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Effects of tea tree (Melaleuca alternifolia) oil on Staphylococcus aureus in biofilms and stationary growth phase
International Journal of Antimicrobial Agents 33 (2009) 343–347
Contents lists available at ScienceDirect
International Journal of Antimicrobial Agents journal homepage: http://www.elsevier.com/locate/ijantimicag
Effects of tea tree (Melaleuca alternifolia) oil on Staphylococcus aureus in biofilms and stationary growth phase a,∗ ´ Jakub Kwiecinski , Sigrun Eick b , Kinga Wójcik a a Department of Microbiology, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, Gronostajowa 7, 30-387 Kraków, Poland b Institute of Medical Microbiology, University Hospital of Jena, Erlanger Allee 101, 07747 Jena, Germany
a r t i c l e
i n f o
Article history: Received 23 July 2008 Accepted 22 August 2008 Keywords: Tea tree oil Staphylococcus aureus Stationary growth phase Biofilm
a b s t r a c t Tea tree oil (TTO) is known for its antimicrobial activity. In this study, we determined whether TTO is effective against Staphylococcus aureus in biofilms and how TTO activity is affected by the S. aureus growth phase. All clinical strains tested were killed by TTO both as planktonic cells and as biofilms. The minimum biofilm eradication concentration was usually two times higher than the minimum bactericidal concentration, yet it was never higher than 1% v/v. The fastest killing of biofilm occurred during the first 15 min of contact with TTO and was not influenced by increasing TTO concentration above 1% v/v. Planktonic stationary phase cells exhibited decreased susceptibility to TTO compared with exponential phase cells. The killing rate for stationary phase cells was also less affected by increasing TTO concentration than that for exponential phase cells. These data show that TTO efficiently kills S. aureus in the stationary growth phase and within biofilms and is therefore a promising tool for S. aureus eradication. © 2008 Elsevier B.V. and the International Society of Chemotherapy. All rights reserved.
1. Introduction Tea tree oil (TTO), an essential oil obtained from Melaleuca alternifolia, has gained the attention of scientists, physicians and consumers because of its broad antimicrobial and antiinflammatory activities. TTO consists of ca. 100 different compounds, including terpinen-4-ol, which is one of the main antibacterial components [1]. Drugs and care products containing TTO are frequently used for treatment of skin, oral, vaginal and airway infections, or as antiseptics and disinfectants [1,2]. Furthermore, there have been reports of efficient eradication of bacteria in deep infections such as abscesses or osteomyelitis [1,3]. Most TTOrelated research is focused on Staphylococcus aureus. Preliminary clinical trials have shown that the efficacy of TTO in eradication of meticillin-resistant S. aureus (MRSA) carriage is comparable with traditional mupirocin treatment [1]. During infections, bacteria are usually present in various growth phases. In chronic infections or when access to nutrition is limited, bacteria reside predominantly in a stationary or dormant phase [4]. Following brief exposure to antibiotics, continuous suppression of
∗ Corresponding author. Present address: Department of Rheumatology and Inflammation Research, University of Göteborg, Guldhedsgatan 10, S-413 46 Göteborg, Sweden. Tel.: +46 31 342 6475; fax: +46 31 823 925. ´ E-mail address: [email protected] (J. Kwiecinski).
microbial growth is also observed [5]. Non-multiplying or slowly growing organisms are generally less susceptible to antibacterial agents [4]. S. aureus is able to form biofilms, which are complex structures consisting of surface-attached bacteria surrounded by a self-produced extracellular polymer matrix. Bacteria in biofilms exhibit elevated resistance both to antibiotics and the host defence systems, which often results in persistent and difficult-to-treat infections [6–8]. S. aureus biofilm formation contributes to its pathogenesis in a number of conditions, such as damaged skin infections, chronic and recurrent airway infections, osteomyelitis and mastitis [6–11], and to its colonisation on the surface of undamaged skin [12]. Additionally, biofilms of staphylococci on surfaces in the food industry pose a serious risk of food contamination [13]. Therefore, for rational treatment of numerous diseases and development of antiseptics and disinfectants, it is important to determine the influence of TTO on bacteria in stationary growth phase and on already established pre-formed biofilms. It was shown that 1 h treatment with 5% TTO eradicates biofilms of S. aureus [12]. However, it is not presently known at what concentrations TTO is effective against biofilms, what the kinetics of TTO biofilm eradication are, how TTO affects the viability of S. aureus biofilms in comparison with planktonic bacteria, and whether the response to TTO is growth phase-dependent. The aim of this study was to answer these questions, which are crucial for the design of effective and rational therapy.
0924-8579/$ – see front matter © 2008 Elsevier B.V. and the International Society of Chemotherapy. All rights reserved. doi:10.1016/j.ijantimicag.2008.08.028
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J. Kwieci´ nski et al. / International Journal of Antimicrobial Agents 33 (2009) 343–347
2. Materials and methods 2.1. Tea tree oil TTO was obtained from Avicenna-Oil (Wrocław, Poland) and complied with the ISO 4730 and European Pharmacopoeia standards. 2.2. Bacterial strains Twenty-seven clinical isolates of S. aureus as well as the reference strain S. aureus NCTC 8325-4 were used in this study. All clinical strains were isolated at the Institute of Medical Microbiology, University Hospital of Jena, Jena, Germany. The strains were isolated from the following locations: upper respiratory tract (n = 5); lower respiratory tract (n = 6); superficial wound infections (n = 9); deep wound infections, e.g. bone (n = 5); and urinary tract infections (n = 2). Two-thirds of them were penicillin-resistant and additionally two strains were MRSA. 2.3. Determination of the minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) MICs and MBCs were determined using the broth microdilution method. Overnight cultures (37 ◦ C, shaking) of S. aureus in tryptic soy broth (TSB) (Fluka, Buchs, Switzerland) were diluted 10-fold in fresh TSB and incubated (37 ◦ C, shaking) until they reached exponential growth phase. Serial two-fold dilutions of TTO (from 2% to 0.125% v/v) in TSB were prepared in a 96-well plate (200 L per well). Wells with no TTO added were used as a positive growth control. All wells contained Tween 20 at a final concentration of 0.1% v/v to increase TTO solubility. A diluted bacterial suspension was added to each well to give a final concentration of 1–5 × 105 colony-forming units (CFU)/mL, confirmed by viable counts. Wells without bacteria added were used as a negative growth control. The plate was incubated for 24 h at 37 ◦ C and growth was assessed turbidometrically. From all wells not showing visible growth (optical density at 600 nm (OD600 ) ≤0.05), 10 L was plated on tryptic soy agar (TSA) (Fluka) and the number of colonies was counted following overnight incubation at 37 ◦ C. The MIC was defined as the lowest TTO concentration without visible growth, and the MBC was defined as the lowest concentration reducing the initial inoculum by ≥99.9%. The MIC and MBC were determined for all 27 clinical strains and S. aureus 8325-4. For each strain, at least three independent determinations were done and the modal value was taken.
After 24 h of biofilm formation, the medium was gently aspirated from the plate and the wells were rinsed three times with phosphate-buffered saline (PBS). Serial two-fold dilutions of TTO (from 2% to 0.125% v/v) in 200 L of TSB were then added to wells. No TTO was added to the positive biofilm control wells. All wells contained Tween 20 at a final concentration of 0.1% v/v. The plate was incubated at 37 ◦ C for 24 h, thereafter the medium was gently aspirated and wells were rinsed three times with PBS. Biofilm viability was assessed with either the 3-(4,5-dimethylthiazol-2yl)-2,5-diphenyltetrazolium bromide (MTT) staining method (13 clinical strains forming the best biofilms in a preliminary experiment and S. aureus 8325-4) or the CFU counting method (S. aureus 8325-4 only). Viability measures were done in duplicate in three independent experiments. MTT staining was done according to Walencka et al. [15], with minor modifications. Briefly, 200 L of 0.05% MTT (Sigma, Steinheim, Germany) in PBS was added to the wells with biofilm. Following incubation at 37 ◦ C for 2 h, MTT solution was replaced by 200 L of 5 mM HCl in isopropanol, all formed formazan crystals were dissolved and the absorbance was measured at 550 nm. The minimum biofilm eradication concentration (MBEC) was defined as the lowest TTO concentration resulting in a decrease in absorbance to <5% of the positive control (cut-off level for negative control, determined in the preliminary experiment). The biofilm CFU counting method was adapted from Pettit et al. [16]. The level of detection was 102 CFU/mL. In addition, scanning electron microscopy (SEM) images were taken. A 24-well tissue culture plate was filled with coverslips. S. aureus 8325-4 biofilms were prepared as described previously. After an incubation time of 24 h, the slips were carefully washed with PBS, then TTO solutions were added and the slips were incubated again for 24 h. After removal of the media and washing, the samples were initially fixed in 2% glutaraldehyde in cacodylate buffer for 30 min, then washed twice with cacodylate buffer and dehydrated for 10 min using a graded ethanol series. A critical point drying procedure followed and the specimens were then sputter-coated with gold. Samples were examined with a LEO 1450 VP scanning electron microscope (Carl Zeiss, Germany). 2.6. Biofilm time-dependent killing assay A biofilm of S. aureus 8325-4 was established as described previously. TTO solutions (0, 0.5, 1, 2, 4 and 8% v/v) in 200 L of TSB with 0.1% v/v Tween 20 were added to the wells. The plates were incubated at 37 ◦ C for 0, 15, 30, 60 and 120 min. The medium was then aspirated, the wells were washed three times with PBS and biofilm
2.4. Planktonic time-dependent killing assay S. aureus 8325-4 suspensions from stationary (overnight culture in TSB, 37 ◦ C, shaking) and exponential (prepared as for MIC determination) growth phases were prepared. In plastic tubes, 1 mL of each was mixed with 100 L of TTO solution to final concentrations of 0% (control), 0.5% or 8% v/v and with Tween 20 (0.1% v/v final concentration in all tubes). Tubes were incubated at 37 ◦ C with shaking. After 15, 30 and 60 min, 10 L was removed from each tube and the number of CFU/mL was counted by plating serial dilutions on TSA. A sample from the 0% TTO tube taken immediately after mixing was used as a ‘time 0’ control. The detection level was 103 CFU/mL and measurements were made in three independent experiments. 2.5. Effects of TTO on established biofilms The method of biofilm formation in 96-well cell culture plates ´ according to Kuzma et al. was used [14].
Fig. 1. Cumulative minimum inhibitory concentration (MIC), minimum bactericidal concentration (MBC) and minimum biofilm eradication concentration (MBEC) of tea tree oil (TTO) against clinical isolates of Staphylococcus aureus, expressed as a percentage of tested strains. MIC and MBC, n = 27; MBEC, n = 13.
J. Kwieci´ nski et al. / International Journal of Antimicrobial Agents 33 (2009) 343–347
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Fig. 2. Effect of different concentrations of tea tree oil (TTO) on the viability of biofilm formed by Staphylococcus aureus 8325-4 following a 24 h exposure. Viability was determined by (a) 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) staining or (b) colony-forming unit (CFU) counting. Mean ± standard error of the mean for three independent experiments performed in duplicate. The limit of detection for CFU counting was 102 CFU/mL.
Fig. 3. Scanning electron microscopy images of Staphylococcus aureus 8325-4 biofilm: (a) control; and (b) after 24 h exposure to 1% tea tree oil.
viability was assessed with the MTT staining method. All measurements were made in duplicate in three independent experiments. 3. Results 3.1. TTO has a bactericidal effect and disrupts biofilms For clinical isolates, the MICs of TTO were in the range 0.125–0.5% (MIC for 50% of the organisms (MIC50 ) = 0.25%; MIC for 90% of the organisms (MIC90 ) = 0.5%). The highest MBC was 1%
(range 0.25–1%; MBC50 = 0.25%; MBC90 = 0.5%). Although the highest MBEC was also 1%, these values were in general one step higher than the MBCs (range 0.5–1%; MBEC50 = 0.5%; MBEC90 = 1%). These results are summarised in Fig. 1. For S. aureus 8325-4, the MIC was equivalent to the MBC (0.5%). The effects of TTO on S. aureus 8325-4 biofilm viability measured by MTT staining and CFU counting are shown in Fig. 2. Both methods exhibit good correlation and showed that 1% TTO effectively eradicated the biofilm. SEM images underline the results, as 1% TTO destroyed the biofilm formed by S. aureus 8325-4 (Fig. 3). The susceptibility of MRSA strains was comparable with other strains. A characteristic pattern of increased biofilm metabolism (assessed by MTT staining) at sub-MBEC concentrations was observed for all strains tested (data not shown).
3.2. Activity of TTO on S. aureus is growth phase-dependent
Fig. 4. Time–kill curves of Staphylococcus aureus 8325-4 from stationary and exponential growth phase after treatment with none, 0.5% or 8% tea tree oil (TTO). Each symbol indicates the mean ± standard error of the mean for three independent experiments. The limit of detection was 103 colony-forming units (CFU)/mL.
The number of viable S. aureus in exponential growth phase was reduced by two log10 steps (99% killed) with 0.5% TTO after 60 min, whilst addition of 8% TTO resulted in reduction by four log10 steps (99.99% killed) (Fig. 4). The rate of killing decreased in a time-dependent manner, with the fastest killing during the first 15 min. S. aureus cells in stationary growth phase were reduced by one log10 step (90% killed) after addition of both 0.5% and 8% TTO for 15 min. Later, only slight killing was observed.
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Fig. 5. Time–kill curves for Staphylococcus aureus 8325-4 biofilm after treatment with 0.5%, 1% or 8% tea tree oil. Each symbol indicates the mean ± standard error of the mean for three independent experiments performed in duplicate.
3.3. TTO rapidly kills bacteria in biofilm An exponential decrease of biofilm metabolism was seen after exposure to TTO (Fig. 5). After 120 min, biofilm metabolic activity was only 40% of control activity for biofilm exposed to 0.5% TTO, and 15% of control activity for 1% TTO exposure. Increasing TTO concentration up to 8% did not have any additional effect on the kinetics of biofilm killing (data for 2% and 4% TTO not shown). The fastest killing occurred during the first 15 min and the rate of killing decreased in a time-dependent manner. Most of the killing was completed within 30–60 min. 4. Discussion These data show that TTO efficiently kills S. aureus both in suspension and in biofilms. The obtained MIC and MBC values confirm the high susceptibility of clinical strains to TTO and are in the range reported in previous studies (0.12–2% for MICs and 0.25–8% for MBCs [12,17–19], although a much lower MIC has also been described [20]). MTT staining is one of the methods used for assessing the viability of bacterial biofilms based on reduction of tetrazolium salts to formazan by viable metabolising bacteria [14–16]. Using this method, our study shows that 1% TTO was able to inhibit the metabolism of biofilms formed by all tested strains, which is only twice the bactericidal dose. Most antibiotics are up to 1000-times less efficient against bacteria in biofilm than in suspension [7], which makes TTO a very promising treatment alternative. SEM images show disruption of biofilm by 1% TTO. It is therefore likely that biofilm eradication by TTO is not only due to bacteria killing but also partly due to extracellular matrix damage and subsequent removal of biofilm from the surface, especially by higher TTO concentrations. Low TTO concentrations (between 0.125% and 0.25%) had no effect on the number of CFU/mL within S. aureus 8325-4 biofilms, but MTT staining showed increased biofilm metabolic activity. This suggests that the activity of the cells within the biofilm increases in response even to these low concentrations of TTO. This might reflect a stress reaction, indicating that the TTO tolerance in S. aureus is an energy-dependent process. An energy-dependent TTO tolerance in Pseudomonas aeruginosa was previously reported [21]. For the potential clinical use of TTO it is important to establish the kinetics of its action against bacteria within biofilms. The rate of killing was concentration-dependent up to 1% TTO, but further increase did not accelerate killing. After 2 h, metabolic activity of the biofilm was still observed even after treatment with 8% TTO. This
finding is in contrast to previous data suggesting complete biofilm killing after 1 h in 5% TTO [12]. This might be partly explained by the fact that in our study different methods of biofilm formation and viability testing were used. The MTT method slightly overestimates biofilm viability, as many already lethally injured cells might still exhibit metabolism. Conversely, methods used in the previous study might underestimate bacterial viability owing to additional damage caused by sonication used for dislodging bacteria [12]. Nevertheless, after 1 h, 1% TTO causes a large (ca. 85%) reduction of S. aureus biofilm viability. Escherichia coli in stationary growth phase were killed by TTO ca. 105 times slower than cells from the exponential phase [22]. Our results show that S. aureus in the stationary phase is also more tolerant to TTO compared with bacteria in the exponential phase. When entering the stationary growth phase, cell membrane protein composition [23], cell membrane fluidity [24] and the surface charge and hydrophobicity [25] of S. aureus change. These alterations might account for the changed TTO tolerance, as the cell membrane is the main target of TTO antistaphylococcal activity [26,27]. Both in biofilm and in the stationary phase, increasing TTO concentration has only limited effects on the killing rate. This might be connected to so-called ‘persister cells’ present in biofilms and stationary phase cultures [28]. These cells, which exhibit tolerance to numerous antimicrobial agents, might be a key to increased resistance in biofilms. Further studies are required to elucidate their possible role in TTO biofilm and stationary phase tolerance. According to previous studies, rapid development of TTO resistance is rather unlikely [20,29]. Therefore, TTO is a promising alternative for S. aureus eradication, both as a disinfectant and as infection treatment. It acts on stationary phase and biofilm bacteria – two states frequently encountered in the environment and in living organisms – and it kills the bacteria quickly and effectively at a concentration of 1%, which is below concentrations used in commercially available preparations (up to 10%). TTO may be a new, cheap and effective weapon for fighting S. aureus infections, including MRSA. Acknowledgments The authors acknowledge Susanne Linde (Department of Ultrastructural Research, University Hospital of Jena, Jena, Germany) for excellent technical assistance in preparing SEM photographs, Inger Gjertsson (Department of Rheumatology and Inflammation Research, Sahlgrenska Academy at Göteborg University, Sweden) for critical reading of the manuscript, and Ewa Grela for assistance in preparation of the figures. Funding: No funding sources. Competing interests: None declared. Ethical approval: Not required. References [1] Carson CF, Hammer KA, Riley TV. Melaleuca alternifolia (tea tree) oil: a review of antimicrobial and other medicinal properties. Clin Microbiol Rev 2006;19:50–62. [2] Price S, Price L. Aromatherapy for health professionals. Edinburgh, UK: Churchill Livingstone; 1999. [3] Kawakami E, Washizu M, Hirano T, Sakuma M, Takano M, Hori T, et al. Treatment of prostatic abscesses by aspiration of the purulent matter and injection of tea tree oil into the cavities in dogs. J Vet Med Sci 2006;68:1215–7. [4] Eng RH, Padberg FT, Smith SM, Tan EN, Cherubin CE. Bactericidal effects of antibiotics on slowly growing and nongrowing bacteria. Antimicrob Agents Chemother 1991;35:1824–8. [5] Fuursted K. Postantibiotic effect in vitro. APMIS Suppl 1999;90:1–23. [6] Kania RE, Lamers GE, Vonk MJ, Dorpmans E, Struik J, Tran Ba Huy P, et al. Characterization of mucosal biofilms on human adenoid tissues. Laryngoscope 2008;118:128–34. [7] Melchior MB, Vaarkamp H, Fink-Gremmels J. Biofilms: a role in recurrent mastitis infections? Vet J 2006;171:398–407.
J. Kwieci´ nski et al. / International Journal of Antimicrobial Agents 33 (2009) 343–347 [8] Vlastarakos PV, Nikolopoulos TP, Maragoudakis P, Tzagaroulakis A, Ferekidis E. Biofilms in ear, nose, and throat infections: how important are they? Laryngoscope 2007;117:668–73. [9] Akiyama H, Huh WK, Yamasaki O, Oono T, Iwatsuki K. Confocal laser scanning microscopic observation of glycocalyx production by Staphylococcus aureus in mouse skin: does S. aureus generally produce a biofilm on damaged skin? Br J Dermatol 2002;147:879–85. [10] Akiyama H, Hamada T, Huh WK, Yamasaki O, Oono T, Fujimoto W, et al. Confocal laser scanning microscopic observation of glycocalyx production by Staphylococcus aureus in skin lesions of bullous impetigo, atopic dermatitis and pemphigus foliaceus. Br J Dermatol 2003;148:526–32. [11] Psaltis AJ, Ha KR, Beule AG, Tan LW, Wormald PJ. Confocal scanning laser microscopy evidence of biofilms in patients with chronic rhinosinusitis. Laryngoscope 2007;117:1302–6. [12] Brady A, Loughlin R, Gilpin D, Kearney P, Tunney M. In vitro activity of tea-tree oil against clinical skin isolates of meticillin-resistant and -sensitive Staphylococcus aureus and coagulase-negative staphylococci growing planktonically and as biofilms. J Med Microbiol 2006;55:1375–80. [13] Gibson H, Taylor JH, Hall KE, Holah JT. Effectiveness of cleaning techniques used in the food industry in terms of the removal of bacterial biofilms. J Appl Microbiol 1999;87:41–8. ´ ˙ ˙ ´ [14] Kuzma Ł, Rózalski M, Walencka E, Rózalska B, Wysokinska H. Antimicrobial activity of diterpenoids from hairy roots of Salvia sclarea L.: salvipisone as a potential anti-biofilm agent active against antibiotic resistant Staphylococci. Phytomedicine 2007;14:31–5. ˙ ˙ [15] Walencka E, Sadowska B, Rózalska S, Hryniewicz W, Rózalska B. Lysostaphin as a potential therapeutic agent for staphylococcal biofilm eradication. Pol J Microbiol 2005;54:191–200. [16] Pettit RK, Weber CA, Kean MJ, Hoffmann H, Pettit GR, Tan R, et al. Microplate Alamar blue assay for Staphylococcus epidermidis biofilm susceptibility testing. Antimicrob Agents Chemother 2005;49:2612–7. [17] Carson CF, Cookson BD, Farrelly HD, Riley TV. Susceptibility of methicillinresistant Staphylococcus aureus to the essential oil of Melaleuca alternifolia. J Antimicrob Chemother 1995;35:421–4. [18] Elsom GKF, Hide D. Susceptibility of methicillin-resistant Staphylococcus aureus to tea tree oil and mupirocin. J Antimicrob Chemother 1999;43:427–8.
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[19] Hammer KA, Carson CF, Riley TV. Susceptibility of transient and commensal skin flora to the essential oil of Melaleuca alternifolia (tea tree oil). Am J Infect Control 1996;24:186–9. [20] Ferrini AM, Mannoni V, Aureli P, Salvatore G, Piccirilli E, Ceddia T, et al. Melaleuca alternifolia essential oil possesses potent anti-staphylococcal activity extended to strains resistant to antibiotics. Int J Immunopathol Pharmacol 2006;19:539–44. [21] Longbottom CJ, Carson CF, Hammer KA, Mee BJ, Riley TV. Tolerance of Pseudomonas aeruginosa to Melaleuca alternifolia (tea tree) oil is associated with the outer membrane and energy-dependent cellular processes. J Antimicrob Chemother 2004;54:386–92. [22] Gustafson JE, Liew YC, Chew S, Markham J, Bell HC, Wyllie SG, et al. Effects of tea tree oil on Escherichia coli. Lett Appl Microbiol 1998;26:194–8. [23] Cullmann W, Schlunegger H. Influence of exogenous factors and antibiotics on the cytoplasmic membrane proteins of Staphylococcus aureus. Chemotherapy 1992;38:211–7. [24] Xiong Z, Ge S, Chamberlain NR, Kapral FA. Growth cycle-induced changes in sensitivity of Staphylococcus aureus to bactericidal lipids from abscesses. J Med Microbiol 1993;39:58–63. [25] Beck G, Puchelle E, Plotkowski C, Peslin R. Effect of growth on surface charge and hydrophobicity of Staphylococcus aureus. Ann Inst Pasteur Microbiol 1988;139:655–64. [26] Cox SD, Mann CM, Markham JL, Bell HC, Gustafson JE, Warmington JR, et al. The mode of antimicrobial action of the essential oil of Melaleuca alternifolia (tea tree oil). J Appl Microbiol 2000;88:170–5. [27] Carson CF, Mee BJ, Riley TV. Mechanism of action of Melaleuca alternifolia (tea tree) oil on Staphylococcus aureus determined by time–kill, lysis, leakage, and salt tolerance assays and electron microscopy. Antimicrobial Agents Chemother 2002;46:1914–20. [28] Lewis K. Persister cells, dormancy and infectious disease. Nat Rev Microbiol 2007;5:48–56. [29] Hammer KA, Carson CF, Riley TV. Frequencies of resistance to Melaleuca alternifolia (tea tree) oil and rifampicin in Staphylococcus aureus, Staphylococcus epidermidis and Enterococcus faecalis. Int J Antimicrob Agents 2008;32: 170–3.
Contents lists available at ScienceDirect
International Journal of Antimicrobial Agents journal homepage: http://www.elsevier.com/locate/ijantimicag
Effects of tea tree (Melaleuca alternifolia) oil on Staphylococcus aureus in biofilms and stationary growth phase a,∗ ´ Jakub Kwiecinski , Sigrun Eick b , Kinga Wójcik a a Department of Microbiology, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, Gronostajowa 7, 30-387 Kraków, Poland b Institute of Medical Microbiology, University Hospital of Jena, Erlanger Allee 101, 07747 Jena, Germany
a r t i c l e
i n f o
Article history: Received 23 July 2008 Accepted 22 August 2008 Keywords: Tea tree oil Staphylococcus aureus Stationary growth phase Biofilm
a b s t r a c t Tea tree oil (TTO) is known for its antimicrobial activity. In this study, we determined whether TTO is effective against Staphylococcus aureus in biofilms and how TTO activity is affected by the S. aureus growth phase. All clinical strains tested were killed by TTO both as planktonic cells and as biofilms. The minimum biofilm eradication concentration was usually two times higher than the minimum bactericidal concentration, yet it was never higher than 1% v/v. The fastest killing of biofilm occurred during the first 15 min of contact with TTO and was not influenced by increasing TTO concentration above 1% v/v. Planktonic stationary phase cells exhibited decreased susceptibility to TTO compared with exponential phase cells. The killing rate for stationary phase cells was also less affected by increasing TTO concentration than that for exponential phase cells. These data show that TTO efficiently kills S. aureus in the stationary growth phase and within biofilms and is therefore a promising tool for S. aureus eradication. © 2008 Elsevier B.V. and the International Society of Chemotherapy. All rights reserved.
1. Introduction Tea tree oil (TTO), an essential oil obtained from Melaleuca alternifolia, has gained the attention of scientists, physicians and consumers because of its broad antimicrobial and antiinflammatory activities. TTO consists of ca. 100 different compounds, including terpinen-4-ol, which is one of the main antibacterial components [1]. Drugs and care products containing TTO are frequently used for treatment of skin, oral, vaginal and airway infections, or as antiseptics and disinfectants [1,2]. Furthermore, there have been reports of efficient eradication of bacteria in deep infections such as abscesses or osteomyelitis [1,3]. Most TTOrelated research is focused on Staphylococcus aureus. Preliminary clinical trials have shown that the efficacy of TTO in eradication of meticillin-resistant S. aureus (MRSA) carriage is comparable with traditional mupirocin treatment [1]. During infections, bacteria are usually present in various growth phases. In chronic infections or when access to nutrition is limited, bacteria reside predominantly in a stationary or dormant phase [4]. Following brief exposure to antibiotics, continuous suppression of
∗ Corresponding author. Present address: Department of Rheumatology and Inflammation Research, University of Göteborg, Guldhedsgatan 10, S-413 46 Göteborg, Sweden. Tel.: +46 31 342 6475; fax: +46 31 823 925. ´ E-mail address: [email protected] (J. Kwiecinski).
microbial growth is also observed [5]. Non-multiplying or slowly growing organisms are generally less susceptible to antibacterial agents [4]. S. aureus is able to form biofilms, which are complex structures consisting of surface-attached bacteria surrounded by a self-produced extracellular polymer matrix. Bacteria in biofilms exhibit elevated resistance both to antibiotics and the host defence systems, which often results in persistent and difficult-to-treat infections [6–8]. S. aureus biofilm formation contributes to its pathogenesis in a number of conditions, such as damaged skin infections, chronic and recurrent airway infections, osteomyelitis and mastitis [6–11], and to its colonisation on the surface of undamaged skin [12]. Additionally, biofilms of staphylococci on surfaces in the food industry pose a serious risk of food contamination [13]. Therefore, for rational treatment of numerous diseases and development of antiseptics and disinfectants, it is important to determine the influence of TTO on bacteria in stationary growth phase and on already established pre-formed biofilms. It was shown that 1 h treatment with 5% TTO eradicates biofilms of S. aureus [12]. However, it is not presently known at what concentrations TTO is effective against biofilms, what the kinetics of TTO biofilm eradication are, how TTO affects the viability of S. aureus biofilms in comparison with planktonic bacteria, and whether the response to TTO is growth phase-dependent. The aim of this study was to answer these questions, which are crucial for the design of effective and rational therapy.
0924-8579/$ – see front matter © 2008 Elsevier B.V. and the International Society of Chemotherapy. All rights reserved. doi:10.1016/j.ijantimicag.2008.08.028
344
J. Kwieci´ nski et al. / International Journal of Antimicrobial Agents 33 (2009) 343–347
2. Materials and methods 2.1. Tea tree oil TTO was obtained from Avicenna-Oil (Wrocław, Poland) and complied with the ISO 4730 and European Pharmacopoeia standards. 2.2. Bacterial strains Twenty-seven clinical isolates of S. aureus as well as the reference strain S. aureus NCTC 8325-4 were used in this study. All clinical strains were isolated at the Institute of Medical Microbiology, University Hospital of Jena, Jena, Germany. The strains were isolated from the following locations: upper respiratory tract (n = 5); lower respiratory tract (n = 6); superficial wound infections (n = 9); deep wound infections, e.g. bone (n = 5); and urinary tract infections (n = 2). Two-thirds of them were penicillin-resistant and additionally two strains were MRSA. 2.3. Determination of the minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) MICs and MBCs were determined using the broth microdilution method. Overnight cultures (37 ◦ C, shaking) of S. aureus in tryptic soy broth (TSB) (Fluka, Buchs, Switzerland) were diluted 10-fold in fresh TSB and incubated (37 ◦ C, shaking) until they reached exponential growth phase. Serial two-fold dilutions of TTO (from 2% to 0.125% v/v) in TSB were prepared in a 96-well plate (200 L per well). Wells with no TTO added were used as a positive growth control. All wells contained Tween 20 at a final concentration of 0.1% v/v to increase TTO solubility. A diluted bacterial suspension was added to each well to give a final concentration of 1–5 × 105 colony-forming units (CFU)/mL, confirmed by viable counts. Wells without bacteria added were used as a negative growth control. The plate was incubated for 24 h at 37 ◦ C and growth was assessed turbidometrically. From all wells not showing visible growth (optical density at 600 nm (OD600 ) ≤0.05), 10 L was plated on tryptic soy agar (TSA) (Fluka) and the number of colonies was counted following overnight incubation at 37 ◦ C. The MIC was defined as the lowest TTO concentration without visible growth, and the MBC was defined as the lowest concentration reducing the initial inoculum by ≥99.9%. The MIC and MBC were determined for all 27 clinical strains and S. aureus 8325-4. For each strain, at least three independent determinations were done and the modal value was taken.
After 24 h of biofilm formation, the medium was gently aspirated from the plate and the wells were rinsed three times with phosphate-buffered saline (PBS). Serial two-fold dilutions of TTO (from 2% to 0.125% v/v) in 200 L of TSB were then added to wells. No TTO was added to the positive biofilm control wells. All wells contained Tween 20 at a final concentration of 0.1% v/v. The plate was incubated at 37 ◦ C for 24 h, thereafter the medium was gently aspirated and wells were rinsed three times with PBS. Biofilm viability was assessed with either the 3-(4,5-dimethylthiazol-2yl)-2,5-diphenyltetrazolium bromide (MTT) staining method (13 clinical strains forming the best biofilms in a preliminary experiment and S. aureus 8325-4) or the CFU counting method (S. aureus 8325-4 only). Viability measures were done in duplicate in three independent experiments. MTT staining was done according to Walencka et al. [15], with minor modifications. Briefly, 200 L of 0.05% MTT (Sigma, Steinheim, Germany) in PBS was added to the wells with biofilm. Following incubation at 37 ◦ C for 2 h, MTT solution was replaced by 200 L of 5 mM HCl in isopropanol, all formed formazan crystals were dissolved and the absorbance was measured at 550 nm. The minimum biofilm eradication concentration (MBEC) was defined as the lowest TTO concentration resulting in a decrease in absorbance to <5% of the positive control (cut-off level for negative control, determined in the preliminary experiment). The biofilm CFU counting method was adapted from Pettit et al. [16]. The level of detection was 102 CFU/mL. In addition, scanning electron microscopy (SEM) images were taken. A 24-well tissue culture plate was filled with coverslips. S. aureus 8325-4 biofilms were prepared as described previously. After an incubation time of 24 h, the slips were carefully washed with PBS, then TTO solutions were added and the slips were incubated again for 24 h. After removal of the media and washing, the samples were initially fixed in 2% glutaraldehyde in cacodylate buffer for 30 min, then washed twice with cacodylate buffer and dehydrated for 10 min using a graded ethanol series. A critical point drying procedure followed and the specimens were then sputter-coated with gold. Samples were examined with a LEO 1450 VP scanning electron microscope (Carl Zeiss, Germany). 2.6. Biofilm time-dependent killing assay A biofilm of S. aureus 8325-4 was established as described previously. TTO solutions (0, 0.5, 1, 2, 4 and 8% v/v) in 200 L of TSB with 0.1% v/v Tween 20 were added to the wells. The plates were incubated at 37 ◦ C for 0, 15, 30, 60 and 120 min. The medium was then aspirated, the wells were washed three times with PBS and biofilm
2.4. Planktonic time-dependent killing assay S. aureus 8325-4 suspensions from stationary (overnight culture in TSB, 37 ◦ C, shaking) and exponential (prepared as for MIC determination) growth phases were prepared. In plastic tubes, 1 mL of each was mixed with 100 L of TTO solution to final concentrations of 0% (control), 0.5% or 8% v/v and with Tween 20 (0.1% v/v final concentration in all tubes). Tubes were incubated at 37 ◦ C with shaking. After 15, 30 and 60 min, 10 L was removed from each tube and the number of CFU/mL was counted by plating serial dilutions on TSA. A sample from the 0% TTO tube taken immediately after mixing was used as a ‘time 0’ control. The detection level was 103 CFU/mL and measurements were made in three independent experiments. 2.5. Effects of TTO on established biofilms The method of biofilm formation in 96-well cell culture plates ´ according to Kuzma et al. was used [14].
Fig. 1. Cumulative minimum inhibitory concentration (MIC), minimum bactericidal concentration (MBC) and minimum biofilm eradication concentration (MBEC) of tea tree oil (TTO) against clinical isolates of Staphylococcus aureus, expressed as a percentage of tested strains. MIC and MBC, n = 27; MBEC, n = 13.
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Fig. 2. Effect of different concentrations of tea tree oil (TTO) on the viability of biofilm formed by Staphylococcus aureus 8325-4 following a 24 h exposure. Viability was determined by (a) 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) staining or (b) colony-forming unit (CFU) counting. Mean ± standard error of the mean for three independent experiments performed in duplicate. The limit of detection for CFU counting was 102 CFU/mL.
Fig. 3. Scanning electron microscopy images of Staphylococcus aureus 8325-4 biofilm: (a) control; and (b) after 24 h exposure to 1% tea tree oil.
viability was assessed with the MTT staining method. All measurements were made in duplicate in three independent experiments. 3. Results 3.1. TTO has a bactericidal effect and disrupts biofilms For clinical isolates, the MICs of TTO were in the range 0.125–0.5% (MIC for 50% of the organisms (MIC50 ) = 0.25%; MIC for 90% of the organisms (MIC90 ) = 0.5%). The highest MBC was 1%
(range 0.25–1%; MBC50 = 0.25%; MBC90 = 0.5%). Although the highest MBEC was also 1%, these values were in general one step higher than the MBCs (range 0.5–1%; MBEC50 = 0.5%; MBEC90 = 1%). These results are summarised in Fig. 1. For S. aureus 8325-4, the MIC was equivalent to the MBC (0.5%). The effects of TTO on S. aureus 8325-4 biofilm viability measured by MTT staining and CFU counting are shown in Fig. 2. Both methods exhibit good correlation and showed that 1% TTO effectively eradicated the biofilm. SEM images underline the results, as 1% TTO destroyed the biofilm formed by S. aureus 8325-4 (Fig. 3). The susceptibility of MRSA strains was comparable with other strains. A characteristic pattern of increased biofilm metabolism (assessed by MTT staining) at sub-MBEC concentrations was observed for all strains tested (data not shown).
3.2. Activity of TTO on S. aureus is growth phase-dependent
Fig. 4. Time–kill curves of Staphylococcus aureus 8325-4 from stationary and exponential growth phase after treatment with none, 0.5% or 8% tea tree oil (TTO). Each symbol indicates the mean ± standard error of the mean for three independent experiments. The limit of detection was 103 colony-forming units (CFU)/mL.
The number of viable S. aureus in exponential growth phase was reduced by two log10 steps (99% killed) with 0.5% TTO after 60 min, whilst addition of 8% TTO resulted in reduction by four log10 steps (99.99% killed) (Fig. 4). The rate of killing decreased in a time-dependent manner, with the fastest killing during the first 15 min. S. aureus cells in stationary growth phase were reduced by one log10 step (90% killed) after addition of both 0.5% and 8% TTO for 15 min. Later, only slight killing was observed.
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Fig. 5. Time–kill curves for Staphylococcus aureus 8325-4 biofilm after treatment with 0.5%, 1% or 8% tea tree oil. Each symbol indicates the mean ± standard error of the mean for three independent experiments performed in duplicate.
3.3. TTO rapidly kills bacteria in biofilm An exponential decrease of biofilm metabolism was seen after exposure to TTO (Fig. 5). After 120 min, biofilm metabolic activity was only 40% of control activity for biofilm exposed to 0.5% TTO, and 15% of control activity for 1% TTO exposure. Increasing TTO concentration up to 8% did not have any additional effect on the kinetics of biofilm killing (data for 2% and 4% TTO not shown). The fastest killing occurred during the first 15 min and the rate of killing decreased in a time-dependent manner. Most of the killing was completed within 30–60 min. 4. Discussion These data show that TTO efficiently kills S. aureus both in suspension and in biofilms. The obtained MIC and MBC values confirm the high susceptibility of clinical strains to TTO and are in the range reported in previous studies (0.12–2% for MICs and 0.25–8% for MBCs [12,17–19], although a much lower MIC has also been described [20]). MTT staining is one of the methods used for assessing the viability of bacterial biofilms based on reduction of tetrazolium salts to formazan by viable metabolising bacteria [14–16]. Using this method, our study shows that 1% TTO was able to inhibit the metabolism of biofilms formed by all tested strains, which is only twice the bactericidal dose. Most antibiotics are up to 1000-times less efficient against bacteria in biofilm than in suspension [7], which makes TTO a very promising treatment alternative. SEM images show disruption of biofilm by 1% TTO. It is therefore likely that biofilm eradication by TTO is not only due to bacteria killing but also partly due to extracellular matrix damage and subsequent removal of biofilm from the surface, especially by higher TTO concentrations. Low TTO concentrations (between 0.125% and 0.25%) had no effect on the number of CFU/mL within S. aureus 8325-4 biofilms, but MTT staining showed increased biofilm metabolic activity. This suggests that the activity of the cells within the biofilm increases in response even to these low concentrations of TTO. This might reflect a stress reaction, indicating that the TTO tolerance in S. aureus is an energy-dependent process. An energy-dependent TTO tolerance in Pseudomonas aeruginosa was previously reported [21]. For the potential clinical use of TTO it is important to establish the kinetics of its action against bacteria within biofilms. The rate of killing was concentration-dependent up to 1% TTO, but further increase did not accelerate killing. After 2 h, metabolic activity of the biofilm was still observed even after treatment with 8% TTO. This
finding is in contrast to previous data suggesting complete biofilm killing after 1 h in 5% TTO [12]. This might be partly explained by the fact that in our study different methods of biofilm formation and viability testing were used. The MTT method slightly overestimates biofilm viability, as many already lethally injured cells might still exhibit metabolism. Conversely, methods used in the previous study might underestimate bacterial viability owing to additional damage caused by sonication used for dislodging bacteria [12]. Nevertheless, after 1 h, 1% TTO causes a large (ca. 85%) reduction of S. aureus biofilm viability. Escherichia coli in stationary growth phase were killed by TTO ca. 105 times slower than cells from the exponential phase [22]. Our results show that S. aureus in the stationary phase is also more tolerant to TTO compared with bacteria in the exponential phase. When entering the stationary growth phase, cell membrane protein composition [23], cell membrane fluidity [24] and the surface charge and hydrophobicity [25] of S. aureus change. These alterations might account for the changed TTO tolerance, as the cell membrane is the main target of TTO antistaphylococcal activity [26,27]. Both in biofilm and in the stationary phase, increasing TTO concentration has only limited effects on the killing rate. This might be connected to so-called ‘persister cells’ present in biofilms and stationary phase cultures [28]. These cells, which exhibit tolerance to numerous antimicrobial agents, might be a key to increased resistance in biofilms. Further studies are required to elucidate their possible role in TTO biofilm and stationary phase tolerance. According to previous studies, rapid development of TTO resistance is rather unlikely [20,29]. Therefore, TTO is a promising alternative for S. aureus eradication, both as a disinfectant and as infection treatment. It acts on stationary phase and biofilm bacteria – two states frequently encountered in the environment and in living organisms – and it kills the bacteria quickly and effectively at a concentration of 1%, which is below concentrations used in commercially available preparations (up to 10%). TTO may be a new, cheap and effective weapon for fighting S. aureus infections, including MRSA. Acknowledgments The authors acknowledge Susanne Linde (Department of Ultrastructural Research, University Hospital of Jena, Jena, Germany) for excellent technical assistance in preparing SEM photographs, Inger Gjertsson (Department of Rheumatology and Inflammation Research, Sahlgrenska Academy at Göteborg University, Sweden) for critical reading of the manuscript, and Ewa Grela for assistance in preparation of the figures. Funding: No funding sources. Competing interests: None declared. Ethical approval: Not required. References [1] Carson CF, Hammer KA, Riley TV. Melaleuca alternifolia (tea tree) oil: a review of antimicrobial and other medicinal properties. Clin Microbiol Rev 2006;19:50–62. [2] Price S, Price L. Aromatherapy for health professionals. Edinburgh, UK: Churchill Livingstone; 1999. [3] Kawakami E, Washizu M, Hirano T, Sakuma M, Takano M, Hori T, et al. Treatment of prostatic abscesses by aspiration of the purulent matter and injection of tea tree oil into the cavities in dogs. J Vet Med Sci 2006;68:1215–7. [4] Eng RH, Padberg FT, Smith SM, Tan EN, Cherubin CE. Bactericidal effects of antibiotics on slowly growing and nongrowing bacteria. Antimicrob Agents Chemother 1991;35:1824–8. [5] Fuursted K. Postantibiotic effect in vitro. APMIS Suppl 1999;90:1–23. [6] Kania RE, Lamers GE, Vonk MJ, Dorpmans E, Struik J, Tran Ba Huy P, et al. Characterization of mucosal biofilms on human adenoid tissues. Laryngoscope 2008;118:128–34. [7] Melchior MB, Vaarkamp H, Fink-Gremmels J. Biofilms: a role in recurrent mastitis infections? Vet J 2006;171:398–407.
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