Pediatric Pulmonology
Detection of Biofilm in Bronchoalveolar Lavage from Children With Non-Cystic Fibrosis Bronchiectasis Robyn L. Marsh, PhD,1* Ruth B. Thornton, PhD,2,3 Heidi C. Smith-Vaughan, Peter Richmond, FRACP, MRCP(UK),2,3 Susan J. Pizzutto, Bsc(Hons),1 and Anne B. Chang, FRACP, PhD1,4
PhD,
1
Summary. Background: The presence of Pseudomonas aeruginosa biofilms in lower airway specimens from cystic fibrosis (CF) patients is well established. To date, biofilm has not been demonstrated in bronchoalveolar lavage (BAL) from people with non-CF bronchiectasis. The aim of this study was to determine (i) if biofilm was present in BAL from children with and without bronchiectasis, and (ii) if biofilm detection differed between sequentially collected BAL. Methods: Testing for biofilm in two sequentially collected BAL from children with and without bronchiectasis was done using BacLightTM live–dead staining and lectin staining for extracellular polymeric biofilm matrices. Bacterial culture and cytological measures were performed on the first and second lavages, respectively. Clinically important BAL infection was defined as >104 cfu of respiratory pathogens/ml BAL. Results: Biofilm was detected in BAL from seven of eight (87.5%) children with bronchiectasis (aged 0.8–6.9 years), but was not detected in any of three controls (aged 1.3–8.6 years). The biofilms contained both live and dead bacteria irrespective of antibiotic use prior to bronchoscopy. Biofilm was detected more frequently in the second lavage than the first. Three of the seven biofilm-positive BAL were culture-positive for respiratory pathogens at clinically important levels. Conclusions: Biofilm is present in BAL from children with non-CF bronchiectasis even when BAL-defined clinically important infection was absent. Studies to characterize lower airway biofilms and determine how biofilm contributes to bronchiectasis disease progression and treatment ß 2014 Wiley Periodicals, Inc. outcomes are necessary. Pediatr Pulmonol. Key words: respiratory infection; chronic suppurative lung disease; confocal microscopy. Funding source: University of Western Australia, State and Commonwealth Governments, Australian National Health and Medical Research Council (NHMRC) Fellowships, Numbers: 1034703, 1024175, 545216; Post-graduate scholarship, Number: 1038415; Centre for Research Excellence in Respiratory Health for Aboriginal and Torres Strait Islander Children, Number: 1040830.
INTRODUCTION
The burden of bronchiectasis unrelated to cystic fibrosis (CF) is increasingly recognized.1 In adults, 1 Child Health Division, Menzies School of Health Research, Charles Darwin University, Darwin, Northern Territory, Australia. 2 School of Paediatrics and Child Health, University of Western Australia, Perth, Western Australia, Australia. 3 Telethon Institute for Child Health Research, Centre for Child Health Research, University of Western Australia, Western Australia, Australia. 4 Queensland Children’s Respiratory Centre, Queensland Children’s Medical Research Institute, Royal Children’s Hospital, Brisbane, Queensland, Australia.
Robyn L. Marsh and Ruth B. Thornton contributed equally to this work. Authors contribution: The authors declare the following competing interests: R.B. Thornton has received travel funding from GlaxoSmithKline; P. Richmond has received Institutional funding from GSK for investigator-
ß 2014 Wiley Periodicals, Inc.
bronchiectasis prevalence and mortality have increased during the last two decades.2,3 In children, bronchiectasis is particularly prevalent in settings with disadvantaged populations such as Indigenous communities in Australia, led epidemiological studies in otitis media and has received travel support from GSK, Pfizer, and other vaccine companies to present scientific data and chair workshops. Other authors have no competing interests to declare. None of the authors have shares or paid employment with any pharmaceutical company.
Correspondence to: Robyn L. Marsh, Menzies School of Health Research, PO Box 41096, Casuarina, NT 0811, Australia. E-mail: [email protected] Received 29 June 2013; Accepted 3 February 2014. DOI 10.1002/ppul.23031 Published online in Wiley Online Library (wileyonlinelibrary.com).
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Alaska, and New Zealand.4 Despite the burden, there are relatively few pathobiological data on bronchiectasis in children. The pathogenesis of bronchiectasis likely involves a combination of host and microbial factors including reduced mucociliary clearance and ongoing inflammation related to endobronchial infection.5 Not surprisingly, antibiotics play a central role in bronchiectasis treatment1,6 with prolonged therapy often required to prevent acute exacerbations and disease progression.7 The chronic and recalcitrant nature of airway infection in children with bronchiectasis is suggestive of a biofilmassociated infection.7,8 Biofilms are defined as clusters of bacteria enclosed in matrices of self-produced and hostderived DNA and exopolymeric substances.8 Bacteria in biofilm exhibit increased resistance to antibiotics and host immune responses,9 and can be difficult to culture. Indeed, biofilm-associated infections are often culturenegative despite clinical signs of infection.8 One recent retrospective Australian study reported culture of pathogenic bacteria at clinically important levels (>104 cfu/ml of BAL) from only 68% of BAL from 113 children with non-CF bronchiectasis;10 however, it is not known if the low recovery rate was due to the presence of biofilm or other factors. Biofilms produced by Pseudomonas aeruginosa, Haemophilus influenzae, and other common respiratory pathogens have been demonstrated in lower airways specimens from CF patients,11–13 bronchus tissue from an adult with bronchiectasis,14 and middle ear specimens from patients with chronic and recurrent otitis media.15–17 In vitro studies have also shown that respiratory pathogens such as Streptococcus pneumoniae, nontypeable H. influenzae, and Moraxella catarrhalis can produce biofilm.13,18,19 Although culture-based studies report these species as the most commonly isolated respiratory pathogens from lower airway specimens of children with non-CF bronchiectasis,20 no studies have determined if bacteria in a biofilm are also present. To date, direct detection of biofilm in bronchoalveolar lavage (BAL) specimens has only been described in CF, which has a different pathobiology to non-CF bronchiectasis. Current recommendations for pediatric BAL analyses are for the first lavage to be used for microbiological assessment and the second lavage (þ/ a third lavage) for cytology and inflammation studies.21 No studies to date have determined whether the first or second lavage should be used for biofilm testing. It is biologically plausible that the second lavage may better demonstrate biofilm, possibly due to improved mucosal sampling following removal of airway mucus by the first lavage. This hypothesis is supported by data from an otitis media animal model that showed increased detection of biofilm following multiple sequential middle ear lavages.22 Thus, the aims of this study are to determine (i) if biofilm is Pediatric Pulmonology
present in lower airway specimens from children with and without non-CF bronchiectasis, and (ii) whether biofilm detection differs between two sequentially collected lavages. MATERIALS AND METHODS Ethics Statement
This study was approved by the Human Research Ethics Committee of the Northern Territory Department of Health and Menzies School of Health Research, Darwin (HREC 07/63), and the Royal Children’s Hospital, Brisbane (HREC 2003/017 and 200800064). Written informed consent was obtained from the parent/ career(s) of each child. Specimen Collection
BAL were collected as part of two prospective studies. BAL from children with non-CF bronchiectasis were selected from a prospective study of pediatric bronchiectasis at the Royal Darwin Hospital, Darwin, Australia, between January 2010 and July 2011.23 BAL for biofilm testing were selected randomly from children with bronchiectasis confirmed by chest high resolution computed tomography (cHRCT).7 As the aim of the study was to provide proof-of-principle demonstrating if biofilm was present in young children with non-CF bronchiectasis, the study was arbitrarily restricted to analysis of BAL from eight children aged 104 cfu/ ml (Table 1). Pediatric Pulmonology
DISCUSSION
In this first study to evaluate direct detection of bacterial biofilm in lower airway specimens from children with non-CF bronchiectasis, we found biofilms in the BAL of 87.5% of eight children with bronchiectasis, and 0% of three controls. Biofilm was also demonstrated in BAL that were culture-negative for respiratory pathogens at clinically important levels. Lavage-2 was better than lavage-1 for biofilm detection. These data support the hypothesis that pediatric bronchiectasis is a biofilm-associated infection 7 and add to the growing body of work demonstrating biofilm in upper and lower airway specimens. Our study has demonstrated biofilm in BAL from children with non-CF bronchiectasis; previously it was reported in bronchus tissue from an adult bronchiectasis patient;14 sputum, BAL, and lung tissue from CF patients;12,29,30 middle ear effusions and biopsies from otitis media patients;15–17 and adenoid tissue from patients with chronic adenotonsillitis, otitis media, or obstructive sleep apnoea.31,32 Collectively, these data suggest an important role for biofilm in the
Lower Airway Biofilm in Pediatric Bronchiectasis
5
Fig. 1. Biofilms and DNA stranding were observed in BAL from children with bronchiectasis. Representative maximum projection images showing biofilm in BAL from children with bronchiectasis. Extensive biofilm was seen throughout lavage-1 and lavage-2 from child 3 (arrows), while microcolonies were observed in lavage-2 (arrow) but not lavage-1 from child 4. BacLightTM staining. Live bacteria are stained green. Dead bacteria are stained red. Scale bar ¼ 10 mm.
development and persistence of chronic respiratory infections. Further studies are now required to determine the prevalence of biofilm-associated lower airway infection in larger cohorts of children with non-CF bronchiectasis, and to test for associations between the presence of lower airway biofilm and important clinical parameters such as age, disease severity, recurrence of infections, and acute exacerbations. Testing for biofilm in lower airway specimens from children with acute and sub-acute infections is also warranted. Further research is also required to understand the bacteriology and mechanisms underlying biofilm development in the lower airways of children with bronchiectasis. Three of the seven biofilm-positive children were culture-positive for the respiratory pathogens H. influenzae, S. pneumoniae, and/or M. catarrhalis at loads indicative of lower airway infection. Earlier in vitro
studies have shown that cultured isolates of each of these species can produce biofilm.13,18,19 As bacteria in biofilm can be difficult to recover by culture,8 it is possible that other species, or bacteria cultured below clinically important levels, may be present in the lower airway biofilms. This limitation may be overcome by using DNAbased methods to characterize lower airway bacteriology when biofilm-associated infections are present. Speciesspecific testing using fluorescent in situ hybridization (FISH) techniques is also required to characterize the bacteriology of lower airway biofilm in children with nonCF bronchiectasis. However, FISH-based studies will be complicated by the polymicrobial nature of lower airway infections in bronchiectasis. Between three and seven different species were cultured from the biofilm-positive children with bronchiectasis. One child was culturepositive for S. pneumoniae, H. influenzae, Pseudomonas Pediatric Pulmonology
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Fig. 2. X-Y-Z images showing biofilm ultrastructure. X-Y-Z images of BacLightTM stained lavage-2 specimens showing biofilm ultrastructure in the microcolony from child 4 and the more mature biofilm from child 3. The side panels indicate the thickness of the collected images stacks. For child 3 the collected stack was 17 mm thick while that from child 4 was 13.5 mm thick. Live bacteria are stained green. Dead bacteria are stained red. Scale bar ¼ 10 mm.
sp., Staphylococcus sp., b-hemolytic streptococci, a-hemolytic streptococci, and other unidentified species. As bacteria in biofilm may not be detected by culture,8 the presence of yet more bacteria in these lower airway biofilms cannot be excluded. Regardless of the species involved, detection of biofilm in BAL has potential therapeutic implications as bacteria
Fig. 3. Bacterial biofilm matrices demonstrated by lectin staining. Concanavalin A (ConA) and wheat germ agglutinin (WGA) lectin staining (green), and propidium iodide staining of DNA (red). Representative maximum projection image from child 8 showing bacterial biofilm with bacteria surrounded by lectins binding the matrix (arrows). Scale bar ¼ 10 mm.
Pediatric Pulmonology
within biofilm show higher levels of resistance to antibiotic therapies and immune clearance.9,13 While the planktonic bacteria responsible for acute exacerbations may be eradicated using standard anti-microbial therapies, biofilm eradication may not be achieved. Failure to eradicate bacteria in lower airway biofilms may result in persistent airway infection stimulating chronic inflammation.33 Biofilms may also provide a reservoir of bacteria that may be released to cause recurrent acute exacerbations. This may be particularly important following viral infection as in vitro assays demonstrate virus-induced release of planktonic bacteria from biofilm.34 Direct demonstration of biofilm in BAL from children with bronchiectasis suggests that development of adjunct anti-biofilm strategies that complement standard therapies may be needed to effectively treat these lower airway infections. One such strategy may include the earlier use of azithromycin that is known to diminish biofilm development in vitro;35 however, this has not been proven in vivo. Biofilms in the children with bronchiectasis were associated with inflammatory cells and extracellular DNA. Overall, lower airway neutrophil percentages were 10% in six of the seven biofilm-positive children. More extensive DNA stranding was observed in BAL from children with bronchiectasis than in the control children, likely reflecting either active immune responses such as neutrophil extracellular traps,36 or the presence of necrotic neutrophils. This finding requires further investigation as neutrophil extracellular traps may also contribute to lower airway inflammation.36 Additionally, neutrophil extracellular traps may also provide a target for new treatment strategies.17
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Fig. 4. No evidence of bacterial biofilms in lavage-1 or lavage-2 from control children. Representative maximum projection images of BacLightTM stained BAL demonstrating intact human cells but no bacteria in lavage-1 and lavage-2 from child 10 and evidence of planktonic bacteria (arrows), but not biofilm in lavage-1 and lavage-2 from child 11. Scale bar ¼ 10 mm.
To our knowledge this is the first study to compare testing for biofilm in sequentially collected BAL. We found that biofilms in the bronchiectasis patients were detected in the second lavage more commonly than in the first. This finding most likely reflects improved mucosal sampling following removal of airway mucus by the first lavage. While this hypothesis remains to be tested in larger studies, a similar finding was reported for multiple middle ear lavages from a chinchilla model of nontypeable H. influenzae otitis media.22 This has potential implications for bacteriological analyses performed only on the first lavage (as per current recommendations),21 as this may lead to under-reporting of biofilm. Thus, we recommend that future studies test for biofilm in both the first and second lavage. Where specimen volume is limited, the second lavage should be used for biofilm testing. Further studies are also required to determine if detection of biofilm in both lavages (as was observed for three of the seven biofilm-positive children) is associated
with more severe disease. Viable bacteria in biofilm formation were observed in BAL that had been stored for up to 12 months at 808C. While this indicates the suitability of long-term storage at 808C for maintaining biofilm integrity, further testing is required to determine if specimens stored at 808C for >12 months are suitable for biofilm testing. As with other lower airway studies,37 we cannot exclude potential oropharyngeal contamination of the lower airway lavages. a-Hemolytic streptococci were present in BAL from all children indicating a degree of oropharyngeal contamination. Detection of biofilmassociated with squamous epithelial cells in lavages from two children with bronchiectasis also suggests a degree of upper respiratory contamination. However, biofilm not associated with squamous cells was also present in each of these children consistent with lower airway sampling. Challenges associated with differentiating upper and lower airway microbiology are common Pediatric Pulmonology
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to many respiratory studies37 and must be considered in future bronchiectasis biofilm research. In conclusion, we have demonstrated the presence of biofilm in BAL from children with non-CF bronchiectasis without culture-based evidence of P. aeruginosa infection. Importantly, some of these occurred even when BAL-defined clinically important infection was absent. Studies to understand the role of biofilm in the pathobiology of bronchiectasis and other chronic lower respiratory infections are required. Evaluation of possible associations between biofilm detection and markers of clinical severity is also required. As biofilm was more commonly detected in the second lavage, in contrast to recommended guidelines for microbiological work,21 we recommend that both lavages are used for biofilm examination. ACKNOWLEDGMENTS
We would like to acknowledge the families and children that have participated in our studies. The authors also acknowledge the facilities, scientific, and technical assistance of the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterisation & Analysis at The University of Western Australia (funded by The University of Western Australia, State and Commonwealth Governments). RLM, HCS-V, and ABC are supported by Australian National Health and Medical Research Council (NHMRC) Fellowships 1034703, 1024175, and 545216. SJP is supported by NHMRC Post-graduate Scholarship 1038415. The work was also supported by the NHMRC Centre for Research Excellence in Respiratory Health for Aboriginal and Torres Strait Islander children (1040830). The views expressed in this publication are those of the authors and do not reflect the views of the NHMRC. Funding organizations had no role in the study design, data collection, and analysis, decision to publish, manuscript preparation or decision to submit the manuscript for publication. REFERENCES 1. Chang AB, Bell SC, Byrnes CA, Grimwood K, Holmes PW, King PT, Kolbe J, Landau LI, Maguire GP, McDonald MI, et al. Chronic suppurative lung disease and bronchiectasis in children and adults in Australia and New Zealand. Med J Aust 2010;193:356–365. 2. Roberts HJ, Hubbard R. Trends in bronchiectasis mortality in England and Wales. Resp Med 2010;104:981–985. 3. Seitz AE, Olivier KN, Steiner CA, Montes de Oca R, Holland SM, Prevots DR. Trends and burden of bronchiectasis-associated hospitalizations in the United States, 1993–2006. Chest 2010; 138:944–949. 4. Chang AB, Marsh RL, Smith-Vaughan HC, Hoffman LR. Emerging drugs for bronchiectasis. Expert Opin Emerg Drugs 2012;17:361–378. 5. King P. Pathogenesis of bronchiectasis. Paediatr Resp Rev 2010;12:104–110.
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6. Pasteur MC, Bilton D, Hill AT. British Thoracic Society guideline for non-CF bronchiectasis. Thorax 2010;65:i1–i58. 7. Chang AB, Byrnes CA, Everard ML. Diagnosing and preventing chronic suppurative lung disease (CSLD) and bronchiectasis. Paediatr Respir Rev 2011;12:97–103. 8. Hall-Stoodley L, Stoodley P, Kathju S, Hoiby N, Moser C, William CJ, Moter A, Bjarnsholt T. Towards diagnostic guidelines for biofilm-associated infections. FEMS Immunol Med Microbiol 2012;65:127–145. 9. Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science 1999;284:1318– 1322. 10. Kapur N, Grimwood K, Masters IB, Morris PS, Chang AB. Lower airway microbiology and cellularity in children with newly diagnosed non-CF bronchiectasis. Pediatr Pulmonol 2012;47: 300–307. 11. Lam J, Chan R, Lam K, Costerton JW. Production of mucoid microcolonies by Pseudomonas aeruginosa within infected lungs in cystic fibrosis. Infect Immun 1980;28:546–556. 12. Bjarnsholt T, Jensen PO, Fiandaca MJ, Pedersen J, Hansen CR, Andersen CB, Pressler T, Givskov M, Hoiby N. Pseudomonas aeruginosa biofilms in the respiratory tract of cystic fibrosis patients. Pediatr Pulm 2009;44:547–558. 13. Starner TD, Zhang N, Kim G, Apicella MA, McCray PB. Haemophilus influenzae forms biofilms on airway epithelia. Am J Resp Crit Care 2006;174:213–220. 14. Ohgaki N. Bacterial biofilm in chronic airway infection. Kansenshogaku Zasshi 1994;68:138–151. 15. Hall-Stoodley L, Hu FZ, Gieseke A, Nistico L, Nguyen D, Hayes J, Forbes M, Greenberg DP, Dice B, Burrows A, et al. Direct detection of bacterial biofilms on the middle-ear mucosa of children with chronic otitis media. JAMA 2006;296:202–211. 16. Thornton R, Rigby P, Wiertsema S, Filion P, Langlands J, Coates H, Vijayasekaran S, Keil A, Richmond P. Multi-species bacterial biofilm and intracellular infection in otitis media. BMC Pediatr 2011;11:94. 17. Thornton RB, Wiertsema SP, Kirkham LA, Rigby PJ, Vijayasekaran S, Coates HL, Richmond PC. Neutrophil extracellular traps and bacterial biofilms in middle ear effusion of children with recurrent acute otitis media––a potential treatment target. PLoS ONE 2013;8:e53837. 18. Moscoso M, Garcia E, Lopez R. Pneumococcal biofilms. Int Microbiol 2009;12:77–85. 19. Matejka KM, Bremer PJ, Tompkins GR, Brooks HJL. Antibiotic susceptibility of Moraxella catarrhalis biofilms in a continuous flow model. Diagn Micr Infec Dis 2012;74:394–398. 20. Grimwood K. Airway microbiology and host defences in paediatric non-CF bronchiectasis. Paediatr Respir Rev 2011;12:111–118. 21. de Blic J, Midulla F, Barbato A, Clement A, Dab I, Eber E, Green C, Grigg J, Kotecha S, Kurland G, et al. Bronchoalveolar lavage in children. ERS Task Force on bronchoalveolar lavage in children. European Respiratory Society. Eur Res J 2000;15:217–231. 22. Leroy M, Cabral H, Figueira M, Bouchet V, Huot H, Ram S, Pelton SI, Goldstein R. Multiple consecutive lavage samplings reveal greater burden of disease and provide direct access to the nontypeable Haemophilus influenzae biofilm in experimental otitis media. Infect Immun 2007;75:4158–4172. 23. Pizzutto SJ, Grimwood K, Bauert P, Schutz KL, Yerkovich ST, Upham JW, Chang AB. Bronchoscopy contributes to the clinical management of indigenous children newly diagnosed with bronchiectasis. Pediatr Pulmonol 2012;48:67–73. 24. Chang AB, Yerkovich ST, Gibson PG, Anderson-James S, Petsky HL, Carroll ML, Masters IB, Marchant JM, Wurzel D, Upham JW. Pulmonary innate immunity in children with protracted bacterial bronchitis. J Pediatr 2012;161:621–625.
Lower Airway Biofilm in Pediatric Bronchiectasis 25. Gibson L, Khoury J. Storage and survival of bacteria by ultrafreeze. Lett Appl Microbiol 1986;3:127–129. 26. Hare KM, Grimwood K, Leach AJ, Smith-Vaughan H, Torzillo PJ, Morris PS, Chang AB. Respiratory bacterial pathogens in the nasopharynx and lower airways of Australian Indigenous children with bronchiectasis. J Pediatr 2010;157:1001–1005. 27. Smith-Vaughan H, Byun R, Nadkarni M, Jacques NA, Hunter N, Halpin S, Morris PS, Leach AJ. Measuring nasal bacterial load and its association with otitis media. BMC Ear Nose Throat Disord 2006;10:10. 28. Hare KM, Marsh RL, Binks MJ, Grimwood K, Pizzutto SJ, Leach AJ, Chang AB, Smith-Vaughan HC. Quantitative PCR confirms culture as the gold standard for detection of lower airway infection by nontypeable Haemophilus influenzae in Australian Indigenous children with bronchiectasis. J Microbiol Methods 2012;92:270– 272. 29. Rudkjobing VB, Thomsen TR, Alhede M, Kragh KN, Nielsen PH, Johansen UR, Givskov M, Hoiby N, Bjarnsholt T. The microorganisms in chronically infected end-stage and non-end-stage cystic fibrosis patients. FEMS Immunol Med Microbiol 2012; 65:236–244. 30. Pusztaszeri M, Aubert JD, Braunschweig R, Mihaescu A. Pseudomonas aeruginosa biofilms in a bronchoalveolar lavage. Diagn Cytopathol 2009;37:825.
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31. Nistico L, Kreft R, Gieseke A, Coticchia JM, Burrows A, Khampang P, Liu Y, Kerschner JE, Post JC, Lonergan S, et al. Adenoid reservoir for pathogenic biofilm bacteria. J Clin Microbiol 2011;49:1411–1420. 32. Winther B, Gross BC, Hendley JO, Early SV. Location of bacterial biofilm in the mucus overlying the adenoid by light microscopy. Arch Otolaryngol Head Neck Surg 2009;135:1239–1245. 33. Cole PJ. Inflammation: a two-edged sword––the model of bronchiectasis. Eur J Respir Dis Suppl 1986;147:6–15. 34. Chattoraj SS, Ganesan S, Jones AM, Helm JM, Comstock AT, Bright-Thomas R, Lipuma JJ, Hershenson MB, Sajjan US. Rhinovirus infection liberates planktonic bacteria from biofilm and increases chemokine responses in cystic fibrosis airway epithelial cells. Thorax 2011;66:333–339. 35. Starner TD, Shrout JD, Parsek MR, Appelbaum PC, Kim G. Subinhibitory concentrations of azithromycin decrease nontypeable Haemophilus influenzae biofilm formation and diminish established biofilms. Antimicrob Agents Chemother 2008;52:137–145. 36. Kaplan MJ, Radic M. Neutrophil extracellular traps: double-edged swords of innate immunity. J Immunol 2012;189:2689–2695. 37. Charlson ES, Bittinger K, Haas AR, Fitzgerald AS, Frank I, Yadav A, Bushman FD, Collman RG. Topographical continuity of bacterial populations in the healthy human respiratory tract. Am J Resp Crit Care 2011;184:957–963.
Pediatric Pulmonology
Detection of Biofilm in Bronchoalveolar Lavage from Children With Non-Cystic Fibrosis Bronchiectasis Robyn L. Marsh, PhD,1* Ruth B. Thornton, PhD,2,3 Heidi C. Smith-Vaughan, Peter Richmond, FRACP, MRCP(UK),2,3 Susan J. Pizzutto, Bsc(Hons),1 and Anne B. Chang, FRACP, PhD1,4
PhD,
1
Summary. Background: The presence of Pseudomonas aeruginosa biofilms in lower airway specimens from cystic fibrosis (CF) patients is well established. To date, biofilm has not been demonstrated in bronchoalveolar lavage (BAL) from people with non-CF bronchiectasis. The aim of this study was to determine (i) if biofilm was present in BAL from children with and without bronchiectasis, and (ii) if biofilm detection differed between sequentially collected BAL. Methods: Testing for biofilm in two sequentially collected BAL from children with and without bronchiectasis was done using BacLightTM live–dead staining and lectin staining for extracellular polymeric biofilm matrices. Bacterial culture and cytological measures were performed on the first and second lavages, respectively. Clinically important BAL infection was defined as >104 cfu of respiratory pathogens/ml BAL. Results: Biofilm was detected in BAL from seven of eight (87.5%) children with bronchiectasis (aged 0.8–6.9 years), but was not detected in any of three controls (aged 1.3–8.6 years). The biofilms contained both live and dead bacteria irrespective of antibiotic use prior to bronchoscopy. Biofilm was detected more frequently in the second lavage than the first. Three of the seven biofilm-positive BAL were culture-positive for respiratory pathogens at clinically important levels. Conclusions: Biofilm is present in BAL from children with non-CF bronchiectasis even when BAL-defined clinically important infection was absent. Studies to characterize lower airway biofilms and determine how biofilm contributes to bronchiectasis disease progression and treatment ß 2014 Wiley Periodicals, Inc. outcomes are necessary. Pediatr Pulmonol. Key words: respiratory infection; chronic suppurative lung disease; confocal microscopy. Funding source: University of Western Australia, State and Commonwealth Governments, Australian National Health and Medical Research Council (NHMRC) Fellowships, Numbers: 1034703, 1024175, 545216; Post-graduate scholarship, Number: 1038415; Centre for Research Excellence in Respiratory Health for Aboriginal and Torres Strait Islander Children, Number: 1040830.
INTRODUCTION
The burden of bronchiectasis unrelated to cystic fibrosis (CF) is increasingly recognized.1 In adults, 1 Child Health Division, Menzies School of Health Research, Charles Darwin University, Darwin, Northern Territory, Australia. 2 School of Paediatrics and Child Health, University of Western Australia, Perth, Western Australia, Australia. 3 Telethon Institute for Child Health Research, Centre for Child Health Research, University of Western Australia, Western Australia, Australia. 4 Queensland Children’s Respiratory Centre, Queensland Children’s Medical Research Institute, Royal Children’s Hospital, Brisbane, Queensland, Australia.
Robyn L. Marsh and Ruth B. Thornton contributed equally to this work. Authors contribution: The authors declare the following competing interests: R.B. Thornton has received travel funding from GlaxoSmithKline; P. Richmond has received Institutional funding from GSK for investigator-
ß 2014 Wiley Periodicals, Inc.
bronchiectasis prevalence and mortality have increased during the last two decades.2,3 In children, bronchiectasis is particularly prevalent in settings with disadvantaged populations such as Indigenous communities in Australia, led epidemiological studies in otitis media and has received travel support from GSK, Pfizer, and other vaccine companies to present scientific data and chair workshops. Other authors have no competing interests to declare. None of the authors have shares or paid employment with any pharmaceutical company.
Correspondence to: Robyn L. Marsh, Menzies School of Health Research, PO Box 41096, Casuarina, NT 0811, Australia. E-mail: [email protected] Received 29 June 2013; Accepted 3 February 2014. DOI 10.1002/ppul.23031 Published online in Wiley Online Library (wileyonlinelibrary.com).
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Alaska, and New Zealand.4 Despite the burden, there are relatively few pathobiological data on bronchiectasis in children. The pathogenesis of bronchiectasis likely involves a combination of host and microbial factors including reduced mucociliary clearance and ongoing inflammation related to endobronchial infection.5 Not surprisingly, antibiotics play a central role in bronchiectasis treatment1,6 with prolonged therapy often required to prevent acute exacerbations and disease progression.7 The chronic and recalcitrant nature of airway infection in children with bronchiectasis is suggestive of a biofilmassociated infection.7,8 Biofilms are defined as clusters of bacteria enclosed in matrices of self-produced and hostderived DNA and exopolymeric substances.8 Bacteria in biofilm exhibit increased resistance to antibiotics and host immune responses,9 and can be difficult to culture. Indeed, biofilm-associated infections are often culturenegative despite clinical signs of infection.8 One recent retrospective Australian study reported culture of pathogenic bacteria at clinically important levels (>104 cfu/ml of BAL) from only 68% of BAL from 113 children with non-CF bronchiectasis;10 however, it is not known if the low recovery rate was due to the presence of biofilm or other factors. Biofilms produced by Pseudomonas aeruginosa, Haemophilus influenzae, and other common respiratory pathogens have been demonstrated in lower airways specimens from CF patients,11–13 bronchus tissue from an adult with bronchiectasis,14 and middle ear specimens from patients with chronic and recurrent otitis media.15–17 In vitro studies have also shown that respiratory pathogens such as Streptococcus pneumoniae, nontypeable H. influenzae, and Moraxella catarrhalis can produce biofilm.13,18,19 Although culture-based studies report these species as the most commonly isolated respiratory pathogens from lower airway specimens of children with non-CF bronchiectasis,20 no studies have determined if bacteria in a biofilm are also present. To date, direct detection of biofilm in bronchoalveolar lavage (BAL) specimens has only been described in CF, which has a different pathobiology to non-CF bronchiectasis. Current recommendations for pediatric BAL analyses are for the first lavage to be used for microbiological assessment and the second lavage (þ/ a third lavage) for cytology and inflammation studies.21 No studies to date have determined whether the first or second lavage should be used for biofilm testing. It is biologically plausible that the second lavage may better demonstrate biofilm, possibly due to improved mucosal sampling following removal of airway mucus by the first lavage. This hypothesis is supported by data from an otitis media animal model that showed increased detection of biofilm following multiple sequential middle ear lavages.22 Thus, the aims of this study are to determine (i) if biofilm is Pediatric Pulmonology
present in lower airway specimens from children with and without non-CF bronchiectasis, and (ii) whether biofilm detection differs between two sequentially collected lavages. MATERIALS AND METHODS Ethics Statement
This study was approved by the Human Research Ethics Committee of the Northern Territory Department of Health and Menzies School of Health Research, Darwin (HREC 07/63), and the Royal Children’s Hospital, Brisbane (HREC 2003/017 and 200800064). Written informed consent was obtained from the parent/ career(s) of each child. Specimen Collection
BAL were collected as part of two prospective studies. BAL from children with non-CF bronchiectasis were selected from a prospective study of pediatric bronchiectasis at the Royal Darwin Hospital, Darwin, Australia, between January 2010 and July 2011.23 BAL for biofilm testing were selected randomly from children with bronchiectasis confirmed by chest high resolution computed tomography (cHRCT).7 As the aim of the study was to provide proof-of-principle demonstrating if biofilm was present in young children with non-CF bronchiectasis, the study was arbitrarily restricted to analysis of BAL from eight children aged 104 cfu/ ml (Table 1). Pediatric Pulmonology
DISCUSSION
In this first study to evaluate direct detection of bacterial biofilm in lower airway specimens from children with non-CF bronchiectasis, we found biofilms in the BAL of 87.5% of eight children with bronchiectasis, and 0% of three controls. Biofilm was also demonstrated in BAL that were culture-negative for respiratory pathogens at clinically important levels. Lavage-2 was better than lavage-1 for biofilm detection. These data support the hypothesis that pediatric bronchiectasis is a biofilm-associated infection 7 and add to the growing body of work demonstrating biofilm in upper and lower airway specimens. Our study has demonstrated biofilm in BAL from children with non-CF bronchiectasis; previously it was reported in bronchus tissue from an adult bronchiectasis patient;14 sputum, BAL, and lung tissue from CF patients;12,29,30 middle ear effusions and biopsies from otitis media patients;15–17 and adenoid tissue from patients with chronic adenotonsillitis, otitis media, or obstructive sleep apnoea.31,32 Collectively, these data suggest an important role for biofilm in the
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Fig. 1. Biofilms and DNA stranding were observed in BAL from children with bronchiectasis. Representative maximum projection images showing biofilm in BAL from children with bronchiectasis. Extensive biofilm was seen throughout lavage-1 and lavage-2 from child 3 (arrows), while microcolonies were observed in lavage-2 (arrow) but not lavage-1 from child 4. BacLightTM staining. Live bacteria are stained green. Dead bacteria are stained red. Scale bar ¼ 10 mm.
development and persistence of chronic respiratory infections. Further studies are now required to determine the prevalence of biofilm-associated lower airway infection in larger cohorts of children with non-CF bronchiectasis, and to test for associations between the presence of lower airway biofilm and important clinical parameters such as age, disease severity, recurrence of infections, and acute exacerbations. Testing for biofilm in lower airway specimens from children with acute and sub-acute infections is also warranted. Further research is also required to understand the bacteriology and mechanisms underlying biofilm development in the lower airways of children with bronchiectasis. Three of the seven biofilm-positive children were culture-positive for the respiratory pathogens H. influenzae, S. pneumoniae, and/or M. catarrhalis at loads indicative of lower airway infection. Earlier in vitro
studies have shown that cultured isolates of each of these species can produce biofilm.13,18,19 As bacteria in biofilm can be difficult to recover by culture,8 it is possible that other species, or bacteria cultured below clinically important levels, may be present in the lower airway biofilms. This limitation may be overcome by using DNAbased methods to characterize lower airway bacteriology when biofilm-associated infections are present. Speciesspecific testing using fluorescent in situ hybridization (FISH) techniques is also required to characterize the bacteriology of lower airway biofilm in children with nonCF bronchiectasis. However, FISH-based studies will be complicated by the polymicrobial nature of lower airway infections in bronchiectasis. Between three and seven different species were cultured from the biofilm-positive children with bronchiectasis. One child was culturepositive for S. pneumoniae, H. influenzae, Pseudomonas Pediatric Pulmonology
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Fig. 2. X-Y-Z images showing biofilm ultrastructure. X-Y-Z images of BacLightTM stained lavage-2 specimens showing biofilm ultrastructure in the microcolony from child 4 and the more mature biofilm from child 3. The side panels indicate the thickness of the collected images stacks. For child 3 the collected stack was 17 mm thick while that from child 4 was 13.5 mm thick. Live bacteria are stained green. Dead bacteria are stained red. Scale bar ¼ 10 mm.
sp., Staphylococcus sp., b-hemolytic streptococci, a-hemolytic streptococci, and other unidentified species. As bacteria in biofilm may not be detected by culture,8 the presence of yet more bacteria in these lower airway biofilms cannot be excluded. Regardless of the species involved, detection of biofilm in BAL has potential therapeutic implications as bacteria
Fig. 3. Bacterial biofilm matrices demonstrated by lectin staining. Concanavalin A (ConA) and wheat germ agglutinin (WGA) lectin staining (green), and propidium iodide staining of DNA (red). Representative maximum projection image from child 8 showing bacterial biofilm with bacteria surrounded by lectins binding the matrix (arrows). Scale bar ¼ 10 mm.
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within biofilm show higher levels of resistance to antibiotic therapies and immune clearance.9,13 While the planktonic bacteria responsible for acute exacerbations may be eradicated using standard anti-microbial therapies, biofilm eradication may not be achieved. Failure to eradicate bacteria in lower airway biofilms may result in persistent airway infection stimulating chronic inflammation.33 Biofilms may also provide a reservoir of bacteria that may be released to cause recurrent acute exacerbations. This may be particularly important following viral infection as in vitro assays demonstrate virus-induced release of planktonic bacteria from biofilm.34 Direct demonstration of biofilm in BAL from children with bronchiectasis suggests that development of adjunct anti-biofilm strategies that complement standard therapies may be needed to effectively treat these lower airway infections. One such strategy may include the earlier use of azithromycin that is known to diminish biofilm development in vitro;35 however, this has not been proven in vivo. Biofilms in the children with bronchiectasis were associated with inflammatory cells and extracellular DNA. Overall, lower airway neutrophil percentages were 10% in six of the seven biofilm-positive children. More extensive DNA stranding was observed in BAL from children with bronchiectasis than in the control children, likely reflecting either active immune responses such as neutrophil extracellular traps,36 or the presence of necrotic neutrophils. This finding requires further investigation as neutrophil extracellular traps may also contribute to lower airway inflammation.36 Additionally, neutrophil extracellular traps may also provide a target for new treatment strategies.17
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Fig. 4. No evidence of bacterial biofilms in lavage-1 or lavage-2 from control children. Representative maximum projection images of BacLightTM stained BAL demonstrating intact human cells but no bacteria in lavage-1 and lavage-2 from child 10 and evidence of planktonic bacteria (arrows), but not biofilm in lavage-1 and lavage-2 from child 11. Scale bar ¼ 10 mm.
To our knowledge this is the first study to compare testing for biofilm in sequentially collected BAL. We found that biofilms in the bronchiectasis patients were detected in the second lavage more commonly than in the first. This finding most likely reflects improved mucosal sampling following removal of airway mucus by the first lavage. While this hypothesis remains to be tested in larger studies, a similar finding was reported for multiple middle ear lavages from a chinchilla model of nontypeable H. influenzae otitis media.22 This has potential implications for bacteriological analyses performed only on the first lavage (as per current recommendations),21 as this may lead to under-reporting of biofilm. Thus, we recommend that future studies test for biofilm in both the first and second lavage. Where specimen volume is limited, the second lavage should be used for biofilm testing. Further studies are also required to determine if detection of biofilm in both lavages (as was observed for three of the seven biofilm-positive children) is associated
with more severe disease. Viable bacteria in biofilm formation were observed in BAL that had been stored for up to 12 months at 808C. While this indicates the suitability of long-term storage at 808C for maintaining biofilm integrity, further testing is required to determine if specimens stored at 808C for >12 months are suitable for biofilm testing. As with other lower airway studies,37 we cannot exclude potential oropharyngeal contamination of the lower airway lavages. a-Hemolytic streptococci were present in BAL from all children indicating a degree of oropharyngeal contamination. Detection of biofilmassociated with squamous epithelial cells in lavages from two children with bronchiectasis also suggests a degree of upper respiratory contamination. However, biofilm not associated with squamous cells was also present in each of these children consistent with lower airway sampling. Challenges associated with differentiating upper and lower airway microbiology are common Pediatric Pulmonology
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to many respiratory studies37 and must be considered in future bronchiectasis biofilm research. In conclusion, we have demonstrated the presence of biofilm in BAL from children with non-CF bronchiectasis without culture-based evidence of P. aeruginosa infection. Importantly, some of these occurred even when BAL-defined clinically important infection was absent. Studies to understand the role of biofilm in the pathobiology of bronchiectasis and other chronic lower respiratory infections are required. Evaluation of possible associations between biofilm detection and markers of clinical severity is also required. As biofilm was more commonly detected in the second lavage, in contrast to recommended guidelines for microbiological work,21 we recommend that both lavages are used for biofilm examination. ACKNOWLEDGMENTS
We would like to acknowledge the families and children that have participated in our studies. The authors also acknowledge the facilities, scientific, and technical assistance of the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterisation & Analysis at The University of Western Australia (funded by The University of Western Australia, State and Commonwealth Governments). RLM, HCS-V, and ABC are supported by Australian National Health and Medical Research Council (NHMRC) Fellowships 1034703, 1024175, and 545216. SJP is supported by NHMRC Post-graduate Scholarship 1038415. The work was also supported by the NHMRC Centre for Research Excellence in Respiratory Health for Aboriginal and Torres Strait Islander children (1040830). The views expressed in this publication are those of the authors and do not reflect the views of the NHMRC. Funding organizations had no role in the study design, data collection, and analysis, decision to publish, manuscript preparation or decision to submit the manuscript for publication. REFERENCES 1. Chang AB, Bell SC, Byrnes CA, Grimwood K, Holmes PW, King PT, Kolbe J, Landau LI, Maguire GP, McDonald MI, et al. Chronic suppurative lung disease and bronchiectasis in children and adults in Australia and New Zealand. Med J Aust 2010;193:356–365. 2. Roberts HJ, Hubbard R. Trends in bronchiectasis mortality in England and Wales. Resp Med 2010;104:981–985. 3. Seitz AE, Olivier KN, Steiner CA, Montes de Oca R, Holland SM, Prevots DR. Trends and burden of bronchiectasis-associated hospitalizations in the United States, 1993–2006. Chest 2010; 138:944–949. 4. Chang AB, Marsh RL, Smith-Vaughan HC, Hoffman LR. Emerging drugs for bronchiectasis. Expert Opin Emerg Drugs 2012;17:361–378. 5. King P. Pathogenesis of bronchiectasis. Paediatr Resp Rev 2010;12:104–110.
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6. Pasteur MC, Bilton D, Hill AT. British Thoracic Society guideline for non-CF bronchiectasis. Thorax 2010;65:i1–i58. 7. Chang AB, Byrnes CA, Everard ML. Diagnosing and preventing chronic suppurative lung disease (CSLD) and bronchiectasis. Paediatr Respir Rev 2011;12:97–103. 8. Hall-Stoodley L, Stoodley P, Kathju S, Hoiby N, Moser C, William CJ, Moter A, Bjarnsholt T. Towards diagnostic guidelines for biofilm-associated infections. FEMS Immunol Med Microbiol 2012;65:127–145. 9. Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science 1999;284:1318– 1322. 10. Kapur N, Grimwood K, Masters IB, Morris PS, Chang AB. Lower airway microbiology and cellularity in children with newly diagnosed non-CF bronchiectasis. Pediatr Pulmonol 2012;47: 300–307. 11. Lam J, Chan R, Lam K, Costerton JW. Production of mucoid microcolonies by Pseudomonas aeruginosa within infected lungs in cystic fibrosis. Infect Immun 1980;28:546–556. 12. Bjarnsholt T, Jensen PO, Fiandaca MJ, Pedersen J, Hansen CR, Andersen CB, Pressler T, Givskov M, Hoiby N. Pseudomonas aeruginosa biofilms in the respiratory tract of cystic fibrosis patients. Pediatr Pulm 2009;44:547–558. 13. Starner TD, Zhang N, Kim G, Apicella MA, McCray PB. Haemophilus influenzae forms biofilms on airway epithelia. Am J Resp Crit Care 2006;174:213–220. 14. Ohgaki N. Bacterial biofilm in chronic airway infection. Kansenshogaku Zasshi 1994;68:138–151. 15. Hall-Stoodley L, Hu FZ, Gieseke A, Nistico L, Nguyen D, Hayes J, Forbes M, Greenberg DP, Dice B, Burrows A, et al. Direct detection of bacterial biofilms on the middle-ear mucosa of children with chronic otitis media. JAMA 2006;296:202–211. 16. Thornton R, Rigby P, Wiertsema S, Filion P, Langlands J, Coates H, Vijayasekaran S, Keil A, Richmond P. Multi-species bacterial biofilm and intracellular infection in otitis media. BMC Pediatr 2011;11:94. 17. Thornton RB, Wiertsema SP, Kirkham LA, Rigby PJ, Vijayasekaran S, Coates HL, Richmond PC. Neutrophil extracellular traps and bacterial biofilms in middle ear effusion of children with recurrent acute otitis media––a potential treatment target. PLoS ONE 2013;8:e53837. 18. Moscoso M, Garcia E, Lopez R. Pneumococcal biofilms. Int Microbiol 2009;12:77–85. 19. Matejka KM, Bremer PJ, Tompkins GR, Brooks HJL. Antibiotic susceptibility of Moraxella catarrhalis biofilms in a continuous flow model. Diagn Micr Infec Dis 2012;74:394–398. 20. Grimwood K. Airway microbiology and host defences in paediatric non-CF bronchiectasis. Paediatr Respir Rev 2011;12:111–118. 21. de Blic J, Midulla F, Barbato A, Clement A, Dab I, Eber E, Green C, Grigg J, Kotecha S, Kurland G, et al. Bronchoalveolar lavage in children. ERS Task Force on bronchoalveolar lavage in children. European Respiratory Society. Eur Res J 2000;15:217–231. 22. Leroy M, Cabral H, Figueira M, Bouchet V, Huot H, Ram S, Pelton SI, Goldstein R. Multiple consecutive lavage samplings reveal greater burden of disease and provide direct access to the nontypeable Haemophilus influenzae biofilm in experimental otitis media. Infect Immun 2007;75:4158–4172. 23. Pizzutto SJ, Grimwood K, Bauert P, Schutz KL, Yerkovich ST, Upham JW, Chang AB. Bronchoscopy contributes to the clinical management of indigenous children newly diagnosed with bronchiectasis. Pediatr Pulmonol 2012;48:67–73. 24. Chang AB, Yerkovich ST, Gibson PG, Anderson-James S, Petsky HL, Carroll ML, Masters IB, Marchant JM, Wurzel D, Upham JW. Pulmonary innate immunity in children with protracted bacterial bronchitis. J Pediatr 2012;161:621–625.
Lower Airway Biofilm in Pediatric Bronchiectasis 25. Gibson L, Khoury J. Storage and survival of bacteria by ultrafreeze. Lett Appl Microbiol 1986;3:127–129. 26. Hare KM, Grimwood K, Leach AJ, Smith-Vaughan H, Torzillo PJ, Morris PS, Chang AB. Respiratory bacterial pathogens in the nasopharynx and lower airways of Australian Indigenous children with bronchiectasis. J Pediatr 2010;157:1001–1005. 27. Smith-Vaughan H, Byun R, Nadkarni M, Jacques NA, Hunter N, Halpin S, Morris PS, Leach AJ. Measuring nasal bacterial load and its association with otitis media. BMC Ear Nose Throat Disord 2006;10:10. 28. Hare KM, Marsh RL, Binks MJ, Grimwood K, Pizzutto SJ, Leach AJ, Chang AB, Smith-Vaughan HC. Quantitative PCR confirms culture as the gold standard for detection of lower airway infection by nontypeable Haemophilus influenzae in Australian Indigenous children with bronchiectasis. J Microbiol Methods 2012;92:270– 272. 29. Rudkjobing VB, Thomsen TR, Alhede M, Kragh KN, Nielsen PH, Johansen UR, Givskov M, Hoiby N, Bjarnsholt T. The microorganisms in chronically infected end-stage and non-end-stage cystic fibrosis patients. FEMS Immunol Med Microbiol 2012; 65:236–244. 30. Pusztaszeri M, Aubert JD, Braunschweig R, Mihaescu A. Pseudomonas aeruginosa biofilms in a bronchoalveolar lavage. Diagn Cytopathol 2009;37:825.
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31. Nistico L, Kreft R, Gieseke A, Coticchia JM, Burrows A, Khampang P, Liu Y, Kerschner JE, Post JC, Lonergan S, et al. Adenoid reservoir for pathogenic biofilm bacteria. J Clin Microbiol 2011;49:1411–1420. 32. Winther B, Gross BC, Hendley JO, Early SV. Location of bacterial biofilm in the mucus overlying the adenoid by light microscopy. Arch Otolaryngol Head Neck Surg 2009;135:1239–1245. 33. Cole PJ. Inflammation: a two-edged sword––the model of bronchiectasis. Eur J Respir Dis Suppl 1986;147:6–15. 34. Chattoraj SS, Ganesan S, Jones AM, Helm JM, Comstock AT, Bright-Thomas R, Lipuma JJ, Hershenson MB, Sajjan US. Rhinovirus infection liberates planktonic bacteria from biofilm and increases chemokine responses in cystic fibrosis airway epithelial cells. Thorax 2011;66:333–339. 35. Starner TD, Shrout JD, Parsek MR, Appelbaum PC, Kim G. Subinhibitory concentrations of azithromycin decrease nontypeable Haemophilus influenzae biofilm formation and diminish established biofilms. Antimicrob Agents Chemother 2008;52:137–145. 36. Kaplan MJ, Radic M. Neutrophil extracellular traps: double-edged swords of innate immunity. J Immunol 2012;189:2689–2695. 37. Charlson ES, Bittinger K, Haas AR, Fitzgerald AS, Frank I, Yadav A, Bushman FD, Collman RG. Topographical continuity of bacterial populations in the healthy human respiratory tract. Am J Resp Crit Care 2011;184:957–963.
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