use Among Youth and Young Adults: A Report of the Surgeon General. Murthy VH, et al, 2016.
  176. 176. Lerner CA, et al. Vapors produced by electronic cigarettes and e-juices with flavorings induce toxicity, oxidative stress, and inflammatory response in lung epithelial cells and in mouse lung. PLoS One. 2015;10(2):e0116732
  177. 177. Sussan TE, et al. Exposure to electronic cigarettes impairs pulmonary anti-bacterial and anti-viral defenses in a mouse model. PLoS One. 2015;10(2):e0116861
  178. 178. Garcia-Arcos I, et al. Chronic electronic cigarette exposure in mice induces features of COPD in a nicotine-dependent manner. Thorax. 2016;71(12):1119-1129
  179. 179. Yoshizaki K, et al. Chronic exposure of diesel exhaust particles induces alveolar enlargement in mice. Respiratory Research. 2015;16:18
  180. 180. Mehra D, et al. A comparison of the inflammatory and proteolytic effects of dung biomass and cigarette smoke exposure in the lung. PLoS One. 2012;7(12):e52889
  181. 181. Shapiro D, et al. Activation of transient receptor potential ankyrin-1 (TRPA1) in lung cells by wood smoke particulate material. Chemical Research in Toxicology. 2013;26(5):750-758
  182. 182. Organization, W.H. Household Air Pollution and Health; WHO Organization; 2016.
  183. 183. Donaldson GC, et al. Factors associated with change in exacerbation frequency in COPD. Respiratory Research. 2013;14:79
  184. 184. Seemungal TA, et al. Detection of rhinovirus in induced sputum at exacerbation of chronic obstructive pulmonary disease. The European Respiratory Journal. 2000;16(4):677-683
  185. 185. Stockley RA, et al. Relationship of sputum color to nature and outpatient management of acute exacerbations of COPD. Chest. 2000;117(6):1638-1645
  186. 186. Sakhatskyy P, et al. Double-hit mouse model of cigarette smoke priming for acute lung injury. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2017;312(1):L56-L67
  187. 187. Mizutani N, et al. Pulmonary emphysema induced by cigarette smoke solution and lipopolysaccharide in guinea pigs. Biological and Pharmaceutical Bulletin. 2009;32(9):1559-1564
  188. 188. Huvenne W, et al. Exacerbation of cigarette smoke-induced pulmonary inflammation by Staphylococcus aureus enterotoxin B in mice. Respiratory Research. 2011;12:69
  189. 189. Kang MJ, et al. Cigarette smoke selectively enhances viral PAMP- and virus-induced pulmonary innate immune and remodeling responses in mice. The Journal of Clinical Investigation. 2008;118(8):2771-2784
  190. 190. Kimura G, et al. Toll-like receptor 3 stimulation causes corticosteroid-refractory airway neutrophilia and hyperresponsiveness in mice. Chest. 2013;144(1):99-105
  191. 191. Kearley J, et al. Cigarette smoke silences innate lymphoid cell function and facilitates an exacerbated type I interleukin-33-dependent response to infection. Immunity. 2015;42(3):566-579
  192. 192. Sajjan U, et al. Elastase- and LPS-exposed mice display altered responses to rhinovirus infection. American Journal of Physiology - Lung Cellular and Molecular Physiology. 2009;297(5):L931-L944
  193. 193. Takahashi S, et al. Pneumococcal infection aggravates elastase-induced emphysema via matrix metalloproteinase 12 overexpression. Journal of Infectious Diseases. 2016;213(6):1018-1030
  194. 194. Potash MJ, et al. A mouse model for study of systemic HIV-1 infection, antiviral immune responses, and neuroinvasiveness. Proceedings of the National Academy of Sciences of the United States of America. 2005;102(10):3760-3765
  195. 195. Diaz PT, et al. Increased susceptibility to pulmonary emphysema among HIV-seropositive smokers. Annals of Internal Medicine. 2000;132(5):369-372
  196. 196. Crothers K, et al. Increased COPD among HIV-positive compared to HIV-negative veterans. Chest. 2006;130(5):1326-1333
  197. 197. Gingo MR, Morris A, Crothers K. Human immunodeficiency virus-associated obstructive lung diseases. Clinics in Chest Medicine. 2013;34(2):273-282
  198. 198. Drummond MB, Kirk GD. HIV-associated obstructive lung diseases: Insights and implications for the clinician. The Lancet Respiratory Medicine. 2014;2(7):583-592
  199. 199. Gaschler GJ, et al. Bacteria challenge in smoke-exposed mice exacerbates inflammation and skews the inflammatory profile. American Journal of Respiratory and Critical Care Medicine. 2009;179(8):666-675
  200. 200. Gaschler GJ, et al. Mechanisms of clearance of nontypeable Haemophilus influenzae from cigarette smoke-exposed mouse lungs. The European Respiratory Journal. 2010;36(5):1131-1142
  201. 201. Sun L, et al. The effect and mechanism of action of carbocysteine on airway bacterial load in rats chronically exposed to cigarette smoke. Respirology. 2010;15(7):1064-1071
  202. 202. Barnes PJ, Celli BR. Systemic manifestations and comorbidities of COPD. The European Respiratory Journal. 2009;33(5):1165-1185
  203. 203. Sin DD, Man SF. Chronic obstructive pulmonary disease as a risk factor for cardiovascular morbidity and mortality. Proceedings of the American Thoracic Society. 2005;2(1):8-11
  204. 204. Chatila WM, et al. Comorbidities in chronic obstructive pulmonary disease. Proceedings of the American Thoracic Society. 2008;5(4):549-555
  205. 205. Chen H, et al. Cigarette smoke exposure reprograms the hypothalamic neuropeptide Y axis to promote weight loss. American Journal of Respiratory and Critical Care Medicine. 2006;173(11):1248-1254
  206. 206. Hu N, et al. Cardiac-specific overexpression of metallothionein rescues against cigarette smoking exposure-induced myocardial contractile and mitochondrial damage. PLoS One. 2013;8(2):e57151
  207. 207. Sussan TE, et al. Targeting Nrf2 with the triterpenoid CDDO-imidazolide attenuates cigarette smoke-induced emphysema and cardiac dysfunction in mice. Proceedings of the National Academy of Sciences of the United States of America. 2009;106(1):250-255
  208. 208. Talukder MA, et al. Chronic cigarette smoking causes hypertension, increased oxidative stress, impaired NO bioavailability, endothelial dysfunction, and cardiac remodeling in mice. American Journal of Physiology - Heart and Circulatory Physiology. 2011;300(1):H388-H396
  209. 209. Antunes MA, et al. Effects of different mesenchymal stromal cell sources and delivery routes in experimental emphysema. Respiratory Research. 2014;15:118
  210. 210. Hoffmann D, Hecht SS, Wynder EL. Tumor promoters and cocarcinogens in tobacco carcinogenesis. Environmental Health Perspectives. 1983;50:247-257
  211. 211. Cosio M, et al. The relations between structural changes in small airways and pulmonary-function tests. The New England Journal of Medicine. 1978;298(23):1277-1281
  212. 212. Jobse BN, et al. Impact of inflammation, emphysema, and smoking cessation on V/Q in mouse models of lung obstruction. Respiratory Research. 2014;15:42

Written By

Lillian Chow, David Smith, Khushboo Chokshi, Wendy Ezegbunam, Prangthip Charoenpong, Kimberly Foley, Adrian Cargill and Patrick Geraghty

Submitted: 01 March 2017 Reviewed: 30 June 2017 Published: 20 December 2017