Abstract
In vitro antifungal activities of three biocides including one biguanide (chlorhexidine) and two quaternary ammonium compounds (benzalkonium chloride and cetrimide) were studied against eight cleanroom fungal isolates by using a microbiological broth dilution technique as per Clinical and Laboratory Standards Institute (CLSI) M38-A guidelines. No data exists on the activity of biocides on pharmaceutical cleanroom fungal isolates. Minimum inhibitory concentrations (MICs) for all three biocides against species of Aspergillus and Penicillium species ranged between 4 and 16 μg/mL. MICs of Curvularia, Cladosporium, and Alternaria species also showed less than 16 μg/mL. To date, susceptibility breakpoints have not been established for biocides, and this is the first study using the CLSI broth microdilution antifungal susceptibility testing to determine the MIC value of biocides. This study clearly demonstrates that the most frequently isolated micro-organisms from an environmental monitoring program may be periodically subjected to microbroth dilution testing with cleanroom disinfectant agents used in the disinfection program to confirm their sensitivity profile. Further work is needed in this field to increase our understanding of biocides against different fungal isolates and to enable to the design of more efficient disinfection and contamination control programs.
LAY ABSTRACT: Increased trend of fungal growth is observed in pharmaceutical and medical device cleanrooms. It is essential to have knowledge of choosing effective disinfectants for minimizing fungal occurrence in cleanrooms. The present study establish minimum cut-offs for specific fungal isolates that are problematic in cleanrooms against commonly used disinfectants (quaternary ammonium compounds and biguanides). Further studies based on this minimum inhibitory concentration of disinfectants, effective time, and cleaning methods could prevent fungal occurrence.
Introduction
Fungi (moulds and yeasts) are an important group of micro-organisms which are responsible for various infections. They are, however, primarily associated with contamination of surfaces and spoilage of pharmaceutical, cosmetic, and food products. Fungal infection or contamination can result in serious economic losses, for example spoilage in pharmaceutical, food, or cosmetic manufacturing and increased duration in hospital (1, 2). Antiseptics and disinfectants are used extensively in hospitals and other health care centres to control the growth of microbes on both living tissues and inanimate objects. They are essential parts of infection control practices and aid in the prevention of nosocomial infections (3). However, a common problem is the selection of appropriate disinfectants and antiseptics because different pathogens vary in their response to different antiseptics or disinfectants (4). In a similar way to pharmaceutical industries, cleaning and disinfection measures are important and decisive process steps for fulfilling the quality requirements of the medicinal product. Nevertheless, maintaining the integrity of a cleanroom is a constant battle.
In order to decide which method or combination of methods are to be employed in disinfecting aseptic processing areas, it is important to understand the types of micro-organisms that are the prime sources of contamination (5, 6). Therefore, knowledge of the microbial diversity of cleanrooms, as well as any extreme characteristics these microbes might possess, is essential to the development of disinfection technologies. The more frequently detected bacterial and fungal contaminants in the pharmaceutical cleanroom environment are species of Staphylococcus, Micrococcus, Bacillus, Penicillium, Cladosporium, and Aspergillus (7, 8). Antiseptics and disinfectants are used extensively to control the growth of microbes on both living tissues and inanimate objects. According to USP 〈1072〉 and the European Commission's good manufacturing practice (GMP) guidelines, monitoring of environmental isolates and checking their susceptibility pattern to disinfectants is very important for cleanroom disinfection programs (9, 10).
Antifungal susceptibility tests are performed on fungal pathogens in clinical microbiology setups, especially if they belong to a species exhibiting resistance to commonly used antifungal agents. Antifungal susceptibility testing is also important in resistance surveillance, epidemiological studies, and in comparison of the in vitro activity of new and existing agents. Dilution methods are used to establish the minimum inhibitory concentrations (MICs) of antimicrobial agents: these are the reference methods for antimicrobial susceptibility testing and are mainly used to establish the activity of a new antifungal agent, to confirm the susceptibility of micro-organisms to the antifungal agent that give equivocal results in routine tests, and to determine the susceptibility on fungi where routine tests may be unreliable.
In dilution tests, fungi are tested for their ability to produce visible growth in microdilution plate wells of broth culture media containing serial dilutions of the antimicrobial agents (broth microdilution). The MIC is defined as the lowest concentration, recorded in milligrams per liter (mg/L) or micrograms per liter (μg/mL) of an antifungal agent that inhibits the growth of a fungus (through exhibiting fungistatic or fungicidal properties). The MIC informs about the susceptibility or resistance of the organism to the antifungal agent and can help in treatment decisions (11–13). The development of microbial resistance to antibiotics is a well-described phenomenon. However, there are no published reports available describing the resistance of pharmaceutical cleanroom fungal isolates against disinfectants. However, in order to test the efficacy of disinfectants the most frequently isolated micro-organisms from a cleanroom environmental monitoring program (such as settle plates, contact plates, swabs, and active air samples) may be periodically subjected to dilution testing with the agents used in the disinfection program to confirm their susceptibility. Notwithstanding the importance of such studies when compared to bacterial cleanroom contaminants, reports on the MIC of fungal isolates against common disinfectants are not available.
Hence, the aim of our present study is to examine the MIC of biocides against fungi isolated from a cleanroom environment. For this, three biocides commonly used in hospitals and pharmaceutical manufacturing environments were selected: a biguanide (chlorhexidine) and two quaternary ammonium compounds (QACs—benzalkonium chloride, cetrimide). Biguanides are polymers supplied in salt form, such as chlorhexidine, alexidine, or hydrochloride. Biguanides have a relatively wide spectrum of activity with the exception of killing endospores. Biguanides target the bacterial cell membrane, enter the cell through diffusion, and cause cell disruption and cytoplasm leakage. QACs are cationic salts of organically substituted ammonium compounds and have a fairly broad range of activity against micro-organisms. The mode of action targets the cell membrane, leading to cytoplasm leakage and cytoplasm coagulation through interaction with phospholipids. These biocides were examined against cleanroom fungal isolates using the microbroth dilution method.
Methods
(a) Isolates
We selected eight cleanroom fungal isolates that included predominant hyaline and dematiaceous cleanroom isolates. All the fungal isolates were collected from environmental monitoring samples collected from change room facilities (Grades C and D) located in Aurolab (Madurai, India). The samples were collected between September 2010 and March 2011. The fungal isolates included predominant cleanroom isolates like species of hyaline fungi Aspergillus, Penicillium, and dematiaceous fungi of Cladosporium, Curvularia, and Alternaria.
Unlike the standardized approaches for disinfectant testing using the quantitative suspension test, there is no standardized end point for the MIC for biocidal agents against fungi; hence, the U.S. Association of Analytical Communities challenge organisms like Aspergillus brasiliensis ATCC 16404 and Candida albicans ATCC 10231 were not included in this study. The antifungal susceptibility against disinfectants was determined according to methods outlined in Clinical and Laboratory Standards Institute (CLSI) documents M38-A (14).
(b) Inoculum Preparation
The inoculum was prepared by overlaying mature slants with sterile distilled water and gently scraping the surface with a wooden applicator stick. The suspension was permitted to sit for 5 min to allow large particles to settle down and then adjusted spectrophotometrically to the correct optical density for each species as outlined in M38-A, providing an inoculum concentration of 0.4–5 × 104 conidia/mL, which was verified by colony count. The inoculum suspensions were diluted (1:50) in RPMI-1640 (Roswell Park Memorial Institute) media buffered with 0.165M morpholinepropanesulfonic acid (34.54 g/L) at pH 7.0.
(c) Antifungal Agents
One biguanide (20% chlorhexidine gluconate, Unilab Chemicals, Mumbai, India), two QACs of 20% benzalkonium chloride (Ubichem Fine Chemicals, Eastleigh, UK), and 100% potency of cetrimide (Unilab Chemicals) were selected as antifungal agents. To prepare, each was dissolved in sterile distilled water following the protocol of CLSI and were prepared in stock solutions of 1000 μg/mL, which subsequently were diluted in RPMI 1640 test medium for further various dilutions preparations.
(d) Preparation of Microdilution Plates
Using sterile plastic, disposable, 96-well microdilution plates with flat-bottom wells with a nominal capacity of approximately 300 μL, 100 μL from each of the tubes containing the corresponding concentration (2× final concentration) of target disinfectants was dispensed into the wells in each column (from 1 to 10). For example, with chlorhexidine, cetrimide, or benzalkonium chloride, to column 1 the medium containing 128 μg/mL (128 mg/L) was dispensed, to column 2 the medium containing 64 μg/mL was dispensed, and so on to column 10 where the medium containing 0.25 μg/mL was dispensed. To each well of columns 11 and 12, 100 μL of RPMI 1640 medium was dispensed. Thus, each well in columns 1–10 contained 100 μL of twice the final antifungal drug concentrations in RPMI medium. Columns 11 and 12 contained double-strength RPMI 1640 medium. The final well concentrations reached were 0.125 μg/mL to 64 μg/mL after addition of inoculum (100 μL). Microdilution plates were stored at −70 °C prior to use.
(e) Antifungal Susceptibility Testing
The MICs were determined following the microdilution method recommended by CLSI, approved standard M38-A (14). This involved testing each fungal isolate in duplicate. Each microdilution well containing 100 μL of the two-fold drug (biocides) concentration was inoculated with 100 μL of diluted inoculum suspension. For each test plate, two drug-free controls were included, one with the medium alone (sterility control) and the other with 100 μL of medium plus 100 μL inoculum suspension (growth control). The biocide concentrations assayed ranged from 0.125 to 64 μg/mL.
(f) Incubation Time and Temperature
The microdilution plates were incubated at 35 °C for up to 48 h or until growth was visible in the drug (biocide)-free control well.
(g) Reading and Interpretation
End point determination values were read visually with the aid of an inverted reading mirror. The MIC was defined as the lowest concentration that exhibited a 100% visual reduction in turbidity when compared with the control well at 48 h. The geometric mean of the MIC for each biocide only was considered a final result.
Results
All fungal isolates tested produced detectable growth after 48–72 h of incubation. The eight fungal isolates were grouped as two categories named hyaline and dematiaceous fungal groups. All Aspergillus species and Penicillium are hyaline fungi, and Cladosporium, Curvularia, and Alternaria, are dematiaceous fungi. Table I summarizes susceptibility data for five hyaline cleanroom fungi and three dematiaceous fungi. Only reproducible results observed in the duplicate plates were considered for the final MIC value. All Aspergillus species tested showed 4–8 μg/mL MIC values for benzalkonium chloride, and a maximum MIC 16 μg/mL was found for Alternaria species. MIC value for all fungal isolates tested for chlorhexidine showed in the range between 2 and 16 μg/mL. In the cetrimide group the MIC range was found to be 4–16 μg/mL.
Antifungal Activity of Bigunides and Quaternary Ammonium Compounds against Hyaline and Dematiaceous Cleanroom Fungal Isolates
Discussion
Antiseptics and disinfectants are broad-spectrum biocidal compounds that inactivate micro-organisms on living tissue and inanimate surfaces. Their mechanisms of action have been extensively studied, as has fungal resistance to them (15). However, limited data are available on the susceptibility and mechanisms of resistance of reference strains of fungi to biocides. The aim of this study was to evaluate the efficacy of selected disinfectant agents against various species and genera of fungal isolates of pharmaceutical environment origin, by using a micro broth dilution technique.
In the present study, biguanides and QACs were tested against environmental fungal isolates. With biguanides, chlorhexidine is probably the most widely used biocide in antiseptic products, in particular in hand washing and oral products but also as a disinfectant and preservative. It interacts with the cell surface and promotes membrane damage, which in turn causes an irreversible loss of cytoplasmic components (16, 17). The killing action of chlorhexidine at relatively low concentrations (e.g., 2–2.5 μg/mL) is similar to the action of some antibiotics. At high concentrations (≥20 μg/mL), chlorhexidine causes coagulation of cytoplasm and precipitation of proteins and nucleic acids. The study found that the MIC range for chlorhexidine against hyaline fungi like species of Penicillium and Aspergillus are 8–16 μg/mL. The MIC range of Cladosporium, Curvularia, and Alternaria was found to be 8–16 μg/mL. This conforms to A. D. Russell's findings where it was reported that chlorhexidine at high concentrations like >20 μg/mL causes coagulation of cytoplasm and precipitation of protein and nucleic acids of bacteria and yeasts (18). Chlorhexidine antifungal activity has been studied by various authors in the context of dental products and the oral environment (19–22).
Cationic agents, as exemplified by QACs, are among the most widespread antiseptics and disinfectants. QACs have been used for a variety of clinical purposes like preoperative disinfection of unbroken skin, application to mucous membranes, and disinfection of non-critical surfaces. The mechanisms of action of benzalkonium chloride are arguably less well elucidated than those of antibiotic agents. Benzalkonium chloride is one of the QACs, membrane-active agents which primarily target the cytoplasmic (inner) membrane of bacteria of the plasma membrane of yeasts (15). The following sequence of events has been proposed in micro-organisms exposed to QACs: adsorption and penetration of the agent into the cell wall, reaction with the cytoplasmic membrane followed by membrane disorganisation, leakage of low-molecular-weight material, degradation of proteins and nucleic acids, and wall lysis caused by autolytic enzymes (15). In vitro testing of the fungicidal activity of benzalkonium chloride against Aspergillus fumigatus showed biocidal activity in less than 5 min of contact time, defined as a 104 or more reduction in the viability of Aspergillus fumigatus strains (23). The study also demonstrated that the MICs of cetrimide and benzalkonium chloride were in the same range between 4 and 16 μg/mL. Four out of eight isolates (50%) showed the same MIC values, three isolates (38%) showed one dilution variation like 8–16 μg/mL and remaining isolate showed two dilution variation (4–16 μg/mL) among QACs tested. This is comparable with our previous reports in which the MIC of benzalkonium chloride against ocular corneal pathogens of Fusarium species were 32–64 μg/mL and against ocular corneal pathogens of Aspergillus species were 16 μg/mL (24).
The dematiaceous fungi contain melanin pigment in their cell walls, with melanins and sporopollenin possibly being involved with cellular resistance to physical and chemical agents (25). However, the study findings showed that there was no significant MIC value difference found between the hyaline and dematiaceous fungal groups. The MICs of both groups of fungi were in the range between 4 and 16 μg/mL. This could be verified with other dematiaceous fungi in further studies.
No previous published studies have examined the MIC value of cleanroom fungi against biguanides and QACs. Therefore, comparison of MIC results with other fungal reports might be difficult. Although several authors have investigated the in vitro effect of biocides against bacterial pathogens, molds are generally more resistant than yeasts and considerably more resistant than non-sporulating bacteria. Mold spores, although more resistant than non-sporulating bacteria, are less resistant than bacterial spores to antiseptics and disinfectants (24). In our study reports, all the fungal isolates MIC range was less than 16 μg/mL which are higher than non-sporulating bacteria such as Staphylococcus aureus and E.coli. But comparing the available literature shown in Table II indicates that MIC data of Pseudomonas aeruginosa is more resistant than fungal spores (26–28).
In regard to fungal resistance against biocides, the development of microbial resistance to antimicrobials is a well-described phenomenon. The development of microbial resistance to disinfectants is less likely, as disinfectants are more powerful biocidal agents than antibiotics and are applied in high concentrations against low populations of micro-organisms usually not growing actively, so the selective pressure for the development of resistance is less profound (29). However, the most frequently isolated micro-organisms from an environmental monitoring program may be periodically subjected to use dilution testing with the agents used in the disinfection program to confirm their susceptibility. To date, susceptibility breakpoints have not been established for biocides, but the MICs obtained in the present study are likely within the satisfactory level. For discussion purposes, the desirable target value <32 μg/mL is used to represent susceptibility. Interestingly, the MICs obtained for all fungal isolates tested susceptible to biguanides and QACs.
Conclusion
Overall, our present study reveals that MICs of hyaline fungi against chlorhexidine, benzalkonium chloride, and cetrimide are 8–16 μg/mL. The MIC range of dematiaceous fungi reported is 8–16 μg/mL for biguanides and QACs. This study clearly demonstrates that the most frequently isolated micro-organisms from an environmental monitoring program may be periodically subjected to microbroth dilution testing with cleanroom disinfectant agents used in the disinfection program confirming their sensitivity profile. In our knowledge this is the first study report using the CLSI broth microdilution antifungal susceptibility testing to determine the MIC value of biocides versus cleanroom fungal isolates.
Further work is needed in this field to increase our understanding of biocides against different fungal isolates and to enable the design of more efficient disinfection and contamination control programs. Additional multi-centre studies should be undertaken to confirm the results presented in this paper using a wide range of fungal isolates. In addition to further research, the test method presented in this paper can be utilized to periodically verify fungi isolated from an environmental monitoring program for their susceptibility pattern. This will provide important information for microbiologists about the effectiveness of in-use disinfectants against the fungi present in hospital and pharmaceutical environments. Through further MIC determinations and activity spectrum classification of commonly used disinfection agents, it will become possible to outline a more effective cleaning and disinfection program for use in the pharmaceutical, food, and cosmetic industrial sectors.
Conflict of Interest Declaration
The authors received no financial support for this work.
Acknowledgments
The authors wish to thank Aurolab for providing the lab facility and support for the research work. The authors would also like to acknowledge the staff of the antiseptic and microbiology division, Aurolab, for their support.
- © PDA, Inc. 2012
