Mucosal Immunity

Abstract United airways disease (UAD) is the concept that the upper and lower airways, which are anatomically and immunologically related, form a single organ. According to this concept, upper and lower airway diseases are frequently comorbid because they reflect manifestations of a single underlying disease at different sites of the respiratory tract. Allergic asthma‐allergic rhinitis is the archetypal UAD, but emerging data indicate that UAD is a heterogeneous condition and consists of multiple phenotypes (observable clinical characteristics) and endotypes (pathobiologic mechanisms). The UAD paradigm also extends to myriad sinonasal diseases (eg, chronic rhinosinusitis with or without nasal polyps) and lower airway diseases (eg, bronchiectasis, chronic obstructive pulmonary disease). Here, we review currently known phenoendotypes of UAD and propose a “treatable traits” approach for the classification and management of UAD, wherein pathophysiological mechanisms and factors contributing to disease are identified and targeted for treatment. Treatable traits in UAD can be analyzed according to a framework comprising airway inflammation (eosinophilic, neutrophilic), impaired airway mucosal defense (impaired mucociliary clearance, antibody deficiency), and exogenous cofactors (allergic sensitizers, tobacco smoke, microbes). Appreciation of treatable traits is necessary in advancing the effort to deliver precise treatments and achieve better outcomes in patients with UAD. 1 INTRODUCTION The respiratory tract is divided into the upper and lower airways at the level of the vocal cords. Despite this artificial distinction, they are anatomically contiguous and immunologically related. United airways disease (UAD) is the concept that upper and lower airways form a single organ, with upper and lower airway diseases co‐occurring frequently because they reflect different manifestations of a single underlying disease process.1 Possible mechanisms linking the upper and lower airways include aspiration of nasal inflammatory cells and mediators into the tracheobronchial tree, hematogenous propagation of inflammatory mediators from one part of the airway to another, stimulation of bone marrow progenitor cells, or the nasobronchial reflex.2 The archetypal UAD is allergic asthma‐rhinitis, but emerging data indicate that UAD is complex and heterogenous, and may present with other clinical phenotypes of sinonasal disease (eg, chronic rhinosinusitis with or without nasal polyps) and lower airway disease (eg, bronchiectasis, chronic obstructive pulmonary disease [COPD]). Even within a given clinical phenotype, there is further variability based on the underlying pathobiologic mechanisms, known as endotypes. Current guideline‐based management approaches (summarized in Table 1) stratify patients according to symptoms, disease control, future risk, and occasionally broad clinical phenotypes, but do not sufficiently consider complexity and variability in other important dimensions such as airway inflammation, airway mucosal defense, and exogenous cofactors. Failure to recognize this complexity or treat both upper and lower compartments of the airways inevitably leads to suboptimal outcomes for patients. Site Disease Approach to stratification of patients and selection of therapies Patient stratification groups Recommended treatments according to patient stratification Other possible treatments Upper airway Allergic rhinitis Depends on symptoms, time course, severity, control, patient preferences, availability of treatment, and cost. One approach is a step‐up/step‐down approach based on assessment of control Step 1: mild symptoms Intranasal or oral non‐sedating H1‐antihistamines Leukotriene antagonists Allergen and irritant avoidance Allergen immunotherapy Nasal filters or saline Step 2: moderate‐to‐severe symptoms or persistent AR Intranasal corticosteroids Step 3: uncontrolled at step 2 (current or historical) Combination intranasal corticosteroids and intranasal H1‐antihistamines Step 4: uncontrolled at step 3 Short course oral steroids Chronic rhinosinusitis Based on clinical phenotype CRSsNP Intranasal corticosteroids Antibiotics (second line) Consider short course of oral steroids Saline irrigation Sinus surgery for medically refractory disease CRSwNP Intranasal corticosteroids Short course of oral steroids Antibiotic if suspicion of infection Consider leukotriene receptor for acetylsalicylic acid sensitivity Lower airway Asthma Assessment of symptoms, exacerbations, and lung function followed by stepwise escalation/de‐escalation of therapy Step 1 Consider low‐dose ICS Leukotriene receptor antagonists Theophylline Allergen avoidance Allergen immunotherapy for patients with allergic rhinitis and sensitized to house dust mites, frequent exacerbations, and FEV1 > 70% Vaccinations Weight reduction for obese asthma patients Step 2 Low‐dose ICS Step 3 Low‐dose ICS + LABA Step 4 Medium‐/high‐dose ICS + LABA Step 5 Low‐dose oral corticosteroids Add‐on treatments: tiotropium, anti‐IgE, anti‐IL5, bronchial thermoplasty Patients may benefit from phenotyping and endotyping at Step 5 Chronic obstructive pulmonary disease Assessment of symptoms (CAT, mMRC), exacerbations, airflow obstruction (FEV1) Group A (CAT < 10, mMRC < 2, infrequent exacerbations) Bronchodilator Smoking cessation (including pharmacologic treatment) Education, self‐management, and pulmonary rehabilitation Pneumococcal and influenza vaccinations Long‐term oxygen therapy in the presence of severe resting hypoxemia Long‐term noninvasive ventilation for patients with severe chronic hypercapnia and a history of hospitalization for acute respiratory failure Bronchoscopic or surgical lung volume reduction surgery for selected patients Group B (CAT ≥ 10, mMRC ≥ 2, infrequent exacerbations) LABA or LAMA Increase to LAMA + LABA if persistent symptoms Group C (CAT < 10, mMRC < 2, frequent exacerbations) LAMA Increase to LAMA + LABA or change to LABA + ICS if further exacerbations Group D (CAT ≥ 10, mMRC ≥ 2, frequent exacerbations) LAMA + LABA Increase to LAMA + LABA + ICS if persistent symptoms/further exacerbations Consider macrolides Consider roflumilast if FEV1 < 50% predicted and patient has chronic bronchitis Bronchiectasis Assessment of microbial infection, exacerbations and airway symptoms Pseudomonas aeruginosa infection with frequent exacerbations Long‐term inhaled antibiotic treatment Escalate to combined oral and inhaled antibiotic treatment if inadequate response Physiotherapy, pulmonary rehabilitation, and/or airway clearance Pneumococcal and influenza vaccinations Long‐acting bronchodilators for patients with significant breathlessness Non‐Pseudomonas aeruginosa infection with frequent exacerbations Long‐term macrolide treatment Escalate to combined oral and inhaled antibiotic treatment if inadequate response Multiple sequential airway treatments Short‐ or long‐acting bronchodilators → Mucolytic/physiotherapy adjuncts → airway clearance → inhaled antibiotics AR, allergic rhinitis; CAT, COPD Assessment Tool; CRSsNP, chronic rhinosinusitis without nasal polyps; CRSwNP, chronic rhinosinusitis with nasal polyps; FEV1, forced expiratory volume in one second; ICS, inhaled corticosteroids; ICS + LABA, combination inhaled corticosteroids and long‐acting beta agonist; LABA, long‐acting beta agonist; LAMA, long‐acting muscarinic antagonist; mMRC, modified Medical Research Council dyspnea score. The treatable traits approach has been described for lower airway diseases,3 and in this review, we extend its application to combined upper and lower airway disease. We summarize the current evidence for known clinicopathobiologic subtypes (“phenoendotypes”) of UAD. We have grouped the clinical manifestations of UAD according to the “treatable traits” of airway inflammation, airway mucosal defense dysfunction, and exogenous cofactors (Figure 1 and Table 2). These traits are not mutually exclusive and multiple traits may contribute to disease in an individual at any one point in time. Treatable traits may also vary over time. It is with the intention of drawing the readers’ attention to traits that are common to both airway compartments that we have artificially classified the upper and lower airway conditions as presented in this review. We believe that targeting common disease processes, when used in combination with current patient stratification approaches, is likely to lead to clinical improvements at both sites of the respiratory tract. Trait Endotype/pathophysiology Method of assessment/biomarker Clinical phenotype Targeted treatment Upper airway Lower airway Inflammation Eosinophilic, high type 2/TH2‐cytokines IL‐4, IL‐5, IL‐13 Blood periostin, high FeNO, sputum eosinophilia, blood eosinophilia CRSwNP Eosinophilic asthma Corticosteroids (systemic, inhaled, intranasal), anti‐IL‐5, IL‐4, IL‐13, anti‐TSLP, ?CRTh2 antagonist Allergic, systemic CD4+ TH2 cells Blood total IgE, blood specific IgEs, skin prick tests Allergic rhinitis Allergic asthma Allergen avoidance, corticosteroids, antileukotrienes, anti‐IgE, immunotherapy Allergic, local Local nasal and bronchial mucosa specific IgE production Blood total IgE not raised, negative blood specific IgEs, negative skin prick tests, local IgE and specific IgE production, nasal allergen provocation test Local allergic rhinitis Intrinsic asthma ?Omalizumab, ?immunotherapy Non‐allergic eosinophilic IL‐25, IL‐33, TSLP, group 2 innate lymphoid cells Blood periostin, high FeNO, sputum eosinophilia, blood eosinophilia, non‐reactive skin prick tests, absent specific IgE in blood Non‐allergic rhinitis with eosinophilia syndrome (NARES) Adult onset eosinophilic asthma Anti‐TSLP, anti‐IL‐4, IL‐5, IL‐13, ?CRTh2 antagonist Neutrophilic TH17 cells, IL‐17, CXCL1, CXCL8 IL‐8, sputum neutrophilia CRSsNP COPD, neutrophilic asthma Smoking cessation, macrolides Impaired airway mucosal defense Cystic fibrosis Autosomal recessive disorder in CFTR gene leading to improper salt balance and viscous airway surface liquid Genetic testing, sweat chloride test CRSwNP (acid mucin) Bronchiectasis CFTR modulators Primary ciliary dyskinesia Autosomal recessive mutations causing dynein and radial spoke defects Low nasal nitric oxide, ciliary motion and ultrastructure CRSsNP Bronchiectasis No validated PCD‐specific therapies Primary antibody deficiency Deficient immunoglobulins or functional antibody response Quantitative serum immunoglobulin levels, functional antibody responses CRS Bronchiectasis Immunoglobulins, vaccinations Exogenous cofactors Occupational exposures High molecular weight sensitizers: IgE‐mediated type 2 inflammation Low molecular weight sensitizers: haptens Irritant‐induced: no immunological reaction Total IgE, skin prick tests Peak flow monitoring Specific bronchoprovocation challenge Occupational rhinitis, reactive upper airways dysfunction syndrome Occupational asthma, reactive airways dysfunction syndrome Exposure avoidance, respiratory protection devices, ?anti‐IgE Aspirin/non‐steroidal anti‐inflammatories Constitutive overexpression of 5‐lipoxygenase and leukotriene C4 synthase Aspirin/NSAID challenge, urinary leukotriene E4 CRSwNP Severe asthma Avoidance, desensitization, antileukotrienes Staphylococcus aureus enterotoxin Superantigens elicit Th1, Th17 and Th2 cytokines, including IL‐4, IL‐5, IL‐13 Staph aureus enterotoxin IgE's CRSwNP Severe asthma ?anti‐IgE Aspergillus/fungal hypersensitivity TH2 cells, IL‐4, IL‐5, IL‐13, IgE, eosinophilia Aspergillus skin test positivity, Aspergillus specific IgE, IgE > 1000 IU/mL, Blood eosinophil >500/μL, precipitating antibodies to Aspergillus Allergic fungal rhinosinusitis Allergic bronchopulmonary aspergillosis (asthma, bronchiectasis) Steroids, antifungal, ?anti‐IgE Upper respiratory tract infection Variable depending on microbe and underlying disease Respiratory cultures and nucleic acid tests Exacerbation of rhinitis/rhinosinusitis Exacerbation of asthma, COPD or bronchiectasis Antivirals, antibiotics, vaccinations Typical upper and lower airway clinical phenotypes associated with underlying treatable traits are listed. Taxonomy based on clinical phenotypes has significant limitations: each phenotype may represent comprise treatable traits, for example, CRSwNP and asthma may be neutrophilic or eosinophilic inflammation depending on the population studied. Hence, a more precise approach is needed to identify contributory treatable trait(s) in each case, which can aid in selection of targeted treatments. Question mark (?) indicates targeted therapies under investigation. Treatable traits and targeted therapies may be used to complement the current approaches for patient stratification and treatment selection—including non‐targeted treatments—shown in Table 1. 2 AIRWAY INFLAMMATION Airway inflammation is important in the pathogenesis of many upper and lower airway diseases. It is a dynamic trait which varies over time and is modulated by factors such as treatment, infection, environmental exposures, and natural history of disease.4 Clinicians commonly assume that asthma and chronic rhinosinusitis with nasal polyps (CRSwNP) are stereotypically eosinophilic in nature, and that chronic rhinosinusitis without nasal polyps (CRSsNP) or COPD is stereotypically neutrophilic. As discussed below, while these generalizations may hold true for many cases there is substantial variability in the underlying pathophysiological mechanisms for each clinical phenotype or disease. Identification and quantification of airway inflammation can help to direct appropriate treatment. Current methods to assess upper and lower airway inflammation are summarized in Table 3. Method Comments Strengths Limitations Nasal lavage Luminal proteins, cytokines and cells Simple and safe Correlates with, but less invasive than induced sputum Variable fluid recovery and dilution with lavages No sampling of the epithelium Nasal brush or scraping Samples the nasal epithelium Simple and minimally invasive Does not sample deeper layers of the mucosa or the nasal lumen Nasal biopsy Identifies tissue inflammatory cellular subtypes of rhinitis or rhinosinusitis Samples the total nasal mucosa Technically difficult No information about nasal lumen Blood periostin Marker of Th2, eosinophilic airway inflammation Predicts severity of chronic rhinosinusitis Predicts chronic rhinosinusitis with nasal polyps phenotype Noninvasive Not widely available Conflicting evidence on whether periostin is associated with sputum eosinophilia Sputum induction Neutrophil count ≥65% or >500 × 104/mL identifies neutrophilic airway inflammation Eosinophil count ≥3% identifies eosinophilic airway inflammation Noninvasive Well‐validated Not widely available Logistically, technically and practically difficult High failure rate Risk of inducing bronchoconstriction Peripheral blood eosinophils Eosinophil count ≥2% identifies eosinophilic airway inflammation Noninvasive, minimal risk Widely available, cheap Predicts with important clinical outcomes such as exacerbation risk and response to treatment Wide diurnal variation of readings (40%) within the same subject May not be applicable when only local mucosal inflammation occurs Fractional exhaled nitric oxide (FeNO) FeNO < 25 ppb indicates low likelihood of airway eosinophilia FeNO > 47 ppb suggests eosinophilic airway inflammation and corticosteroid responsiveness Noninvasive Quick, easy, and safe to perform Readings may be affected by smoking, respiratory tract infections Sputum eosinophilia and FeNO are not always concordant Urinary leukotriene E4 (LTE4) Marker of cysteinyl leukotriene activity Higher levels observed in aspirin‐exacerbated respiratory disease Collection of urine is simple and of no risk Special equipment and training are required to measure LTE4 2.1 Allergic asthma‐rhinitis The link between allergic asthma and allergic rhinitis (AR) is well established. Up to 40% of AR patients have asthma and almost all patients with asthma have AR.5 Many patients with AR also exhibit nonspecific bronchial hyperresponsiveness.6 IgE‐mediated hypersensitivity to aeroallergens results in type 2/eosinophilic inflammation in the respiratory tract, manifesting as rhinitis in the upper airways and asthma in the lower airways. Upon allergen exposure, sensitization occurs whereby specific IgE (sIgE) are generated, circulate in the blood, and attach to the surface of mast cells and basophils, including those that reside in the airway mucosa. Subsequent exposure to allergen triggers mast cells degranulation, release of chemical mediators, and synthesis of leukotrienes and cytokines, resulting in an immediate allergic airway reaction. The late phase response involves Th2 lymphocyte recruitment with production of inflammatory cytokines such as IL‐4, IL‐5, and IL‐13, as well as activation of eosinophils. The central role of sIgE in allergic asthma‐rhinitis is highlighted by a population‐based birth cohort study showing that presence of peripheral blood sIgE directed against common allergens is associated with increased odds of asthma and rhinitis in childhood.7 In adults with AR, experimental nasal provocation with house dust mite allergens induces not only nasal eosinophilia but also bronchial eosinophilic inflammation, bronchoconstriction, and bronchial hyperresponsiveness,8 suggesting inflammatory crosstalk between upper and lower airways. The allergic asthma‐rhinitis phenoendotype usually begins in childhood and can be identified by measuring peripheral blood total IgE which is elevated, plus demonstration of atopy by the presence of raised sIgE levels in blood or skin prick test positivity. Omalizumab, an anti‐IgE monoclonal antibody, is efficacious for both severe allergic asthma9 and uncontrolled AR.10 Therapeutic response to omalizumab is predicted by higher baseline periostin levels and reduced free serum IgE levels during period of treatment.11 In some cases, specific causative allergens can be identified and targeted with allergen immunotherapy. In both AR12 and allergic asthma,13 immunotherapy improves symptoms and reduces the need for pharmacotherapy in the short term, albeit with limited evidence for longer‐term efficacy. Treatment of AR patients with grass pollen sublingual immunotherapy tablets was associated with slower AR progression, less frequent asthma onset, and slower asthma progression.14 Candidate biomarkers that can identify and/or predict response to allergen immunotherapy include the sIgE:total IgE ratio, specific IgG4 and IgE‐FAB.15 2.2 Local allergic rhinitis‐non‐allergic asthma: local mucosal IgE production A subset of asthma and rhinitis patients demonstrate type 2/eosinophilic allergic airway responses mediated by sIgE which is produced and sequestered in the airway mucosa, with skin prick test non‐reactivity and no detectable circulating sIgE in peripheral blood. In the upper airway, this phenoendotype is known as local allergic rhinitis (LAR), a distinct entity characterized by local production of sIgE in the nasal mucosa without evidence of systemic atopy. LAR does not appear to represent a forme fruste of AR,16 and is associated with increased risk of asthma compared to healthy individuals without LAR.17 The lower airway analog of LAR may correspond to a subtype of non‐allergic asthma which is currently not well defined, but whose existence is supported by studies (Supporting Information: References S1‐S4). In the aforementioned entity, intramucosal IgE synthesis is observed in the lower airways, and functional sIgE can be found in the sputum despite negative skin prick tests and blood sIgE levels. Bronchial inhalational challenge with allergen induces a clinical lower airway allergic response characterized by sputum eosinophilia and bronchial hyperresponsiveness. It is not known why individuals with allergic asthma‐rhinitis demonstrate positive skin prick tests and elevated sIgE in blood, whereas non‐allergic asthmatics and patients with LAR do not although local mucosal IgE synthesis occurs in all those conditions. One possibility is that local mucosal IgE synthesis may not reach sufficient quantities to spill over into peripheral blood. Alternatively, IgE may be sequestered within the bronchial mucosa. Regardless, efficacy of omalizumab has been shown in non‐atopic asthma with evidence of local mucosal IgE production. Pillai et al. (Supporting Information: References S5) randomized 18 such subjects and obtained bronchial biopsies from subjects before and after treatment with 12‐14 weeks of omalizumab. The patients subsequently underwent supervised dose reduction of their regular asthma therapy. Compared to placebo, omalizumab therapy significantly reduced total bronchial mucosal IgE+ cells. Lung function declined in the placebo‐treated group but improved in the omalizumab‐treated group. This small but important study serves as proof‐of‐concept that anti‐IgE can target disease associated with local mucosal IgE production. 2.3 Non‐allergic rhinitis with eosinophilia syndrome‐non‐allergic eosinophilic asthma Type 2 (eosinophilic) inflammation can occur in UAD without evidence of IgE‐mediated hypersensitivity. In the upper airways, this endotype has been termed non‐allergic rhinitis with eosinophilia syndrome (NARES) and is characterized by persistent rhinitis and marked eosinophilia of the nasal mucosa in the absence of local or systemic IgE production, suggesting allergen‐independent induction of eosinophilic airway inflammation.18 Almost half of patients with NARES who do not report asthmatic symptoms nevertheless demonstrate bronchial hyperresponsiveness and sputum eosinophilia, supporting a link between NARES and asthma.19 The corresponding asthma phenoendotype is non‐allergic eosinophilic asthma, in which eosinophilic inflammation and type 2 cytokine production are not driven by allergen‐activated Th2 lymphocytes, but is instead postulated to be initiated by type 2 innate lymphoid cells (ILC2).20 ILC2s are a recently discovered population of lymphocytes capable of producing large amounts of type 2 cytokines (IL‐5 and IL‐13) and orchestrate eosinophilic inflammation in the absence of adaptive immunity, including Th2 cells, B cells, and IgE antibody.21 ILC2s are activated by epithelium‐derived cytokines IL‐25, IL‐33, and thymic stromal lymphopoietins (TSLP), which are released by epithelial cells in response to epithelial damage caused by a wide range of stimuli including microbiota and pollutants. ILC2s are increased in peripheral blood in eosinophilic asthmatics compared to non‐eosinophilic asthmatics and their numbers in sputum correlate with eosinophil counts.22 Increased ILC2s has also been observed in eosinophilic nasal polyps,23 but evidence for the involvement of ILC2 cells in NARES and non‐allergic eosinophilic asthma is lacking and should be the focus of future studies. Potential targeted treatments for this pathway include monoclonal antibodies targeting any of the type 2 cytokines, discussed in the section below. TSLP, targeted by the monoclonal antibody tezepelumab, was shown in a recent dose‐ranging proof‐of‐concept study to reduce exacerbations in patients with uncontrolled asthma, including in those with low IgE (<100 IU/mL).24 Antagonists of CRTH2, the receptor for prostaglandin D2 on ILC2 cells, are currently undergoing evaluation to assess clinical efficacy (Supporting Information: References S6, S7). 2.4 Asthma‐chronic rhinosinusitis Chronic rhinosinusitis (CRS) is broadly defined as an inflammatory disorder of the paranasal sinuses and mucosa of the nasal passages that lasts 12 weeks or longer. The condition is heterogeneous, with 2 broad clinical phenotypes, with nasal polyps (CRSwNP) and without nasal polyps (CRSsNP), as well as diverse endotypes.25 Epidemiologic studies have demonstrated higher prevalence of CRS in asthmatics compared to non‐asthmatics across different populations.26, 27 However, the inflammatory endotype of CRS varies according to the population being studied. In Caucasian subjects, CRSwNP is predominantly eosinophilic and CRSsNP is predominantly neutrophilic, whereas in Chinese subjects CRSwNP is mainly neutrophilic, with fewer Chinese subjects demonstrating eosinophilic nasal polyps.28 Various studies have elucidated the detailed phenoendotypes of either CRS or asthma separately, but there is a paucity of data about phenoendotypes of combined asthma‐CRS. Despite this, phenoendotypes of asthma and CRS are remarkably analogous to each other. Based on sputum inflammatory cells, asthma can be eosinophilic, neutrophilic, mixed eosinophilic and neutrophilic, or paucigranulocytic.29 Based on bronchoepithelial cell gene expression profiles, asthma inflammatory signatures can be partitioned into “Th2 high” (associated with elevated IL‐4, IL‐5, IL‐13, sputum, and blood eosinophilia) or “Th2 low”.30 CRS endotypes can be similarly classified into “IL‐5 high” and “IL‐5 low” clusters.31 Subjects with IL‐5 high CRS have high eosinophilic cationic protein and IgE from nasal specimens, and present clinically with CRSwNP and asthma. Low IL‐5 clusters clinically exhibit the CRSsNP phenotype. “IL‐5 high” CRS appears to be the upper airway analog of “Th2‐high” asthma. The inflammatory pattern of the upper and lower airways in patients with nasal polyps and comorbid asthma was evaluated in a recent study using nasal polyp biopsies and exhaled nitric oxide.32 Approximately, two‐thirds of patients were eosinophilic and one‐third was non‐eosinophilic, with concordance demonstrated between the upper and lower airways in individual subjects. More studies are needed to clarify concordance between upper and lower airway phenoendotypes across the spectrum of inflammatory endotypes in asthma and CRS. Systemic, inhaled, or intranasal corticosteroids are cost‐effective treatments for eosinophilic upper and lower airway inflammation. In cases that are refractory to corticosteroids or in patients who are unable to tolerate high doses of corticosteroids, several biologics targeting eosinophilic inflammation can be considered, such as anti‐IL‐13 (tralokinumab), anti‐IL‐5 (mepolizumab and reslizumab), antibodies against alpha chain of IL‐5 receptor (benralizumab), and alpha subunit of the IL‐4 receptor (dupilumab). Mepolizumab is efficacious in both eosinophilic nasal polyposis and severe eosinophilic asthma,33, 34 reducing polyp burden and need for surgery in the former and reducing exacerbations in the latter. Similarly, dupilumab was demonstrated in small phase 2 trials to reduce polyp burden in patients with chronic sinusitis and nasal polyposis,35 and was found to decrease exacerbations and improve forced expiratory volume in one second (FEV1) in patients with severe eosinophilic asthma.36 Patients with asthma and nasal polyps show greater improvement in Asthma Control Questionnaire score with reslizumab compared to asthmatics without nasal polyps.37 2.5 Chronic obstructive pulmonary disease‐chronic rhinosinusitis Chronic obstructive pulmonary disease is a disease characterized by chronic respiratory symptoms and persistent airflow limitation caused by significant exposure to noxious particles or gases, most commonly tobacco smoke. Although COPD has traditionally been thought to be a purely lower airway disease, there is increasing evidence to suggest frequent concomitant involvement of the upper airways. Multiple epidemiologic studies have shown a high prevalence of nasal symptoms among COPD patients. An early population‐based epidemiologic study conducted in the Swedish general population in 1992 found that 40.1% of individuals with self‐reported chronic bronchitis or emphysema had symptoms of rhinosinusitis, compared with 32.7% in the total study sample.38 In a follow‐up study in 2000, those who reported thick, yellow, nasal discharge and nasal blockage in 1992 had a higher incidence of COPD diagnosis over an 8‐year follow‐up period.39 Another cross‐sectional population‐based Canadian study also found an association between CRS and COPD, both self‐reported.40 Self‐report may lead to misclassification errors of COPD and sinonasal disease but are an inherent limitation of large‐scale epidemiologic studies. Medical record diagnoses, which are considered more reliable than self‐reported medical history, were used in one study from Taiwan which extracted International Classification of Diseases diagnostic codes from a national insurance database and found an increased risk of CRSsNP among COPD patients.41 Recent studies have utilized more stringent and objective criteria for classifying COPD and rhinosinusitis. Bergqvist et al.42 used spirometry for identifying COPD, defined as FEV1/forced vital capacity (FVC) ratio <0.7, and found that the 5‐year incidence of nasal symptoms without having a cold was higher in subjects who demonstrated obstruction on spirometry compared to subjects without baseline spirometric evidence of airflow limitation. Hansen et al43 evaluated the prevalence of paranasal sinus opacification at magnetic resonance imaging in COPD and asthma patients compared to a control group. COPD was defined using similar spirometric criteria as the Bergqvist et al study, whereas asthma was self‐reported. COPD and asthma patients were, respectively, found to be 6 and 2 times more likely than healthy controls to demonstrate paranasal sinus opacification. These findings support the link between upper and lower airway diseases in both asthma and COPD, but we cannot conclude from this study that the association with sinonasal disease is stronger for COPD than for asthma because misclassification bias may have occurred among asthma patients, who were diagnosed based on self‐report instead of objective evidence of variable airflow limitation. Similarly, uncontrolled COPD cohort studies have also found a high prevalence of up to 75% of sinonasal disease, when using spirometry for COPD diagnosis and CT paranasal sinuses for diagnosis of CRS.44, 45 An important limitation of all the aforementioned studies is the use of prebronchodilator, instead of postbronchodilator, FEV1/FVC ratio for identifying cases of COPD. Such a definition could result in other obstructive lung diseases (such as asthma or bronchiectasis) being misclassified as COPD. Only one small study thus far has evaluated COPD using postbronchodilator spirometry and found higher nasal symptoms scored on a visual analogue scale compared to healthy age‐matched controls.46 Although prevalence of sinonasal disease does not increase with declining FEV1,44, 45 the presence of sinonasal disease has profound effects on other important COPD‐related outcomes such as symptom morbidity and exacerbations, which are known to be relatively independent of FEV1. Upper airway symptoms worsen over the natural history of COPD,47 and are associated with lower airway symptoms44 and impaired quality of life.48 Sinonasal symptoms also feature prominently during COPD exacerbations, and are associated with frequent exacerbations47 as well as treatment failure of acute exacerbation of COPD.49 Tobacco smoke forms the common link between COPD and CRS,50 likely by inducing neutrophilic inflammation in the nasal and bronchial mucosa. Compared with non‐smoking healthy controls, smokers without COPD present with nasal inflammation characterized by increased neutrophils and CD8 T lymphocytes51 although this has not been consistently demonstrated.52 Nasal and bronchial IL‐8 levels, neutrophil counts, and CD8 Th1 cells are even higher in smoking COPD patients than in smoking “healthy” controls matched for pack‐year smoking history.51, 53 Nasal and sputum IL‐8 concentrations correlate in COPD patients,53 indicating concordance of neutrophilic inflammation between upper and lower airways in keeping with the UAD hypothesis. Release of neutrophil‐derived proteases such as elastase and matrix metalloproteinase‐9 causes alveolar wall destruction and mucous hypersecretion characteristic of COPD, and may contribute to tissue destruction and remodeling of the sinonasal mucosa seen in CRS. Neutrophilic inflammation results in airflow limitation in the lower airways in COPD, which is mirrored in the upper airways by reduced nasal patency54 and higher nasal resistance.55 Severity of lower airway obstruction, as measured by FEV1, correlates with severity of CRS symptoms.45 Smoking cessation alters the progressive clinical course of COPD by reducing lung function decline and mortality, hence remaining a cornerstone of COPD treatment in addition to other medical therapies. Additionally, in CRS, smoking cessation improves symptoms, quality of life and decreases CRS‐related antibiotic use.56Neutrophilic airway inflammation per se may potentially be targeted for treatment using macrolides, which exert a repertory of effects that suppress neutrophilic inflammation, including downregulation of IL‐8, elastase and matrix metalloproteinase‐9, and reduction of neutrophil chemotaxis and survival.57 Macrolides have proven efficacy in decreasing the frequency of COPD exacerbations,58 and in a small double‐blind randomized controlled trial was associated with improvement in symptoms, endoscopic scores, and nasal lavage IL‐8 levels specifically in Th1, neutrophilic CRS patients.59 Further studies are needed to confirm the efficacy of macrolides. In addition, long‐term macrolides should only be used in carefully selected patients after optimization of other standard medical therapies because of potential cardiac and otological adverse effects, as well as risk of inducing antibiotic resistance. 3 IMPAIRED AIRWAY MUCOSAL DEFENCE Major components of the airway mucosal defense mechanism include the mucociliary escalator, immunoglobulins, antimicrobial peptides, the epithelial barrier, and cells of the innate and adaptive immune system. Impaired mucosal airway defense has a causal role in several phenoendotypes of UAD. The predominant lower airway phenotype in UAD due to impaired mucosal defense is bronchiectasis, which is characterized by abnormal and irreversible dilatation of the lower airways. Frequent coexistence of bronchiectasis and sinonasal disease has long been recognized.60 Patients with bronchiectasis have a high prevalence of CRS when assessed using European Position Paper on Rhinosinusitis and Nasal Polyps criteria, that is, presence of symptoms and positive findings on either nasoendoscopy or CT of paranasal sinuses.61, 62 Conversely, patients with CRS more frequently demonstrate CT evidence of dilated airways compared to individuals without CRS.63 3.1 Cystic fibrosis: bronchiectasis‐chronic rhinosinusitis due to airway surface liquid viscosity Airway surface liquid (ASL) viscosity is regulated in part by the cystic fibrosis transmembrane regulator (CFTR) protein, which mediates ion transport at the airway epithelial surface. Pathological processes which impair CFTR activity can result in abnormal ion flux at the epithelial surface, increased sodium reabsorption, periciliary liquid depletion, and increased mucous viscosity, thereby impairing mucociliary clearance.64 Loss of CFTR‐mediated bicarbonate secretion in the airway epithelium also results in acidification of the ASL, which impairs the action of antimicrobial peptides in the ASL, rendering the airway more susceptible to colonization and infection by microbes.64 Cystic fibrosis (CF) is due to heterogeneous autosomal recessive mutations in the CFTR gene and affects mucociliary clearance at the sinoepithelial lining, lower airways, gastrointestinal tract, and exocrine organs. Sinonasal disease is ubiquitous in CF and characterized by neutrophilic polyps with acid mucin glands, mucoceles, or pseudomucoceles.65 Lower airway disease is characterized by intense neutrophilic inflammation resulting in bronchiectasis, which occurs in 70%‐75% of children by 5 years of age and represents the major cause of morbidity and death in this condition. Treatment of CF has been transformed by the discovery of novel CFTR modulators aimed at addressing the specific defects causing reduced CFTR activity. The most common gene mutation in CF is F508del which causes a processing defect that severely reduces CFTR levels at the epithelial surface and also disrupts channel opening. The latter gating defect is also seen in other mutations, such as the G551D mutation. Lumacaftor and tezacaftor are both CFTR correctors, which improve trafficking of CFTR to the epithelial cell surface. Ivacaftor is a CFTR potentiator and acts by increasing the probability of the channel remaining open. Randomized controlled trials have proven that treatment with these compounds is associated with improved lung function in CF,66-69 and improvement of CF‐associated CRS following treatment with CFTR modulators has been reported anecdotally.70, 71 There is emerging evidence that CFTR dysfunction is a trait in other UADs and may be a target for treatment. CF carrier mutations are implicated in CRS patients who do not have CF.72 Constituents in tobacco smoke such as cadmium, arsenic, and acrolein can cause CFTR dysfunction,73 either by reducing CFTR expression or by inducing gating defects. CFTR decrements are also seen in COPD, including reduced nasal and lower airway potential difference74 and elevated sweat chloride,75 and the degree of CFTR impairment correlates with severity of symptoms and airflow limitation.74, 75 Ivacaftor reverses the acute reduction of CFTR activity after exposure to cigarette smoke in human bronchoepithelial cells,76 and has been demonstrated to ameliorate chronic bronchitis symptoms and improve CFTR activity in a preliminary study.77 Roflumilast, a phosphodiesterase‐4 inhibitor, can stimulate CFTR by increasing cyclic adenosine monophosphate and reverses smoke‐induced inhibition of CFTR activity.78 Roflumilast is approved for reducing exacerbations in severe COPD associated with chronic bronchitis. 3.2 Primary ciliary dyskinesia (PCD): bronchiectasis‐chronic rhinosinusitis due to ciliary dysfunction Chronic rhinosinusitis and bronchiectasis are universal in adults with PCD. The underlying pathophysiology is absent or impaired ciliary motility due to heterogeneous recessive mutations affecting the biogenesis of cilia lining the upper and lower airways.79 In turn, this leads to impaired mucociliary clearance manifesting in the neonatal period as difficulty feeding due to nasal blockage/rhinitis, neonatal respiratory distress syndrome and later in life as recurrent sinopulmonary infections, and bronchiectasis. Nasal nitric oxide is an excellent screening test for patients with a history suggestive of PCD.80 Other tests to assess for ciliary dyskinesia include analysis of ciliary ultrastructure by electron microscopy, assessment of ciliary beat frequency and pattern by high resolution, high‐speed video microscopy, and immunofluorescent staining of ciliary proteins. Ciliary dysfunction occurs in other airway diseases but is usually acquired, for example, due to smoking or infection. In CRS, blunted ciliary response to environmental stimuli has been demonstrated, and this is reversed once the tissue is removed from infected/inflamed environment.81 In COPD, cigarette smoke is associated with ciliary abnormalities in both bronchial82 and nasal mucosae.83 Sputum from atopic and non‐atopic asthmatics have ciliary inhibitory effect on human bronchoepithelial cell cultures, which was reversible upon removal of the sputum.84 Bronchial biopsies from asthma patients demonstrate reduced ciliary number which recovers after treatment with inhaled corticosteroids.85 There are currently no validated effective therapies for primary ciliary dyskinesia and this should be a focus for future research. In acquired cases, removal of the causative factor may reverse ciliary impairment. 3.3 Primary antibody deficiency The most common immunodeficiencies associated with CRS and bronchiectasis are primary antibody deficiencies,86 of which common variable immunodeficiency (CVID), characterized by reduced serum IgG concentrations, is the most common in adults. One‐third and two‐thirds of patients with CVID have bronchiectasis and recurrent sinusitis, respectively.87 Specific antibody deficiency, which is associated with normal immunoglobulin and IgG subclass levels but impaired antibody response to encapsulated respiratory pathogens such as Streptococcus pneumoniae and Hemophilus influenza, is also associated with a high prevalence of CRS and bronchiectasis, accounting for as many as 50% of idiopathic bronchiectasis.88 Patients with primary antibody deficiency most likely require lifelong immunoglobulin replacement, vaccinations, and prophylactic antibiotics. There is also some evidence that antibody deficiency may be a treatable underlying contributor in other airway diseases. For example, low circulating IgG levels are associated with higher rate of exacerbations and shorter time to next exacerbation in two COPD cohorts.89 Treatment with immunoglobulin replacement is associated with significant reduction in exacerbations and rescue antibiotic use in a small case series of 14 COPD patients (Supporting Information: References S8), warranting future prospective, randomized clinical trials to assess the efficacy of IVIG in preventing exacerbations in selected COPD patients with antibody deficiency. A potential contributing role of antibody deficiency in asthma has also been suggested by a small case‐control study which found that asthma patients are more likely to have a diagnosis of CVID or selective IgA deficiency than non‐asthmatic individuals, potentially accounting for the increased risk of bacterial infections in some individuals with asthma.90 A preliminary proof‐of‐concept study found that IVIG significantly reduces the frequency of asthma exacerbations and improved asthma control in adults with IgG subclass deficiency.91 4 EXOGENOUS COFACTORS The respiratory tract is continuously in contact with air from the environment which contains particulates such as microbes, pollutants, and other noxious agents that can cause or aggravate UAD. Aeroallergens such as house dust mites, pollen, and animal dander are the archetypal examples of how specific exposures can cause and worsen allergic asthma‐rhinitis. There are several other well‐delineated phenoendotypes of UAD in which specific inciting or aggravating cofactors are implicated. 4.1 Occupational asthma‐rhinitis More than 90% of individuals with occupational asthma (OA) also have occupational rhinitis (OR).92 Different occupational exposures can cause OA‐OR by different mechanisms. High molecular weight sensitizers, such as proteins and glycopeptides, often cause symptoms after a latency period via IgE‐mediated hypersensitivity.93 In contrast, the mechanism by which low molecular weight agents induces OA‐OR is poorly understood. Single or multiple exposures to high concentrations of irritants can lead to nonspecific inflammation in both upper and lower airways, clinically recognized as reactive upper airways dysfunction syndrome and reactive airways dysfunction syndrome, respectively. In addition to usual care for asthma and rhinitis, the management of OA‐OR is centered around exposure avoidance and use of appropriate respiratory protection equipment. The efficacy of omalizumab for sensitizing agents with IgE‐dependent reactions requires future studies. 4.2 Aspirin‐exacerbated respiratory disease The clinical phenotype of aspirin‐exacerbated respiratory disease (AERD) consists of a triad of asthma, CRSwNP, and exacerbation of upper and lower respiratory tract symptoms within 30 minutes to 3 hours after exposure to aspirin and other cyclooxygenase‐1‐inhibiting drugs. Nasal polyps in AERD can be distinguished from CRSwNP in individuals without AERD by the presence of enhanced eosinophilia, GM‐CSF, and decreased tissue plasminogen activator.94 The endotype of AERD is not fully understood but current available data suggest an underlying dysregulation of arachidonic acid metabolism, with constitutive overproduction of proinflammatory cysteinyl leukotrienes and decreased levels of anti‐inflammatory mediators such as prostaglandin E2.95 This dysregulation is aggravated by cyclooxygenase inhibition following exposure to aspirin and other non‐steroidal anti‐inflammatory drugs in susceptible individuals. The gold standard for diagnosing AERD is aspirin challenge, but challenge testing carries the risk of severe bronchospasm. Urinary leukotriene E4, the end‐product of arachidonic acid metabolism, is raised in AERD and may have a role in diagnosing AERD.96 Treatment options include avoidance of aspirin and NSAIDs, desensitization, and leukotriene receptor antagonists. Omalizumab and mepolizumab may be helpful in AERD as suggested by small trials involving subsets of patients with AERD (Supporting Information: References S9, S10). Whether the observed clinical efficacy of these biologics can be attributed to target AERD‐specific endotype above and beyond their effect on type‐2 eosinophilic inflammation is unclear. 4.3 Asthma‐CRS: hypersensitivity to Staphylococcus aureus enterotoxins Staphylococcus aureus is consistently found in nasal tissues of CRS patients, and evidence supports a role for S. aureus in the pathogenesis of CRS‐asthma. S. aureus colonizes nasal polyps and secretes proteins, including enterotoxins and serine protease‐like proteins in the nasal mucosa, which have the ability to elicit a polyclonal IgE response resulting in an intense type 2, IL‐5 high, eosinophilic inflammation which is likely responsible for chronic severe inflammation of the airways. Presence of sIgE to S. aureus enterotoxins in nasal polyp tissue is strongly associated with asthma in patients with CRSwNP,31, 97 providing an etiological link between asthma‐CRS. In a small placebo‐controlled, proof‐of‐concept study, omalizumab was found to ameliorate nasal polyps and airway symptoms in a small cohort of patients with asthma‐CRSwNP who demonstrated sIgE toward S. aureus enterotoxins.98 4.4 Allergic bronchopulmonary aspergillosis‐allergic aspergillus rhinosinusitis: fungal hypersensitivity IgE‐mediated hypersensitivity to Aspergillus hyphae is the underlying endotype for allergic bronchopulmonary aspergillosis (ABPA) and allergic aspergillus rhinosinusitis (AARS). Upper airway disease is characterized by pansinusitis, passage of nasal plugs, and recurrent nasal polyposis. Lower airway disease is characterized by mucus inspissation within the bronchi and severe asthma, central bronchiectasis, mucous plugging, and fleeting pulmonary infiltrates. In a case series, 22/95 patients of ABPA had also concomitant AARS. All these patients had positive skin tests, positive precipitins, and raised total and sIgE (Supporting Information: References S11). This combination of ABPA and AAS has been termed “sinobronchial allergic mycosis” to recognize their shared etiology and association. The optimal treatment of ABPA and AAS is not known but includes the use of steroids, antifungals, or omalizumab. 5 CONCLUSION The evidence for UAD is strongest for asthma and AR/CRS. However, close associations between upper airway diseases and other lower respiratory tract disorders can also be appreciated. UAD is complex and heterogeneous but a “treatable traits” paradigm, based on assessment of airway inflammation, mucosal defense dysfunction, and exogenous cofactors can help to identify pathophysiological mechanisms involved in the development of disease and potential targets for intervention in individuals presenting with combined upper and lower airway disease. A given individual with disease can demonstrate multiple traits which may vary over time. We have summarized the various treatable traits in UAD and their proven or notional targeted treatments in Table 2. Appreciation of these treatable traits is necessary toward advancing the precision medicine approach in UAD. CONFLICT OF INTEREST The authors declare that they have no conflicts of interest. Supporting Information REFERENCES Notes : AR, allergic rhinitis; CAT, COPD Assessment Tool; CRSsNP, chronic rhinosinusitis without nasal polyps; CRSwNP, chronic rhinosinusitis with nasal polyps; FEV1, forced expiratory volume in one second; ICS, inhaled corticosteroids; ICS + LABA, combination inhaled corticosteroids and long‐acting beta agonist; LABA, long‐acting beta agonist; LAMA, long‐acting muscarinic antagonist; mMRC, modified Medical Research Council dyspnea score. Typical upper and lower airway clinical phenotypes associated with underlying treatable traits are listed. Taxonomy based on clinical phenotypes has significant limitations: each phenotype may represent comprise treatable traits, for example, CRSwNP and asthma may be neutrophilic or eosinophilic inflammation depending on the population studied. Hence, a more precise approach is needed to identify contributory treatable trait(s) in each case, which can aid in selection of targeted treatments. Question mark (?) indicates targeted therapies under investigation. Treatable traits and targeted therapies may be used to complement the current approaches for patient stratification and treatment selection—including non‐targeted treatments—shown in Table 1. Citing Literature Number of times cited: 2 Ana Margarida Pereira, Cristina Jácome, Rute Almeida and João Almeida Fonseca, How the Smartphone Is Changing Allergy Diagnostics, Current Allergy and Asthma Reports, 10.1007/s11882-018-0824-4, 18, 12, (2018). Crossref Umut Can Kucuksezer, Cevdet Ozdemir, Mubeccel Akdis and Cezmi A. Akdis, Precision/Personalized Medicine in Allergic Diseases and Asthma, Archivum Immunologiae et Therapiae Experimentalis, 10.1007/s00005-018-0526-6, (2018). Crossref

Via Krishan Maggon

Background: The mechanism underlying transient increases in immunoglobulin (Ig) A concentrations in the cecal contents of rats fed fructo-oligosaccharide (FOS) is unclear.
Objective: This study was designed to test whether increased IgA concentrations represent one aspect of the inflammatory response to increased permeability induced by FOS in the cecum.
Methods: Seven-week-old male Wistar rats were fed a fiber-free semipurified diet (FFP) with or without supplemental FOS (60 g/kg diet) for 9 or 58 d [experiment (expt.) 1], 7 d (expt. 2), or 7 or 56 d (expt. 3). In addition to measuring IgA concentrations in cecal content, we assessed gut permeability, inflammatory responses (expt. 1), the number of IgA plasma cells in the cecal lamina propria, polymeric Ig receptor (pIgR) expression in the cecal mucosa (expt. 2), and the condition of the cecal mucus layer (expt. 3).
Results: The cecal IgA concentration in the FOS-fed rats was 15-fold higher than that of the rats fed FFP for 9 d (P < 0.05). Gut permeability estimated by urinary chromium-EDTA excretion, bacterial translocation to mesenteric lymph nodes, myeloperoxidase activity, and expression of inflammatory cytokine genes in the cecal mucosa was greater in the FOS-fed rats than in the rats fed FFP for 9 d. These effects were not observed in the rats fed FOS for 58 d (expt. 1). Accompanying the higher cecal IgA concentration, pIgR protein and the number of IgA plasma cells in the cecal mucosa were higher in the FOS-fed rats than in the rats fed FFP for 7 d (expt. 2). Destruction of the mucus layer on the epithelial surface, as evidenced by Alcian blue staining in the cecal sections, was evident in the rats fed FOS for 7 d, but the mucus layer appeared normal in the rats fed FOS for 56 d (expt. 3).
Conclusions: These findings suggest that transient increases in cecal IgA concentrations induced by FOS in rats are associated with mucosal inflammation in response to increased gut permeability; these are presumably evoked by disruption of the cecal mucus barrier. The observed responses could contribute to the maturation of the gut immune system.

Mucosal immunity is now one of the hottest topics among fish scientists working in a wide range of disciplines. So far, in several teleosts species it has been clearly demonstrated the presence of a full set of immune cells necessary to elicit strong immune responses at the mucosal sites like the gills, gonads, gut and skin. These sites are collectively regarded as mucosal associated lymphoid tissues (MALT). However, the study of such structures in fish need special attention due to the fact they are functionally different from the more commonly studied mammalian counterparts. Particularly, at the intestine, a recently described fish specific immunoglobulin isotype IgT/IgZ has turned even more attractive the immune output of this organ. The IgT/IgZ, which resembles the mammalian IgA, is of particular interest to perform basic, developmental and comparative fish gut functional analyses. Nevertheless, it is suspected that IgT may have a relevant role in different areas dominated by MALTs, but further investigation is required. Among, the main functions of such active immunological sites are to protect the host against exposure to pathogens by means of the immune properties of the mucus, but also to avoid hypersensitivity reactions to food proteins, commensal microflora or to immune modulators (pre/probiotics, immunostimulants and vaccine adjuvants), which are frequently used in aquaculture as means to increase disease resistance. Several research papers claim the effectiv

Abstract Background Nonspecific induction of local innate immune responses by mucosally administered immunotherapy is a new approach to protection from upper respiratory tract infections. Therefore, a new liposome‐toll‐like receptor complex (LTC) immune stimulant was developed and investigated for its ability to activate innate immune responses in cats, both in vitro and in vivo, as part of an initial evaluation of LTC for use as an immunotherapeutic agent in cats. Objectives We hypothesized that LTC could activate innate immune responses in cats after topical application to nasal and oropharyngeal mucosal surfaces. Animals Mucosal immune responses to topical administration of LTC were assessed in 7 healthy, purpose‐bred cats, and in vitro responses were assessed using blood samples from healthy cats. Methods Cytokine and cellular immune responses to LTC were evaluated in blood samples, nasal lavage specimens, and pharyngeal swabs from cats, using reverse transcriptase polymerase chain reaction assays, ELISA assays, and flow cytometry. Results Liposome‐TLR complexes rapidly activated leukocytes in vitro, including upregulation of costimulatory molecule expression and cytokine production. Topical administration of LTC in healthy cats triggered rapid recruitment of monocytes to the nasal and oropharyngeal mucosa. Conclusions and Clinical Importance Liposome‐TLR complexes were found to effectively activate innate immune responses in cats after mucosal administration. These findings suggest that LTC have potential for use as a new mucosally administered immunotherapy for nonspecific protection from viral and bacterial respiratory tract infections. Abbreviations FBS fetal bovine serum IL interleukin INF interferon LTC liposome‐TLR complexes NALT nasopharyngeal‐associated lymphoid tissue NK natural killer OX40 tumor necrosis factor receptor superfamily, member 4 PBMC peripheral blood mononuclear cells PBS phosphate‐buffered saline RT‐PCR reverse transcriptase polymerase chain reaction TLR toll‐like receptor TNF tumor necrosis factor 1 INTRODUCTION Upper respiratory infection (URI) is a common and serious problem in cats in boarding facilities, animal hospitals, and shelters. Unfortunately, few quickly effective options are available for preventing these infections in newly exposed cats. Immunoprophylaxis may be most effective if focused at the likely portal of pathogen entry. The upper respiratory tract is the site of entry for several important respiratory pathogens of cats, consisting of a diverse array of viral and bacterial organisms, including feline herpesvirus 1 (FHV‐1), calicivirus, Mycoplasma sp., Bordetella bronchiseptica, and Chlamydia felis.1-6 Activation of host innate immune defenses, including airway epithelial cells and immune cells in oropharyngeal lymphoid tissues, results in production of antiviral and antibacterial cytokines, including type I (INF‐α) and II (INF‐γ) interferons, tumor necrosis factor alpha (TNF‐α), and interleukin 12 (IL‐12) and chemokines such as monocyte chemoattractant protein 1 and IL‐8 that recruit monocytes, T cells, and neutrophils. Local activation of innate immune responses may create an environment that is less susceptible to viral entry and replication or to bacterial invasion and colonization.7 Clinically, such an immune protective effect could be manifested as either partial or complete protection from infection or as a decrease in the severity and duration of clinical signs of illness, along with decreased viral or bacterial shedding. One potential solution to the problem of complex upper respiratory tract infections in cats would be an immunotherapeutic agent capable of inducing nonspecific immune activation at mucosal surfaces in the oropharynx, such as liposome‐toll‐like receptor complexes (LTC). These complexes have been shown previously to induce strong nonspecific protection from lethal bacterial and viral respiratory tract infections after intranasal administration in several different mouse infection models.8-12 In addition, we demonstrated induction of therapeutic immune responses in cats with chronic rhinitis after parenteral administration of LTC.13 Therefore, we reasoned that immune stimulatory complexes could be further optimized for mucosal delivery and generation of local mucosal immune responses in the upper respiratory tract of cats. Therefore, the original LTC formulation was modified to improve its innate immune stimulatory potency and to improve adhesion and contact time with mucosal surfaces. In the present study, the immune activating properties of modified LTC were evaluated in cats.12, 14 The modified LTC consisted of cationic liposomes formulated with 2 TLR agonists: polyinosinic‐polycytidylic acid (poly I:C), as a TLR3, retinoic acid‐inducible gene I, melanoma differentiation‐associated protein 5 agonist, and noncoding plasmid DNA (pDNA), as a TLR9 agonist rich in cytosine‐guanine. In addition, the modified LTC contained a mucosal adhesive agent (carboxymethylcellulose) to increase adhesion to mucosal cell surfaces. We also assessed cytokine and cell activation responses by feline leukocytes in vitro and also assessed the local mucosal immune responses to topical administration of LTC in healthy cats. 2 MATERIALS AND METHODS 2.1 Preparation of liposome‐TLR agonist complexes for mucosal immune stimulation Liposomes were prepared by dissolving the cationic lipid, 1,2‐dioleoyl‐3‐trimentylammonium‐propane (DOTAP) and cholesterol (Avanti Polar Lipids, Alabaster, AL) in chloroform, and the solution was dried overnight in a vacuum desiccator to a thin film. The lipids were rehydrated in 5% sterile dextrose in water at 50°C as described previously.12, 14 Poly I:C (InVivoGen, San Diego, California) and pDNA (noncoding commercial plasmid PCR2.1) Thermo Fisher Scientific, Waltham, MA were added to form liposome complexes, as reported previously.12, 14 The endotoxin content of the plasmid was between 0.04 and 0.25 EU/μg. Carboxymethylcellulose (Sigma‐Aldrich, St. Louis, Missouri) was added to the LTCs to increase mucosal adhesion. In studies in which fluorescent LTCs were delivered, the liposomes were formulated to contain 10% (vol/vol) Topfluor‐labeled cholesterol (Avanti Polar Lipids). 2.2 Animal studies Animal protocols were approved by the Institutional Animal Care and Use Committee of an independent contract research facility in Fort Collins, Colorado. Purpose‐bred cats for the studies were purchased from Liberty Research (Waverly, New York). All animals used in the studies were adopted to owners in the Fort Collins area after completion of the studies, and no animals were euthanized as part of these studies. For collection of nasal lavage samples, cats were restrained manually in a head‐down position, and 0.5 mL of prewarmed sterile phosphate‐buffered saline (PBS) solution was administered into each nostril, and the resulting fluid backflow was collected from the nostrils using a 15 mL conical tube. The procedure was repeated once for each nostril, and all of the fluid collected was pooled for later analysis and stored on ice. For oropharyngeal samples, alginate swabs were gently rubbed and rolled against the mucosa of the caudal oropharyngeal region and placed in tubes with 1 mL PBS and stored on ice before analysis. Nasal and oropharyngeal samples were collected 24 hours before LTC administration, 6 hours after LTC administration, and again 20 and 72 hours after LTC administration. Cats were treated with LTC administered on day 0. Control cats were treated with an equivalent volume of sterile PBS. Each cat was given 1 mL of LTC (0.2 mL intranasally in each nostril and 0.6 mL PO) while being gently restrained. In a separate experiment designed to track LTC uptake by mucosal cells in the nostril and oropharynx of cats, 1 mL of LTC labeled with TopFluor fluorescent dye was administered by instilling 0.2 mL intranasally in each nostril and 0.6 mL (equivalent to 60 μg of Poly I:C and PCR2.1) PO. Average weight of the cats was approximately 3 kg; thus, the intranasal dose of each TLR ligand was approximately 20 μg per kilogram body weight. No adverse effects, either acute or longer term, were observed after intranasal LTC administration (data not shown). 2.3 Preparation of peripheral blood mononuclear cells Whole blood was obtained by jugular venipuncture from a separate group healthy young adult cats and collected into EDTA tubes. For separation of peripheral blood mononuclear cell (PBMC), blood was diluted 1:2 with sterile PBS, then layered over a Ficoll (GE Healthcare, Uppsala, Sweden) gradient and centrifuged. The PBMC were collected and washed twice in PBS and then resuspended in complete tissue culture medium before flow cytometric analysis. 2.4 Cell culture and PBMC activation with LTC Peripheral blood mononuclear cells were cultured in complete medium, which consisted of Dulbecco's modified eagle medium (Thermo Fisher Scientific, Watham, Massachusetts) containing 15% fetal bovine serum (FBS; Avanti Polar Lipids), essential and nonessential amino acids, penicillin, and streptomycin (Gibco‐Thermo Fisher Scientific, Pittsburgh, Pennsylvania). Cells were plated in 96‐well flat bottom plates (Celltreat, Pepperell, Massachusetts) at a density of 1 × 106 cells/well in 200 μL medium. For assays involving LTC activation of PBMC, LTC were added at 3 different dilutions (1 μL per well, 0.3 μL per well, and 0.1 μL per well) in triplicate wells of PBMC in 100 μL complete medium, with careful mixing, and then the cells were incubated for an additional 48 hours. Conditioned medium was collected for IFN‐γ assay, and cells were collected for flow cytometric analysis of activation markers (see below) and for analysis of cytokine gene expression by quantitative reverse transcriptase polymerase chain reaction (qRT‐PCR) (see below). Assays were repeated at least twice, using PBMC from different donor animals. 2.5 Quantitative RT‐PCR analysis of cytokine gene expression in vitro Ribonucleic acid was extracted from PBMC incubated for 48 hours with LTC, as noted above, and expression of IL‐1, IL‐12, IFN‐α, IFN‐γ, and TNF‐α genes was determined using quantitative RT‐PCR. Briefly, RNA was isolated and reverse transcribed using a commercial kit (Qiagen, Germantown, Maryland), and the complementary DNA then was amplified using Cyber green primers (Bio‐Rad, Hercules, California) using a qPCR MX3000p system instrument (Agilent, Santa Clara, California). Cytokine transcript copy numbers were determined by quantitative RT‐PCR, as noted previously.13 2.6 IFN‐γ ELISA Cell culture supernatants from PBMC cultures were analyzed for IFN‐γ concentrations, using a feline IFN‐γ specific ELISA (DuoSet Feline IFN‐γ kit; R&D systems, Minneapolis, Minnesota) according to the manufacturer's instructions. 2.7 Flow cytometry Cells were harvested after 48 hours of in vitro stimulation and immunostained with the following antibodies conjugated with fluorochromes: CD5‐PE (clone f43; Southern Biotech, Birmingham, Alabama), CD21‐APC (clone CA2.1Dg; Bio‐Rad), and OX40‐FITC (clone 7D6; Bio‐Rad), or CD14‐APC (clone TÜK4; Bio‐Rad) and MHCII‐FITC (clone TÜ39; BD Biosciences, San Jose, California). Cells were immunostained with primary antibodies for 20 minutes at 4°C in FACs buffer (PBS with 2% FBS and 0.05% sodium azide) after a 5‐minute incubation with normal cat serum (Jackson ImmunoResearch) to block nonspecific binding. Cells obtained by nasal lavage and oropharyngeal swabs were washed once with PBS, then immunostained in 96‐well plates with the directly conjugated antibodies noted above. Isotype‐matched antibodies were used as control for each primary antibody. Cells from in vitro activated PBMC cultures were collected 48 hours after stimulation and measured using the same panel of antibodies, with appropriate controls. Cells were analyzed using a Beckman Coulter Gallios flow cytometer (Beckman Coulter, Indianapolis, Indiana), and data were analyzed using FlowJo software (Tree Star, Ashland, Oregon). 2.8 Preparation and analysis of nasal and pharyngeal samples Swabs were collected from the distal oropharynx and immediately placed in 2 mL of complete tissue culture medium in 15 mL polypropylene tubes on ice. The samples were lightly vortexed to free cells from the swabs. Fluid was transferred to a new tube, and 2 mL of PBS was added to the empty tube containing the swab, vortexed again, and pooled with the first sample. The sample then was filtered using a 70 μm cell strainer (Celltreat, Pepperell, Massachusetts) to remove debris and mucus. Samples were centrifuged, the cell pellets were resuspended in 0.5 mL of PBS, and the cells were counted. Nasal lavage samples were filtered, washed, centrifuged, resuspended, and counted in the same fashion as for oropharyngeal swab samples. Cells were stained in the same manner as described for PBMC, and equal numbers were analyzed by flow cytometry to determine the cell type and activation. 2.9 Confocal microscopy Monocyte‐derived macrophages were generated from cat PBMC by incubation of plastic adherent monocytes with 10 ng/mL human recombinant macrophage colony stimulating factor for 5 days. The macrophages that were generated were then incubated with 0.1, 1, or 2 μL per well of TopFluor‐labeled LTC in complete medium at 37°C for 4 hours, then washed gently 3 times, fixed with 2% paraformaldehyde for 30 minutes, washed again with distilled water, cover‐slipped, and analyzed by confocal microscopy. Confocal images of feline monocyte‐derived macrophages (untreated control cells or cells incubated with labeled LTC) were obtained using an Olympus IX3 confocal microscope (Waltham, Massachusetts). Images were processed and analyzed using Olympus cellSens software. 2.10 Statistical methods Statistical comparisons between data sets with 2 treatment groups were performed using the nonparametric Mann‐Whitney test. Comparisons among 3 or more groups were performed using analysis of variance, followed by the Tukey multiple means post‐test. Analyses were performed using Prism6 software (GraphPad, La Jolla, California). For all analyses, statistical significance was set at P < .05. 3 RESULTS 3.1 B cells, T cells, and monocytes from feline PBMC activated by LTC in vitro Feline lymphoid tissues express TLRs such as TLR3 and TLR9, and feline immune effector cells (T cells and B cells) express TNF receptor superfamily, member 4 (OX40). When bound to its ligand OX40L, OX40 augments immune responses by prolonging survival of T cells, generation of immune memory, preventing emergence of T‐cell tolerance, and mediating decreased immunosuppressive activity of regulatory T cells. Thus, we chose to examine the effect of in vitro treatment of PBMC with LTC on OX40 expression in these cells.13, 15-17 Unfractionated PBMC were used because LTC may have both a direct effect on leukocytes as well as an indirect effect through crosstalk among various cell types in the mixture. B lymphocytes that were CD21+ were found to exhibit significant upregulation of expression of the costimulatory molecule OX40, in a dose‐dependent fashion with respect to the amount of added LTC (Figure 1, panel A). For T‐cell responses to LTC stimulation, the percentage of CD5+ T cells expressing OX40 also increased significantly as a function of LTC concentration (Figure 1, panel B). Finally, a significant increase in expression of MHCII by CD14+ monocytes was observed in cultures of PBMC treated with LTC (Figure 1, panel C). Similar results were observed in 3 separate studies. 3.2 LTC treatment activates cytokine gene expression by PBMC Quantitative RT‐PCR analysis was used to assess induction of cytokine gene expression by PBMC treated with LTC (Figure 2). Significant upregulation of expression of mRNA for INF‐α as well as upregulation of IL‐1β, IL‐12, and TNF‐α expression after LTC stimulation was identified in most cats. 3.3 LTC treatment triggers IFN‐γ secretion by PBMC Supernatants were collected from PBMC after 48 hours stimulation with LTC, for analysis of IFN‐γ production (Figure 3). Liposome‐TLR complexes treatment stimulated a dose‐dependent, significant secretion of IFN‐γ secretion, as determined by ELISA analysis of supernatants. 3.4 Uptake of labeled LTC by macrophages in vitro Monocyte‐derived macrophages were generated from PBMC of cats by plastic adherence, followed by culture for 4 days in complete medium, after which macrophage uptake of LTC was assessed. Extensive uptake of LTC complexes by macrophages was observed (Figure 4), consistent with the known targeting of liposome complexes to monocytes and macrophages and other phagocytic immune cells. 3.5 Retention of labeled LTC by cells in the oropharynx and nasal cavity Seven healthy adult cats were given 0.1 mL of labeled LTC bilaterally into the nostrils and 0.6 mL PO. Nasal lavage and throat swab samples were obtained before administration, and 6, 20, and 72 hours after LTC administration; cell populations were analyzed by flow cytometry, as described in the Materials and Methods section. Compared to pretreatment samples, samples obtained from the nose and oropharynx at 6 and 20 hours after LTC administration contained a readily identifiable population of cells containing fluorescently‐labeled LTC (Figure 5). The LTC‐containing cells were detected in both nasal lavage cells (Figure 5A) and in throat swab cells (oropharynx, Figure 5B). Peak uptake of LTC‐labeled cells in the nasal lavage samples occurred 6 hours after LTC administration, whereas in the oropharynx peak of LTC uptake occurred 20 hours after administration. 3.6 Intranasal and PO administration of LTC triggers influx of inflammatory leukocytes to mucosal surfaces Cells from the nasal cavity or oropharynx were obtained by nasal lavage or swabbing the back of the throat, respectively, as described above. Cells were immunostained with fluorescently conjugated antibodies against specific cell markers for T cells (CD5+) and monocytes (CD14+) and analyzed by flow cytometry. Neutrophils were identified by their characteristic forward versus side scatter profiles. Pretreatment samples from the nasal cavity of 7 healthy cats contained an average of 7% CD5+ T cells, whereas the average percentage of CD5+ T cells in the oropharynx of cats before treatment was 73%. The percentage of CD14+ monocytes in the nasal cavity before treatment was 2.5%, similar to the 3% of CD14+ monocytes found in the oropharynx (Figure 6). Percentages of neutrophils found in the nasal cavity were 10%, and 2% in the oropharynx of untreated animals. Six hours after topical administration of LTC, a marked increase in the percentage of CD14+ monocytes was observed in both the nasal mucosa (Figure 6A, increasing from an average of 2.5% to an average of 14%) and in the oropharynx (Figure 6B, increasing from an average of 3% to an average of 11%). The percentage of CD14+ monocytes from both nasal and oropharynx samples had decreased by 20 and 72 hours compared to the 6 hours samples but still remained higher than in pretreatment samples. Interestingly, the percentages of CD5+ T cells from the nasal mucosa decreased significantly 6 hours post‐LTC treatment (decreasing from 7% to 3%) and from 73% to 51% in the oropharynx. Nasal CD5+ T cells returned to pretreatment values by 72 hours after treatment. In the oropharynx, the decrease in CD5+ cells observed at 6 hours was also transient, with percentages of CD5+ T cells returning to nearly baseline values (67%) at 20 hours post‐LTC treatment. The neutrophil population was essentially unchanged throughout the study time course in both the nose and the oropharynx after LTC administration (Figure 6). 4 DISCUSSION An effective, broad‐spectrum immunotherapeutic agent capable of activating protective mucosal immune responses would provide a valuable means of inducing rapid protection from a variety of upper respiratory tract pathogens in cats. Therefore, we investigated innate immune responses to LTC, a new immune therapeutic agent designed for mucosal application. We found that LTC induced activation of feline leukocytes, including upregulation of costimulatory molecule (OX40 and MHCII) expression. In addition, LTC induced production of cytokines associated with innate immune responses, including INF‐α, INF‐γ, TNF‐α, and IL‐12. in vivo studies in healthy cats demonstrated uptake of LTC by immune cells and epithelial cells in the upper airways, as well as recruitment of monocytes to the upper airways. Our studies indicated that cats were capable of responding to mucosal immune activation by LTC. Significant upregulation of IFN‐γ along with increases in lymphocyte expression of OX40 and monocyte expression of MHCII all would be associated with innate immune system activation for better viral and bacterial pathogen clearance.7, 18, 19 These findings are consistent with LTC activation of mucosal innate immune responses in cats and induction of partial protection against inhalational challenge with FHV‐1.19 Nasopharyngeal‐associated lymphoid tissue (NALT) is a network of immune cells located in the oropharyngeal region that is primed for immune recognition of viral, bacterial, and fungal pathogens.20-23 When activated by infections, innate immune effector cells in the NALT (including monocytes, neutrophils, innate lymphoid cells, and natural killer [NK] cells) respond by producing cytokines and chemokines, and by activating other immune effector cells including tissue macrophages, epithelial cells, and T and B cells. In our study, several key antiviral and antibacterial cytokines were produced after LTC treatment, including significant increases in secretion of IFN‐γ protein, and increases in INF‐α and IL‐12 transcription, as assessed by qRT‐PCR. These results together suggest broad activation of innate immune responses in nasal and oropharyngeal tissues after administration of LTC in cats. Distinct cellular trafficking responses to LTC treatment were noted when monocyte and T‐cell responses were compared in the nose and oropharynx. For example, LTC administration triggered strong recruitment of CD14+ monocytes to both the nasal cavity and the oropharynx, while at the same time triggering significant, transient disappearance of T cells from these mucosal sites (see Figure 6). Interestingly, minimal neutrophil responses to LTC administration were observed at any time point evaluated, in either the nose or oropharynx. The recruitment of monocytes by LTC could be attributed to production of the chemokine CCL2, which has been noted previously after treatment with LTC.11 In contrast, the transient disappearance of T cells from mucosal sites after LTC administration most likely could be attributed to transient redistribution of T cells after adherence to capillary blood vessels or migration into tissues. A similar effect has been observed previously in dogs treated systemically with LTC.24 In support of transient redistribution, we observed that CD5+ T lymphocytes were rapidly restored in mucosal sites within 72 hours after LTC administration. We also observed distinct differences in resident immune cell populations in the nasal cavity and oropharynx, especially with respect to the density of mucosal T cells. For example, the oropharynx was found to be highly enriched in mucosal CD5+ T cells, whereas T cells were much less numerous in the nasal cavity (see Figure 6). These differences are considered real (and not simply a reflection of differences in sampling techniques), because the percentages of unmarked cells (assumed to represent primarily mucosal epithelial cells, see Figure 6) were similar in samples from the oropharynx, compared to nasal lavage samples. Thus, the oropharynx in cats appears to be enriched in superficially located T cells compared to the nasal cavity and thus well suited to induction of mucosal immune responses after PO administration of nonspecific immune stimuli. The ability of LTC to rapidly induce mucosal immune responses in the upper airway and oropharynx of cats suggests potential use as a nonspecific immune therapeutic agent for prevention of viral and bacterial infections in cats. Administration of LTC significantly decreased signs of FHV‐1 infection and viral shedding in experimentally inoculated cats.19 In addition, we have found that topical application of LTC in dogs can induce mucosal activation and cellular recruitment, similar to the responses seen in cats (Dow S, unpublished data). Thus, topical treatment in the upper airways with LTC may have utility for preventing or mitigating infections with common respiratory tract pathogens in both dogs and cats. The present study had several limitations. For example, the lack of available reagents for full phenotyping of feline leukocytes precluded a more complete assessment of cellular responses to topical LTC administration. In addition, the relatively small numbers of cells collected by nasal lavage or oral swabs precluded cytokine analysis after LTC administration. The in vitro studies using PBMC did not determine the cellular source of specific cytokine production after LTC activation. However, previous studies of LTC in mice indicated that dendritic cells were the primary cellular source of IL‐12 production after LTC administration, and that NK cells were the primary cellular source of IFN‐γ.25, 26 In summary, our studies describe the innate immune activation properties of a novel mucosally active immune stimulant in cats. Innate immune activation by LTC was associated with production of key antiviral and antibacterial cytokines, and with upregulation of important costimulatory molecules on immune effector cells. These findings suggest further evaluation of the LTC mucosal immune stimulant for prevention of URI in cats and for treatment of established URI refractory to treatment with antibiotics or other medications such as lysine. ACKNOWLEDGMENTS The authors acknowledge the assistance of Ms. Jade Kurihara with preparation of LTC used in the study, and Ms. Jennifer Hawley for performing RT‐PCR assays. Portions of this research were presented at the 2017 American College of Veterinary Internal Medicine Forum, National Harbor, Maryland. This study was supported by a grant from the state of Colorado Office of Economic Development and International Trade. CONFLICT OF INTEREST DECLARATION Drs. Lappin and Dow both hold stock options and corporate positions in Poudre Canyon Therapeutics, a Fort Collins company developing the LTC immunotherapy platform technology. OFF‐LABEL ANTIMICROBIAL DECLARATION Authors declare no off‐label use of antimicrobials. INSTITUTIONAL ANIMAL AND CARE AND USE COMMITTEE (IACUC) OR OTHER APPROVAL DECLARATION The cats were managed by a contract research facility in Fort Collins under an approved IACUC protocol. HUMAN ETHICS APPROVAL DECLARATION Authors declare human ethics approval was not needed for this study. REFERENCES