Journal of Microbiological Methods 112 (2015) 49–54
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Journal of Microbiological Methods journal homepage: www.elsevier.com/locate/jmicmeth
An in situ antimicrobial susceptibility testing method based on in vivo measurements of chlorophyll α fluorescence Nikolaos S. Heliopoulos a, Angeliki Galeou b, Sergios K. Papageorgiou a,⁎, Evangelos P. Favvas a, Fotios K. Katsaros a, Kostas Stamatakis b a b
Institute of Nanoscience and Nanotechnology, N.C.S.R. Demokritos, Terma Patriarchou Grigoriou & Neapoleos, Ag.Paraskevi Attikis, 15310 Athens, Greece Institute of Biosciences and Applications, N.C.S.R. Demokritos, Terma Patriarchou Grigoriou & Neapoleos, Ag. Paraskevi Attikis, 15310 Athens, Greece
a r t i c l e
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Article history: Received 24 December 2014 Received in revised form 11 March 2015 Accepted 12 March 2015 Available online 13 March 2015 Keywords: Antibacterial properties Chlorophyll fluorescence Cyanobacteria Textiles
a b s t r a c t Up to now antimicrobial susceptibility testing (AST) methods are indirect and generally involve the manual counting of bacterial colonies following the extraction of microorganisms from the surface under study and their inoculation in a separate procedure. In this work, an in situ, direct and instrumental method for the evaluation and assessment of antibacterial properties of materials and surfaces is proposed. Instead of indirectly determining antibacterial activity using the typical gram(−) test organisms with the subsequent manual colony count or inhibition zone measurement, the proposed procedure, employs photosynthetic gram(−) cyanobacteria deposited directly onto the surface under study and assesses cell proliferation and viability by a quick, accurate and reproducible instrumental chlorophyll fluorescence spectrophotometric technique. In contrast with existing methods of determination of antibacterial properties, it produces high resolution and quantitative results and is so versatile that it could be used to evaluate the antibacterial properties of any compound (organic, inorganic, natural or man-made) under any experimental conditions, depending on the targeted application. © 2015 Elsevier B.V. All rights reserved.
1. Introduction The increasing concern of human societies about hygiene and healthcare has intensified efforts for the development and production of materials and surfaces with antibacterial properties. In order to evaluate the bactericidal properties of such materials, several testing methods have been developed and are being implemented, both in research and in industrial applications. Based on the type of evaluation of antibacterial properties, methods can be classified into two major categories: qualitative and quantitative. The standard methods of the first category (Bauer et al., 1959; Jorgensen and Turnidge, 2007; Clinical Laboratory Standards Institute, 2006) evaluate antibacterial action based on the inhibition of growth around the sample under study. Even though sometimes the diameter of the zones of inhibition are measured and recorded they cannot be considered quantitative as the actual sizes of the colonies cannot be measured. In addition, this kind of approach has a number of drawbacks. Antibacterials that leach out into the aqueous agar matrix, such as silver ions, show better test performance than when they are fixed on the substrate or they are not water soluble, while antibacterials that react with the agar cannot be tested. Moreover, the relative ⁎ Corresponding author. E-mail address: [email protected] (S.K. Papageorgiou).
http://dx.doi.org/10.1016/j.mimet.2015.03.011 0167-7012/© 2015 Elsevier B.V. All rights reserved.
efficiency of different antimicrobial compounds cannot be evaluated as their rate of diffusion after plating the bacteria in agar differs greatly. Finally, diffusion as observed in the test does not represent antimicrobial behavior in actual use (Dickert et al., 1981; Wilkins and Thiel, 1973; Gouveia, 2010; Klancnik et al., 2010). On the other hand, quantitative methods are the most widely used and provide results based on the measurement of bacterial populations. These are inoculation and recovery methods that measure reduction in surviving cell populations in relation either to an initial inoculation level of bacteria or against an untreated control. However, quantitative methods have also their own limitations. Firstly, different methods must be used for porous, non-porous and uneven surfaces. Moreover, the bacterial growth conditions used are too favorable (e.g., rich nutrients in the inoculum and saturating moisture in the testing surfaces) to realistically simulate real life conditions (AATCC 100-2004; ASTM E2149-10). To date, very few quantitative studies have examined the antimicrobial effects under conditions of normal material use. For example ISO 20743:2007 that determines the antibacterial activity of antibacterial finished textile products based on the intended application and on the environment in which the textile product is to be to used, uses a lower number of nutrients but is time consuming and costly as it requires incubations, sterilization of the samples, humidity regulation of the test samples, filtering of test bacteria, printing of test bacteria with a special printing apparatus, etc. (Gao and Cranston, 2008).
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In general, quantitative methods are long, are expensive to apply and require time consuming work while a number of treatments must be performed to both the sample and the organisms used making them strongly dependent on the operator's interpretation and technique. Furthermore, modifications are often made according to standard protocols while often details of the assays are not fully disclosed. Recently, several commercial systems have been developed that provide conveniently prepared and formatted microdilution panels as well as instrumentation and automated reading of plates, intended to reduce technical errors and lengthy preparation times. Most of these automated antimicrobial susceptibility testing systems provide automated inoculation, reading and interpretation. Some examples of these include: Vitek System (bioMerieux, France), Walk-Away System (Dade International, Sacramento, Calif.), Sensititre ARIS (Trek Diagnostic Systems, East Grinstead, UK), Avantage Test System (Abbott Laboratories, Irving, Texas), Micronaut (Merlin, Bornheim-Hersel, Germany), Phoenix (BD Biosciences, Maryland) and many more. In these systems, although they are fast and convenient, the cost entailed in initial purchase, operation and maintenance of the machinery is very high for most laboratories. Cyanobacteria are prokaryotic organisms that perform oxygenic photosynthesis in a manner remarkably similar to green plants. Because they are bacteria, they are usually unicellular, though they often grow in colonies large enough to see. They are the oldest and one of the largest and most important groups of bacteria on earth (Herrero and Flores, 2008). Absorbed light energy is used almost quantitatively for photosynthesis; about 3% of it is re-emitted as fluorescence. A complementarity relation between photosynthesis and fluorescence could potentially provide a handy tool for estimating photosynthetic yields of cyanobacteria, by an easy, non-destructive and relatively inexpensive Chl α fluorescence measurement (Papageorgiou and Govindjee, 2004). Copper is known to inhibit growth on both photosynthetic and nonphotosynthetic organisms. In fact the antimicrobial effect of copper (Borkow and Gabbay, 2009) has been attributed to mechanisms such as the generation of hydrogen peroxide with subsequent release of hydroxyl radicals in a Fenton-type reaction (Grass et al., 2011): þ
þ
2Cu þ 2H þ O2 →2Cu þ
2þ
Cu þ H2 O2 →Cu
2þ
þ H2 O2
ð1Þ
−
þ HO þ HO
ð2Þ
or to the reaction of thiol-groups in a cycle between Eqs. (1) and (3) (Magnani and Solioz, 2007): 2þ
2Cu
þ
þ
þ 2RSH→2Cu þ RS SR þ 2H :
ð3Þ
There have also been reports indicating that copper destroys lipopolysaccharides in bacterial cell walls altering potassium permeability that results in a fatal decrease of potassium concentration (Nan et al., 2008). Moreover, it has been suggested that copper can damage dehydratase enzymes containing iron-sulfur clusters by oxidizing or displacing the iron atom (Macomber and Imlay, 2009). Especially for Escherichia coli, it has recently been shown that copper has a bacteriolytic effect after binding with electronegative groups on the outer membrane of the bacteria. On the other hand for photosynthetic organisms, copper has been documented to affect photosynthesis, suppressing the activities and electron transport rate of Photosystem II at the secondary quinone acceptor for both terrestrial and aquatic organisms (Mohanty et al., 1989; Deng et al., 2014). In this work, we propose a universal, accurate, easy and versatile AST method for the evaluation of antibacterial properties of various surfaces on the bases of the emission of Chl α fluorescence by the gram(−) cyanobacterium Synechococcus sp. Wool fabric modified with copper ions was used as a model antibacterial surface in order to compare the proposed method and the AATCC 100-2004.
2. Materials and methods Commercial, plain weave, undyed, 100% wool fabric (weight, 155 g·m− 2) was used for the antimicrobial testing. Cu(NO3)2·5H2O was purchased from Sigma-Aldrich, Taufkirchen, Germany; Levantin LNB from BASF, Athens, Greece; acetic acid 100% from Merck, Whitehouse Station, U.S.A.; DCMU (3-(3′,4′-dichlorophenyl)-1,1-dimethylurea) from Sigma-Aldrich, Taufkirchen, Germany; and tetrachloroethylene from Panreac, Barcelona, Spain. Ampicillin, Kanamycin and Cycloheximide powders were purchased from Sigma-Aldrich. The unicellular cyanobacterium Synechococcus sp. PCC7942, obtained from the Collection Nationale de Cultures de Microorganismes (CNCM), Institut Pasteur, Paris, France, was used in all experiments. The cyanobacterial cells were cultured in BG11 (Rippka et al., 1979) that additionally contained 20 mM HEPES NaOH, pH 7.5 (basal medium). The cultures were incubated under white fluorescent light (100 μE·m−2·s−1), in an orbital incubator (Galenkamp INR-400) at 31 °C (Stamatakis and Papageorgiou, 2001). Culture growth was monitored in terms of concentration of Chl α, determined in N,N-dimethylformamide extracts of cell pellets (Moran, 1982). All chemicals were of analytical grade and were used without any further purification. 2.1. Test sample preparation The wool fabric was washed in a bath containing 1.0% w/v of the non-ionic washing agent Levantin LNB at a liquor-to-fabric ratio of 30 mL:1 g for 15 min at 40 °C. The pH was adjusted at 4.5 by the addition of acetic acid solution (10 g·L−1). The fabric was subsequently rinsed with warm, twice-distilled water (40 °C) for 3 min and then with cold, twice-distilled water (25 °C) for 9 min. The samples were then dried at room temperature. The wool/copper ion fabrics (WCFs) were prepared as reported previously (Heliopoulos et al., 2013). Specifically, Cu(NO3)2·5H2O was dissolved in twice-distilled water and stock solutions ranging from Cu2 + 100 to 5000 mg·L− 1 were prepared. A known quantity of dry wool fabric was placed in 100 mL of copper cation solution in 200-mL Erlenmeyer flasks under shaking at 180 rpm, at 25 °C. Known amounts of 0.5 M HNO3 were added until the pH measured on a Metrohm 744 pH meter was stabilized at 4.5. The samples were then left shaking for 24 h in a Julabo SW22 shaking bath, for complete equilibration and were consequently rinsed with cold twice-distilled water and were dried at room temperature. The total WCF Cu content of was quantitatively determined by measuring the remaining copper concentration on the liquid using a GBC GF
Fig. 1. PEA-fluorometer and measuring clip (PEA, Hansatech Instruments LTD, Norfolk, UK).
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Fig. 2. Mι evolution curves of cyanobacteria, cyanobacteria with DCMU and Chl α on raw wool.
300 Avanta atomic absorption spectrometer (AAS) using the following equation:
qm ¼
C in −C f V ms
ð4Þ
where qm (mg·g−1) is the sorbed copper, Cin (mg·L−1) the copper concentration in the initial solution, Cf (mg·L−1) the copper concentration after sorption, V (L) the volume of the solution and ms (g) the mass of the wool fabric sample used. 2.2. Proposed AST method basis A suitable quantity of cyanobacterial cells was harvested from the culture suspensions by centrifugation (5000 rpm, 5 min) and was resuspended in buffered BG11, so that the concentration of Chl α was 52.0 μ g·mL−1. Untreated and treated as above wool fabric samples were used as test specimens for the evaluation of the antibacterial activity. A drop of the abovementioned solution was pipetted (0.05 mL) on
each test specimen producing a stain of less than 3.0 mm diameter on the fabric. These samples were dark-adapted for 15 min in a suitable clip and Chl α fluorescence was measured using a PEA-fluorometer (PEA, Hansatech Instruments Ltd, Norfolk, UK) (Fig. 1). Upon excitation of a dark-adapted photosynthetic sample, Chl α fluorescence decays from a higher to a lower steady level within ns (Haworth et al., 1983; Holzwarth et al., 1985). This transiently steady fluorescence, the first recorded signal lasting a few μs, is labeled as O while its intensity is usually denoted as F0. Often, the Fo is also designated by the equivalent terms constant fluorescence, initial fluorescence, or dark-level fluorescence (Papageorgiou et al., 2007). The measuring area is circular with a diameter of 3.0 mm and the measurements were made at 25 °C. The actinic light (peak at 650 nm) is supplied by an array of three light emitting diodes and is focused on the sample surface to provide a homogeneous irradiation of the exposed area. The PEA fluorometer provides continuous excitation at 650 nm (3000 μE·m−2·s− 1; Δλ = 22 nm). It detects fluorescence at wavelengths above 700 nm (50% transmission at 720 nm) and records it continuously from 0.01 ms to 30 s with data acquisition every
Fig. 3. Mι evolution curves of cyanobacteria (control), cyanobacteria with Ampicillin, cyanobacteria with Kanamycin and cyanobacteria with Cycloeximide at different concentrations on raw wool.
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Fig. 4. Copper sorption isotherm on wool fabric samples.
0.01 ms for the first 0.3 ms, every 0.1 ms between 0.3 ms and 3 ms, every 1 ms between 3 ms and 30 ms, every 10 ms between 30 ms and 300 ms, every 100 ms between 300 ms and 3000 ms and every 1000 ms thereafter. In this study, raw fluorescence data collected during the first 10 s of illumination were used. The first reliable fluorescence value in the transient (at 20 μs) is used as the initial fluorescence value F0, for the samples that had been kept in darkness for 15 min. Cyanobacterial growth on wool and growth inhibition on antimicrobial samples was monitored by measurement of the Chl α fluorescence every 24 h for seven days. Throughout the course of the seven days the samples were kept in an incubator under white fluorescent light (100 μ E·m−2·s−1) at 31 °C. The % change (Mι) in Chl α fluorescence after i days of incubation is calculated using Eq. (5)
Mι ¼
F 0i − F 00 F 00
100
ð5Þ
where F 00 is the value of Chl α fluorescence of cyanobacteria at zero contact time and F 0i is the value of Chl α fluorescence of cyanobacteria after 1, 2, …, i days. The material's antibacterial action is represented by the Bacterial Protection Index (BPI) Π7, given by the equation: Π7 ¼
MU7 −MT7 M U7
100
ð6Þ
where MU7 , is the change in the cyanobacterial Chl α F0 value on the untreated sample after 7 days of incubation and MT7 , is the change in
the cyanobacterial Chl α F0 value on the treated sample after 7 days of incubation. In order to validate the method it is necessary to confirm that the measured quantity F0 is directly related to the bacterial population and that the test organism is sensitive to gram(−) antimicrobials. To this end, daily cyanobacterial Chl α monitoring was performed on untreated wool samples with: 1) control cyanobacterial culture, 2) cyanobacterial culture after the addition of 20 μM DCMU (Metz et al., 1986), 3) Chl α extracted from cyanobacteria (Moran, 1982) and 4) cyanobacterial culture after the addition of the known prokaryote (Kanamycin and Ampicillin) and one eukaryote (Cycloeximide) antibiotic incubated at concentrations ranging from 1 to 4 mg mL−1 for 6 h. DCMU is a very specific and sensitive inhibitor of photosynthesis. With Photosystem II (PS II) representing the first component of the photosynthetic electron transfer chain where water oxidation and reduction of the plastoquinone pool occur, DCMU blocks the electron transport by the displacement of the bound plastoquinone called Q B from its binding site on the D1 protein and stops the electron flow from where it is generated, in PS II, to plastoquinone. This interrupts the linear photosynthetic electron transport chain in photosynthesis, the cyclic photosynthetic pathway since electron shuttling is associated with proton pumping across the membrane into the lumen and thus blocks the ability of the plant to turn light energy into chemical energy (ATP and reductant potential). DCMU only blocks electron flow from PS II, and it has no effect on PS I or other reactions in photosynthesis, such as light absorption or carbon fixation in the Calvin cycle (Hsu et al., 1986; Trebst, 2007). Using DCMU will confirm whether or not Chl α fluorescence measurement is correlated to cell viability. On the other hand, extracted Chl α measurements over time will confirm that the presence of Chl α independent of growth does not affect the method's applicability. Furthermore, the use of antibiotics will confirm that the test organism is sensitive to antimicrobials targeting gram(−) bacteria and this sensitivity can be measured as proposed. Fig. 2 shows that Mι values remain constant for extracted Chl α as well as for cells in the presence of DCMU as their photosynthetic functions, hence their ability to proliferate, have been inhibited. It must be noted that even though poisoned with DCMU and cyanobacterial photosynthetic functions are inhibited leading to the eventual death of the organism, Chl α is not destroyed and continues fluorescing (for the time period of measurements). On the other hand an Mι increase only occurs for the culture of living cells. Such an increase is directly correlated with the cyanobacterial cell population. Only active bacteria have the ability to increase their photosynthetic devices via proliferation, with the number of active PS II photosynthetic centers increasing with time leading in turn to an increase in Mι values. The cyanobacterial population follows the typical bacterial growth curve with the small decrease of Mι during the first 24 h attributed to the adaptation of the bacteria to the new conditions during the lag phase. The exponential growth phase starts after the 1st day and ends on the 6th day where the stationary phase is reached. Furthermore, as seen in Fig. 3, antibiotic concentration is correlated to bacterial growth, as increasing antibiotic concentration results in increasing growth inhibition revealing that the test organism is sensitive to known non-photosynthetically active agents that affect gram(−)
Table 1 Π7 (Bacterial Protection Index) and R24 (AATCC 100-2004) values for the examined WCF samples. Initial Cu2+ concentration (mg·L−1) 100
200
300
500
750
1000
2000
3000
4000
5000
6.15 83.1 100
6.28 87 100
6.21 88.4 100
6.33 87.9 100
Cu2+ concentration on WCF (mg·g−1) Π7 R24
1.92 33 77.7
2.66 42.8 97
3.21 46.9 98
3.94 55.5 99.3
4.45 59.3 100
4.85 68.6 100
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bacteria. The eukaryote antibiotic (Cycloeximide) had no effect on cyanobacterial growth as expected. Thus, it can be concluded that the proposed method is applicable, as it was established that: a) the test organism has dose dependent sensitivity to known antimicrobials targeting gram(−) bacteria, b) the reduction of the cyanobacterial population has a proportional effect on Chl α fluorescence, c) Chl α fluorescence can be accurately and effectively monitored by the proposed instrument. However, in order to assess the quantitative accuracy of the proposed method, a way to mathematically quantify the relationship between bacterial reduction/Chl α measurements and antimicrobial concentration has to be determined in order to allow comparison with other already established AST methods. To this end, a wool fabric surface modified with copper ions was used as a model antibacterial surface, was evaluated using the proposed method and was compared to the results obtained by evaluation with the AATCC 100-2004 standard. Fig. 6. Mι evolution curves on raw wool and WCF samples.
3. Results and discussion 3.1. Wool copper content Copper uptake from the wool fabric samples is depicted in the sorption isotherm in Fig. 4. Increasing equilibrium concentration, results in higher copper content on the wool as indicated in Table 1. The copper ion has been previously reported to be sorbed onto the active anionic sites of wool fibers such as carboxylate, amine disulfide, and thiol (Balkose and Baltacioglu, 1992). These keratin sites are saturated at around 6 mg·g−1 copper content so increasing the equilibrium concentration over 2000 mg·L−1 has little or no effect on the copper uptake by the fabric. 3.2. Antimicrobial susceptibility testing The AATCC 100-2004 method was used as a standard for the evaluation of the antibacterial properties of the WCF samples (Fig. 5). The resulting R24 values can be seen in Table 1, and represent the % reduction of the bacterial colonies after inoculation on a growth medium in 24 h compared to the control. The antimicrobial action of copper is revealed as expected, with R24 values reaching 100 – classified as providing full antibacterial protection – obtained for all samples with copper concentrations over 750 mg·L−1 (copper content ≈ 4 mg·g−1). For comparison, the same samples were examined for their antibacterial properties using the proposed antimicrobial susceptibility testing method as described in the previous section. All experiments were performed three times using different cyanobacterial cell cultures. The F0 value of the cyanobacterial colony stain on each wool sample was recorded every 24 h for a period of 7 days and the corresponding Mι values were calculated using of Eq. (5).
The results are presented in Fig. 6, where it is obvious that a cyanobacterial population increase, results in a constant increase of the Mι value for the colony stain on the wool samples examined. Increasing copper concentration on the wool samples results in a decrease in bacterial growth depicted by the corresponding decrease in Mι values in the whole timescale under study. Synechococcus sp. as a photosynthetic organism is certainly affected by the photosynthesis inhibitory action of copper; however, as a gram(−) bacterium, it is also affected by copper via the mechanisms affecting all gram(−) bacteria. The antibacterial effect of copper on E. coli confirms its influence on the metabolic functions of gram(−) bacteria; thus, it is highly unlikely that the results presented in Fig. 6 represent the photosynthetic component of Synechococcus sp. vital functions. The Mι values obtained on the 7th day were used for the calculation of the Π7 factor since at that time the cyanobacterial growth reaches the stationary phase. The Π7 values calculated for each of the examined samples are presented in Table 1. As Π7 represents the level of antibacterial protection it can be deduced that increasing amount of copper on the samples under study results in increasing antibacterial protection, with the Π7 value being almost constant when copper concentration exceeds 2000 mg·L−1 (copper content ≈ 6 mg·g−1). In comparison, the same trend is observed for both methods for increasing the copper content on wool as increasing the copper content results in higher antibacterial protection. R24 values obtained by the AATCC 100-2004 standard method reach 100 for all samples with copper content over 4 mg·g−1, as no bacterial growth can be detected for copper content exceeding that amount. On the other hand, for the same samples, the introduced Π7 index increases with increasing copper content well over 4 mg·g−1 and up to 6 mg·g−1 where it remains constant indicating that the resolution of the proposed method is significantly higher in providing information on antibacterial
Fig. 5. Antibacterial effect of WCFs as per the AATCC 100-2004 standard method.
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activity well beyond the AATCC 100-2004 detection limit correlating to a value of Π7 ≈ 60. This can be attributed to the protocol of the AATCC 100-2004 itself which employs a number of dilutions before incubation and colony counts that can introduce negative errors in the results. Even though copper was used as a model antibacterial agent, this method could be used successfully with any other antibacterial that inhibits the growth of gram(−) bacteria such as the Synechococcus sp. The PEA fluorometer is an accurate, small, low cost and low maintenance instrument that can be used on various substrates with a suitable measuring clip making the method highly versatile and its application very cost effective compared to the existing instrumental methods. As the method involves neither the removal of the bacteria from the material under study, nor the addition of growth medium or special control of the experimental conditions, it can be used to evaluate the antibacterial properties of agents on substrates and under the conditions of their final use thus providing results closer to the material's intended application. 4. Conclusions The method proposed in this work, gives quantitative results and could be used to evaluate the antibacterial properties of any compound (organic, inorganic, natural or man-made) in real life conditions. The method can be applied on textile surfaces but could also be very easily modified and standardized in order to be applied in a variety of other materials or substrates. It provides very high resolution and can be applied simultaneously to more than one sample in real application conditions while remaining quick and cost effective. Contrary to qualitative methods, it does not require special conditions for bacterial growth while there are no compatibility issues of the antibacterial compound and the nutrient. The proposed method uses bacteria without the use of growth medium and uses an instrument for measurement resulting in more accurate readings in less than 10 s per sample; all measurements can be performed on the same sample, while bacteria are in contact with the antibacterial surface during the whole experimental procedure. Moreover, the proposed method can be applied under various conditions allowing materials to be tested under the conditions of the targeted application. References AATCC Technical Manual, Vol. 85, 2010, pp. 142–144. Clinical Laboratory Standards Institute, 2006. Performance standards for antimicrobial disk susceptibility tests, approved standard. 9th ed. CLSI Document M2-A9. 26:1 Clinical Laboratory Standards Institute, Wayne, PA. ASTM E2149-10, 2010. Standard Test Method for Determining the Antimicrobial Activity of Immobilized Antimicrobial Agents Under Dynamic Contact Conditions. ASTM International, West Conshohocken, PA www.astm.org. Balkose, D., Baltacioglu, H., 1992. Adsorption of heavy metal cations from aqueous solutions by wool fibers. J. Chem. Technol. Biotechnol. 54 (4), 393–397.
Bauer, A.W., Perry, D.M., Kirby, W.M.M., 1959. Single-disk antibiotic-sensitivity testing of staphylococci; an analysis of technique and results. A.M.A. Arch. Intern. Med. 104, 208–216. Borkow, G., Gabbay, J., 2009. Copper, an ancient remedy returning to fight microbial, fungal and viral infections. Curr. Chem. Biol. 8, 272–278. Deng, C., Pan, X., Wang, S., Zhang, D., 2014. Cu2+ inhibits photosystem II activities but enhances photosystem I quantum yield of Microcystis aeruginosa. Biol. Trace Elem. Res. 160, 268–275. Dickert, H., Machka, K., Braveny, I., 1981. The uses and limitations of disc diffusion in the antibiotic sensitivity testing of bacteria. Infection 9, 18–24. Gao, Y., Cranston, R., 2008. Recent advances in antimicrobial treatments of textiles. Text. Res. J. 78, 60. Gouveia, I.C., 2010. Nanobiotechnology: A New Strategy to Develop Non-toxic Antimicrobial Textiles. FORMATEX, pp. 407–414. Grass, G., Rensing, C., Solioz, M., 2011. Metallic copper as an antimicrobial surface. Appl. Environ. Microbiol. 77 (5), 1541–1547. Haworth, P., Karukstis, K.K., Sauer, K., 1983. Picosecond fluorescence kinetics in spinach chloroplasts at room temperature effects of phosphorylation. Biochim. Biophys. Acta 725, 261–271. Heliopoulos, N.S., Papageorgiou, S.K., Galeou, A., Favvas, E.P., Katsaros, F.K., Stamatakis, K., 2013. Effect of copper and copper alginate treatment on wool fabric. Study of textile and antibacterial properties. Surf. Coat. Technol. 235, 24–31. Herrero, A., Flores, E., 2008. The Cyanobacteria: Molecular Biology, Genomics and Evolution. Caister Academic Press, Norfolk, England. Holzwarth, A.R., Wendler, J., Haehnel, W., 1985. Time-resolved picosecond fluorescence spectra of the antenna chlorophylls in Chlorella vulgaris. Resolution of Photosystem I fluorescence. Biochim. Biophys. Acta 807, 155–167. Hsu, B.-D., Lee, J.-Y., Pan, R.-L., 1986. The two binding sites for DCMU in photosystem II. Biochem. Biophys. Res. Commun. 141, 682–688. ISO 20743, 2007. Textiles—Determination of Antibacterial Activity of Antibacterial Finished Products. 1st ed. . Jorgensen, J.H., Turnidge, J.D., 2007. Manual of Clinical Microbiology. 9th ed. ASM Press, Washington, D.C., pp. 1152–1172. Klancnik, A., Piskernik, S., Jersek, B., Mozina, S.S., 2010. Evaluation of diffusion and dilution methods to determine the antibacterial activity of plant extracts. J. Microbiol. Methods 81, 121–126. Macomber, L., Imlay, J., 2009. The iron–sulfur clusters of dehydratases are primary intracellular targets of copper toxicity. Proc. Natl. Acad. Sci. U. S. A. 106 (20), 8344–8349. Magnani, D., Solioz, M., 2007. How bacteria handle copper. In: Nies, D.H., Silver, S. (Eds.), Molecular Microbiology of Heavy Metals. Springer-Verlag, Berlin Heidelberg. Metz, J.G., Pakrasi, H.B., Seibert, M., Arntzer, C.J., 1986. Evidence for a dual function of the herbicide-binding D1 protein in photosystem II. FEBS Lett. 205, 269–274. Mohanty, N., Vass, I., Demeter, S., 1989. Copper toxicity affects photosystem II electron transport at the secondary quinone acceptor, QB1. Plant Physiol. 90, 175–179. Moran, P., 1982. Formulae for determination of chlorophyllous pigments extracted with N,N-dimethylformamide. Plant Physiol. 69, 1376–1381. Nan, L., Liu, Y., Lü, M., 2008. Study on antimicrobial mechanism of copper-bearing austenitic antibacterial stainless steel by atomic force microscopy. J. Mater. Sci. Mater. Med. 19, 3057–3062. Papageorgiou, G.C., Govindjee, 2004. Chlorophyll Fluorescence: A Signature of Photosynthesis. Springer, Dordrecht, The Netherlands. Papageorgiou, G.C., Tsimilli-Michael, M., Stamatakis, K., 2007. The fast and slow kinetics of chlorophyll α fluorescence induction in plants, algae and cyanobacteria: a viewpoint. Photosynth. Res. 94, 275–290. Rippka, R., Deruelles, J., Waterbury, J.B., Herdman, M., Stanier, R.T., 1979. Generic assignments, strain histories and properties of pure cultures of cyanobacteria. J. Gen. Microbiol. 111, 1–61. Stamatakis, K., Papageorgiou, G.C., 2001. The osmolality of the cell suspension regulates phycobilisome-to-photosystem I transfers in cyanobacteria. Biochim. Biophys. Acta 1506, 172–181. Trebst, A., 2007. Inhibitors in the functional dissection of the photosynthetic electron transport system. Photosynth. Res. 92, 217–224. Wilkins, T.D., Thiel, T., 1973. Modified broth-disk method for testing the antibiotic susceptibility of anaerobic bacteria. Am. Soc. Microbiol. 3, 350–356.
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Journal of Microbiological Methods journal homepage: www.elsevier.com/locate/jmicmeth
An in situ antimicrobial susceptibility testing method based on in vivo measurements of chlorophyll α fluorescence Nikolaos S. Heliopoulos a, Angeliki Galeou b, Sergios K. Papageorgiou a,⁎, Evangelos P. Favvas a, Fotios K. Katsaros a, Kostas Stamatakis b a b
Institute of Nanoscience and Nanotechnology, N.C.S.R. Demokritos, Terma Patriarchou Grigoriou & Neapoleos, Ag.Paraskevi Attikis, 15310 Athens, Greece Institute of Biosciences and Applications, N.C.S.R. Demokritos, Terma Patriarchou Grigoriou & Neapoleos, Ag. Paraskevi Attikis, 15310 Athens, Greece
a r t i c l e
i n f o
Article history: Received 24 December 2014 Received in revised form 11 March 2015 Accepted 12 March 2015 Available online 13 March 2015 Keywords: Antibacterial properties Chlorophyll fluorescence Cyanobacteria Textiles
a b s t r a c t Up to now antimicrobial susceptibility testing (AST) methods are indirect and generally involve the manual counting of bacterial colonies following the extraction of microorganisms from the surface under study and their inoculation in a separate procedure. In this work, an in situ, direct and instrumental method for the evaluation and assessment of antibacterial properties of materials and surfaces is proposed. Instead of indirectly determining antibacterial activity using the typical gram(−) test organisms with the subsequent manual colony count or inhibition zone measurement, the proposed procedure, employs photosynthetic gram(−) cyanobacteria deposited directly onto the surface under study and assesses cell proliferation and viability by a quick, accurate and reproducible instrumental chlorophyll fluorescence spectrophotometric technique. In contrast with existing methods of determination of antibacterial properties, it produces high resolution and quantitative results and is so versatile that it could be used to evaluate the antibacterial properties of any compound (organic, inorganic, natural or man-made) under any experimental conditions, depending on the targeted application. © 2015 Elsevier B.V. All rights reserved.
1. Introduction The increasing concern of human societies about hygiene and healthcare has intensified efforts for the development and production of materials and surfaces with antibacterial properties. In order to evaluate the bactericidal properties of such materials, several testing methods have been developed and are being implemented, both in research and in industrial applications. Based on the type of evaluation of antibacterial properties, methods can be classified into two major categories: qualitative and quantitative. The standard methods of the first category (Bauer et al., 1959; Jorgensen and Turnidge, 2007; Clinical Laboratory Standards Institute, 2006) evaluate antibacterial action based on the inhibition of growth around the sample under study. Even though sometimes the diameter of the zones of inhibition are measured and recorded they cannot be considered quantitative as the actual sizes of the colonies cannot be measured. In addition, this kind of approach has a number of drawbacks. Antibacterials that leach out into the aqueous agar matrix, such as silver ions, show better test performance than when they are fixed on the substrate or they are not water soluble, while antibacterials that react with the agar cannot be tested. Moreover, the relative ⁎ Corresponding author. E-mail address: [email protected] (S.K. Papageorgiou).
http://dx.doi.org/10.1016/j.mimet.2015.03.011 0167-7012/© 2015 Elsevier B.V. All rights reserved.
efficiency of different antimicrobial compounds cannot be evaluated as their rate of diffusion after plating the bacteria in agar differs greatly. Finally, diffusion as observed in the test does not represent antimicrobial behavior in actual use (Dickert et al., 1981; Wilkins and Thiel, 1973; Gouveia, 2010; Klancnik et al., 2010). On the other hand, quantitative methods are the most widely used and provide results based on the measurement of bacterial populations. These are inoculation and recovery methods that measure reduction in surviving cell populations in relation either to an initial inoculation level of bacteria or against an untreated control. However, quantitative methods have also their own limitations. Firstly, different methods must be used for porous, non-porous and uneven surfaces. Moreover, the bacterial growth conditions used are too favorable (e.g., rich nutrients in the inoculum and saturating moisture in the testing surfaces) to realistically simulate real life conditions (AATCC 100-2004; ASTM E2149-10). To date, very few quantitative studies have examined the antimicrobial effects under conditions of normal material use. For example ISO 20743:2007 that determines the antibacterial activity of antibacterial finished textile products based on the intended application and on the environment in which the textile product is to be to used, uses a lower number of nutrients but is time consuming and costly as it requires incubations, sterilization of the samples, humidity regulation of the test samples, filtering of test bacteria, printing of test bacteria with a special printing apparatus, etc. (Gao and Cranston, 2008).
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In general, quantitative methods are long, are expensive to apply and require time consuming work while a number of treatments must be performed to both the sample and the organisms used making them strongly dependent on the operator's interpretation and technique. Furthermore, modifications are often made according to standard protocols while often details of the assays are not fully disclosed. Recently, several commercial systems have been developed that provide conveniently prepared and formatted microdilution panels as well as instrumentation and automated reading of plates, intended to reduce technical errors and lengthy preparation times. Most of these automated antimicrobial susceptibility testing systems provide automated inoculation, reading and interpretation. Some examples of these include: Vitek System (bioMerieux, France), Walk-Away System (Dade International, Sacramento, Calif.), Sensititre ARIS (Trek Diagnostic Systems, East Grinstead, UK), Avantage Test System (Abbott Laboratories, Irving, Texas), Micronaut (Merlin, Bornheim-Hersel, Germany), Phoenix (BD Biosciences, Maryland) and many more. In these systems, although they are fast and convenient, the cost entailed in initial purchase, operation and maintenance of the machinery is very high for most laboratories. Cyanobacteria are prokaryotic organisms that perform oxygenic photosynthesis in a manner remarkably similar to green plants. Because they are bacteria, they are usually unicellular, though they often grow in colonies large enough to see. They are the oldest and one of the largest and most important groups of bacteria on earth (Herrero and Flores, 2008). Absorbed light energy is used almost quantitatively for photosynthesis; about 3% of it is re-emitted as fluorescence. A complementarity relation between photosynthesis and fluorescence could potentially provide a handy tool for estimating photosynthetic yields of cyanobacteria, by an easy, non-destructive and relatively inexpensive Chl α fluorescence measurement (Papageorgiou and Govindjee, 2004). Copper is known to inhibit growth on both photosynthetic and nonphotosynthetic organisms. In fact the antimicrobial effect of copper (Borkow and Gabbay, 2009) has been attributed to mechanisms such as the generation of hydrogen peroxide with subsequent release of hydroxyl radicals in a Fenton-type reaction (Grass et al., 2011): þ
þ
2Cu þ 2H þ O2 →2Cu þ
2þ
Cu þ H2 O2 →Cu
2þ
þ H2 O2
ð1Þ
−
þ HO þ HO
ð2Þ
or to the reaction of thiol-groups in a cycle between Eqs. (1) and (3) (Magnani and Solioz, 2007): 2þ
2Cu
þ
þ
þ 2RSH→2Cu þ RS SR þ 2H :
ð3Þ
There have also been reports indicating that copper destroys lipopolysaccharides in bacterial cell walls altering potassium permeability that results in a fatal decrease of potassium concentration (Nan et al., 2008). Moreover, it has been suggested that copper can damage dehydratase enzymes containing iron-sulfur clusters by oxidizing or displacing the iron atom (Macomber and Imlay, 2009). Especially for Escherichia coli, it has recently been shown that copper has a bacteriolytic effect after binding with electronegative groups on the outer membrane of the bacteria. On the other hand for photosynthetic organisms, copper has been documented to affect photosynthesis, suppressing the activities and electron transport rate of Photosystem II at the secondary quinone acceptor for both terrestrial and aquatic organisms (Mohanty et al., 1989; Deng et al., 2014). In this work, we propose a universal, accurate, easy and versatile AST method for the evaluation of antibacterial properties of various surfaces on the bases of the emission of Chl α fluorescence by the gram(−) cyanobacterium Synechococcus sp. Wool fabric modified with copper ions was used as a model antibacterial surface in order to compare the proposed method and the AATCC 100-2004.
2. Materials and methods Commercial, plain weave, undyed, 100% wool fabric (weight, 155 g·m− 2) was used for the antimicrobial testing. Cu(NO3)2·5H2O was purchased from Sigma-Aldrich, Taufkirchen, Germany; Levantin LNB from BASF, Athens, Greece; acetic acid 100% from Merck, Whitehouse Station, U.S.A.; DCMU (3-(3′,4′-dichlorophenyl)-1,1-dimethylurea) from Sigma-Aldrich, Taufkirchen, Germany; and tetrachloroethylene from Panreac, Barcelona, Spain. Ampicillin, Kanamycin and Cycloheximide powders were purchased from Sigma-Aldrich. The unicellular cyanobacterium Synechococcus sp. PCC7942, obtained from the Collection Nationale de Cultures de Microorganismes (CNCM), Institut Pasteur, Paris, France, was used in all experiments. The cyanobacterial cells were cultured in BG11 (Rippka et al., 1979) that additionally contained 20 mM HEPES NaOH, pH 7.5 (basal medium). The cultures were incubated under white fluorescent light (100 μE·m−2·s−1), in an orbital incubator (Galenkamp INR-400) at 31 °C (Stamatakis and Papageorgiou, 2001). Culture growth was monitored in terms of concentration of Chl α, determined in N,N-dimethylformamide extracts of cell pellets (Moran, 1982). All chemicals were of analytical grade and were used without any further purification. 2.1. Test sample preparation The wool fabric was washed in a bath containing 1.0% w/v of the non-ionic washing agent Levantin LNB at a liquor-to-fabric ratio of 30 mL:1 g for 15 min at 40 °C. The pH was adjusted at 4.5 by the addition of acetic acid solution (10 g·L−1). The fabric was subsequently rinsed with warm, twice-distilled water (40 °C) for 3 min and then with cold, twice-distilled water (25 °C) for 9 min. The samples were then dried at room temperature. The wool/copper ion fabrics (WCFs) were prepared as reported previously (Heliopoulos et al., 2013). Specifically, Cu(NO3)2·5H2O was dissolved in twice-distilled water and stock solutions ranging from Cu2 + 100 to 5000 mg·L− 1 were prepared. A known quantity of dry wool fabric was placed in 100 mL of copper cation solution in 200-mL Erlenmeyer flasks under shaking at 180 rpm, at 25 °C. Known amounts of 0.5 M HNO3 were added until the pH measured on a Metrohm 744 pH meter was stabilized at 4.5. The samples were then left shaking for 24 h in a Julabo SW22 shaking bath, for complete equilibration and were consequently rinsed with cold twice-distilled water and were dried at room temperature. The total WCF Cu content of was quantitatively determined by measuring the remaining copper concentration on the liquid using a GBC GF
Fig. 1. PEA-fluorometer and measuring clip (PEA, Hansatech Instruments LTD, Norfolk, UK).
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Fig. 2. Mι evolution curves of cyanobacteria, cyanobacteria with DCMU and Chl α on raw wool.
300 Avanta atomic absorption spectrometer (AAS) using the following equation:
qm ¼
C in −C f V ms
ð4Þ
where qm (mg·g−1) is the sorbed copper, Cin (mg·L−1) the copper concentration in the initial solution, Cf (mg·L−1) the copper concentration after sorption, V (L) the volume of the solution and ms (g) the mass of the wool fabric sample used. 2.2. Proposed AST method basis A suitable quantity of cyanobacterial cells was harvested from the culture suspensions by centrifugation (5000 rpm, 5 min) and was resuspended in buffered BG11, so that the concentration of Chl α was 52.0 μ g·mL−1. Untreated and treated as above wool fabric samples were used as test specimens for the evaluation of the antibacterial activity. A drop of the abovementioned solution was pipetted (0.05 mL) on
each test specimen producing a stain of less than 3.0 mm diameter on the fabric. These samples were dark-adapted for 15 min in a suitable clip and Chl α fluorescence was measured using a PEA-fluorometer (PEA, Hansatech Instruments Ltd, Norfolk, UK) (Fig. 1). Upon excitation of a dark-adapted photosynthetic sample, Chl α fluorescence decays from a higher to a lower steady level within ns (Haworth et al., 1983; Holzwarth et al., 1985). This transiently steady fluorescence, the first recorded signal lasting a few μs, is labeled as O while its intensity is usually denoted as F0. Often, the Fo is also designated by the equivalent terms constant fluorescence, initial fluorescence, or dark-level fluorescence (Papageorgiou et al., 2007). The measuring area is circular with a diameter of 3.0 mm and the measurements were made at 25 °C. The actinic light (peak at 650 nm) is supplied by an array of three light emitting diodes and is focused on the sample surface to provide a homogeneous irradiation of the exposed area. The PEA fluorometer provides continuous excitation at 650 nm (3000 μE·m−2·s− 1; Δλ = 22 nm). It detects fluorescence at wavelengths above 700 nm (50% transmission at 720 nm) and records it continuously from 0.01 ms to 30 s with data acquisition every
Fig. 3. Mι evolution curves of cyanobacteria (control), cyanobacteria with Ampicillin, cyanobacteria with Kanamycin and cyanobacteria with Cycloeximide at different concentrations on raw wool.
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Fig. 4. Copper sorption isotherm on wool fabric samples.
0.01 ms for the first 0.3 ms, every 0.1 ms between 0.3 ms and 3 ms, every 1 ms between 3 ms and 30 ms, every 10 ms between 30 ms and 300 ms, every 100 ms between 300 ms and 3000 ms and every 1000 ms thereafter. In this study, raw fluorescence data collected during the first 10 s of illumination were used. The first reliable fluorescence value in the transient (at 20 μs) is used as the initial fluorescence value F0, for the samples that had been kept in darkness for 15 min. Cyanobacterial growth on wool and growth inhibition on antimicrobial samples was monitored by measurement of the Chl α fluorescence every 24 h for seven days. Throughout the course of the seven days the samples were kept in an incubator under white fluorescent light (100 μ E·m−2·s−1) at 31 °C. The % change (Mι) in Chl α fluorescence after i days of incubation is calculated using Eq. (5)
Mι ¼
F 0i − F 00 F 00
100
ð5Þ
where F 00 is the value of Chl α fluorescence of cyanobacteria at zero contact time and F 0i is the value of Chl α fluorescence of cyanobacteria after 1, 2, …, i days. The material's antibacterial action is represented by the Bacterial Protection Index (BPI) Π7, given by the equation: Π7 ¼
MU7 −MT7 M U7
100
ð6Þ
where MU7 , is the change in the cyanobacterial Chl α F0 value on the untreated sample after 7 days of incubation and MT7 , is the change in
the cyanobacterial Chl α F0 value on the treated sample after 7 days of incubation. In order to validate the method it is necessary to confirm that the measured quantity F0 is directly related to the bacterial population and that the test organism is sensitive to gram(−) antimicrobials. To this end, daily cyanobacterial Chl α monitoring was performed on untreated wool samples with: 1) control cyanobacterial culture, 2) cyanobacterial culture after the addition of 20 μM DCMU (Metz et al., 1986), 3) Chl α extracted from cyanobacteria (Moran, 1982) and 4) cyanobacterial culture after the addition of the known prokaryote (Kanamycin and Ampicillin) and one eukaryote (Cycloeximide) antibiotic incubated at concentrations ranging from 1 to 4 mg mL−1 for 6 h. DCMU is a very specific and sensitive inhibitor of photosynthesis. With Photosystem II (PS II) representing the first component of the photosynthetic electron transfer chain where water oxidation and reduction of the plastoquinone pool occur, DCMU blocks the electron transport by the displacement of the bound plastoquinone called Q B from its binding site on the D1 protein and stops the electron flow from where it is generated, in PS II, to plastoquinone. This interrupts the linear photosynthetic electron transport chain in photosynthesis, the cyclic photosynthetic pathway since electron shuttling is associated with proton pumping across the membrane into the lumen and thus blocks the ability of the plant to turn light energy into chemical energy (ATP and reductant potential). DCMU only blocks electron flow from PS II, and it has no effect on PS I or other reactions in photosynthesis, such as light absorption or carbon fixation in the Calvin cycle (Hsu et al., 1986; Trebst, 2007). Using DCMU will confirm whether or not Chl α fluorescence measurement is correlated to cell viability. On the other hand, extracted Chl α measurements over time will confirm that the presence of Chl α independent of growth does not affect the method's applicability. Furthermore, the use of antibiotics will confirm that the test organism is sensitive to antimicrobials targeting gram(−) bacteria and this sensitivity can be measured as proposed. Fig. 2 shows that Mι values remain constant for extracted Chl α as well as for cells in the presence of DCMU as their photosynthetic functions, hence their ability to proliferate, have been inhibited. It must be noted that even though poisoned with DCMU and cyanobacterial photosynthetic functions are inhibited leading to the eventual death of the organism, Chl α is not destroyed and continues fluorescing (for the time period of measurements). On the other hand an Mι increase only occurs for the culture of living cells. Such an increase is directly correlated with the cyanobacterial cell population. Only active bacteria have the ability to increase their photosynthetic devices via proliferation, with the number of active PS II photosynthetic centers increasing with time leading in turn to an increase in Mι values. The cyanobacterial population follows the typical bacterial growth curve with the small decrease of Mι during the first 24 h attributed to the adaptation of the bacteria to the new conditions during the lag phase. The exponential growth phase starts after the 1st day and ends on the 6th day where the stationary phase is reached. Furthermore, as seen in Fig. 3, antibiotic concentration is correlated to bacterial growth, as increasing antibiotic concentration results in increasing growth inhibition revealing that the test organism is sensitive to known non-photosynthetically active agents that affect gram(−)
Table 1 Π7 (Bacterial Protection Index) and R24 (AATCC 100-2004) values for the examined WCF samples. Initial Cu2+ concentration (mg·L−1) 100
200
300
500
750
1000
2000
3000
4000
5000
6.15 83.1 100
6.28 87 100
6.21 88.4 100
6.33 87.9 100
Cu2+ concentration on WCF (mg·g−1) Π7 R24
1.92 33 77.7
2.66 42.8 97
3.21 46.9 98
3.94 55.5 99.3
4.45 59.3 100
4.85 68.6 100
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bacteria. The eukaryote antibiotic (Cycloeximide) had no effect on cyanobacterial growth as expected. Thus, it can be concluded that the proposed method is applicable, as it was established that: a) the test organism has dose dependent sensitivity to known antimicrobials targeting gram(−) bacteria, b) the reduction of the cyanobacterial population has a proportional effect on Chl α fluorescence, c) Chl α fluorescence can be accurately and effectively monitored by the proposed instrument. However, in order to assess the quantitative accuracy of the proposed method, a way to mathematically quantify the relationship between bacterial reduction/Chl α measurements and antimicrobial concentration has to be determined in order to allow comparison with other already established AST methods. To this end, a wool fabric surface modified with copper ions was used as a model antibacterial surface, was evaluated using the proposed method and was compared to the results obtained by evaluation with the AATCC 100-2004 standard. Fig. 6. Mι evolution curves on raw wool and WCF samples.
3. Results and discussion 3.1. Wool copper content Copper uptake from the wool fabric samples is depicted in the sorption isotherm in Fig. 4. Increasing equilibrium concentration, results in higher copper content on the wool as indicated in Table 1. The copper ion has been previously reported to be sorbed onto the active anionic sites of wool fibers such as carboxylate, amine disulfide, and thiol (Balkose and Baltacioglu, 1992). These keratin sites are saturated at around 6 mg·g−1 copper content so increasing the equilibrium concentration over 2000 mg·L−1 has little or no effect on the copper uptake by the fabric. 3.2. Antimicrobial susceptibility testing The AATCC 100-2004 method was used as a standard for the evaluation of the antibacterial properties of the WCF samples (Fig. 5). The resulting R24 values can be seen in Table 1, and represent the % reduction of the bacterial colonies after inoculation on a growth medium in 24 h compared to the control. The antimicrobial action of copper is revealed as expected, with R24 values reaching 100 – classified as providing full antibacterial protection – obtained for all samples with copper concentrations over 750 mg·L−1 (copper content ≈ 4 mg·g−1). For comparison, the same samples were examined for their antibacterial properties using the proposed antimicrobial susceptibility testing method as described in the previous section. All experiments were performed three times using different cyanobacterial cell cultures. The F0 value of the cyanobacterial colony stain on each wool sample was recorded every 24 h for a period of 7 days and the corresponding Mι values were calculated using of Eq. (5).
The results are presented in Fig. 6, where it is obvious that a cyanobacterial population increase, results in a constant increase of the Mι value for the colony stain on the wool samples examined. Increasing copper concentration on the wool samples results in a decrease in bacterial growth depicted by the corresponding decrease in Mι values in the whole timescale under study. Synechococcus sp. as a photosynthetic organism is certainly affected by the photosynthesis inhibitory action of copper; however, as a gram(−) bacterium, it is also affected by copper via the mechanisms affecting all gram(−) bacteria. The antibacterial effect of copper on E. coli confirms its influence on the metabolic functions of gram(−) bacteria; thus, it is highly unlikely that the results presented in Fig. 6 represent the photosynthetic component of Synechococcus sp. vital functions. The Mι values obtained on the 7th day were used for the calculation of the Π7 factor since at that time the cyanobacterial growth reaches the stationary phase. The Π7 values calculated for each of the examined samples are presented in Table 1. As Π7 represents the level of antibacterial protection it can be deduced that increasing amount of copper on the samples under study results in increasing antibacterial protection, with the Π7 value being almost constant when copper concentration exceeds 2000 mg·L−1 (copper content ≈ 6 mg·g−1). In comparison, the same trend is observed for both methods for increasing the copper content on wool as increasing the copper content results in higher antibacterial protection. R24 values obtained by the AATCC 100-2004 standard method reach 100 for all samples with copper content over 4 mg·g−1, as no bacterial growth can be detected for copper content exceeding that amount. On the other hand, for the same samples, the introduced Π7 index increases with increasing copper content well over 4 mg·g−1 and up to 6 mg·g−1 where it remains constant indicating that the resolution of the proposed method is significantly higher in providing information on antibacterial
Fig. 5. Antibacterial effect of WCFs as per the AATCC 100-2004 standard method.
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activity well beyond the AATCC 100-2004 detection limit correlating to a value of Π7 ≈ 60. This can be attributed to the protocol of the AATCC 100-2004 itself which employs a number of dilutions before incubation and colony counts that can introduce negative errors in the results. Even though copper was used as a model antibacterial agent, this method could be used successfully with any other antibacterial that inhibits the growth of gram(−) bacteria such as the Synechococcus sp. The PEA fluorometer is an accurate, small, low cost and low maintenance instrument that can be used on various substrates with a suitable measuring clip making the method highly versatile and its application very cost effective compared to the existing instrumental methods. As the method involves neither the removal of the bacteria from the material under study, nor the addition of growth medium or special control of the experimental conditions, it can be used to evaluate the antibacterial properties of agents on substrates and under the conditions of their final use thus providing results closer to the material's intended application. 4. Conclusions The method proposed in this work, gives quantitative results and could be used to evaluate the antibacterial properties of any compound (organic, inorganic, natural or man-made) in real life conditions. The method can be applied on textile surfaces but could also be very easily modified and standardized in order to be applied in a variety of other materials or substrates. It provides very high resolution and can be applied simultaneously to more than one sample in real application conditions while remaining quick and cost effective. Contrary to qualitative methods, it does not require special conditions for bacterial growth while there are no compatibility issues of the antibacterial compound and the nutrient. The proposed method uses bacteria without the use of growth medium and uses an instrument for measurement resulting in more accurate readings in less than 10 s per sample; all measurements can be performed on the same sample, while bacteria are in contact with the antibacterial surface during the whole experimental procedure. Moreover, the proposed method can be applied under various conditions allowing materials to be tested under the conditions of the targeted application. References AATCC Technical Manual, Vol. 85, 2010, pp. 142–144. Clinical Laboratory Standards Institute, 2006. Performance standards for antimicrobial disk susceptibility tests, approved standard. 9th ed. CLSI Document M2-A9. 26:1 Clinical Laboratory Standards Institute, Wayne, PA. ASTM E2149-10, 2010. Standard Test Method for Determining the Antimicrobial Activity of Immobilized Antimicrobial Agents Under Dynamic Contact Conditions. ASTM International, West Conshohocken, PA www.astm.org. Balkose, D., Baltacioglu, H., 1992. Adsorption of heavy metal cations from aqueous solutions by wool fibers. J. Chem. Technol. Biotechnol. 54 (4), 393–397.
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