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Biofilm formation on tracheostomy tubes

Ear, Nose & Throat Journal, Sept, 2002 by William A. Jarrett, Julie Ribes, Jose M. Manaligod
An increased awareness of biofilms and their mechanisms has led to a better understanding of bacterial infections that occur following the placement of tracheostomy tubes and other implanted devices and prostheses.

One aspect of biofilm formation that is still subject to debate is whether the specific material that is used to manufacture a tube has any bearing in the incidence of infection. We conducted a test of four different tube materials--polyvinyl chloride, silicone, stainless steel, and sterling silver--to ascertain how bacterial biofilms form on tracheostomy tubes and to determine if there is a material-dependent difference in biofilm formation. Scanning electron microscopy demonstrated that Pseudomonas aeruginosa and Staphylococcus epidermidis both formed bacterial biofilms on tracheostomy tubes in vitro. We also found that there was no difference in susceptibility to biofilm formation among the four tube materials tested.

Introduction

The practice of surgically placing prostheses and other devices is widespread and beneficial, although these products are susceptible to harboring bacterial infection. Chronic infections that are resistant to antibiotic therapy can ultimately necessitate the removal and replacement of a device or implant, including tracheostomy tubes, which are associated with nosocomial respiratory infections.

In the past few years, our increasing understanding of biofilms has emerged as a key factor in treating and preventing antibiotic-resistant implant-associated infections. Biofilms are organized matrices of bacterial colonies. A significant proportion of the respiratory infections related to chronic intubation and tracheostomy are caused by Pseudomonas aeruginosa and Staphylococcus epidermidis. (1) Both of these pathogens form polysaccnrinde matrices, which facilitate biofilm formation.

The goal of this study was to determine whether bacterial biofilms form on tracheostomy tubes in vitro and to ascertain if there is a material-dependent difference in bacterial biofilm formation.

Materials and methods

We followed a protocol established by Biedlingmaier et al to prepare bacteria to a concentration of [10.sup.6] colony-forming units (CFU) per ml (the standard for biofilm formation in previous studies). (2) Standardized laboratory strains of P. aeruginosa and S. epidermidis were inoculated in trypticase soy broth (TSB), a nonselective, enriched, liquid growth medium. Each bacterial strain was grown to a logarithmic phase over 24 hours at 37[degrees] C. The bacteria were harvested by centrifugation at 2,000 rpm at 25[degrees] C for 15 minutes. Three cycles of centrifugation were performed to assure thorough bacterial isolation. Bacterial isolates were redispersed in 20 ml of TSB in sterile polystyrene tubes and serially diluted to a concentration of [10.sup.6] CFU/ml based on McFarland turbidity.

Four types of sterile tracheostomy tube were used in this study: polyvinyl chloride (Mallinckrodt; St. Louis), silicone (BivonaMedical Technologies; Gary, Ind.), stainless steel (Pilling Weck; Markham, Ont.), sterling silver (Pilling Weck). A 1-cm section was taken at the same location (2 cm from the tip) from each tube because the size and shape of an entire tube made its complete immersion in culture tubes impossible.

For negative controls, samples of each tracheostomy tube were placed in TSB solution without bacteria. The experimental arm consisted of samples of each tracheostomy tube in separate solutions of P. aeruginosa alone and S. epidermidis alone. All culture tubes were maintained at 37[degrees] C for 6 days to promote biofilm formation.

The samples were removed and rinsed 10 times with 35 ml of sterile water to remove free bacteria and debris that were not attached to the samples. To ensure the presence of viable bacteria in experimental culture tubes and to ensure that no bacterial contamination was present in the TSB control tubes, a sterile loop was dipped into the TSB control tubes and into the bacteria-containing experimental tubes and streaked over sheep-blood agar medium. No growth was seen from the negative controls, while the experimental sample cultures contained numerous bacterial colonies within 48 hours.

In preparation for scanning electron microscopy (SEM), each sample was fixed in a formaldehyde and glutaraldehyde solution. Samples were washed in phosphatebuffered saline solution, and then a second fixative, thiocarbohydrazide and osmium tetroxide, was applied to preserve lipids and large molecules. The samples were silver-coated and dried to complete the preparation for SEM. A scanning electron microscope (model S450; Hitachi; Tokyo) was used to analyze each sample of both the control and bacteria-exposed tubes for the presence of bacterial biofilm formation.

Results

All of the control tubes yielded no evidence of biofilm formation on gross inspection and SEM examination. All tubes exposed to P. aeruginosa and S. epidermidis were coated with a brown, slimy, translucent film on gross examination, and all had nearly identical biofilm coverage on SEM (figure). The P. aeruginosa biofilms appeared as confluent sheets of cells within a dense extracellular matrix. The S. epidermidis biofilms appeared as mounds of tightly adherent cells within a relatively less-extracellular matrix.

Discussion

In contrast to early theories that bacteria are primitive free-floating organisms, we now know that bacterial populations typically exist as sessile colonies attached to surfaces. These organized communities, known as biofilms, exist as a result of a complex network of molecular and cellular interactions that have only recently been discovered. Much of the work on bacterial biofilms has been focused on P. aeruginosa, which readily forms biofilms under almost any culture condition or environment. As seen in studies of P. aeruginosa, free-swimming bacteria use flagella and pili to sense surfaces and initiate contact. (3) Surface contact induces critical changes in gene expression. One important gene activated by surface adhesion is algC, which encodes exopolysaccharide alginate, the key component of P. aeruginosa extracellular matrices. (4) This attachment and the exopolysaccharide alginate formation result in a secondary cascade of gene up-regulation that governs antibiotic resistance, the exchange of genetic information, and other changes that contribute to the maturation of the biofilm community.

S. epidermidis forms biofilms in a similar manner--one in which cell-surface interactions play a key role. Capsular polysaccharide/adhesin and cell-surface-localized autolysin (atlE) are two of the key proteins that enable the surface attachment of S. epidermidis. (5,6) Polysaccharide/adhesin is also a major component of a polysaccharide matrix that induces the accumulative phase of S. epidermidis biofilm formation, which involves an increase in the cell-cell exchange of genetic information and an up-regulation of antibiotic resistance. In addition to the cellular cooperation that is possible in a polysaccharide matrix, another interesting feature of biofilms is their surprisingly complex architecture. Optical sectioning has demonstrated the presence of water channels in biofilms as well as a significant structural heterogeneity. (7)

With the greater use of mechanical ventilation in the care of critically ill patients, nosocomial respiratory infections have become more prevalent. In premature infants, purulent tracheal aspirations increase with prolonged intubation. (8) A similar pattern of nosocomial respiratory infections in association with prolonged intubation during mechanical ventilation is also seen in the adult population. (9) An important factor in such infections is the endotracheal tube itself. Inglis et al reported that biofilms have been identified in endotracheal tubes of patients with ventilator-associated pneumonia. (10) In a separate report, Inglis et al also demonstrated that fragments of endotracheal tube biofilms are propelled toward the tracheobronchial tree by positive pressure ventilation. (11)

Compared with endotracheal intubation, tracheostomy is associated with a lesser degree of respiratory bacterial colonization and therefore results in faster recovery from ventilator-associated pneumonia. (12) Although critically ill patients with respiratory failure who undergo tracheostomy have generally favorable outcomes as a result of the procedure, (13) a significant association with nosocomial infections still remains. (9) Of the bacteria associated with tracheostomy and endotracheal infections, P. aeruginosa and S. epidermidis are two of the most commonly identified pathogens. (1,14)

Our study found that P. aeruginosa and S. epidermidis both formed bacterial biofilms on tracheostomy tube sections in vitro. Among the four different materials tested-polyvinyl chloride, silicone, stainless steel, and sterling silver--no difference was noted in the susceptibility to biofilm formation. Although no difference was noted among these materials, the selection of tracheostomy tube material has historically not been focused on infection resistance. Different materials have been chosen because of their respective merits with regard to flexibility, ease of use, and skin reaction.

Previous work by Berry et al on tympanostomy tubes demonstrated that phosphorylcholine-coated fluoroplastic tubes were more resistant to biofilm formation than were silicone tubes and silver-oxide-treated silicone tubes. (15) Materials that distribute antibiotics in a controlled-release fashion are also more resistant to biofilm formation. (16,17) These findings might lead to the development of a tracheostomy tube that is resistant to biofilm formation.

In vivo studies of patients with tracheostomy tubes might shed further light on biofilm development. Future work on biofilms and tracheostomy tubes is also being planned to study the effect of other species of bacteria--Staphylococcus aureus and Escherichia coli, among others--that are common in critically ill patients. The study of multiorganism biofilms would also be fruitful, because involvement of more than one organism represents a more accurate clinical picture of most infections.

References

(1.) Stratton CW. Topics in clinical microbiology: Tracheostomy and endotracheal tube specimens. Infect Control 1981;2:162-4.

(2.) Biedlingmaier JF, Samaranayake R, Whelan P. Resistance to biofilm formation on otologic implant materials. Otolaryngol Head Neck Surg 1998;l18:444-51.

(3.) O'Toole GA, Kolter R. Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol Microbiol 1998;30:295-304.

(4.) Davies DG, Chakrabarty AM, Geesey GG. Exopolysaceharide production in biofilms: Substratum activation of alginate gene expression by Pseudomonas aeruginosa. Appl Environ Microbiol 1993;59:1181-6.

(5.) McKenney D, Hubner J, Muller E, et al. The ica locus of Staphylococcus epidermidis encodes production of the capsular polysaceharide/adhesin. Infect Immun 1998;66:4711-20.

(6.) Heilmann C, Hussain M, Peters G, Gotz F. Evidence for autolysin-mediated primary attachment of Staphylococcus epidermidis to a polystyrene surface. Mol Microbiol 1997;24:1013-24.

(7.) Lawrence JR, Korber DR, Hoyle BD, et al. Optical sectioning of microbial biofilms, J Bacteriol 1991;173:6558-67.

(8.) Cordero L, Sananes M, Dedhiya P. Ayers LW. Purulence and gram-negative bacilli in tracheal aspirates of mechanically ventilated very low birth weight infants. J Perinatol 2001;21:376-81.

(9.) Tejada Artigas A, Bello Dronda S, Chacon Valles E, et al. Risk factors for nosocomial pneumonia in critically ill trauma patients. Crit Care Med 2001;29:304-9.

(10.) Inglis TJ, Lim TM, Ng ML, et al. Structural features of tracheal tube biofilm formed during prolonged mechanical ventilation. Chest 1995;108:1049-52.

(11.) Inglis TJ, Jones JG. Paxton S. Penetration of an aerosol, produced by film atomization, through the carinal bifurcation. Br J Anaesth 1993;70:527-31.

(12.) Curtis JJ, Clark NC, McKenney CA, et al. Tracheostomy: A risk factor for mediastinitis after cardiac operation. Ann Thorac Surg 2001;72:731-4.

(13.) Kollef MH, Ahrens TS, Shannon W. Clinical predictors and outcomes for patients requiring tracheostomy in the intensive care unit, Crit Care Med 1999;27:1714-20.

(14.) Teoh N, Parr MJ, Finfer SR. Bacteraemia following percutaneous dilational tracheostomy. Anaesth Intensive Care 1997;25:354-7.

(15.) Berry JA, Biedlingmaier JF, Whelan PJ. In vitro resistance to bacterial biofilm formation on coated fluoroplastic tympanostomy tubes. Otolaryngol Head Neck Surg 2000;123:246-51.

(16.) Hendricks SK, Kwok C, Shen M, et al. Plasma-deposited membranes for controlled release of antibiotic to prevent bacterial adhesion and biofilm formation. J Biomed Mater Res 2000;50:160-70.

(17.) Owusu-Ababio G, Rogers JA, Morck DW, Olson ME. Efficacy of sustained release ciprofloxacin microspheres against device-associated Pseudomonas aeruginosa biofilm infection in a rabbit peritoneal model. J Med Microbiol 1995;43:368-76.

From the Department of Otolaryngology--Head and Neck Surgery, University of Iowa Hospital and Clinics, Iowa City.

Reprint requests: Jose M. Manaligod, MD, Assistant Professor, Department of Otolaryngology--Head and Neck Surgery, University of Iowa Hospital and Clinics, 200 Hawkins Dr., Iowa City, IA 52242. Phone: (319) 353-5837; fax: (319) 356-4547; e-mail: This email address is being protected from spambots. You need JavaScript enabled to view it.

 

 

 

 

 

 

 

 

 

 

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