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Growth inhibitory effect on bacteria of chitosan membranes regulated with deacetylation degree
Biochemical Engineering Journal 40 (2008) 485–491
Contents lists available at ScienceDirect
Biochemical Engineering Journal journal homepage: www.elsevier.com/locate/bej
Growth inhibitory effect on bacteria of chitosan membranes regulated with deacetylation degree Tomoki Takahashi a,1 , Masanao Imai a,∗ , Isao Suzuki a , Jun Sawai b a b
Course in Bioresource Utilization Sciences, Graduate School of Bioresource Sciences, Nihon University, 1866 Kameino, Fujisawa, Kanagawa 252-8510, Japan Department of Applied Bioscience, Faculty of Engineering, Kanagawa Institute of Technology, 1030 Shimo-ogino, Atsugi, Kanagawa 243-0292, Japan
a r t i c l e
i n f o
Article history: Received 30 August 2007 Received in revised form 6 February 2008 Accepted 8 February 2008 Keywords: Chitosan membrane Deacetylation degree Bactometer Conductimetric assay Antibacterial activity
a b s t r a c t Antibacterial activity of chitosan membranes was investigated by a conductimetric assay using a Bactometer. The purpose of this investigation was to produce a practical, high-performance membrane for separation engineering. The antibacterial activity of powdered chitosan membrane was evaluated by the minimal inhibitory concentration (MIC). The MIC for Escherichia coli was almost 200 (mg-chitosan/mlbacterial suspension), and for Staphylococcus aureus it was 40 (mg-chitosan/ml-bacterial suspension). Growth of the gram-positive sample (S. aureus) was more strongly inhibited by chitosan than the gramnegative sample (E. coli). This inhibitory effect was recognized as a bactericidal effect. Antibacterial activity was also observed and depended on the shape and the specific surface area of the powdered chitosan membrane. The influence of the deacetylation degree (DD) of the chitosan on inhibiting the growth of S. aureus was investigated by two methods: incubation using a mannitol salt agar medium, and a conductimetric assay. By both methods, chitosan with a higher DD successfully inhibited growth of S. aureus. Our findings regarding the dominant role of the DD of chitosan will be useful for designing long-life, hygienic, membrane-based processes. © 2008 Elsevier B.V. All rights reserved.
1. Introduction For years, material development focused on growth inhibition of bacteria was expected to lead to long-life, hygienic, membrane-based processes. The growth of microorganisms and the accumulation of colloids or organic compounds were major causes of membrane fouling, generally called “biofouling” [1]. Membrane fouling negatively influenced the permeation flux and some aspects of membrane performance (e.g., reduced salt rejection and elevated operational pressure [2–5]). The development of membrane materials with antibacterial activity is important from both the economic viewpoint and for the hygienic management of practical membrane processes. Chitosan produced from crustacean shells is an attractive material for reduc-
Abbreviations: BHI, brain heart infusion; BPU, Bactometer Processing Unit; CFU, colony-forming unit; DD, deacetylation degree; DT, detection time; MIC, minimal inhibitory concentration; MPCA, modified plate count agar; MW, molecular weight; PEG, polyethylene glycol; PVS-K, potassium polyvinyl sulfate; PVC, polyvinylidene chloride; SEM, scanning electron microscope. ∗ Corresponding author. Tel.: +81 466 84 3978; fax: +81 466 84 3978. E-mail address: [email protected] (M. Imai). 1 Present address: Department of Industrial Chemistry, Faculty of Engineering Division 1, Tokyo University of Science, 1-3 Kagurazaka, Shinjuku-ku, Tokyo 1628601, Japan. 1369-703X/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.bej.2008.02.009
ing biofouling. The authors previously reported a typical molecular characteristic of chitosan membranes, namely, the water permeability of these membranes when used to control the deacetylation degree (DD) [6]. An examination of the practical aspects of using chitosan membranes is necessary for developing membrane processing techniques. Chitosan and its resolvent inhibit the growth of mold with plant pathogenicity [7,8]. In contrast, chitin does not inhibit the growth of mold [9]. The effect of chitosan concentration on the growth of mold (Fusarium solani, Fusarium oxysporum) was investigated. Mold growth was completely inhibited by 0.1% chitosan. A higher DD has a stronger growth inhibitory effect on mold [10]. According to Uchida [10], chitosan inhibited bacterial growth not only of gram-negative bacteria (Escherichia coli, Pseudomonas aeruginosa) but also of gram-positive bacteria (Bacillus subtilis, Staphylococcus aureus). For example, after 4 days of incubation, bacterial growth was completely inhibited in broth containing a chitosan concentration of 0.02% at pH 6.0. Lower viscosity (i.e., lower molecular weight) chitosan had a stronger growth inhibitory effect on bacteria. According to No et al. [11], antibacterial activities of six chitosans and six chitosan oligomers with different molecular weights were examined against four gram-negative (including E. coli) and seven gram-positive bacteria (including S. aureus). Chitosans showed higher antibacterial activities than chitosan oligomers,
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T. Takahashi et al. / Biochemical Engineering Journal 40 (2008) 485–491
Nomenclature A CFU DD DT0 DTi MIC
additive volume of acetic anhydride (l/50 gchitosan solution of 2 wt%) colony-forming unit (−) deacetylation degree (%) referred from Ref. [20] detection time at an antibacterial agent concentration of zero (h) detection time at an antibacterial agent concentration of i (h) minimal inhibitory concentration (mg-chitosan/mlbacterial suspension)
and markedly inhibited growth of most of the bacteria tested. Chitosan generally showed stronger bactericidal effects towards gram-positive bacteria than gram-negative bacteria. In addition, antibacterial activity of chitosan was inversely affected by pH (in the range tested, pH 4.5–5.9), with higher activity at lower pH values. These results were reported within the context of potential applications of liquid-soluble chitosan. Various rapid examination techniques for antibacterial activity have been suggested, including turbidimetry [12], an ATP assay [13], calorimetry [14], an impedance assay [15], and conductimetric assays [16–19]. The present study employed the conductance method to evaluate the antibacterial activity of chitosan membranes; the method was based on the detection time of electric conductance. Bacteria are generally classified according to gram dyeing. E. coli 745 (gram-negative) and S. aureus 9779 (gram-positive) were employed to evaluate the antibacterial activity of chitosan. These bacteria are representative of the bacterial species that are important in public sanitation and food hygiene. The antibacterial activity of chitosan membranes as a solid system for regulating the deacetylation degree was investigated by a direct conductimetric assay using a Bactometer. In this paper, chitosan was examined as a solid, not as a solute in solution as in previous papers. This approach will contribute to the design of a biopolymer membrane with high performance and long life for use in practical separation processing applications.
Fig. 1. Effect of acetic anhydride on N-acetylation of chitosan. Acetic anhydride was added to 2 wt% chitosan solution (50 g). Original data are quoted from one of the authors’ previous publications [6].
During preparation of the membranes, the deacetylation degree (DD) can be regulated by changing the volume of acetic anhydride added (A (l)). Fig. 1 depicts the linear relationship between the DD and A. The correlation equation (1) has been reported previously by the authors [6]. DD = −0.1045A + 92.22
(1)
We successfully employed empirical equation (1) for controlling the DD of chitosan membranes. 2.2. Preparation of chitosan powder from membranes The water contained in a chitosan membrane was displaced by organic solvents in the order: ethanol aqueous solution, ethanol, and acetone. The membrane was then dried in a thermostatic oven at 333 K for 24 h. The dried membrane pieces were powdered by using a grinder (MILLSER IFM-77G, Iwatani Int. Co., Tokyo). The powder was classified into four classes according to the size of the particles: between 37 and 63 m, between 74 and 105 m, between 250 and 297 m, and between 420 and 500 m. These classified chitosan powders were stored in a glass vessel in a desiccator.
2. Materials and methods
2.3. Microorganisms
2.1. Preparation of chitosan membranes
E. coli 745 and S. aureus 9779 were obtained from the Tokyo Metropolitan Institute of Public Health. The bacterial strains were stored at 193 K. They were thawed and incubated in brain heart infusion broth (BHI) (Eiken Chemicals, Tokyo) at 310 K for 20 h. The cells were in stationary phase and were washed once with sterile saline (0.85%, w/v) and then resuspended in saline at approximately 103 CFU/ml.
Chitosan (low molecular weight, Sigma–Aldrich Japan K.K., Tokyo) and polyethylene glycol (MW7500, Wako, Osaka) were dissolved in 10% acetic acid. The chitosan solution was diluted to 2% (w/w) with methanol. Acetic anhydride (97.0%, Wako, Osaka) was added to the chitosan solution after vacuum filtration. The resultant casting solution was dried in a petri dish for 12 h at 333 K and subsequently gelled by immersing it in 4% NaOH. The resultant product was washed with distilled water, and the membrane was obtained by putting it into hot water (353 K). The DD of the chitosan membrane was analyzed by the colloidal titration method. The terminal point of each titration was clearly confirmed by a change in color from blue to claret. Potassium polyvinyl sulfate solution was used as the titrant, and toluidine blue was used as the indicator. Colloidal titration is a method of measuring free amino groups in a chitosan solution, and DD provides the molar percentage of glucosamine in the molecular chains of chitosan.
2.4. Conductimetric assay The antibacterial activity of the powder samples was judged by measuring the change in electrical conductivity with bacterial growth. In this paper, the direct conductimetric assay was employed [17]. The Bactometer Microbial Monitoring System, Model 64 (bioMerieux, Tokyo) was used for measuring the electric conductivity of broth (Fig. 2). The Bactometer consisted of a monitor, a computer, and a Bactometer Processing Unit (BPU, Fig. 2a). It has a dedicated module. One BPU can accommodate four modules. The standard module for the Bactometer was divided into
T. Takahashi et al. / Biochemical Engineering Journal 40 (2008) 485–491
Fig. 2. Schematic drawing of the Bactometer system: (a) system—(1) display, (2) computer, and (3) BPU; (b) module; (c) well—(4) cap, (5) partition, and (6) electrodes. This figure is reproduced from one of the authors’ previous publications [18].
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Fig. 3. Effect of chitosan particles (DD 92.2%, 74–105 m in size) on the growth of Escherichia coli.
2.6. Mannitol salt agar medium 16 individual wells, and was made of plastic (Fig. 2b). A pair of stainless-steel electrodes was inserted into the bottom of each well (Fig. 2c). The wells have a cap to prevent evaporation of the liquid phase and to keep out atmospheric dust and pollutants. Modified plate count agar (MPCA, yeast extract 20 g, dextrose 4 g, tryptone 20 g, agar 10 g, water 1000 ml: bioMerieux) was poured into the wells until the electrodes were covered (0.5 ml). After the agar solidified, sterile saline (0.1 ml), the chitosan particles (0–150 mg), and the bacterial suspension (0.1 ml) were introduced into the wells. The chitosan particles employed ranged in size from 74 to 105 m. The module was capped tightly and set in the Bactometer incubator. The conductivity change with bacterial growth was monitored during incubation at 308 K for 48 h.
Mannitol salt agar medium includes a large quantity of salt. Therefore, it restrains general bacterial growth. This medium can be used to selectively incubate S. aureus because the primary nutrient is mannitol. The color of the medium turns yellow due to the presence of phenol red when the medium is acidified by the metabolic products of S. aureus. The medium initially is a weak base and is reddish purple. The S. aureus culture fluid prepared in Section 2.3 was diluted with sterile saline and then suspended in saline at approximately 103 CFU/ml. This suspension (50 l) was applied to mannitol salt agar medium, and a 35-mm chitosan membrane was placed on top. The medium was incubated for 48 h at 308 K. 3. Results and discussion 3.1. Blank test for initial electrical conductivity
2.5. Judging bactericidal action or bacteriostasis After measurement by conductimetry, the samples were gathered from each well using a platinum wire and then incubated with BHI broth (10 ml) at 310 K for 24 h. The influence of contamination by antibacterial agents in the BHI broth can be ignored because the broth was diluted 1000-fold. After incubation, bactericidal action or bacteriostasis was determined from the turbidity of the fluid culture. The turbidity of the fluid culture indicates a bacteriostasis effect; the clear fluid culture indicates a bactericidal action.
Blank tests are necessary for judging the reproducibility and reliability of the conductimetric assay using the Bactometer employed in this paper. Table 1 presents the blank initial conductance values for various experimental well conditions in the modules. The electrical conductivity for the initial 48 h in bacteria-free systems was almost constant. Table 2 presents the conductivity for different DDs of chitosan. The conductivity did not depend on the DD of chitosan. These blank tests indicate that chitosan has no effect on conductivity. Therefore, elevated electrical conductivity would indicate an increasing bacterial population in the well.
Table 1 Blank test of conductance method Well conditions
Membrane
Concentration (mgsamp. /mlbact.soln. )
Bacteria
Initial value (S/cm)
Detection time (h)
Free Free Water 0.1 ml Water 0.1 ml NaCl 0.1 ml NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml
Free Free Free Free Free Free Free Free DD 92.2% DD 92.2%
– – – – – – – – 50 50
Free Free Free Free Free Free Free Free Free Free
1579 1661 1772 1792 3001 2997 2431 2498 2309 2389
– – – – – – – – – –
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Table 2 Blank test of conductance method Membrane
Concentration (mgsamp. /mlbact.soln. )
Bacteria
Initial value (S/cm)
Detection time (h)
Free Free DD 92.2% DD 92.2% DD 88.0% DD 88.0% DD 86.0% DD 81.8% DD 81.8% DD 71.3% DD 71.3% DD 60.9% DD 50.4% DD 50.4%
– – 100 100 100 100 100 100 100 100 100 100 100 100
Free Free Free Free Free Free Free Free Free Free Free Free Free Free
2107 2144 2102 2168 2054 2100 2069 2033 2051 2091 2092 1993 2229 2069
– – – – – – – – – – – – – –
3.2. Effect of chitosan concentration on growth of bacteria Figs. 3 and 4 depict the effect of chitosan concentration on the growth of E. coli and S. aureus. When the bacterial concentration in the medium reached 107 CFU/ml, the conductance change rate increased remarkably. The detection time (DT) was automatically indicated from the derivative of conductance change per unit time by the Bactometer. Table 3 shows the initial value of conductance and the detection time (DT) in Figs. 3 and 4. The DT was determined from the experimental data in Table 3. The reproducibility of the detection time was sufficient for detection of the DT. The authors employed the smaller value as a DT datum between the pair of DT presented in Table 3. DT0 means the detection time for the control medium. DTi means the detection time for the sample medium. A decrease in DT0 /DTi means enhancement of growth inhibitory effect. DT0 /DTi equal to zero indicates that bacterial growth is completely inhibited, and this concentration determines the minimal inhibitory concentration (MIC). In this paper, the MIC was determined from the intersection of the abscissa in Figs. 5 and 6. A smaller MIC indicates stronger antibacterial activity. Elevation of the conductance curve of E. coli (103 CFU/ml) in the chitosan-free control was observed at 7.7 h. Elevation for S. aureus started at 13.4 h, when the bacterial concentration reached 107 CFU/ml. The detection time (DT0 ) was determined based on when conductance began to increase.
Fig. 4. Effect of chitosan particles (DD 92.2%, 74–105 m in size) on the growth of Staphylococcus aureus.
Fig. 5. Effect of chitosan (DD 92.2%, 74–105 m in size) concentration on the growth of E. coli and S. aureus.
For the system with added chitosan, the DTi of S. aureus was prolonged by increasing the amount of chitosan added. The slope of the increasing electric conductivity became moderate. The prolongation of the detection time indicated that the addition of chitosan restrained bacterial growth. In contrast, the slopes of the conduc-
Fig. 6. Determination of the minimal inhibition concentration of chitosan on the growth of S. aureus. Data were classified according to the size distribution of the powders.
T. Takahashi et al. / Biochemical Engineering Journal 40 (2008) 485–491
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Table 3 Effect of chitosan particles on growth of bacteria Membrane
Concentration (mgsamp. /mlbact.soln. )
Bacteria
Initial value (S/cm)
Detection time (h)
DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2%
0 0 100 100 50 50 25 25 0 0 100 100 50 50 25 25
Escherichia coli E. coli E. coli E. coli E. coli E. coli E. coli E. coli Staphylococcus aureus S. aureus S. aureus S. aureus S. aureus S. aureus S. aureus S. aureus
2132 2152 2146 2177 2166 2149 2181 2172 2117 2144 2124 2103 2123 2128 2184 2155
7.9 7.7 9.4 9.6 8.8 9.0 7.7 7.8 14.1 13.4 – – – – 16.8 17.2
tance curves of E. coli increased in the same manner as the control regardless of the chitosan concentration. Chitosan therefore has a growth inhibitory effect that depends on the bacterial species. Fig. 5 shows a regression analysis for the evaluation of the MIC. The MIC was almost 200 (mg-chitosan/ml-bacterial suspension) for E. coli and 40 (mg-chitosan/ml-bacterial suspension) for S. aureus. Chitosan evidently has a greater growth inhibitory effect on S. aureus than on E. coli. After the conductance measurement, the activity of chitosan on the bacteria was judged to be bactericidal or bacteriostasis by BHI broth incubation. The growth inhibitory effect on bacteria of chitosan indicated as the bactericidal action on E. coli and S. aureus. Physicochemical interaction between the bacteria and the chitosan polymer caused bactericidal action towards the bacteria. Differences in the antibacterial activity of the cationic chitosan polymer were induced by differences between the gram-negative (E. coli), and gram-positive samples (S. aureus).
A decisive structural difference between gram-negative and gram-positive bacteria can be recognized from the molecular structure of the outer cell membrane. In gram-negative bacteria, phospholipids, lipopolysaccharides and proteins exist in the outer membrane. Therefore, electric interaction hardly influences the inner membrane. In contrast, in gram-positive bacteria, the electric charge of chitosan acts directly on the inner membrane of the cell. The electric interaction between chitosan and the cell membrane was one factor inducing bacterial growth inhibition, cell destruction, and activity depression.
3.3. Effect of particle size and particle shape on bacterial growth inhibition In Section 3.2, the growth inhibitory effect of chitosan on S. aureus appeared stronger than that on E. coli. Therefore, further investigations were focused primarily on S. aureus.
Fig. 7. Scanning electron microscope photographs of the chitosan particles. The particle size was determined by crushing and sieving dried chitosan membrane (DD 92%): (a) 420–500 m; (b) 250–297 m; (c) 74–105 m; (d) 37–63 m.
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Fig. 8. Optical photographs of mannitol salt agar with chitosan membrane added: (a) DD 92.2%; (b) DD 90.1%; (c) DD 88.0%; (d) DD 83.9%; (e) DD 79.7%; (f) DD 75.5%; (g) PVC; (h) control.
The influence of particle size and shape of powdered chitosan membrane on S. aureus was examined. As shown in Fig. 6, decreased particle size improved antibacterial activity. However, the activity of 37- to 63-m particles was less than that of 74- to 105-m particles. Therefore, there is an optimal particle size for antibacterial activity. The morphology of each particle size is depicted in Fig. 7. Powdered chitosan membrane in the range 74–500 m looks like a flake or board, whereas in the range 37–63 m, it looks like a sphere. The antibacterial activity of the powdered chitosan membrane depends on the shape as well as the specific surface area. Quantitative characterization of sharp aspect on growth inhibition of S. aureus will be necessary for future discussions. 3.4. Effect of deacetylation degree of chitosan on growth inhibition of S. aureus The influence of the DD of a chitosan membrane on growth inhibition of S. aureus was investigated by using mannitol salt agar medium; a photograph of the petri dish is shown in Fig. 8. A CFU count of around 150 was observed in the control (chitosan-free) medium, and the medium was yellow. Medium containing polyvinylidene chloride (PVC) was also yellow. PVC was employed as an inert material which has no antibacte-
rial activity. Yellow-colored medium indicates the growth of S. aureus. In contrast, the medium apparently remained red in the system containing chitosan membrane. The trend was clearer as DD increased from 83.9% to 92.2%. Higher DD was especially successful in inhibiting the growth of S. aureus. The number of colonies on the membrane was fewer than that of the circumference area of membrane. The growth inhibitory effect was consistent based both on the color of the medium and the number of colonies on the membrane. The above discussion of bacterial growth inhibition focuses mainly on color changes occurring in the medium. Fig. 9 depicts the MIC of each DD for S. aureus. The MIC for a DD of 92% was about 40 (mg-chitosan/ml-bacterial suspension), and the MIC for a DD of 86% was about 100 (mg-chitosan/ml-bacterial suspension). The antibacterial activity of chitosan towards S. aureus was enhanced with increasing DD. The growth inhibitory effect was not observed at a DD of 82% or a DD of 50%, for reasons that presently are not clear. Other major factors affecting growth inhibition must be investigated in the future. 4. Conclusion Direct conductimetric assays using a Bactometer clearly demonstrated the antibacterial activity of chitosan membranes differing in their deacetylation degree. The influence of chitosan membranes on E. coli and S. aureus, which are food hygiene index bacteria, was investigated. Chitosan inhibits the growth of S. aureus more strongly than it inhibits the growth of E. coli. Chitosan inhibited the growth of gram-positive samples more strongly than that of gram-negative samples, and the effect was bactericidal. At higher DD, chitosan membrane sufficiently inhibits bacterial growth. The shape of the chitosan membrane particles are apparently a major factor influencing antibacterial activity. The shape of the powder particles and their antibacterial activity must be quantitatively correlated in the future. It is anticipated that our findings regarding the dominant role of DD will benefit the production of chitosan membranes for use in a variety of applications. Acknowledgements
Fig. 9. Determination of minimal inhibition concentration of chitosan on the growth of S. aureus. Data were classified according to the deacetylation degree of the particles.
The authors express sincere gratitude to Professor Mikio Kikuchi of the Kanagawa Institute of Technology for conducting the bacterial experiments and for useful discussions. The authors sincerely appreciate the Keyence Corporation for their photographing technique through the Scanning Electron Microscope Model VE-7800.
T. Takahashi et al. / Biochemical Engineering Journal 40 (2008) 485–491
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[12] H.L. Jorgensen, E. Schuz, Turbidimetric measurement as a rapid method for the determination on the bacteriological quality of minced meat, Int. J. Food Microbiol. 2 (1985) 177–183. [13] R. Bossuyt, Determination of bacteriological quality of raw milk by ATP assay technique, Milchwissenschaft 36 (1981) 257–262. [14] F. Okada, A. Kobayashi, N. Fujiwara, K. Takahashi, Microbial calorimetry of supported cultures and its application to the study antibacterial action, Biocontrol Sci. 3 (1998) 79–85. [15] P. Silly, S. Forsythe, Impedance microbiology—a rapid change for microbiologists, J. Appl. Bacteriol. 80 (1996) 233–243. [16] D. Hardy, S.J. Kraeger, S.W. Dufour, P. Cady, Rapid detection of microbial contamination in frozen vegetables by automated impedance measurements, Appl. Environ. Microbiol. 34 (1977) 14–17. [17] J. Sawai, H. Igarashi, A. Hashimoto, T. Kokugan, M. Shimizu, Evaluation of growth inhibitory effect of ceramic powder slurry on bacteria by conductance method, J. Chem. Eng. Jpn. 28 (1995) 288–293. [18] J. Sawai, Y. Maekawa, R. Doi, H. Kojima, H. Igarashi, Indirect conductimetric assay for detection of bacteria using a bactometer, J. Antibact. Antifung. Agents 28 (2000) 617–622. [19] J. Sawai, Quantitative evaluation of antibacterial activities of metallic oxide powders (ZnO, MgO and CaO) by conductimetric assay, J. Microbiol. Methods 54 (2003) 177–182. [20] T. Takahashi, M. Imai, I. Suzuki, High-potential molecular properties of chitosan and reaction conditions for removal of p-quinone from the aqueous phase, Biochem. Eng. J. 25 (2005) 7–13.
Contents lists available at ScienceDirect
Biochemical Engineering Journal journal homepage: www.elsevier.com/locate/bej
Growth inhibitory effect on bacteria of chitosan membranes regulated with deacetylation degree Tomoki Takahashi a,1 , Masanao Imai a,∗ , Isao Suzuki a , Jun Sawai b a b
Course in Bioresource Utilization Sciences, Graduate School of Bioresource Sciences, Nihon University, 1866 Kameino, Fujisawa, Kanagawa 252-8510, Japan Department of Applied Bioscience, Faculty of Engineering, Kanagawa Institute of Technology, 1030 Shimo-ogino, Atsugi, Kanagawa 243-0292, Japan
a r t i c l e
i n f o
Article history: Received 30 August 2007 Received in revised form 6 February 2008 Accepted 8 February 2008 Keywords: Chitosan membrane Deacetylation degree Bactometer Conductimetric assay Antibacterial activity
a b s t r a c t Antibacterial activity of chitosan membranes was investigated by a conductimetric assay using a Bactometer. The purpose of this investigation was to produce a practical, high-performance membrane for separation engineering. The antibacterial activity of powdered chitosan membrane was evaluated by the minimal inhibitory concentration (MIC). The MIC for Escherichia coli was almost 200 (mg-chitosan/mlbacterial suspension), and for Staphylococcus aureus it was 40 (mg-chitosan/ml-bacterial suspension). Growth of the gram-positive sample (S. aureus) was more strongly inhibited by chitosan than the gramnegative sample (E. coli). This inhibitory effect was recognized as a bactericidal effect. Antibacterial activity was also observed and depended on the shape and the specific surface area of the powdered chitosan membrane. The influence of the deacetylation degree (DD) of the chitosan on inhibiting the growth of S. aureus was investigated by two methods: incubation using a mannitol salt agar medium, and a conductimetric assay. By both methods, chitosan with a higher DD successfully inhibited growth of S. aureus. Our findings regarding the dominant role of the DD of chitosan will be useful for designing long-life, hygienic, membrane-based processes. © 2008 Elsevier B.V. All rights reserved.
1. Introduction For years, material development focused on growth inhibition of bacteria was expected to lead to long-life, hygienic, membrane-based processes. The growth of microorganisms and the accumulation of colloids or organic compounds were major causes of membrane fouling, generally called “biofouling” [1]. Membrane fouling negatively influenced the permeation flux and some aspects of membrane performance (e.g., reduced salt rejection and elevated operational pressure [2–5]). The development of membrane materials with antibacterial activity is important from both the economic viewpoint and for the hygienic management of practical membrane processes. Chitosan produced from crustacean shells is an attractive material for reduc-
Abbreviations: BHI, brain heart infusion; BPU, Bactometer Processing Unit; CFU, colony-forming unit; DD, deacetylation degree; DT, detection time; MIC, minimal inhibitory concentration; MPCA, modified plate count agar; MW, molecular weight; PEG, polyethylene glycol; PVS-K, potassium polyvinyl sulfate; PVC, polyvinylidene chloride; SEM, scanning electron microscope. ∗ Corresponding author. Tel.: +81 466 84 3978; fax: +81 466 84 3978. E-mail address: [email protected] (M. Imai). 1 Present address: Department of Industrial Chemistry, Faculty of Engineering Division 1, Tokyo University of Science, 1-3 Kagurazaka, Shinjuku-ku, Tokyo 1628601, Japan. 1369-703X/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.bej.2008.02.009
ing biofouling. The authors previously reported a typical molecular characteristic of chitosan membranes, namely, the water permeability of these membranes when used to control the deacetylation degree (DD) [6]. An examination of the practical aspects of using chitosan membranes is necessary for developing membrane processing techniques. Chitosan and its resolvent inhibit the growth of mold with plant pathogenicity [7,8]. In contrast, chitin does not inhibit the growth of mold [9]. The effect of chitosan concentration on the growth of mold (Fusarium solani, Fusarium oxysporum) was investigated. Mold growth was completely inhibited by 0.1% chitosan. A higher DD has a stronger growth inhibitory effect on mold [10]. According to Uchida [10], chitosan inhibited bacterial growth not only of gram-negative bacteria (Escherichia coli, Pseudomonas aeruginosa) but also of gram-positive bacteria (Bacillus subtilis, Staphylococcus aureus). For example, after 4 days of incubation, bacterial growth was completely inhibited in broth containing a chitosan concentration of 0.02% at pH 6.0. Lower viscosity (i.e., lower molecular weight) chitosan had a stronger growth inhibitory effect on bacteria. According to No et al. [11], antibacterial activities of six chitosans and six chitosan oligomers with different molecular weights were examined against four gram-negative (including E. coli) and seven gram-positive bacteria (including S. aureus). Chitosans showed higher antibacterial activities than chitosan oligomers,
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T. Takahashi et al. / Biochemical Engineering Journal 40 (2008) 485–491
Nomenclature A CFU DD DT0 DTi MIC
additive volume of acetic anhydride (l/50 gchitosan solution of 2 wt%) colony-forming unit (−) deacetylation degree (%) referred from Ref. [20] detection time at an antibacterial agent concentration of zero (h) detection time at an antibacterial agent concentration of i (h) minimal inhibitory concentration (mg-chitosan/mlbacterial suspension)
and markedly inhibited growth of most of the bacteria tested. Chitosan generally showed stronger bactericidal effects towards gram-positive bacteria than gram-negative bacteria. In addition, antibacterial activity of chitosan was inversely affected by pH (in the range tested, pH 4.5–5.9), with higher activity at lower pH values. These results were reported within the context of potential applications of liquid-soluble chitosan. Various rapid examination techniques for antibacterial activity have been suggested, including turbidimetry [12], an ATP assay [13], calorimetry [14], an impedance assay [15], and conductimetric assays [16–19]. The present study employed the conductance method to evaluate the antibacterial activity of chitosan membranes; the method was based on the detection time of electric conductance. Bacteria are generally classified according to gram dyeing. E. coli 745 (gram-negative) and S. aureus 9779 (gram-positive) were employed to evaluate the antibacterial activity of chitosan. These bacteria are representative of the bacterial species that are important in public sanitation and food hygiene. The antibacterial activity of chitosan membranes as a solid system for regulating the deacetylation degree was investigated by a direct conductimetric assay using a Bactometer. In this paper, chitosan was examined as a solid, not as a solute in solution as in previous papers. This approach will contribute to the design of a biopolymer membrane with high performance and long life for use in practical separation processing applications.
Fig. 1. Effect of acetic anhydride on N-acetylation of chitosan. Acetic anhydride was added to 2 wt% chitosan solution (50 g). Original data are quoted from one of the authors’ previous publications [6].
During preparation of the membranes, the deacetylation degree (DD) can be regulated by changing the volume of acetic anhydride added (A (l)). Fig. 1 depicts the linear relationship between the DD and A. The correlation equation (1) has been reported previously by the authors [6]. DD = −0.1045A + 92.22
(1)
We successfully employed empirical equation (1) for controlling the DD of chitosan membranes. 2.2. Preparation of chitosan powder from membranes The water contained in a chitosan membrane was displaced by organic solvents in the order: ethanol aqueous solution, ethanol, and acetone. The membrane was then dried in a thermostatic oven at 333 K for 24 h. The dried membrane pieces were powdered by using a grinder (MILLSER IFM-77G, Iwatani Int. Co., Tokyo). The powder was classified into four classes according to the size of the particles: between 37 and 63 m, between 74 and 105 m, between 250 and 297 m, and between 420 and 500 m. These classified chitosan powders were stored in a glass vessel in a desiccator.
2. Materials and methods
2.3. Microorganisms
2.1. Preparation of chitosan membranes
E. coli 745 and S. aureus 9779 were obtained from the Tokyo Metropolitan Institute of Public Health. The bacterial strains were stored at 193 K. They were thawed and incubated in brain heart infusion broth (BHI) (Eiken Chemicals, Tokyo) at 310 K for 20 h. The cells were in stationary phase and were washed once with sterile saline (0.85%, w/v) and then resuspended in saline at approximately 103 CFU/ml.
Chitosan (low molecular weight, Sigma–Aldrich Japan K.K., Tokyo) and polyethylene glycol (MW7500, Wako, Osaka) were dissolved in 10% acetic acid. The chitosan solution was diluted to 2% (w/w) with methanol. Acetic anhydride (97.0%, Wako, Osaka) was added to the chitosan solution after vacuum filtration. The resultant casting solution was dried in a petri dish for 12 h at 333 K and subsequently gelled by immersing it in 4% NaOH. The resultant product was washed with distilled water, and the membrane was obtained by putting it into hot water (353 K). The DD of the chitosan membrane was analyzed by the colloidal titration method. The terminal point of each titration was clearly confirmed by a change in color from blue to claret. Potassium polyvinyl sulfate solution was used as the titrant, and toluidine blue was used as the indicator. Colloidal titration is a method of measuring free amino groups in a chitosan solution, and DD provides the molar percentage of glucosamine in the molecular chains of chitosan.
2.4. Conductimetric assay The antibacterial activity of the powder samples was judged by measuring the change in electrical conductivity with bacterial growth. In this paper, the direct conductimetric assay was employed [17]. The Bactometer Microbial Monitoring System, Model 64 (bioMerieux, Tokyo) was used for measuring the electric conductivity of broth (Fig. 2). The Bactometer consisted of a monitor, a computer, and a Bactometer Processing Unit (BPU, Fig. 2a). It has a dedicated module. One BPU can accommodate four modules. The standard module for the Bactometer was divided into
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Fig. 2. Schematic drawing of the Bactometer system: (a) system—(1) display, (2) computer, and (3) BPU; (b) module; (c) well—(4) cap, (5) partition, and (6) electrodes. This figure is reproduced from one of the authors’ previous publications [18].
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Fig. 3. Effect of chitosan particles (DD 92.2%, 74–105 m in size) on the growth of Escherichia coli.
2.6. Mannitol salt agar medium 16 individual wells, and was made of plastic (Fig. 2b). A pair of stainless-steel electrodes was inserted into the bottom of each well (Fig. 2c). The wells have a cap to prevent evaporation of the liquid phase and to keep out atmospheric dust and pollutants. Modified plate count agar (MPCA, yeast extract 20 g, dextrose 4 g, tryptone 20 g, agar 10 g, water 1000 ml: bioMerieux) was poured into the wells until the electrodes were covered (0.5 ml). After the agar solidified, sterile saline (0.1 ml), the chitosan particles (0–150 mg), and the bacterial suspension (0.1 ml) were introduced into the wells. The chitosan particles employed ranged in size from 74 to 105 m. The module was capped tightly and set in the Bactometer incubator. The conductivity change with bacterial growth was monitored during incubation at 308 K for 48 h.
Mannitol salt agar medium includes a large quantity of salt. Therefore, it restrains general bacterial growth. This medium can be used to selectively incubate S. aureus because the primary nutrient is mannitol. The color of the medium turns yellow due to the presence of phenol red when the medium is acidified by the metabolic products of S. aureus. The medium initially is a weak base and is reddish purple. The S. aureus culture fluid prepared in Section 2.3 was diluted with sterile saline and then suspended in saline at approximately 103 CFU/ml. This suspension (50 l) was applied to mannitol salt agar medium, and a 35-mm chitosan membrane was placed on top. The medium was incubated for 48 h at 308 K. 3. Results and discussion 3.1. Blank test for initial electrical conductivity
2.5. Judging bactericidal action or bacteriostasis After measurement by conductimetry, the samples were gathered from each well using a platinum wire and then incubated with BHI broth (10 ml) at 310 K for 24 h. The influence of contamination by antibacterial agents in the BHI broth can be ignored because the broth was diluted 1000-fold. After incubation, bactericidal action or bacteriostasis was determined from the turbidity of the fluid culture. The turbidity of the fluid culture indicates a bacteriostasis effect; the clear fluid culture indicates a bactericidal action.
Blank tests are necessary for judging the reproducibility and reliability of the conductimetric assay using the Bactometer employed in this paper. Table 1 presents the blank initial conductance values for various experimental well conditions in the modules. The electrical conductivity for the initial 48 h in bacteria-free systems was almost constant. Table 2 presents the conductivity for different DDs of chitosan. The conductivity did not depend on the DD of chitosan. These blank tests indicate that chitosan has no effect on conductivity. Therefore, elevated electrical conductivity would indicate an increasing bacterial population in the well.
Table 1 Blank test of conductance method Well conditions
Membrane
Concentration (mgsamp. /mlbact.soln. )
Bacteria
Initial value (S/cm)
Detection time (h)
Free Free Water 0.1 ml Water 0.1 ml NaCl 0.1 ml NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml Water 0.1 ml + NaCl 0.1 ml
Free Free Free Free Free Free Free Free DD 92.2% DD 92.2%
– – – – – – – – 50 50
Free Free Free Free Free Free Free Free Free Free
1579 1661 1772 1792 3001 2997 2431 2498 2309 2389
– – – – – – – – – –
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Table 2 Blank test of conductance method Membrane
Concentration (mgsamp. /mlbact.soln. )
Bacteria
Initial value (S/cm)
Detection time (h)
Free Free DD 92.2% DD 92.2% DD 88.0% DD 88.0% DD 86.0% DD 81.8% DD 81.8% DD 71.3% DD 71.3% DD 60.9% DD 50.4% DD 50.4%
– – 100 100 100 100 100 100 100 100 100 100 100 100
Free Free Free Free Free Free Free Free Free Free Free Free Free Free
2107 2144 2102 2168 2054 2100 2069 2033 2051 2091 2092 1993 2229 2069
– – – – – – – – – – – – – –
3.2. Effect of chitosan concentration on growth of bacteria Figs. 3 and 4 depict the effect of chitosan concentration on the growth of E. coli and S. aureus. When the bacterial concentration in the medium reached 107 CFU/ml, the conductance change rate increased remarkably. The detection time (DT) was automatically indicated from the derivative of conductance change per unit time by the Bactometer. Table 3 shows the initial value of conductance and the detection time (DT) in Figs. 3 and 4. The DT was determined from the experimental data in Table 3. The reproducibility of the detection time was sufficient for detection of the DT. The authors employed the smaller value as a DT datum between the pair of DT presented in Table 3. DT0 means the detection time for the control medium. DTi means the detection time for the sample medium. A decrease in DT0 /DTi means enhancement of growth inhibitory effect. DT0 /DTi equal to zero indicates that bacterial growth is completely inhibited, and this concentration determines the minimal inhibitory concentration (MIC). In this paper, the MIC was determined from the intersection of the abscissa in Figs. 5 and 6. A smaller MIC indicates stronger antibacterial activity. Elevation of the conductance curve of E. coli (103 CFU/ml) in the chitosan-free control was observed at 7.7 h. Elevation for S. aureus started at 13.4 h, when the bacterial concentration reached 107 CFU/ml. The detection time (DT0 ) was determined based on when conductance began to increase.
Fig. 4. Effect of chitosan particles (DD 92.2%, 74–105 m in size) on the growth of Staphylococcus aureus.
Fig. 5. Effect of chitosan (DD 92.2%, 74–105 m in size) concentration on the growth of E. coli and S. aureus.
For the system with added chitosan, the DTi of S. aureus was prolonged by increasing the amount of chitosan added. The slope of the increasing electric conductivity became moderate. The prolongation of the detection time indicated that the addition of chitosan restrained bacterial growth. In contrast, the slopes of the conduc-
Fig. 6. Determination of the minimal inhibition concentration of chitosan on the growth of S. aureus. Data were classified according to the size distribution of the powders.
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Table 3 Effect of chitosan particles on growth of bacteria Membrane
Concentration (mgsamp. /mlbact.soln. )
Bacteria
Initial value (S/cm)
Detection time (h)
DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2% DD 92.2%
0 0 100 100 50 50 25 25 0 0 100 100 50 50 25 25
Escherichia coli E. coli E. coli E. coli E. coli E. coli E. coli E. coli Staphylococcus aureus S. aureus S. aureus S. aureus S. aureus S. aureus S. aureus S. aureus
2132 2152 2146 2177 2166 2149 2181 2172 2117 2144 2124 2103 2123 2128 2184 2155
7.9 7.7 9.4 9.6 8.8 9.0 7.7 7.8 14.1 13.4 – – – – 16.8 17.2
tance curves of E. coli increased in the same manner as the control regardless of the chitosan concentration. Chitosan therefore has a growth inhibitory effect that depends on the bacterial species. Fig. 5 shows a regression analysis for the evaluation of the MIC. The MIC was almost 200 (mg-chitosan/ml-bacterial suspension) for E. coli and 40 (mg-chitosan/ml-bacterial suspension) for S. aureus. Chitosan evidently has a greater growth inhibitory effect on S. aureus than on E. coli. After the conductance measurement, the activity of chitosan on the bacteria was judged to be bactericidal or bacteriostasis by BHI broth incubation. The growth inhibitory effect on bacteria of chitosan indicated as the bactericidal action on E. coli and S. aureus. Physicochemical interaction between the bacteria and the chitosan polymer caused bactericidal action towards the bacteria. Differences in the antibacterial activity of the cationic chitosan polymer were induced by differences between the gram-negative (E. coli), and gram-positive samples (S. aureus).
A decisive structural difference between gram-negative and gram-positive bacteria can be recognized from the molecular structure of the outer cell membrane. In gram-negative bacteria, phospholipids, lipopolysaccharides and proteins exist in the outer membrane. Therefore, electric interaction hardly influences the inner membrane. In contrast, in gram-positive bacteria, the electric charge of chitosan acts directly on the inner membrane of the cell. The electric interaction between chitosan and the cell membrane was one factor inducing bacterial growth inhibition, cell destruction, and activity depression.
3.3. Effect of particle size and particle shape on bacterial growth inhibition In Section 3.2, the growth inhibitory effect of chitosan on S. aureus appeared stronger than that on E. coli. Therefore, further investigations were focused primarily on S. aureus.
Fig. 7. Scanning electron microscope photographs of the chitosan particles. The particle size was determined by crushing and sieving dried chitosan membrane (DD 92%): (a) 420–500 m; (b) 250–297 m; (c) 74–105 m; (d) 37–63 m.
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Fig. 8. Optical photographs of mannitol salt agar with chitosan membrane added: (a) DD 92.2%; (b) DD 90.1%; (c) DD 88.0%; (d) DD 83.9%; (e) DD 79.7%; (f) DD 75.5%; (g) PVC; (h) control.
The influence of particle size and shape of powdered chitosan membrane on S. aureus was examined. As shown in Fig. 6, decreased particle size improved antibacterial activity. However, the activity of 37- to 63-m particles was less than that of 74- to 105-m particles. Therefore, there is an optimal particle size for antibacterial activity. The morphology of each particle size is depicted in Fig. 7. Powdered chitosan membrane in the range 74–500 m looks like a flake or board, whereas in the range 37–63 m, it looks like a sphere. The antibacterial activity of the powdered chitosan membrane depends on the shape as well as the specific surface area. Quantitative characterization of sharp aspect on growth inhibition of S. aureus will be necessary for future discussions. 3.4. Effect of deacetylation degree of chitosan on growth inhibition of S. aureus The influence of the DD of a chitosan membrane on growth inhibition of S. aureus was investigated by using mannitol salt agar medium; a photograph of the petri dish is shown in Fig. 8. A CFU count of around 150 was observed in the control (chitosan-free) medium, and the medium was yellow. Medium containing polyvinylidene chloride (PVC) was also yellow. PVC was employed as an inert material which has no antibacte-
rial activity. Yellow-colored medium indicates the growth of S. aureus. In contrast, the medium apparently remained red in the system containing chitosan membrane. The trend was clearer as DD increased from 83.9% to 92.2%. Higher DD was especially successful in inhibiting the growth of S. aureus. The number of colonies on the membrane was fewer than that of the circumference area of membrane. The growth inhibitory effect was consistent based both on the color of the medium and the number of colonies on the membrane. The above discussion of bacterial growth inhibition focuses mainly on color changes occurring in the medium. Fig. 9 depicts the MIC of each DD for S. aureus. The MIC for a DD of 92% was about 40 (mg-chitosan/ml-bacterial suspension), and the MIC for a DD of 86% was about 100 (mg-chitosan/ml-bacterial suspension). The antibacterial activity of chitosan towards S. aureus was enhanced with increasing DD. The growth inhibitory effect was not observed at a DD of 82% or a DD of 50%, for reasons that presently are not clear. Other major factors affecting growth inhibition must be investigated in the future. 4. Conclusion Direct conductimetric assays using a Bactometer clearly demonstrated the antibacterial activity of chitosan membranes differing in their deacetylation degree. The influence of chitosan membranes on E. coli and S. aureus, which are food hygiene index bacteria, was investigated. Chitosan inhibits the growth of S. aureus more strongly than it inhibits the growth of E. coli. Chitosan inhibited the growth of gram-positive samples more strongly than that of gram-negative samples, and the effect was bactericidal. At higher DD, chitosan membrane sufficiently inhibits bacterial growth. The shape of the chitosan membrane particles are apparently a major factor influencing antibacterial activity. The shape of the powder particles and their antibacterial activity must be quantitatively correlated in the future. It is anticipated that our findings regarding the dominant role of DD will benefit the production of chitosan membranes for use in a variety of applications. Acknowledgements
Fig. 9. Determination of minimal inhibition concentration of chitosan on the growth of S. aureus. Data were classified according to the deacetylation degree of the particles.
The authors express sincere gratitude to Professor Mikio Kikuchi of the Kanagawa Institute of Technology for conducting the bacterial experiments and for useful discussions. The authors sincerely appreciate the Keyence Corporation for their photographing technique through the Scanning Electron Microscope Model VE-7800.
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