The antimicrobial effects of a coating of molybdenum trioxide (MoO3) has been recently described. The metalloacid material produces oxonium ions (H3O+), which creates an acidic pH that is an effective, non specific antimicrobial. We determined the in vitro antimicrobial activity of molybdenum trioxide metalloacid-coated surfaces.
Methods
Metalloacid-coated and non-coated (control) surfaces were contaminated by exposing them for 15 minutes to microbial suspensions containing 105 cfu/mL. Eleven microorganisms responsible for nosocomial infections were tested: two Staphylococcus aureus strains (the hetero-vancomycin intermediate MRSA Mu50 strain and a ST80-PVL-producing MRSA strain); a vancomycin-resistant van A Enterococcus faecium strain; three extended-spectrum beta-lactamase-producing Enterobacteriaceae strains; a MBL-producing Pseudomonas aeruginosa strain; a multidrug-resistant Acinetobacter baumannii strain; a toxin-producing Clostridium difficile strain; and two fungi (Candida albicans and Aspergillus fumigatus). The assay tested the ability of the coated surfaces to kill microorganisms.
Results
Against all non-sporulating microorganisms tested, metalloacid-coated surfaces exhibited significant antimicrobial activity relative to that of the control surfaces within two to six hours after contact with the microorganisms (p < 0.001). Microorganism survival on the coated surfaces was greatly impaired, whereas microorganism survival on control surfaces remained substantial.
Conclusions
We suggest that, facing the continuing shedding of microorganisms in the vicinity of colonized or infected patients, the continuous biocidal effect of hydroxonium oxides against multidrug-resistant microorganisms may help limit environmental contamination between consecutive cleaning procedures.
Keywords
Metalloacid-coated surfaceBiocidal effectInfection control
Background
Nosocomial infections are a major cause of patient morbidity and mortality. Those associated with contaminated surfaces and the inadequate hand hygiene of healthcare workers (HCWs) are avoidable by cleaning and/or disinfecting environmental surfaces and by appropriate hygiene practices[1–4]. The transmission of microorganisms responsible for contaminated surface-associated nosocomial infections is well understood. Briefly, patients and HCWs carry and shed microorganisms around them. Surviving microorganisms contaminate near-patient items, which then act as reservoirs for microbial pathogens and become sources of contamination for the hands of HCWs and of colonization for new patients. Microbial colonization of surfaces can be restricted by cleaning procedures that employ effective chemical products. Cleaning reduces the prevalence of microorganisms in the environment and has been demonstrated to be a key-measure in the control of outbreaks associated with methicillin-resistant S. aureus (MRSA), vancomycin-resistant enterococci (VRE) and A. baumannii[1–7].
In fact, the continuing shedding of microorganisms between consecutive cleansing interventions, and the difficulties of cleaning particular medical devices, frequently result in residual environmental contamination. An approach to reducing the levels of microbial contamination on inanimate surfaces consists of using anti-adhesive coatings to prevent microorganism adhesion[8, 9]. A second approach is to coat the surfaces of materials with disinfectants or with inorganic antimicrobials such as silver or copper ions[10, 11]. Currently being used for catheters and dressings[12], these technologies are now being frequently applied to other types of medical equipment. The rapid release of the adsorbed biocide after implantation[13], cytotoxicity on mammalian cells[14] and the emergence of resistant microorganisms[15] may limit the biomedical device applications of these technologies. Zollfrank el al. recently reported the antimicrobial effects of a newly developed antimicrobial coating of molybdenum trioxide (MoO3) against two reference bacterial strains, S. aureus ATCC 25923 and Pseudomonas aeruginosa ATCC 10145[16]. The metalloacid material produces oxonium ions (H3O+), which create an acidic pH (pH 5) that is an effective antimicrobial[17]. To assess the value of such coatings in preventing the generation of environmental microbial reservoirs on medical equipment and in diminishing the risk of exposing patients to microorganisms, we studied the biocidal activity of surfaces coated with the MoO3 metalloacid material. We tested eight multidrug-resistant bacteria strains representative of those that cause nosocomial infections and outbreaks worldwide, a spore-producing, toxin-producing strain of Clostridium difficile, and two fungi.
Methods
Microbial strains
Nine bacterial strains were used for the assay: two MRSA strains (the Mu50 reference strain and a ST80-PVL-producing strain); a vancomycin-resistant van A Enterococcus faecium strain; a toxin-producing Clostridium difficile strain, three extended-spectrum beta-lactamase (ESBL)-producing Enterobacteriaceae strains (Escherichia coli, Klebsiella pneumoniae and Enterobacter cloacae); a MBL-producing Pseudomonas aeruginosa strain and a MDR Acinetobacter baumannii strain. All strains except the Mu50 strain were isolated from nosocomial outbreaks in French healthcare institutions during the 12-month period preceding the study. We also tested two fungi: Candida albicans and Aspergillus fumigatus.
Wires
Metalloacid-coated and non-coated EKG leadwires were cut into sections 2 cm long. Groups of six sections of coated and non-coated leadwires were placed into sterile boxes. Before each microbial contamination, the prepacked wires were pasteurized at 65°C for one hour.
Contamination of leadwires
Microbial suspensions were prepared as follows: one colony of a fresh culture on an agar plate was used to inoculate brain-heart infusion broth. After incubation for 24 hours at 37°C, the final microbial suspension was prepared by adding 50 μL of the cultured broth to 100 mL of sterile saline solution. Ten mL of each microbial suspension was added to one test box containing the coated leadwire sections and one control box containing non-coated leadwire sections. The boxes were hermetically sealed and then agitated gently for 15 minutes at ambient temperature. The leadwire sections were then transferred to sterile tubes (one per tube) and this was defined as time 0 (t0). The samples were stored at 37°C for various times.
Semi-quantitative assessment of microorganism survival
In account of the form of the leadwire sections used for the study, we applied the semiquantative culture method of Maki that is well-recognized method for evaluating the microbial density colonization on catheter surface[18]. At t0, and after two, four, six, 24 and 48 hours of storage at 37°C, Maki’s technique was performed by transferring each contaminated leadwire section to a blood agar plate and by rolling the section back and forth across the surface (one section per plate). The plates were then incubated for 48 hours at 37°C. After incubation, the plates were photographed and colonies were counted. For each of the tested microorganisms, the overall assay was repeated five times, and the bacterial counts were used to calculate average counts.
Statistical analysis
Wilcoxon signed-rank tests were used to compare continuous paired data. For all calculations, a two-tailed P value of less than 0.05 was considered to indicate statistical significance. The software package Stata, version 10.0 (Stata Corp., College Station, TX, USA) was used for analysis.
Results and discussion
The data obtained for the contaminated coated and non-coated leadwire sections are reported in Table1. Examples of plate cultures obtained from Mu50 S. aureus-coated leadwires are shown in Figure1. The average colony counts obtained for the non-coated control wires after they were incubated for 2 − 48 hours showed that all the microorganisms tested were able to survive on the surface of the non-coated wires for prolonged periods of time (Table1) (Figure2). Culture of these wires six hours after contamination revealed high bacteria counts in all cases. At 24 hours, very high bacteria counts were detected for Gram-negative organisms. At 48 hours, substantial numbers of bacteria colonies were cultured from wires contaminated with most of the Gram-negative microorganisms (i.e. the ESBL-producing Enterobacteriaceae and P. aeruginosa) and with the two spore-forming microorganisms, A. fumigatus and C. difficile. In contrast, we detected survival of the two staphylococci, E. faecium and C. albicans 24 hours after contamination but never after 48 hours. Overall, these first data confirmed the non-coated leadwire surface as a stand for nosocomial pathogens, capable of harboring viable microorganisms.
Table 1
Colony counts obtained from coated and non-coated surfaces contaminated with microorganisms for 0 − 48 hours
Microorganisms
Colony counts in average cfu (standard deviation) obtained from wires 0 − 48 hours after contamination with various microorganisms1. Data are from five replicate experiments.
P2
Wires
0 hr
2 hr
4 hr
6 hr
24 hr
48 hr
Pseudomonas aeruginosa
coated
> 2000 (nc)
1140 (89)
464 (42)
114 (60)
17 (12)
2 (1)
non-coated
> 2000 (nc)
1540 (89)
1140 (167)
1060 (89)
220 (81)
33 (21)
< 0.001
Acinetobacter baumannii
coated
1440 (89)
430 (110)
72 (38)
69 (53)
21 (30)
< 1 (nc)
non-coated
1420 (84)
1000 (0)
590 (55)
520 (45)
67 (49)
< 1 (nc)
< 0.001
Escherichia coli
coated
> 2000 (nc)
790 (55)
440 (47)
556 (38)
46 (8)
< 1 (nc)
non-coated
> 2000 (nc)
1760 (152)
1500 (0)
1620 (164)
90 (70)
2 (1)
< 0.001
Klebsiella pneumoniae
coated
1140 (nc)
860 (89)
760 (42)
630 (60)
< 1 (nc)
< 1 (nc)
non-coated
1140 (114)
1040 (55)
940 (55)
880 (45)
700 (71)
2 (2)
< 0.001
Enterobacter cloacae
coated
> 2000 (nc)
1940 (89)
1500 (0)
1020 (45)
810 (22)
2 (1)
non-coated
> 2000 (nc)
> 2000 (nc)
> 2000 (nc)
1480 (45)
569 (321)
2 (2)
0.031
Mu50 Staphylococcus aureus
coated
1260 (55)
438 (36)
5 (2)
3 (7)
< 1 (nc)
< 1 (nc)
non-coated
1280 (84)
1060 (55)
980 (45)
760 (55)
11 (8)
< 1 (nc)
< 0.001
PVL Staphylococcus aureus
coated
> 1500 (nc)
770 (45)
540 (89)
108 (16)
< 1 (nc)
< 1 (nc)
non-coated
> 1500 (nc)
1240 (55)
880 (27)
770 (45)
15 (12)
< 1 (nc)
< 0.001
Enterococcus faecium
coated
1220 (110)
124 (33)
42 (26)
11 (5)
< 1 (nc)
< 1 (nc)
non-coated
1480 (45)
470 (76)
350 (35)
306 (26)
3 (2)
< 1 (nc)
< 0.001
Clostridium difficile
coated
280 (45)
88 (12)
86 (15)
54 (15)
46 (12)
55 (22)
non-coated
300 (0)
100 (17)
71 (10)
62 (15)
55 (4)
43 (9)
0.328
Candida albicans
coated
458 (62)
319 (23)
143 (38)
130 (25)
11 (7)
< 1 (nc)
non-coated
710 (86)
482 (63)
338 (24)
336 (32)
4 (4)
< 1 (nc)
< 0.001
Aspergillus fumigatus
coated
123 (22)
101 (9)
43 (12)
28 (7)
11 (4)
4 (4)
non-coated
162 (44)
96 (18)
41 (9)
41 (9)
34 (5)
2 (1)
0.061
Nc: not calculable; 1at t0 and after 2, 4, 6, 24 or 48 hours of storage, contaminated leadwire sections were rolled on blood agar plates. Plate colonies were counted after incubation for 48 hours at 37°C; 2Colony counts were compared using the Wilcoxon signed-rank test to determine statistical significance.
Figure 1
Colonies obtained from contaminated leadwires. At t0 and after 2, 4, 6, 24 and 48 hours of storage at 37°C, coated and non-coated leadwire sections contaminated with Mu50 Staphylococcus aureus were rolled on blood agar plates and then incubated for 48 hours at 37°C.
Figure 2
Schematic representation of the antimicrobial activity of coated and non-coated surfaces contaminated with microorganisms. Colony counts in average cfu obtained from wires 0 − 48 hours after contamination with various microorganisms. Data are from five replicate experiments.
In contrast, the coated surfaces exhibited marked biocidal activity, relative to that of the control surface, against all non-spore-forming microorganisms tested (Table1). The biocidal effect of the coated surface differed with respect to the microorganisms in terms of kinetics, intensity and final outcome. Nevertheless, bacterial counts after six to 24 hours of contact with the coated surface were significantly lower than those observed with non-coated surface (p <0.001). The biocidal effect was the fastest against staphylococci and E. faecium, as marked effects were detected after two hours of contact. In contrast, the biocidal effect on Gram-negative microorganisms took longer and was generally evident after four hours. Of note, the spore-forming microorganisms were completely unaffected by the coated surfaces.
The acidic surface reaction is thought to be related to the diffusion of H3O+ ions through the microbial membranes, resulting in altered enzymatic transport systems and inhibition of key metabolic activities[17]. Thus, the acidic-pH-associated biocidal effect produced by the metalloacid-coated surface would be expected to have a universal effect. The variations in susceptibility between Gram-positive and Gram-negative microorganisms, and the resistance exhibited by the spore-forming ones, suggest that the differences among microorganisms with respect to their susceptibility to the biocidal surface may be due to the H3O+ ion permeability of their cell wall and/or cell membrane. In account of a biological cost of antimicrobial resistance, MDR microorganisms may be less fit and have slower growth rates than sensitive strains[19]. Thus, further studies should be done with wild isolates, to guaranty a biocide effect on them. Despite these limitations, our data show that the survival of non-spore-forming multidrug resistant microorganisms was severely diminished on coated surfaces compared to that on control surfaces. Considering the continuous spread of microorganisms in the surroundings of patients, the usual rhythm of cleaning procedures in hospital wards, and the continuous biocidal effect of the metalloacid-coated surface against the multidrug resistant nosocomial pathogens, we suggest that coated device surfaces may provide an permanent means of minimizing microbial contamination between two cleaning procedures.
The mechanism of microorganism killing at pH values <4.0 is non-specific[1]. Thus, in contrast to disinfectants and antibiotics, microbial resistance to this mode of action may not emerge as a result of exposure to MoO3. In addition, cytotoxicity tests (MTT assays) and thrombogenicity tests of the powder components suggest that they should be safe for human use[16].
Conclusions
In view of these promising in vitro results, we suggest that MoO3 coating may be an effective means of minimizing microorganism contamination on certain hospital surfaces. Further studies are needed to evaluate the benefits of this coating on medical devices that are frequently touched by HCWs and to determine whether it can be used as a complementary measure for the preventing of the spread of microorganisms to sites near patients in hospital settings.
Declarations
Acknowledgements
This work was supported by the Centre de Coordination de la Lutte Contre les Infections Nosocomiales de l’Ouest de la France (CCLIN Ouest), the Agence Régionale de Santé du Centre, and the Centre Universitaire de Tours, France. Metalloacid-coated and non-coated leadwires were provided free of charge by Integral Process and Leoni.
Authors’ Affiliations
(1)
Service de Bactériologie et Hygiène, Centre Hospitalier Universitaire de Tours
(2)
Service d’Hygiène Hospitalière, Centre Hospitalier Universitaire de Besançon
(3)
Réseau des Hygiénistes de la région Centre, Hôpital Trousseau, Centre Hospitalier Universitaire de Tours
References
Brady RRW, Verran J, Damani NN, Gibb AP: Review of mobile communication device as potential reservoirs of nosocomial pathogens. J Hosp Infect. 2009, 71: 295-300. 10.1016/j.jhin.2008.12.009.View ArticlePubMedGoogle Scholar
Dancer SJ: Importance of the environment in methicillin-resistant Staphylococcus aureus acquisition: the case for hospital cleaning. Lancet Infect Dis. 2008, 8: 101-113. 10.1016/S1473-3099(07)70241-4.View ArticlePubMedGoogle Scholar
Dumford DM, Nerandzic MM, Eckstein BC, Donskey CJ: What is on that keyboard? Detecting hidden environmental reservoirs of Clostridium difficile during an outbreak associated with North American pulsed-field gel electrophoresis type I strains. Am J Infect Control. 2009, 37: 15-19. 10.1016/j.ajic.2008.07.009.View ArticlePubMedGoogle Scholar
Weber DJ, Rutala WA, Miller MB, Huslage K, Sickbert-Bennett E: Role of hospital surfaces in the transmission of emerging health care-associated pathogens: Norovirus, Clostridium difficile, and Acinetobacter species. Am J Infect Control. 2010, 38: S25-S33. 10.1016/j.ajic.2010.04.196.View ArticlePubMedGoogle Scholar
Falk PS, Winnike J, Woodmansee C, Desai M, Mayhall CG: Outbreak of vancomycin-resistant enterococci in a burn unit. Infect Control Hosp Epidemiol. 2000, 21: 575-582. 10.1086/501806.View ArticlePubMedGoogle Scholar
Goodman ER, Platt R, Bass R, Onderdonk AB, Yokoe DS, Huang SS: Impact of environmental cleaning intervention on the presence of methicillin-resistant Staphylococcus aureus and vancomycin-resistant enterococci on surfaces in intensive care units. Infect Control Hosp Epidemiol. 2008, 29: 593-599. 10.1086/588566.PubMed CentralView ArticlePubMedGoogle Scholar
van der Mee-Marquet N, Girard S, Lagarrigue F, Leroux I, Voyer I, Bloc D, Besnier JM, Quentin R: Multiresistant Enterobacter cloacae outbreak in an intensive care unit associated with therapeutic beds. Crit Care. 2006, 10: 405-406. 10.1186/cc4835.PubMed CentralView ArticlePubMedGoogle Scholar
Hinüber C, Kleemann C, Friederichs RJ, Haubold L, Scheibe HJ, Schuelke T, Boehlert C, Baumann MJ: Biocompatibility and mechanical properties of diamond-like coatings on cobalt-chromium-molybdenum steel and titanium-aluminum-vanadium biomedical alloys. J Biomed Mater Res A. 2010, 95: 388-400.View ArticlePubMedGoogle Scholar
Rose SF, Okere S, Hanlon GW, Lloyd AW, Lewis AL: Bacterial adhesion to phosphorylcholine-based polymers with varying cationic charge and the effect of heparin pre-adsorption. J Mater Sci Mater Med. 2005, 16: 1003-1015. 10.1007/s10856-005-4755-y.View ArticlePubMedGoogle Scholar
Darouiche RO, Mansouri MD, Gawande PV, Madhyastha S: Antimicrobial and antibiofilm efficacy of triclosan and DispersinB combination. J Antimicrob Chemother. 2009, 64: 88-93. 10.1093/jac/dkp158.View ArticlePubMedGoogle Scholar
Domek MJ, LeChevallier MW, Cameron SC, McFeters GA: Evidence for the role of copper in the injury process of coliform bacteria in drinking water. Appl Environ Microbiol. 1984, 48: 289-293.PubMed CentralPubMedGoogle Scholar
Maki DG: In vitro studies of a novel antimicrobial luer-activated needless connector for prevention of catheter-related bloodstream infection. Clin Infect Dis. 2010, 50: 1580-1587. 10.1086/652764.View ArticlePubMedGoogle Scholar
Atiyeh BS, Costagliola M, Hayek SN, Dibo SA: Effect of silver on burn wound infection control and healing: review of the literature. Burns. 2007, 33: 139-148. 10.1016/j.burns.2006.06.010.View ArticlePubMedGoogle Scholar
Silver S: Bacterial silver resistance: molecular biology and uses and misuses of silver compounds. FEMS Microbiol Rev. 2003, 27: 341-353. 10.1016/S0168-6445(03)00047-0.View ArticlePubMedGoogle Scholar
Plumridge A, Hesse SJ, Watson AJ, Lowe KC, Stratford M, Archer DB: The weak acid preservative sorbic acid inhibits conidial germination and mycelial growth of Aspergillus niger through intracellular acidification. Appl Environ Microbiol. 2004, 70: 3506-3511. 10.1128/AEM.70.6.3506-3511.2004.PubMed CentralView ArticlePubMedGoogle Scholar
Zollfrank C, Gutbrod K, Wechsler P, Guggenbichler JP: Antimicrobial activity of transition metal acid MoO3 prevents microbial growth on material surfaces. Mat Sci Engin. 2011, 32: 47-54.View ArticleGoogle Scholar
Maki DG, Weise CE, Sarafin HW: A semiquantative culture method for identifying intravenous catheter-related-infection. N Engl J Med. 1977, 296: 1305-1309. 10.1056/NEJM197706092962301.View ArticlePubMedGoogle Scholar
Gillespie SH, McHugh TD: The biological cost of antimicrobial resistance. Trends Microbiol. 1997, 5: 337-339. 10.1016/S0966-842X(97)01101-3.View ArticlePubMedGoogle Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
