Comparative analysis of atrophic rhinitis and bronchitis: Etiology, clinical features and therapeutic approaches

Introduction Atrophic Rhinitis

Atrophic rhinitis (AR) is a chronic degenerative disorder of the nasal mucosa characterized by progressive architectural loss of the turbinates, excessive crust formation, and the development of a distinctive malodor (ozaena). Although the precise etiological trigger remains incompletely understood, the disease manifests through mucosal desiccation, accumulation of thick secretions, and an enlarged nasal cavity with paradoxical airflow disturbances (Zohar et al., 1990) [25] . In swine, the progressive form traditionally referred to as turbinate atrophy is primarily associated with toxigenic Pasteurella multocida (capsular types A or D), which produces clinical signs such as facial distortion, sneezing, nasal exudation, and growth impairment. AR has historically been described under several synonymous terms, including rhinitis atrophicans, coryza foetida, acute necrotizing rhinitis, and rhinitis chronica foetida (Loch & Reiriz, 1978; Ruskin, 1932) [14,23] .

Severe forms, often labeled "ozaena," denote cases in which fetid odor is a predominant feature (Menzio, 1950) [16] . The condition may occur as simple, subacute, or ozenous variants, reflecting differences in clinical progression and mucosal pathology.

Bronchitis

Respiratory disease in swine reflects a multifactorial interplay of infectious agents viral and bacterial as well as environmental stressors. Among the diverse pathogens capable of initiating bronchial inflammation, Porcine reproductive and respiratory syndrome virus (PRRSV), Swine influenza virus (SIV), and Porcine respiratory coronavirus (PRCV) are considered major contributors, often acting synergistically with bacterial agents such as Bordetella bronchiseptica (Brockmeier et al., 2008) [2] . This microbial complexity frequently culminates in chronic respiratory compromise and variable clinical severity.

Etiology Atrophic Rhinitis

The earliest pathogenic event in AR typically involves colonization of the nasal mucosa by Bordetella bronchiseptica, which induces acute mucosal inflammation and mild turbinate remodeling. This initial insult facilitates subsequent colonization by toxigenic strains of Pasteurella multocida, whose dermonecrotoxin exerts direct cytotoxic effects on osteoclasts and osteoblasts, accelerating turbinate atrophy and septal deviation (De Jong, 1999; Rimler & Brogden, 1986; Pedersen & Barfod, 1982) [5,20,19] . Experimental observations confirm that coinfections rather than P. multocida alone produce more severe and persistent lesions and significantly impair growth performance (Cowart et al., 1989) [3] .

Bronchitis

The etiology of bronchitis in swine involves a combination of primary and secondary pathogens, including P. multocida, A. pleuropneumoniae, B. bronchiseptica, and Mycoplasma hyopneumoniae. Although each microorganism is capable of inducing respiratory pathology independently, concurrent infections tend to amplify tissue damage, prolong disease, and increase economic losses. Despite general recognition of the multifactorial nature of porcine respiratory disease, the specific immunological and microbiological interactions that underpin disease exacerbation remain only partially defined (Brockmeier et al., 2008) [2] .

Epidemiology Atrophic Rhinitis

Atrophic rhinitis is widely recognized as a prevalent condition in swine-producing regions, particularly in the Midwestern United States. Abattoir-based investigations have consistently reported a high burden of turbinate lesions, with some surveys indicating that up to half of market-weight pigs and sows demonstrate varying degrees of conchal atrophy at slaughter (Gardner et al., 1994) [7] . The clinical incidence, typically ranging between 5-30%, varies considerably depending on detection method, herd management, and implementation of control programs. Evaluation of snout sections from slaughtered pigs frequently yields lesion rates between 14-50%, although values may be underestimated or overrepresented based on the sourcing of the animals and prior disease-control interventions.

Comparative analyses across farms and production systems have demonstrated substantial variability in lesion scoring, partly due to the historical absence of standardized grading protocols. Recent adoption of structured scoring methodologies has improved diagnostic consistency. Seasonal influences and facility type also contribute to epidemiological differences: a two-season assessment of 21 herds revealed that AR lesions were generally more severe in pigs slaughtered during summer months, while pneumonic lesions predominated in winter (Cowart et al., 1992) [4] . Environmental and housing factors play an important role. Pigs reared in enclosed, mechanically ventilated buildings especially when farrowed in centralized farrowing units tend to develop more pronounced turbinate atrophy than pigs raised in outdoor systems such as sow huts or dirt lots. These findings suggest that the severity of observed lesions may reflect not only the conditions present at slaughter but also the historical rearing environment during early developmental stages.

Pathogenesis Atrophic Rhinitis

Initial colonization of the nasal epithelium by B. bronchiseptica results in close association with ciliated epithelial cells. The organism secretes a heat-labile cytotoxin that induces ciliary dysfunction, epithelial hyperplasia and metaplasia, mucociliary impairment, and early-stage remodeling of the conchal bone within 2-4 weeks of infection. Although this non-progressive turbinate atrophy may partially resolve, toxigenic strains of P. multocida can subsequently colonize the altered mucosa more effectively. Once established, P. multocida elaborates a dermonecrotoxin that profoundly disrupts bone homeostasis by stimulating osteoclastic activity and suppressing osteoblastic repair mechanisms (Rozengurt et al., 1990; Mullan et al., 1996) [22,17] . Experimental studies have demonstrated that intranasal exposure of gnotobiotic pigs to dermonecrotoxin-producing type D P. multocida results in rapid and severe bilateral turbinate destruction, with irreversible changes occurring within days and extensive conchal loss evident within 10-14 days (Jordan et al., 2003) [9] . The toxin also exerts systemic immunomodulatory effects: pigs harboring toxigenic P. multocida exhibit suppressed antigen-specific IgG responses, suggesting altered immune regulation and increased susceptibility to co-infecting pathogens (Jordan et al., 2003) [9] . The degenerative process is characteristically noninflammatory, a unique feature supported by findings that parenteral administration of the toxin alone can reproduce turbinate atrophy in the absence of local infection. Severe cases are associated with facial deformities, including dorsal displacement of the nasal bones, twisting of the snout, and varying degrees of maxillofacial malalignment. These structural changes can impair prehension and mastication, contributing to compromised growth performance. Controlled studies further indicate that AR reduces feed intake and activity levels, particularly under challenging environmental conditions (Van Diemen et al., 1995) [24] . Robust experimental models using sequential exposure to B. bronchiseptica and toxigenic P. multocida in gnotobiotic piglets have proven valuable for elucidating disease mechanisms and evaluating vaccine effectiveness (Ackermann et al., 1991) [1] .

Bronchitis

The innate immune response of the lung orchestrates both the initial defense against pathogens and the subsequent activation of adaptive immunity. Recognition of pathogenassociated molecular patterns triggers rapid secretion of proinflammatory mediators such as TNF-α, IL-1, IL-6, IL-8, and MCP-1, which facilitate leukocyte recruitment and containment of infection (Brockmeier et al., 2008) [2] . Coinfection with multiple respiratory pathogens alters these cytokine expression patterns, resulting in more pronounced inflammatory cascades than those observed in single-agent infections. In co-infection models involving B. bronchiseptica and PRCV, the combined pathogen burden produces a distinct cytokine signature characterized by amplified proinflammatory signaling and enhanced tissue injury. This synergistic response likely contributes to the more severe histopathological lesions observed during dual infection, including extensive interstitial thickening, epithelial necrosis, and heightened cellular infiltration (Brockmeier et al., 2008) [2] .

Clinical Findings Atrophic Rhinitis

In controlled experimental settings, pigs infected with P. multocida or B. bronchiseptica generally maintain normal appetite and exhibit unaltered behavior, although sporadic sneezing may occur. The clinical spectrum depends largely on the stage and severity of turbinate degeneration. During the acute phase commonly affecting piglets between 3 and 9 weeks of age irritation of the nasal mucosa leads to frequent sneezing, intermittent coughing, and mild serous to mucopurulent nasal discharge. Transient unilateral or bilateral epistaxis may develop as mucosal fragility increases. Sneezing frequency has been proposed as an indirect indicator of disease severity and is reduced markedly in piglets derived from sows vaccinated with combined B. bronchiseptica and P. multocida bacterins compared with piglets receiving B. bronchiseptica vaccine alone. Mild ocular epiphora may accompany nasal irritation, sometimes resulting in streaking of dried debris beneath the medial canthus. Growth rates may be negatively affected during this stage. In infections restricted to B. bronchiseptica, clinical signs often resolve spontaneously over several weeks. In more advanced or progressive cases driven by toxigenic P. multocida, respiratory compromise intensifies, with thickened nasal secretions, repeated episodes of bleeding, and difficulty nursing in young piglets due to obstruction. Chronic cases exhibit accumulation of inspissated material within the nasal passage that may be forcibly expelled during paroxysms of sneezing. As maxillofacial deformities evolve, dorsal displacement of the nasal bones, distortion of the premaxilla, and misalignment of the incisors become apparent. Brachygnathia superior, protrusion of lower incisors, and impaired mastication frequently contribute to poor body condition. Severe cases produce characteristic facial "dishing" with overlying skin wrinkling (Hamilton et al., 1996) [8] .

Macroscopic & Microscopic Lesions Atrophic Rhinitis

Gross examination of affected nasal cavities reveals turbinate structures with pale, mottled surfaces and varying degrees of atrophy. Rostral sections typically show epithelial metaplasia with limited neutrophilic infiltration in both the epithelium and submucosa. Occasional epithelial necrosis may be present, though microabscesses are uncommon. Mucous glands in the submucosa may appear compressed or reduced in volume. Histologically, osteoclast numbers are elevated and frequently observed within resorption lacunae along the osseous core of the conchae, reflecting active bone remodelling. As atrophy progresses, conchal architecture becomes severely reduced or absent, often without a significant inflammatory reaction (Hamilton et al., 1996) [8] .

Bronchitis

Clinical signs of bronchitis caused by bacterial or viral agents tend to be mild, presenting mainly as sporadic coughing, sneezing, transient lethargy, and exercise intolerance. Despite the modest clinical appearance, pathological lesions may be more extensive.

Macroscopic Pulmonary Lesions

Across infectious groups, lungs typically exhibit multifocal to coalescing areas of dark red consolidation, predominantly affecting the cranioventral lobes. Coinfected pigs those experimentally inoculated with both B. bronchiseptica and PRCV show a greater proportion of lung involvement compared with pigs infected with either pathogen alone (Brockmeier et al., 2008) [2] . Regardless of the pathogen involved, the predominant gross lung changes consisted of multiple areas of red consolidation affecting both lungs, most commonly located in the cranial and ventral regions (Figure 1). In the first experiment, pigs infected simultaneously with B. bronchiseptica and PRCV showed a greater average proportion of lung affected compared with those infected with either organism alone (Brockmeier et al., 2008) [2] . Figure 1 [2] . pneumocyte hyperplasia, and the presence of syncytial cells as defining microscopic lesions (Brockmeier et al., 2008) [2] .

Diagnosis Atrophic Rhinitis Microbiological Assessment

Diagnosis often begins with microbiological evaluation of nasal swabs or tissue samples to identify the presence of pathogenic bacteria. Isolation of Pasteurella multocida and Bordetella bronchiseptica the principal organisms associated with AR provides important etiological evidence (KB & Brockmeier, 2019) [2] .

Serological Analysis

Serological assays, including ELISA-based detection of antibodies against P. multocida and B. bronchiseptica, can help determine prior exposure or ongoing infection. Although serology does not distinguish acute from past infections, it contributes valuable epidemiological information (KB & Brockmeier, 2019) [2] .

Molecular Diagnostics

PCR assays targeting bacterial DNA offer rapid, highly sensitive detection of toxigenic strains, particularly those producing dermonecrotoxin toxin. PCR testing from nasal or tonsillar swabs is effective for large-scale herd screening (Kamp et al., 1996) [11] .

Histopathology

Microscopic evaluation of nasal tissues can reveal epithelial metaplasia, conchal atrophy, glandular compression, and osteoclastic activity, providing definitive pathological confirmation of AR (KB & Brockmeier, 2019) [2] .

Bacterial Isolation and Culture

Culturing P. multocida or B. bronchiseptica from nasal samples allows for strain characterization and may support antimicrobial susceptibility testing (KB & Brockmeier, 2019) [2] .

Clinical and Epidemiological Evaluation

Understanding herd history, introduction of new stock, housing conditions, and the age distribution of affected animals enhances diagnostic accuracy. Epidemiological patterns, such as the presence of carrier animals, also contribute to interpretation (Cowart et al., 1991) [4] .

Given the economic implications of AR, a comprehensive diagnostic approach incorporating clinical, microbiological, and molecular data is essential to guide prevention and control strategies (Gardner et al., 1994) [7] .

Bronchitis Diagnosis of Bordetella bronchiseptica

 Clinical Assessment: Signs such as coughing, nasal discharge, and respiratory distress suggest possible infection but are non-specific (Brockmeier et al., 2008) [2] .  Bacterial Culture: Isolation from nasal or tracheal swabs provides definitive identification but requires specialized laboratory techniques.  PCR: Molecular detection of B. bronchiseptica DNA offers high specificity and sensitivity (Brockmeier et al., 2008) [2] .  Serology: Detection of antibodies can confirm exposure but cannot reliably indicate active infection.

Diagnosis of Porcine Respiratory Coronavirus (PRCV):

 PCR: The primary diagnostic tool for detecting PRCV RNA in nasal or lung tissues (Brockmeier et al., 2008) [2] .  Serological Tests: ELISAs can detect anti-PRCV antibodies but do not differentiate active from prior infection.  Histopathology: Viral pneumonia typically shows epithelial necrosis, syncytia formation, and interstitial inflammation (Brockmeier et al., 2008) [2] .

Treatment Atrophic Rhinitis Antibiotic Therapy

Antimicrobials, such as tetracyclines and macrolides, are commonly administered to manage secondary bacterial infections and reduce the pathogen load. Appropriate drug selection and dosing must be tailored to herd-level sensitivity patterns. Tilmicosin, when used as a feed additive at 363 g/ton for three weeks, has been reported to reduce lesion severity and improve growth performance in pigs challenged with multiple respiratory pathogens (Olson & Bä, 2000) [18] .

Anti-inflammatory Intervention:

Nonsteroidal anti-inflammatory drugs (NSAIDs) may be used to moderate inflammation and improve animal comfort (Olson & Bä, 2000) [18] .

Supportive Management

Nutritional support, clean water access, proper ventilation, and a low-stress environment are essential adjuncts to pharmacological treatment (Olson & Bä, 2000) [18] .

Bronchitis Antibiotics

For bacterial bronchitis caused by B. bronchiseptica, antimicrobials such as tetracyclines, macrolides, and fluoroquinolones may be employed. These treatments primarily target secondary bacterial components of the disease rather than viral etiologies (Brockmeier et al., 2008) [2] .

Prevention and Control Atrophic Rhinitis Biosecurity Strategies

Strict implementation of biosecurity including controlled access to facilities, sanitation protocols, and quarantine of new arrivals is essential to prevent the introduction and spread of AR pathogens (Kobisch, 1989) [12] . Movement of personnel, equipment, and vehicles must be managed to minimize cross-contamination.

Vaccination

Vaccination against B. bronchiseptica and toxigenic P. multocida is a primary preventive measure. Vaccine formulations derived from herd-specific isolates and inactivated with glutaraldehyde, adjuvanted with aluminum hydroxide, have proven effective.

Standard protocols include:- Sows vaccinated twice late in gestation  Piglets vaccinated at 7 and 28 days  Combined sow and piglet vaccination programs (Ross, 2006) [21] .  Herd Management Practices:  Stress reduction, optimal stocking densities, and maintaining appropriate climatic conditions (temperature, humidity, ventilation) are critical for reducing respiratory disease risk (Kobisch, 1989) [12] .

Genetic Selection

Long-term mitigation may be achieved through selective breeding for AR-resistant lines (Kobisch, 1989) [12] .

Sanitation

Routine cleaning, disinfection, and proper waste management help reduce environmental pathogen load.

Environmental Control

Adequate ventilation systems reduce ammonia accumulation and airborne microbial exposure, limiting respiratory tract irritation and disease susceptibility (Kobisch, 1989) [12] .

Monitoring and Surveillance

Routine health assessments and periodic diagnostic testing allow early detection of AR, facilitating timely interventions (Kobisch, 1989) [12] .

Training and Education

Farm personnel must be well-trained in biosecurity, vaccination procedures, and early recognition of respiratory disease.

Bronchitis Biosecurity

Limiting farm access, segregating age groups, and controlling movement of animals and equipment reduce the introduction of respiratory pathogens (Brockmeier et al., 2008) [2] .

Vaccination

Use of vaccines targeting specific bacterial or viral respiratory pathogens should be aligned with regional disease prevalence and veterinary recommendations (Brockmeier et al., 2008) [2] .

Environmental and Husbandry Management

Adequate ventilation, reduced stocking densities, and effective hygiene practices are vital for preventing respiratory stress and cross-infection (Brockmeier et al., 2008) [2] .

Diagnostic Monitoring

Routine respiratory evaluation at the herd level permits early detection of subclinical infection and guides evidence-based interventions (Brockmeier et al., 2008) [2] .

Nutritional Support

Balanced diets that promote immune function enhance herd resilience to respiratory disease challenges.

Conflict of Interest Not available

Financial Support

References

1. Ackermann MR, Register KB, Gentry-Weeks C, Gwaltney SM, Magyar T. A porcine model for assessing the virulence of Bordetella bronchiseptica. J Comp Pathol. 1997;116(1):55-61.
2. Brockmeier SL, Loving CL, Nicholson TL, Palmer MV. Coinfection of pigs with porcine respiratory coronavirus and Bordetella bronchiseptica. Vet Microbiol. 2008;128(1-2):36-47.
3. Cowart RP, Bäckström L, Brim TA. Pasteurella multocida and Bordetella bronchiseptica in atrophic rhinitis and pneumonia in swine. Can J Vet Res. 1989;53(3):295.
4. Cowart RP, Boessen CR, Kliebenstein JB. Seasonal and facility-related patterns associated with atrophic rhinitis and pneumonia in slaughter swine. J Am Vet Med Assoc. 1992;200(2):190-3.
5. Jong DMF. Progressive and nonprogressive atrophic rhinitis. In: Diseases of Swine. 9 th Ed; 1999, p. 577-602.
6. Dutt SN, Kameswaran M. Etiology and management of atrophic rhinitis. J Laryngol Otol. 2005;119(11):843-52.
7. Gardner IA, Kasten R, Eamens GJ, Snipes KP, Anderson RJ. Molecular fingerprinting of Pasteurella multocida associated with progressive atrophic rhinitis in swine herds. J Vet Diagn Invest. 1994;6(4):442-7.
8. Hamilton TD, Roe JM, Webster AJ. Synergistic effects of gaseous ammonia in the pathogenesis of Pasteurella multocida-induced atrophic rhinitis in swine. J Clin Microbiol. 1996;34(9):2185.
9. Jordan RW, Roe JM. An experimental mouse model for studying progressive atrophic rhinitis of swine. Vet Microbiol. 2004;103(3-4):201-7.
10. K. B., Brockmeier SL. Pasteurellosis. In: Diseases of Swine; 2019, p. 884-97.
11. Kamp EM, Bokken GC, Vermeulen TM, Jong DMF, Buys HE, Reek FH, et al. A PCR assay for large-scale detection of toxigenic Pasteurella multocida in nasal and tonsillar swabs of pigs. J Vet Diagn Invest. 1996;8(3):304-309.
12. Kobisch M, Pennings AMMA. Evaluation of a commercial and an experimental atrophic rhinitis vaccine containing P. multocida dermonecrotoxin and B. bronchiseptica. Vet Rec. 1989;124(3):57-61.
13. Lobo CJ, Hartley C, Farrington WT. Closure of the nasal vestibule in atrophic rhinitis: A new non-surgical technique. J Laryngol Otol. 1998;112:543-6.
14. Loch WE, Reiriz HM. Acute necrotizing rhinitis in humans. Rhinology. 1978;16(4):235-42.
15. Martineau-Doizé B, Dumas G, Larochelle R, Frantz JC, Martineau GP. Morphometric analysis of atrophic rhinitis caused by type D Pasteurella multocida. Can J Vet Res. 1991;55(3):224.
16. Menzio P. Vasomotor responses to nicotinic acid in simple and ozenous atrophic rhinitis. Arch Ital Otol Rhinol Laringol. 1950;61(Suppl 4):19-27.
17. Mullan PB, Lax AJ. Pasteurella multocida toxin as a mitogen for primary bone cell cultures. Infect Immun. 1996;64(3):959-65.
18. Olson LB, Bä LR. Effects of tilmicosin on reducing atrophic rhinitis, pneumonia, and pleuritis in swine. J Swine Health Prod. 2000;8(6):263-8.
19. Pedersen KB, Barfod K. Influence of sow vaccination with P. multocida toxin on the incidence of atrophic rhinitis. Nord Vet Med. 1982;34(7-9):293-302.
20. Rimler RB, Brogden KA. P. multocida isolates from rabbits and swine: Serologic typing and toxin production. Am J Vet Res. 1986;47(4):730-7.
21. Ross RF. Pasteurella multocida and its role in porcine pneumonia. Anim Health Res Rev. 2006;7(1-2):13-29.
22. Rozengurt E, Higgins T, Chanter N, Lax AJ, Staddon JM. P. multocida toxin is a potent mitogen for fibroblasts in vitro. Proc Natl Acad Sci USA. 1990;87(1):123-127.
23. Ruskin JL. Differential diagnosis and management of atrophic rhinitis and ozena. Arch Otolaryngol. 1932;15:222-257.
24. Diemen VPM, Schrama JW, Van der Hel W, Verstegen MWA, Noordhuizen JPTM. Influence of atrophic rhinitis and environmental climate on pig performance. Livest Prod Sci. 1995;43(3):275-284.
25. Zohar Y, Talmi YP, Strauss M, Finkelstein Y, Shvilli Y. Reappraisal of ozena. J Otolaryngol. 1990;19(5):345-349.