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Practical Laboratory Aspects of Cystic Fibrosis Microbiology: an Update, Part I
Clinical Microbiology Newsletter Vol. 34, No. 4
www.cmnewsletter.com
Stay Current... Stay Informed.
February 15, 2012
Practical Laboratory Aspects of Cystic Fibrosis Microbiology: an Update, Part I Deanna L. Kiska, Ph.D., D(ABMM) and Scott W. Riddell, Ph.D., MT (ASCP), D(ABMM), Department of Clinical Pathology, SUNY Upstate Medical University, Syracuse, New York
Abstract Cystic fibrosis (CF) microbiology occupies a unique niche in the clinical microbiology laboratory. It is notable for the diversity of potential pathogens, the labor-intensive nature of culture evaluation, and the complexity of organism identification and susceptibility testing. This article provides an update of the major CF pathogens and offers a practical laboratory guide that addresses some of the issues encountered with these cultures. Part I of this two-part article reviews the new and most current information gathered from the literature since 2006 with respect to Pseudomonas aeruginosa, Burkholderia cepacia complex, and other glucose non-fermenting gram-negative rods. Part II of this article will complete the review of the key organisms associated with CF infections, including Staphylococcus aureus, nontuberculous mycobacteria, and fungi. Issues regarding susceptibility testing will also be addressed. Finally, a practical laboratory guide will be provided to address some of the confounding issues associated with CF microbiology. Cystic fibrosis (CF) is the most common hereditary disease in the Caucasian population. The genetic defect occurs in the CF transmembrane conductance regulator (CFTR) gene, a gene that codes for a protein that regulates the transport of electrolytes across epithelial cell membranes. Mutations in the CFTR gene affect sodium and chloride ion transport, resulting in the disruption of the ionic composition and volume of airway surface fluid. This fluid is normally thin to allow removal of inhaled microorganisms via ciliary action; however, in the presence of CFTR mutations, it increases in volume and becomes viscous, clogging the airways. As a result, microorganisms entering the distal airEditor‘s Note: Part II of this article will appear in the March 1, 2012 issue of CMN (Vol. 34, No. 5). Corresponding Author: Deanna L. Kiska, Department of Clinical Pathology, SUNY Upstate Medical University, 750 East Adams St., Syracuse, NY 13210. Tel.: 315-464-6713. Fax: 315-464-6836. E-mail: [email protected]
Clinical Microbiology Newsletter 34:4,2012
ways are not cleared and can cause chronic infections with progressive inflammation and respiratory insufficiency. Chronologically, these infections develop early in life, with Staphylococcus aureus and Haemophilus influenzae as the predominant pathogens. Infection with various morphotypes of Pseudomonas aeruginosa soon follows, resulting in significant morbidity as the inflammatory response progresses. Other organisms associated with chronic infection in CF patients include the Burkholderia cepacia complex (Bcc) and non-tuberculous mycobacteria. As molecular tools are employed to study the microbial diversity of the CF lung, the plethora of organisms is staggering (1). Although some of these microorganisms have been linked to poor outcomes, the role of others as significant players in disease progression is uncertain. For a thorough review of the major CF pathogens, the reader is referred to Cumitech 43: Cystic Fibrosis Microbiology (2) and a recent publication on CF micro© 2012 Elsevier
bial epidemiology (3). The present article incorporates new information gathered from the literature since Cumitech 43 was published in 2006 and provides an update on practical laboratory aspects of CF microbiology.
Pseudomonas aeruginosa P. aeruginosa is the most important and frequent pathogen in CF patients, although some studies report that the annual prevalence is declining (4,5). Infection may be established early in life, with the prevalence increasing with age, reaching 80% in the adult
0196-4399/00 (see frontmatter)
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CF population. Non-mucoid strains predominate in early infection but eventually give rise to the classic mucoid variants, which are nearly impossible to eradicate. Isolation of P. aeruginosa is not difficult, as these organisms grow well on routine media used for culture of CF specimens. Mixed morphotypes with different susceptibility patterns are often present on culture, with one study demonstrating a mean of 4 morphotypes per sputum specimen (6). However, detecting these morphotypes is problematic, particularly when the mucoid phenotype is present, as it may obscure other variant forms (7). Observations from several studies have revealed methods which may help to resolve this issue. In a quality assurance study of diagnostic CF microbiology laboratories, those laboratories that used a quantitative culture technique involving sputum homogenization correctly detected both mucoid and non-mucoid isolates in a simulated sample (7). Along the same lines, employing a commercially available mucolytic agent (COPAN SL; COPAN, Murrieta, CA) for processing CF sputum samples allowed detection of 19% more P. aeruginosa isolates than standard plating (8). In another study, a selective chromogenic agar (ChromID P. aeruginosa; bioMérieux, France) afforded greater detection and differentiation of morphotypes because of decreased polysaccharide production on the medium, preventing highly mucoid isolates from obscuring other variants (9). These methods are currently not standard practice for CF microbiology, but they clearly show potential for greater detection of P. aeruginosa. P. aeruginosa is an oxidase-positive, glucose non-fermenting gram-negative bacillus whose identification from CF patients is relatively straightforward, but a few caveats deserve mention. In the largest study to date on identification of CF P. aeruginosa isolates,
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the rate of misidentification was low (2.3%) (10) and occurred primarily with three species, Achromobacter xylosoxidans, Stenotrophomonas maltophilia, and Inquilinus limosus. Two-thirds of the misidentified isolates were from laboratories that used basic phenotypic tests, such as colony morphology and oxidase, for presumptive identification. A. xylosoxidans and S. maltophilia could have been ruled out by careful attention to colony morphology (i.e., pigment production) and stringent reading of the oxidase reaction. All misidentified A. xylosoxidans isolates were non-pigmented, and all S. maltophilia isolates were oxidase-negative within 10 to 15 seconds. Therefore, an accurate presumptive identification of non-mucoid P. aeruginosa should be based on a positive oxidase reaction (10 to 15 seconds), no lactose fermentation on MacConkey agar, and the presence of a green pigment or metallic sheen. For mucoid P. aeruginosa, presumptive identification has relied on colonial morphology, a positive oxidase reaction, and lack of fermentation on MacConkey agar. Since very few non-fermenting gramnegative bacilli display this characteristic mucoid morphology, these simple phenotypic tests have been sufficient for identification. However, I. limosus is a mucoid glucose nonfermenter that can resemble P. aeruginosa. It is infrequently isolated from CF patients, although one study noted a prevalence similar to that of the Bcc (11). Its mucoid colonial morphology, positive oxidase reaction, and ability to grow at 42°C can cause confusion with P. aeruginosa; however, it exhibits poor or no growth on MacConkey agar and is colistin resistant (12,13). The last two characteristics in an organism resembling P. aeruginosa should arouse suspicion, and definitive identification should be pursued, since prognoses and treatment protocols differ with these two organisms. I. limosus
© 2012 Elsevier
is not included in the databases of most commercial identification systems and is frequently misidentified by these systems as Sphingomonas paucimobilis or Agrobacterium radiobacter. Definitive identification can be achieved by 16S rRNA gene sequencing or matrixassisted laser desorption ionization-timeof-flight mass spectrometry (MALDITOF MS) (14,15). Identification of atypical isolates of P. aeruginosa is often problematic, since they lack typical morphologic features, grow slowly, and are often assacharolytic. One study noted that 19% of P. aeruginosa isolates were atypical, with only 25% of these giving an acceptable identification (score, ≥90%) with a commercial system (API20NE; bioMérieux, Durham, NC) (16). Most of the isolates (75%) fell into an ambiguous/unacceptable category or were “acceptably” identified (score, ≥90%) as various infrequently isolated, non-fermenting species. In these scenarios, further definitive/confirmatory identification should be pursued.
Burkholderia cepacia Complex The Bcc is comprised of at least 17 different closely related species (Table 1), all of which have been isolated from CF patients, except B. ubonensis (3,17,18). The species B. cepacia, B. multivorans, B. cenocepacia, B. stabilis, and B. vietnamiensis account for the great majority of clinical isolates, with B. cenocepacia and B. multivorans predominating (3,19-21); the prevalence of the Bcc in the U.S. CF population is approximately 3 to 4% (3,5,21). The clinical course following colonization with these organisms is variable, with some patients developing fever, bacteremia, and necrotizing pneumonia (cepacia syndrome) while others apparently experience little change or decline (20,21). It had been thought that once CF patients became colonized with Bcc species they always remained
Clinical Microbiology Newsletter 34:4,2012
colonized, but for some patients, this colonization is transient (3). Recent studies have attempted to determine whether some members of the Bcc are associated with greater morbidity and mortality than others. While most commonly associated with B. cenocepacia, fatal infection has also been observed with B. multivorans, B. stabilis, and B. dolosa (1,18,21-23). In particular, specific strains of B. cenocepacia seem to be associated with high transmissibility and poor patient outcomes (19,22). The severity of disease may be related to the infecting strain rather than to a particular species, and host factors may also play a role (18,24-26). Indeed, infection with different strains of the same species may result in various outcomes in patients with the same CFTR defect (18). Most cases of person-to-person transmission have been linked to B. cenocepacia (24), and measures to prevent the spread of Bcc organisms both inside and outside of health care facilities have greatly reduced infection and mortality rates (25,26). Other Burkholderia species causing infection in the CF population are B. gladioli, B. fungorum, and B. pseudomallei (3,27-29). B. gladioli is isolated more frequently than most Bcc species and therefore accounts for a significant percentage of Burkholderia sp. isolates from CF patients (3). A recent retrospective review of 33 CF patients with at least one positive culture for B. gladioli found that infection with the organism was transient in most cases and had a variable clinical impact (30). The accurate and timely identification of Bcc members is critical and has both epidemiological and clinical implications (1,25,26). The closely related of the Bcc are difficult to identify using conventional methods, and the use of commercial biochemical phenotyping systems may lead to inaccurate identification (1,31,32). Previously, species within the Bcc have been incorrectly identified by these systems as B. gladioli, Ralstonia pickettii, Pandoraea spp., Alcaligenes spp., Pseudomonas spp., S. maltophilia, Flavobacterium spp., and Chryseobacterium spp. (1,21,22,31,3335). The opposite may also occur, that is, members of these other genera may be misidentified as Bcc (1,33). More recently, Zbinden et al. (36) found that the colorimetric Vitek 2 (bioMéreiux) Clinical Microbiology Newsletter 34:4,2012
gram-negative identification card (IDGN) correctly identified 12 of 13 Bcc isolates, and almost all tested strains of A. xylosoxidans, Acinetobacter spp., P. aeruginosa, and S. maltophilia but did misidentify one A. xylosoxidans isolate as B. cepacia group. From these findings, the authors concluded that any glucose non-fermenter giving an identification other than the above-mentioned organisms should be subjected to molecular analysis. Currently, 16S rRNA gene sequencing has acceptable discriminatory power and is capable of accurately distinguishing Bcc from other glucose non-fermenters; however, this method has limited species resolution within the complex. Species discrimination is also limited for Ralstonia, Pandoraea, and Bordetella (1). The use of Stewart’s medium has been proposed as a simple screening test to reduce the number of false-positive isolates from B. cepacia selective media (37). Using this medium, 91% of 72 Bcc reference strains demonstrated the oxidation of glucose (yellow slant) and lack of arginine dihydrolase activity (green butt) after 48 hours of incubation, thus distinguishing the isolates from most other glucose non-fermenters, including P. aeruginosa, Stenotrophomonas spp., Achromobacter spp., and Pandoraea spp. The accuracy of Bcc identification by commercial systems can be improved by further testing such strains for growth on BCSA, lysine and ornithine decarboxylase activity, oxidation of sucrose and adonitol, hemolysis, pigment production, and growth at 42°C (33,35). A combination of phenotypic and molecular methods (“polyphasic” approach) may be required to identify Burkholderia spp. accurately (1,17,31). Alternative methods for identification include chromatography and mass spectrometry. Krejcí et al. (34) used cellular fatty acid analysis by high-performance liquid chromatography to identify B. ambifaria, B. pyrrocinia, B. vietnamiensis, Pandoraea spp. and Ralstonia spp. but were unable to resolve other Bcc members. MALDI-TOF MS has also recently been applied to the identification of Bcc species. Vanlaere et al. (38) analyzed 75 Bcc strains and a panel of commonly misidentified species using this technology. Bcc members were distinguished from non-Bcc species and from each other, with the exception of © 2012 Elsevier
Table 1. Current members of the Burkholderia cepacia complexa Former genomovar designation
Species B. cepacia B. multivorans B. cenocepacia B. stabilis B. vietnamiensis B. dolosa B. ambifaria B. anthina B. pyrrocinia B. ubonensis B. latens B. diffusa B. arboris B. seminalis B. metallica B. contaminans B. lata
I II III IV V VI VII VIII IX
a
From references 3, 24, and 38.
B. anthina and B. pyrrocinia, which clustered together. In a similar study (15), MALDI-TOF MS was able to correctly identify 98% of 558 strains of glucose non-fermenting gram-negative rods representing 58 separate species, including 52 Bcc isolates. One author has suggested that MALDI-TOF MS be considered the gold standard for CF isolate identification, particularly for isolates from BCSA, with confirmation by 16S rRNA gene sequencing when necessary (1). 16S rRNA gene sequencing has acceptable discriminatory power and is capable of accurately distinguishing Bcc from other glucose non-fermenters; however, the method has limited species resolution within the complex. Species discrimination is also limited for Ralstonia, Pandoraea, and Bordetella (1). While there are phenotypic tests that help separate Bcc members from other genera, it is critical that clinical isolates phenotypically identified as Bcc be sent to a specialized laboratory (e.g., the B. cepacia Refersence Laboratory) for definitive identification (for a list of laboratories, see http://go.to/cepacia). Good laboratory practice also includes the submission to a reference laboratory of any glucose non-fermenters that grow on B. cepacia selective agars or for which phenotypic identification fails or is in doubt. Strong consideration should also be given to referring glucose non0196-4399/00 (see frontmatter)
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fermenting gram-negative rods exhibiting resistance to colistin (Table 2).
Table 2. Glucose non-fermenting gram-negative bacilli that may demonstrate colistin resistancea
Other Gucose Non-fermenting Gram-Negative Rods
Species Achromobacter xylosoxidans subsp. xylosoxidans Achromobacter piechaudii Acinetobacter calcoaceticus/baumannii Bergeyella zoohelcum Brevundimonas diminuta Brevundimonas vesicularis Burkholderia cepacia Burkholderia pseudomallei Chryseobacterium indologenes Chryseobacterium meningosepticum Delftia acidovorans Empedobacter brevis Myroides odoratus/odoratimimus Ochrobactrum anthropi/intermedium Ralstonia pickettii Ralstonia mannitolilytica Rhizobium radiobacter Sphingobacterium multivorum Sphingomonas paucimobilis Stenotrophomonas maltophilia
Glucose non-fermenting gram-negative rods other than P. aeruginosa and B. cepacia appear to be increasing in frequency, perhaps as a result of clinical and microbiological factors (3). The aging CF population may afford more opportunities for infection with infrequent and unusual organisms, particularly in the setting of chronic suppressive therapy and early-eradication protocols. These treatment regimens may be contributing to the decreasing incidence of P. aeruginosa and the selection of emerging pathogens, many of which are multi-drug resistant and can persist in the airways (3). The two most common isolates are S. maltophilia and A. xylosoxidans; however, the list of additional organisms continues to grow (1). The pathogenic roles of these organisms remain unclear, but they can be problematic from a laboratory perspective. Their identification is often hampered, because most commercial identification system databases do not contain some of the genera (Inquilinus, Herbaspirillum, and Pandoraea) or may only include a limited number of species (Cupriavidus, Ralstonia, and Bordetella); the Biolog (Biolog, Inc., Hayward, CA) and Sherlock MIDI (MIDI, Inc., Newark, DE) systems are exceptions. To further compound the problem, many of these organisms grow on BCSA and are misidentified as Bcc by some commercial identification systems. These issues should be taken into account in the laboratory’s B. cepacia protocol (see above). Editor’s note: Part II of this article will appear in the March 1, 2012 issue of CMN (vol. 34, No. 5). References 1. Bittar, F. and J.M. Rolain. 2010. Detection and accurate identification of new or emerging bacteria in cystic fibrosis patients. Clin. Microbiol. Infect. 16:809-820. 2. Gilligan, P.H. and D.L. Kiska. 2006. Cumitech 43, Cystic Fibrosis Microbiology. M.D. Appleman (Coordinating ed.). American Society for Microbiology, Washington, DC. 3. Lipuma, J.J. 2010. The changing microbial epidemiology in cystic fibrosis.
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a
% Resistance 31 14 2 100 100 100 100 100 100 100 100 100 100 7 100 100 47 100 81 62
Zone diameter ≤ 6 mm (10-μg colistin disk on Mueller-Hinton agar). Table adapted from reference 102. Not all species have been isolated from CF patients.
Clin. Microbiol. Rev. 23:299-323. 4. Emerson, J. et al. 2010. Changes in cystic fibrosis sputum microbiology in the United States between 1995 and 2008. Pediatr. Pulmonol. 45:363-370. 5. Razvi, S. et al. 2009. Respiratory microbiology of patients with cystic fibrosis in the United States, 1995 to 2005. Chest 136:1554-1560. 6. Foweraker, J.E. et al. 2005. Phenotypic variability of Pseudomonas aeruginosa in sputa from patients with acute infective exacerbation of cystic fibrosis and its impact on the validity of antimicrobial susceptibility testing. J. Antimicrob. Chemother. 55:921-927. 7. Balke, B. et al. 2008. A German external quality survey of diagnostic microbiology of respiratory tract infections in patients with cystic fibrosis. J. Cyst. Fibros. 7:7-14. 8. Young, C. et al. 2011. Use of COPAN SL solution for processing sputum from patients with and without cystic fibrosis. Abstr. C-2149. 111th Gen. Meet. Am. Soc. Microbiol. American Society for Microbiology, Washington, DC. 9. Laine, L. et al. 2008. A novel chromogenic medium for isolation of Pseudomonas aeruginosa from the sputa of cystic fibrosis patients. J. Cyst. Fibros. 8:143-149.
© 2012 Elsevier
10. Kidd, T.J. et al. 2009. Low rates of Pseudomonas aeruginosa misidentification in isolates from cystic fibrosis patients. J. Clin. Microbiol. 47:15031509. 11. Bittar, F. et al. 2006. Inquilinus limosus and cystic fibrosis. Emerg. Infect. Dis. 14:993-994. 12. Chiron, R. et al. 2005. Clinical and microbiological features of Inquilinus sp. isolates from five patients with cystic fibrosis. J. Clin. Microbiol. 43:39383943. 13. Cooke, R.P. et al. 2007. Inquilinus limosus isolated from a cystic fibrosis patient: first UK report. Br. J. Biomed. Sci. 64:127-129. 14. Coenye, T. et al. 2002. Characterization of unusual bacteria isolated from respiratory secretions of cystic fibrosis patients and description of Inquilinus limosus gen. nov., sp. nov. J. Clin. Microbiol. 40:2062-2069. 15. Degand, N. et al. 2008. Matrix-assisted laser desorption ionization-time of flight mass spectrometry for identification of nonfermenting gram-negative bacilli isolated from cystic fibrosis patients. J. Clin. Microbiol. 46:3361-3367. 16. Wellinghausen, N. et al. 2005. Superiority of molecular techniques for identification of gram-negative, oxidase-positive
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rods, including morphologically nontypical Pseudomonas aeruginosa, from patients with cystic fibrosis. J. Clin. Microbiol. 43:4070-4075.
Clin. Microbiol. 45:3105-3108.
22. Lipuma, J.J. 2005. Update on the Burkholderia cepacia complex. Curr. Opin. Pulm. Med. 11:528-533.
30. Kennedy, M.P. et al. 2007. Burkholderia gladioli: five year experience in a cystic fibrosis and lung transplantation center. J. Cyst. Fibros. 6:267-273.
23. Turton, J.F. et al. 2007. Revised approach for identification of isolates within the Burkholderia cepacia complex and description of clinical isolates not assigned to any of the known genomovars. J.
31. Payne, G.W. et al. 2005. Development of a recA gene-based identification approach for the entire Burkholderia genus. Appl. Environ. Microbiol. 71:3917-3927.
32. Mahenthiralingam, E., A. Baldwin, and C.G. Dowson. 2008. Burkholderia cepacia complex bacteria: opportunistic pathogens with important natural biology. J. Appl. Microbiol. 104:1539-1551. 33. Coenye, T. et al. 2001. Taxonomy and identification of the Burkholderia cepacia complex. J. Clin. Microbiol. 39:3427-3436. 34. Krejcí, E. and R.M. Kroppenstedt. 2006. Differentiation of species combined into the Burkholderia cepacia complex and related taxa on the basis of their fatty acid patterns. J. Clin. Microbiol. 44:1159-1164. 35. de Vrankrijker, A.M., T.F. Wolfs, and C.K. van der Ent. 2010. Challenging and emerging pathogens in cystic fibrosis. Paediatr. Respir. Rev. 11:246-254. 36. Zbinden, A. et al. 2007. Evaluation of the colorimetric VITEK 2 card for identification of gram-negative nonfermentative rods: comparison to 16S rRNA gene sequencing. J. Clin. Microbiol. 45:2270-2273. 37. Vanlaere, E. et al. 2006. Growth in Stewart’s medium is a simple, rapid and inexpensive screening tool for the identification of Burkholderia cepacia complex. J. Cyst. Fibros. 5:137-139. 38. Vanlaere, E. et al. 2008. Matrix-assisted laser desorption ionisation-time-of offlight mass spectrometry of intact cells allows rapid identification of Burkholderia cepacia complex. J. Microbiol. Methods 75:279-286.
© 2012 Elsevier
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17. Papaleo, M.C. et al. 2010. Identification of species of the Burkholderia cepacia complex by sequence analysis of the hisA gene. J. Med. Microbiol. 59:11631170. 18. Zlosnik, J.E. et al. 2011. Mucoid and nonmucoid Burkholderia cepacia complex bacteria in cystic fibrosis infections. Am. J. Respir. Crit. Care Med. 183:67-72. 19. Vonberg, R.P. et al. 2006. Identification of Burkholderia cepacia complex pathogens by rapid-cycle PCR with fluorescent hybridization probes. J. Med. Microbiol. 55:721-727. 20. Kalish, L.A. et al. 2006. Impact of Burkholderia dolosa on lung function and survival in cystic fibrosis. Am. J. Respir. Crit. Care Med. 173:421-425. 21. Davies, J.C. and B.K. Rubin. 2007. Emerging and unusual gram-negative infections in cystic fibrosis. Semin. Respir. Crit. Care Med. 28:312-321.
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24. Mahenthiralingam, E., T.A. Urban, and J.B. Goldberg. 2005. The multifarious, multireplicon Burkholderia cepacia complex. Nat. Rev. Microbiol. 3:144-156. 25. Govan, J.R., A.R. Brown, and A.M. Jones. 2007. Evolving epidemiology of Pseudomonas aeruginosa and the Burkholderia cepacia complex in cystic fibrosis lung infection. Future Microbiol. 2:153-164. 26. Foweraker, J. 2009. Recent advances in the microbiology of respiratory tract infection in cystic fibrosis. Br. Med. Bull. 89:93-110. 27. Asiah, K. et al. 2006. Unrecognised infection in a cystic fibrosis patient. J. Paediatr. Child Health 42:217-218. 28. Barth, A.L. et al. 2007. Cystic fibrosis patient with Burkholderia pseudomallei infection acquired in Brazil. J. Clin. Microbiol. 45:4077-4080. 29. Corral, D.M. et al. 2008. Burkholderia pseudomallei infection in a cystic fibrosis patient from the Caribbean: a case report. Can. Respir. J. 15:237-239.
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www.cmnewsletter.com
Stay Current... Stay Informed.
February 15, 2012
Practical Laboratory Aspects of Cystic Fibrosis Microbiology: an Update, Part I Deanna L. Kiska, Ph.D., D(ABMM) and Scott W. Riddell, Ph.D., MT (ASCP), D(ABMM), Department of Clinical Pathology, SUNY Upstate Medical University, Syracuse, New York
Abstract Cystic fibrosis (CF) microbiology occupies a unique niche in the clinical microbiology laboratory. It is notable for the diversity of potential pathogens, the labor-intensive nature of culture evaluation, and the complexity of organism identification and susceptibility testing. This article provides an update of the major CF pathogens and offers a practical laboratory guide that addresses some of the issues encountered with these cultures. Part I of this two-part article reviews the new and most current information gathered from the literature since 2006 with respect to Pseudomonas aeruginosa, Burkholderia cepacia complex, and other glucose non-fermenting gram-negative rods. Part II of this article will complete the review of the key organisms associated with CF infections, including Staphylococcus aureus, nontuberculous mycobacteria, and fungi. Issues regarding susceptibility testing will also be addressed. Finally, a practical laboratory guide will be provided to address some of the confounding issues associated with CF microbiology. Cystic fibrosis (CF) is the most common hereditary disease in the Caucasian population. The genetic defect occurs in the CF transmembrane conductance regulator (CFTR) gene, a gene that codes for a protein that regulates the transport of electrolytes across epithelial cell membranes. Mutations in the CFTR gene affect sodium and chloride ion transport, resulting in the disruption of the ionic composition and volume of airway surface fluid. This fluid is normally thin to allow removal of inhaled microorganisms via ciliary action; however, in the presence of CFTR mutations, it increases in volume and becomes viscous, clogging the airways. As a result, microorganisms entering the distal airEditor‘s Note: Part II of this article will appear in the March 1, 2012 issue of CMN (Vol. 34, No. 5). Corresponding Author: Deanna L. Kiska, Department of Clinical Pathology, SUNY Upstate Medical University, 750 East Adams St., Syracuse, NY 13210. Tel.: 315-464-6713. Fax: 315-464-6836. E-mail: [email protected]
Clinical Microbiology Newsletter 34:4,2012
ways are not cleared and can cause chronic infections with progressive inflammation and respiratory insufficiency. Chronologically, these infections develop early in life, with Staphylococcus aureus and Haemophilus influenzae as the predominant pathogens. Infection with various morphotypes of Pseudomonas aeruginosa soon follows, resulting in significant morbidity as the inflammatory response progresses. Other organisms associated with chronic infection in CF patients include the Burkholderia cepacia complex (Bcc) and non-tuberculous mycobacteria. As molecular tools are employed to study the microbial diversity of the CF lung, the plethora of organisms is staggering (1). Although some of these microorganisms have been linked to poor outcomes, the role of others as significant players in disease progression is uncertain. For a thorough review of the major CF pathogens, the reader is referred to Cumitech 43: Cystic Fibrosis Microbiology (2) and a recent publication on CF micro© 2012 Elsevier
bial epidemiology (3). The present article incorporates new information gathered from the literature since Cumitech 43 was published in 2006 and provides an update on practical laboratory aspects of CF microbiology.
Pseudomonas aeruginosa P. aeruginosa is the most important and frequent pathogen in CF patients, although some studies report that the annual prevalence is declining (4,5). Infection may be established early in life, with the prevalence increasing with age, reaching 80% in the adult
0196-4399/00 (see frontmatter)
27
CF population. Non-mucoid strains predominate in early infection but eventually give rise to the classic mucoid variants, which are nearly impossible to eradicate. Isolation of P. aeruginosa is not difficult, as these organisms grow well on routine media used for culture of CF specimens. Mixed morphotypes with different susceptibility patterns are often present on culture, with one study demonstrating a mean of 4 morphotypes per sputum specimen (6). However, detecting these morphotypes is problematic, particularly when the mucoid phenotype is present, as it may obscure other variant forms (7). Observations from several studies have revealed methods which may help to resolve this issue. In a quality assurance study of diagnostic CF microbiology laboratories, those laboratories that used a quantitative culture technique involving sputum homogenization correctly detected both mucoid and non-mucoid isolates in a simulated sample (7). Along the same lines, employing a commercially available mucolytic agent (COPAN SL; COPAN, Murrieta, CA) for processing CF sputum samples allowed detection of 19% more P. aeruginosa isolates than standard plating (8). In another study, a selective chromogenic agar (ChromID P. aeruginosa; bioMérieux, France) afforded greater detection and differentiation of morphotypes because of decreased polysaccharide production on the medium, preventing highly mucoid isolates from obscuring other variants (9). These methods are currently not standard practice for CF microbiology, but they clearly show potential for greater detection of P. aeruginosa. P. aeruginosa is an oxidase-positive, glucose non-fermenting gram-negative bacillus whose identification from CF patients is relatively straightforward, but a few caveats deserve mention. In the largest study to date on identification of CF P. aeruginosa isolates,
28
0196-4399/00 (see frontmatter)
the rate of misidentification was low (2.3%) (10) and occurred primarily with three species, Achromobacter xylosoxidans, Stenotrophomonas maltophilia, and Inquilinus limosus. Two-thirds of the misidentified isolates were from laboratories that used basic phenotypic tests, such as colony morphology and oxidase, for presumptive identification. A. xylosoxidans and S. maltophilia could have been ruled out by careful attention to colony morphology (i.e., pigment production) and stringent reading of the oxidase reaction. All misidentified A. xylosoxidans isolates were non-pigmented, and all S. maltophilia isolates were oxidase-negative within 10 to 15 seconds. Therefore, an accurate presumptive identification of non-mucoid P. aeruginosa should be based on a positive oxidase reaction (10 to 15 seconds), no lactose fermentation on MacConkey agar, and the presence of a green pigment or metallic sheen. For mucoid P. aeruginosa, presumptive identification has relied on colonial morphology, a positive oxidase reaction, and lack of fermentation on MacConkey agar. Since very few non-fermenting gramnegative bacilli display this characteristic mucoid morphology, these simple phenotypic tests have been sufficient for identification. However, I. limosus is a mucoid glucose nonfermenter that can resemble P. aeruginosa. It is infrequently isolated from CF patients, although one study noted a prevalence similar to that of the Bcc (11). Its mucoid colonial morphology, positive oxidase reaction, and ability to grow at 42°C can cause confusion with P. aeruginosa; however, it exhibits poor or no growth on MacConkey agar and is colistin resistant (12,13). The last two characteristics in an organism resembling P. aeruginosa should arouse suspicion, and definitive identification should be pursued, since prognoses and treatment protocols differ with these two organisms. I. limosus
© 2012 Elsevier
is not included in the databases of most commercial identification systems and is frequently misidentified by these systems as Sphingomonas paucimobilis or Agrobacterium radiobacter. Definitive identification can be achieved by 16S rRNA gene sequencing or matrixassisted laser desorption ionization-timeof-flight mass spectrometry (MALDITOF MS) (14,15). Identification of atypical isolates of P. aeruginosa is often problematic, since they lack typical morphologic features, grow slowly, and are often assacharolytic. One study noted that 19% of P. aeruginosa isolates were atypical, with only 25% of these giving an acceptable identification (score, ≥90%) with a commercial system (API20NE; bioMérieux, Durham, NC) (16). Most of the isolates (75%) fell into an ambiguous/unacceptable category or were “acceptably” identified (score, ≥90%) as various infrequently isolated, non-fermenting species. In these scenarios, further definitive/confirmatory identification should be pursued.
Burkholderia cepacia Complex The Bcc is comprised of at least 17 different closely related species (Table 1), all of which have been isolated from CF patients, except B. ubonensis (3,17,18). The species B. cepacia, B. multivorans, B. cenocepacia, B. stabilis, and B. vietnamiensis account for the great majority of clinical isolates, with B. cenocepacia and B. multivorans predominating (3,19-21); the prevalence of the Bcc in the U.S. CF population is approximately 3 to 4% (3,5,21). The clinical course following colonization with these organisms is variable, with some patients developing fever, bacteremia, and necrotizing pneumonia (cepacia syndrome) while others apparently experience little change or decline (20,21). It had been thought that once CF patients became colonized with Bcc species they always remained
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colonized, but for some patients, this colonization is transient (3). Recent studies have attempted to determine whether some members of the Bcc are associated with greater morbidity and mortality than others. While most commonly associated with B. cenocepacia, fatal infection has also been observed with B. multivorans, B. stabilis, and B. dolosa (1,18,21-23). In particular, specific strains of B. cenocepacia seem to be associated with high transmissibility and poor patient outcomes (19,22). The severity of disease may be related to the infecting strain rather than to a particular species, and host factors may also play a role (18,24-26). Indeed, infection with different strains of the same species may result in various outcomes in patients with the same CFTR defect (18). Most cases of person-to-person transmission have been linked to B. cenocepacia (24), and measures to prevent the spread of Bcc organisms both inside and outside of health care facilities have greatly reduced infection and mortality rates (25,26). Other Burkholderia species causing infection in the CF population are B. gladioli, B. fungorum, and B. pseudomallei (3,27-29). B. gladioli is isolated more frequently than most Bcc species and therefore accounts for a significant percentage of Burkholderia sp. isolates from CF patients (3). A recent retrospective review of 33 CF patients with at least one positive culture for B. gladioli found that infection with the organism was transient in most cases and had a variable clinical impact (30). The accurate and timely identification of Bcc members is critical and has both epidemiological and clinical implications (1,25,26). The closely related of the Bcc are difficult to identify using conventional methods, and the use of commercial biochemical phenotyping systems may lead to inaccurate identification (1,31,32). Previously, species within the Bcc have been incorrectly identified by these systems as B. gladioli, Ralstonia pickettii, Pandoraea spp., Alcaligenes spp., Pseudomonas spp., S. maltophilia, Flavobacterium spp., and Chryseobacterium spp. (1,21,22,31,3335). The opposite may also occur, that is, members of these other genera may be misidentified as Bcc (1,33). More recently, Zbinden et al. (36) found that the colorimetric Vitek 2 (bioMéreiux) Clinical Microbiology Newsletter 34:4,2012
gram-negative identification card (IDGN) correctly identified 12 of 13 Bcc isolates, and almost all tested strains of A. xylosoxidans, Acinetobacter spp., P. aeruginosa, and S. maltophilia but did misidentify one A. xylosoxidans isolate as B. cepacia group. From these findings, the authors concluded that any glucose non-fermenter giving an identification other than the above-mentioned organisms should be subjected to molecular analysis. Currently, 16S rRNA gene sequencing has acceptable discriminatory power and is capable of accurately distinguishing Bcc from other glucose non-fermenters; however, this method has limited species resolution within the complex. Species discrimination is also limited for Ralstonia, Pandoraea, and Bordetella (1). The use of Stewart’s medium has been proposed as a simple screening test to reduce the number of false-positive isolates from B. cepacia selective media (37). Using this medium, 91% of 72 Bcc reference strains demonstrated the oxidation of glucose (yellow slant) and lack of arginine dihydrolase activity (green butt) after 48 hours of incubation, thus distinguishing the isolates from most other glucose non-fermenters, including P. aeruginosa, Stenotrophomonas spp., Achromobacter spp., and Pandoraea spp. The accuracy of Bcc identification by commercial systems can be improved by further testing such strains for growth on BCSA, lysine and ornithine decarboxylase activity, oxidation of sucrose and adonitol, hemolysis, pigment production, and growth at 42°C (33,35). A combination of phenotypic and molecular methods (“polyphasic” approach) may be required to identify Burkholderia spp. accurately (1,17,31). Alternative methods for identification include chromatography and mass spectrometry. Krejcí et al. (34) used cellular fatty acid analysis by high-performance liquid chromatography to identify B. ambifaria, B. pyrrocinia, B. vietnamiensis, Pandoraea spp. and Ralstonia spp. but were unable to resolve other Bcc members. MALDI-TOF MS has also recently been applied to the identification of Bcc species. Vanlaere et al. (38) analyzed 75 Bcc strains and a panel of commonly misidentified species using this technology. Bcc members were distinguished from non-Bcc species and from each other, with the exception of © 2012 Elsevier
Table 1. Current members of the Burkholderia cepacia complexa Former genomovar designation
Species B. cepacia B. multivorans B. cenocepacia B. stabilis B. vietnamiensis B. dolosa B. ambifaria B. anthina B. pyrrocinia B. ubonensis B. latens B. diffusa B. arboris B. seminalis B. metallica B. contaminans B. lata
I II III IV V VI VII VIII IX
a
From references 3, 24, and 38.
B. anthina and B. pyrrocinia, which clustered together. In a similar study (15), MALDI-TOF MS was able to correctly identify 98% of 558 strains of glucose non-fermenting gram-negative rods representing 58 separate species, including 52 Bcc isolates. One author has suggested that MALDI-TOF MS be considered the gold standard for CF isolate identification, particularly for isolates from BCSA, with confirmation by 16S rRNA gene sequencing when necessary (1). 16S rRNA gene sequencing has acceptable discriminatory power and is capable of accurately distinguishing Bcc from other glucose non-fermenters; however, the method has limited species resolution within the complex. Species discrimination is also limited for Ralstonia, Pandoraea, and Bordetella (1). While there are phenotypic tests that help separate Bcc members from other genera, it is critical that clinical isolates phenotypically identified as Bcc be sent to a specialized laboratory (e.g., the B. cepacia Refersence Laboratory) for definitive identification (for a list of laboratories, see http://go.to/cepacia). Good laboratory practice also includes the submission to a reference laboratory of any glucose non-fermenters that grow on B. cepacia selective agars or for which phenotypic identification fails or is in doubt. Strong consideration should also be given to referring glucose non0196-4399/00 (see frontmatter)
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fermenting gram-negative rods exhibiting resistance to colistin (Table 2).
Table 2. Glucose non-fermenting gram-negative bacilli that may demonstrate colistin resistancea
Other Gucose Non-fermenting Gram-Negative Rods
Species Achromobacter xylosoxidans subsp. xylosoxidans Achromobacter piechaudii Acinetobacter calcoaceticus/baumannii Bergeyella zoohelcum Brevundimonas diminuta Brevundimonas vesicularis Burkholderia cepacia Burkholderia pseudomallei Chryseobacterium indologenes Chryseobacterium meningosepticum Delftia acidovorans Empedobacter brevis Myroides odoratus/odoratimimus Ochrobactrum anthropi/intermedium Ralstonia pickettii Ralstonia mannitolilytica Rhizobium radiobacter Sphingobacterium multivorum Sphingomonas paucimobilis Stenotrophomonas maltophilia
Glucose non-fermenting gram-negative rods other than P. aeruginosa and B. cepacia appear to be increasing in frequency, perhaps as a result of clinical and microbiological factors (3). The aging CF population may afford more opportunities for infection with infrequent and unusual organisms, particularly in the setting of chronic suppressive therapy and early-eradication protocols. These treatment regimens may be contributing to the decreasing incidence of P. aeruginosa and the selection of emerging pathogens, many of which are multi-drug resistant and can persist in the airways (3). The two most common isolates are S. maltophilia and A. xylosoxidans; however, the list of additional organisms continues to grow (1). The pathogenic roles of these organisms remain unclear, but they can be problematic from a laboratory perspective. Their identification is often hampered, because most commercial identification system databases do not contain some of the genera (Inquilinus, Herbaspirillum, and Pandoraea) or may only include a limited number of species (Cupriavidus, Ralstonia, and Bordetella); the Biolog (Biolog, Inc., Hayward, CA) and Sherlock MIDI (MIDI, Inc., Newark, DE) systems are exceptions. To further compound the problem, many of these organisms grow on BCSA and are misidentified as Bcc by some commercial identification systems. These issues should be taken into account in the laboratory’s B. cepacia protocol (see above). Editor’s note: Part II of this article will appear in the March 1, 2012 issue of CMN (vol. 34, No. 5). References 1. Bittar, F. and J.M. Rolain. 2010. Detection and accurate identification of new or emerging bacteria in cystic fibrosis patients. Clin. Microbiol. Infect. 16:809-820. 2. Gilligan, P.H. and D.L. Kiska. 2006. Cumitech 43, Cystic Fibrosis Microbiology. M.D. Appleman (Coordinating ed.). American Society for Microbiology, Washington, DC. 3. Lipuma, J.J. 2010. The changing microbial epidemiology in cystic fibrosis.
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a
% Resistance 31 14 2 100 100 100 100 100 100 100 100 100 100 7 100 100 47 100 81 62
Zone diameter ≤ 6 mm (10-μg colistin disk on Mueller-Hinton agar). Table adapted from reference 102. Not all species have been isolated from CF patients.
Clin. Microbiol. Rev. 23:299-323. 4. Emerson, J. et al. 2010. Changes in cystic fibrosis sputum microbiology in the United States between 1995 and 2008. Pediatr. Pulmonol. 45:363-370. 5. Razvi, S. et al. 2009. Respiratory microbiology of patients with cystic fibrosis in the United States, 1995 to 2005. Chest 136:1554-1560. 6. Foweraker, J.E. et al. 2005. Phenotypic variability of Pseudomonas aeruginosa in sputa from patients with acute infective exacerbation of cystic fibrosis and its impact on the validity of antimicrobial susceptibility testing. J. Antimicrob. Chemother. 55:921-927. 7. Balke, B. et al. 2008. A German external quality survey of diagnostic microbiology of respiratory tract infections in patients with cystic fibrosis. J. Cyst. Fibros. 7:7-14. 8. Young, C. et al. 2011. Use of COPAN SL solution for processing sputum from patients with and without cystic fibrosis. Abstr. C-2149. 111th Gen. Meet. Am. Soc. Microbiol. American Society for Microbiology, Washington, DC. 9. Laine, L. et al. 2008. A novel chromogenic medium for isolation of Pseudomonas aeruginosa from the sputa of cystic fibrosis patients. J. Cyst. Fibros. 8:143-149.
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10. Kidd, T.J. et al. 2009. Low rates of Pseudomonas aeruginosa misidentification in isolates from cystic fibrosis patients. J. Clin. Microbiol. 47:15031509. 11. Bittar, F. et al. 2006. Inquilinus limosus and cystic fibrosis. Emerg. Infect. Dis. 14:993-994. 12. Chiron, R. et al. 2005. Clinical and microbiological features of Inquilinus sp. isolates from five patients with cystic fibrosis. J. Clin. Microbiol. 43:39383943. 13. Cooke, R.P. et al. 2007. Inquilinus limosus isolated from a cystic fibrosis patient: first UK report. Br. J. Biomed. Sci. 64:127-129. 14. Coenye, T. et al. 2002. Characterization of unusual bacteria isolated from respiratory secretions of cystic fibrosis patients and description of Inquilinus limosus gen. nov., sp. nov. J. Clin. Microbiol. 40:2062-2069. 15. Degand, N. et al. 2008. Matrix-assisted laser desorption ionization-time of flight mass spectrometry for identification of nonfermenting gram-negative bacilli isolated from cystic fibrosis patients. J. Clin. Microbiol. 46:3361-3367. 16. Wellinghausen, N. et al. 2005. Superiority of molecular techniques for identification of gram-negative, oxidase-positive
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22. Lipuma, J.J. 2005. Update on the Burkholderia cepacia complex. Curr. Opin. Pulm. Med. 11:528-533.
30. Kennedy, M.P. et al. 2007. Burkholderia gladioli: five year experience in a cystic fibrosis and lung transplantation center. J. Cyst. Fibros. 6:267-273.
23. Turton, J.F. et al. 2007. Revised approach for identification of isolates within the Burkholderia cepacia complex and description of clinical isolates not assigned to any of the known genomovars. J.
31. Payne, G.W. et al. 2005. Development of a recA gene-based identification approach for the entire Burkholderia genus. Appl. Environ. Microbiol. 71:3917-3927.
32. Mahenthiralingam, E., A. Baldwin, and C.G. Dowson. 2008. Burkholderia cepacia complex bacteria: opportunistic pathogens with important natural biology. J. Appl. Microbiol. 104:1539-1551. 33. Coenye, T. et al. 2001. Taxonomy and identification of the Burkholderia cepacia complex. J. Clin. Microbiol. 39:3427-3436. 34. Krejcí, E. and R.M. Kroppenstedt. 2006. Differentiation of species combined into the Burkholderia cepacia complex and related taxa on the basis of their fatty acid patterns. J. Clin. Microbiol. 44:1159-1164. 35. de Vrankrijker, A.M., T.F. Wolfs, and C.K. van der Ent. 2010. Challenging and emerging pathogens in cystic fibrosis. Paediatr. Respir. Rev. 11:246-254. 36. Zbinden, A. et al. 2007. Evaluation of the colorimetric VITEK 2 card for identification of gram-negative nonfermentative rods: comparison to 16S rRNA gene sequencing. J. Clin. Microbiol. 45:2270-2273. 37. Vanlaere, E. et al. 2006. Growth in Stewart’s medium is a simple, rapid and inexpensive screening tool for the identification of Burkholderia cepacia complex. J. Cyst. Fibros. 5:137-139. 38. Vanlaere, E. et al. 2008. Matrix-assisted laser desorption ionisation-time-of offlight mass spectrometry of intact cells allows rapid identification of Burkholderia cepacia complex. J. Microbiol. Methods 75:279-286.
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17. Papaleo, M.C. et al. 2010. Identification of species of the Burkholderia cepacia complex by sequence analysis of the hisA gene. J. Med. Microbiol. 59:11631170. 18. Zlosnik, J.E. et al. 2011. Mucoid and nonmucoid Burkholderia cepacia complex bacteria in cystic fibrosis infections. Am. J. Respir. Crit. Care Med. 183:67-72. 19. Vonberg, R.P. et al. 2006. Identification of Burkholderia cepacia complex pathogens by rapid-cycle PCR with fluorescent hybridization probes. J. Med. Microbiol. 55:721-727. 20. Kalish, L.A. et al. 2006. Impact of Burkholderia dolosa on lung function and survival in cystic fibrosis. Am. J. Respir. Crit. Care Med. 173:421-425. 21. Davies, J.C. and B.K. Rubin. 2007. Emerging and unusual gram-negative infections in cystic fibrosis. Semin. Respir. Crit. Care Med. 28:312-321.
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24. Mahenthiralingam, E., T.A. Urban, and J.B. Goldberg. 2005. The multifarious, multireplicon Burkholderia cepacia complex. Nat. Rev. Microbiol. 3:144-156. 25. Govan, J.R., A.R. Brown, and A.M. Jones. 2007. Evolving epidemiology of Pseudomonas aeruginosa and the Burkholderia cepacia complex in cystic fibrosis lung infection. Future Microbiol. 2:153-164. 26. Foweraker, J. 2009. Recent advances in the microbiology of respiratory tract infection in cystic fibrosis. Br. Med. Bull. 89:93-110. 27. Asiah, K. et al. 2006. Unrecognised infection in a cystic fibrosis patient. J. Paediatr. Child Health 42:217-218. 28. Barth, A.L. et al. 2007. Cystic fibrosis patient with Burkholderia pseudomallei infection acquired in Brazil. J. Clin. Microbiol. 45:4077-4080. 29. Corral, D.M. et al. 2008. Burkholderia pseudomallei infection in a cystic fibrosis patient from the Caribbean: a case report. Can. Respir. J. 15:237-239.
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