pH Effects on solubility, zeta potential, and correlation between antibacterial activity and molecular weight of chitosan

Accepted Manuscript Title: pH Effects on solubility, zeta potential, and correlation between antibacterial activity and molecular weight of chitosan Author: Shun-Hsien Chang Hong-Ting Victor Lin Guan-James Wu Guo Jane Tsai PII: DOI: Reference:

S0144-8617(15)00701-8 http://dx.doi.org/doi:10.1016/j.carbpol.2015.07.072 CARP 10174

To appear in: Received date: Revised date: Accepted date:

9-6-2015 17-7-2015 21-7-2015

Please cite this article as: Chang, Shun-Hsien., Lin, Hong-Ting Victor., Wu, GuanJames., & Tsai, Guo Jane., pH Effects on solubility, zeta potential, and correlation between antibacterial activity and molecular weight of chitosan.Carbohydrate Polymers http://dx.doi.org/10.1016/j.carbpol.2015.07.072 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

pH Effects on solubility, zeta potential, and correlation between antibacterial activity and molecular weight of chitosan

a,b,

Shun-Hsien Changa, Hong-Ting Victor Lina,b, Guan-James Wuc, and Guo Jane Tsai * a

Department of Food Science, National Taiwan Ocean University, Keelung, Taiwan,

R.O.C b

Center for Marine Bioenvironment and Biotechnology, National Taiwan Ocean

University, Keelung, Taiwan, R. O. C. c

Department of Food Science, National Penghu University of Science and Technology,

Penghu, Taiwan, R.O.C.

*Dr.Guo-Jane Tsai Department of Food Science, National Taiwan Ocean University, 2 Pei-Ning Road, Keelung, 202, Taiwan, ROC Tel: +886-2-2462-2192#5150 Fax: +886-2-2462-7954 E-mail: [email protected]

Highlights

Six chitosans with molecular weights ranging from 3.3-300 kDa were prepared. Combined effects of chitosan MW, temperature, pH on bacterial growth were studied. The pH effects on water solubility and zeta potential of chitosans were examined. Positive correlation between chitosan ZP and antibacterial activity was obtained.

Abstract Six chitosans with molecular weights (MWs) of 300, 156, 72.1, 29.2, 7.1, and 3.3

kDa were prepared by cellulase degradation of chitosan (300 kDa) and ultrafiltration techniques. We examined the correlation between activity against Escherichia coli and Staphylococcus aureus and chitosan MW, and provided the underlying explanation. In acidic pH conditions, the chitosan activity increased with increasing MW, irrespective of the temperature and bacteria tested. However, at neutral pH, chitosan activity increased as the MW decreased, and little activity was observed for chitosans with MW >29.2 kDa. At pH 5.0 and 6.0, chitosans exhibited good water solubility and zeta potential (ZP) decreased with the MW, whereas the solubility and ZP of the chitosans decreased with increasing MW at pH 7.0. Particularly, low solubility and negative ZP values were determined for chitosans with MW >29.2 kDa, which may explain the loss of their antibacterial activity at pH 7.0. Keywords: antibacterial activity; chitosan; molecular weight; solubility; zeta potential 1. Introduction Chitosan, a partially deacetylated chitin (poly-β-(1→4) N-acetyl-D-glucosamine) is a biocompatible polysaccharide with strong antimicrobial activities, which has been employed in the preservation of various foods, including fish (Jeon, Kamil & Shahidi, 2002; Shahidi et al., 2002), shrimp (Tsai et al., 2002), oyster (Cao, Xue & Liu, 2009), cooked rice (Tsai et al., 2006), sausages (Garcia et al., 2010; Moradi et al., 2011; Siripatrawan & Noipha, 2012), and fruits (Campaniello et al., 2008). Chitosan treatment of salmon filets (Tsai et al., 2002) and oyster (Cao et al., 2009) can extend their shelf life at 4°C by 4 days and 6 days, respectively. In addition to increasing the shelf life, chitosan treatments have beneficial effects on firmness, as well as the anthocyanin and vitamin C contents of strawberries and raspberries during storage (Campaniello et al., 2008). Furthermore, chitosan can replace the sanitizing agent chlorine to decontaminate pathogenic bacteria in seafood factories (Chaiyakosa et al.,

2007). Factors including chitosan molecular weight (MW) (Tikhonov et al., 2006), the deacetylation degree (DD), and positive charge content (Takahashia et al., 2008), as well as the temperature and pH of the reaction conditions (Tsai & Su, 1999), individual microbial structural characteristics (Roller & Covill, 1999), and cell age (Yang, Li & Chou, 2007), have been reported to influence the antibacterial activity of chitosan. In general, the antibacterial activity of chitosan increases with its DD and positive charge (Chantarasataporn et al., 2014; Takahashia et al., 2008). Acidic pH conditions and higher temperature are also favorable for the antibacterial activity of chitosans (Georgantelis et al., 2007; Krajewska, Wydro & Janczyk, 2011). However, the effect of chitosan MW on antibacterial activity is still unclear, and some contradictory results have been obtained. Hirano, Tsuchida and Nagao (1989) indicated that chitosans with MWs of 1.5–4.5 kDa exhibited better inhibitory activities than those with higher MWs (6.5–12.0 kDa). However, Tokura et al. (1997) showed that a 9.3-kDa chitosan inhibited the growth of Escherichia coli, whereas a 2.2-kDa chitosan increased bacterial growth. Similarly, Jeon and Kim (2000) found that the MW of chitosans should be >10 kDa to exhibit antibacterial activity. No et al. (2002a) found that the antibacterial activity of chitosans with MWs of 28–1671 kDa increased with the MW. Tsai et al. (2006) first noted that both MW and pH affect the antibacterial activity of chitosans. Thus, the antibacterial activity of a 20-kDa chitosan was greater than that of a 220-kDa chitosan at pH 7.0, whereas the opposite result was obtained at pH 6.0. In this study we prepared a wide range of chitosans with MWs ranging from 3.3 to 300 kDa via cellulase degradation of chitosan, followed by ultrafiltration separation techniques. A comprehensive study was conducted by examining the combined effects

of chitosan MW, reaction temperature, and pH on bacterial growth. We found that the correlation between the antibacterial activity and chitosan MW was opposite between acidic pH and neutral pH. The underlying reason for this difference was proposed by measuring the water solubility and zeta potential (ZP) of chitosans at various pH values.

2. Material and Methods 2.1. Materials Chitin powder was purchased from Applied Chemical Co., Ltd (Taiwan). Acetic acid, acetonitrile, methanol, and glycerol were purchased from Fluka (Garage Gmbh, Switzerland). Sodium azide (NaN3) and sodium bicarbonate (NaHCO3) were purchased from Sigma Chemical Co. (Gillingham, UK). Technical grade cellulase (3000U/g) from Trichoderma viride was purchased from Challenge Bioproducts Co., Ltd (Taichung, Taiwan). Ultrafiltration membrane filters with MW cut off values of 5, 10, 30, 50, and 100 kDa (Amicon Ultra PL-5, 10, 30, 50, and 100) were purchased from Millipore (Billerica, MA, USA). Escherichia coli BCRC 10675 and Staphylococcus aureus BCRC 10780 were purchased from the Bioresources Collection and Research Center (Hsinchu, Taiwan). Nutrient agar (NA) and nutrient broth (NB) and were supplied by Becton Dickinson (Sparks, MD, USA). Molecular size detection columns packed with TSKgel G4000 and G5000 PWXL were obtained from Tosoh Co. Ltd (Tokyo, Japan).

2.2. Chitosan preparation Based on the method of Tsai et al. (2006), a commercial shrimp chitin powder was added to 50% NaOH (1 g of chitin per 13 ml of NaOH) and heated at 140°C in an

oil bath for 1 h to obtain chitosan with 95% DD, which was measured by the colloid titration method (Toei & Kohara, 1976).

2.3. Enzymatic hydrolysis of chitosan and component separation using ultrafiltration techniques Chitosan with 95% DD was hydrolyzed with cellulase (10 U/g chitosan) in 0.5 N acetate-bicarbonate buffer (pH 5.2) at 55°C (Tsai, Zhang & Shieh, 2004). After digestion for 1, 3, 6, 9, and 18h, the hydrolysates were boiled for 15 min, adjusted to pH 7.0, and then centrifuged (12,000×g, 30 min). The supernatants were separated using various ultrafiltration films (UFMs) with MW cutoff values of 100, 50, 10, 5, and 1 kDa (designated as UFM100, UFM50, UFM10, UFM5, and UFM1, respectively). Except for the chitosan obtained from the 1-h digested hydrolysate, each of the other chitosan samples produced from 3-, 6-, 9-, and 12-h digested hydrolysates was prepared using UFMs with two specific cutoff values. For example, to prepare a chitosan sample from the 3-h digested hydrolysate, the neutralized supernatant mentioned earlier was filtered through UFM100. The filtrate was then passed through UFM50, and the residue from UFM50 was retained for further methanol fractionation. Similarly, each pair of UFM50/UFM10, UFM10/UFM5, and UFM5/UFM1 was used to prepare chitosans from the 6-, 9-, and 18-h digested hydrolysates, respectively. The chitosan obtained from the 1-h digested hydrolysate was prepared only using UFM100, and the residue was collected. After adding an equal volume of methanol, each residue was employed to remove the chitooligosaccharides in the supernatant, and chitosan in the precipitate was collected, as described by Tsai et al. (2004). The DD of the five chitosans obtained was approximately 90%, according to the colloid titration method.

2.4. MW determination The MWs of the chitosans were determined by size-exclusion high-performance liquid chromatography (SE-HPLC)(Tsai, Bai & Chen, 2008) using a column packed with TSKgel G4000 PWXL and G5000 PWXL. The mobile phase comprised acetonitrile and distilled water at a ratio of 70:30, and the elution speed was 0.6 ml/min. The elution peak was detected with an RI detector (Model M132, Gilson, Middleton, WI, USA). The MWs of the chitosans were calculated from pullulan standards (5.00 mg/ml, Shodex, standard kit P-1, 2, 3, 5, 10, 20, 50, 100, 200, 400, and 800) using a MW (1.5–800 kDa)(Shodex, Kawasaki, Japan) calibration curve, with SISC-LAB software (Scientific Information Service Co., Taipei, Taiwan).

2.5. Solubility test The solubility of each chitosan was defined as the water-soluble solid content (w/v) in the test solution (Lin, Lin & Chen, 2009). Thus, 0.05 g of chitosan was added to 5 ml of 200 mM phosphate buffer at pH values of 5.0, 6.0, or 7.0, and stirred for 24 h. After centrifugation (6000×g, 30 min) the precipitate (insoluble solid) was collected, washed three times with distilled water, and dried in an oven at 60°C until a constant weight was obtained. Solubility (mg/ml) = (50 mg –insoluble solid dried weight)/sample volume (5 ml)

2.6. ZP measurements The ZP was measured for 150 ppm chitosan in NB with pH values of 5.0, 6.0, and 7.0 using a Zetasizer 2000 system (Malvern Instruments Ltd, Worcestershire, UK), which was equipped with a photon correlation spectroscopy system (Chung et al., 2004). The voltage applied to the driving electrodes of the capillary

electrophoresis cell was 150 V. The Zetasizer 2000 system was calibrated using a standard (DTS5050, Malvern instruments) with a standard ZP of –50 ± 5 mV at 25°C. The Zetasizer 2000 cell was rinsed with HPLC-grade water to ensure the stability of the measurements before each use. All the experiments were conducted at 25°C.

2.7. Culture conditions E. coli BCRC 10675 and S. aureus BCRC 10780 were stored in NB containing 50% sterile glycerol at –80°C. To prepare the bacterial cultures, the strains stored at –80°C were inoculated into 50 ml NB and incubated at 37°C for 18 h, with rotation at 125 rpm. The strains were subcultured twice at 37°C for 18h, and the cultures were diluted to 108CFU/ml with sterile peptone water to conduct antibacterial experiments.

2.8. Antibacterial test A 1% chitosan stock solution was prepared by adding 0.2 g of chitosan to 10 ml of distilled water, sterilizing at 121°C for 15 min, and then adding 10 ml sterile 0.2 N HCl. In 50-ml flasks containing 10 ml of NB, chitosan in 0.1 N HCl was added to obtain a final concentration of 150 ppm, and the pH of NB was then adjusted to 5.0, 6.0, and 7.0 using 0.1N NaOH. Next, 100 µl of E. coli or S. aureus culture was added to the flask to obtain an initial cell density of ca. 106 CFU/ml. After incubating at 4, 15, 37, and 45°C with shaking at 125 rpm for 24 h, 0.1 ml aliquots of 10-fold dilutions of NB culture were spread onto NA plates and incubated at 37°C for 2 days, before the colonies were counted. This experiment was run triplicate Reduction count (Log CFU/ml) = Final cell count (Log CFU/ml) without chitosan–Final cell count (Log CFU/ml) with chitosan

2.9. Statistical analysis The data were analyzed using SPSS Version 12.0 (SPSS Inc., Chicago, IL, USA). One-way analysis of variance was used to determine significant differences between sample means, where the level of significance was set at p < 0.05. Multiple comparisons of means were conducted using Tukey’s test.

3. Results and Discussion 3.1. Effects of chitosan MW on the antibacterial activities at various temperatures and pH Although several studies have investigated the chitosan MW effect on its antibacterial activity, the contradictory results in terms of the correlation between the antibacterial activity and the MWs of chitosans were found. Some studies concluded that the antibacterial activities of chitosans increased as their MWs decreased (Cao & Sun, 2009; Ganan, Carrascosa & Martinez-Rodriguez, 2009); whereas, some reports demonstrated that larger MW chitosans have higher antibacterial activity (Lin et al., 2009; No et al., 2002b; Qin et al., 2006). In order to clarify the equivocal argument mentioned above, a comprehensive study was conducted in this study by investigation of the combined effects of pH, temperature and MW on antibacterial activity of chitosan. A commercial shrimp chitosan with 95% DD and 300 kDa was degraded with cellulase for various periods. The MWs of the chitosans separated from the hydrolysates after cellulase digestion for 1, 3, 6, 9, and 18 h were 156, 72.1, 29.2, 7.1, and 3.3 kDa, respectively. The DD of the five obtained degraded chitosans was around 90%. The original chitosan (300 kDa) and the five degraded chitosan samples were used to determine their antibacterial activities against E. coli BCRC 10675 and S.

aureus BCRC 10780 at pH values of 5, 6, and 7, and at temperatures of 4, 15, 37, and 45°C. After reacting for 24 h, the reduction counts (the difference in the final cell count between the control group and the experimental group) for E. coli, as shown in Figure 1, were measured in the presence of 150 ppm chitosan samples and expressed as the antibacterial activity of the chitosans. As expected, the antibacterial activity of each chitosan sample against E. coli increased with increasing of temperature, irrespective of pH tested (Fig. 1).

In the acidic pH values of 5.0 (Fig.1A) and 6.0 (Fig. 1B), the o

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larger MWs (300, 156, and 72.1 kDa) chitosan at 4 C, 15 C and 37 C generally had higher antibacterial activity, compared to the smaller MWs (29.2, 7.7, and 3.3 kDa) of o

chitosans. At 45 C, the activity for all chitosan samples was similar, based on reduction count after 24 h reaction (Fig. 1A, 1B). However, at pH 7.0 chitosan activity o

increased as the MW decreased, and except at 45 C little activity was observed for chitosans with MW >29.2 kDa (Fig. 1C). The 3.3 kD chitosan at pH 7.0 had the highest antibacterial activity among the 6 tested chitosans.

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The antibacterial activities of the chitosans against S. aureus at various temperatures and pH values are shown in Figure 2. Similar as those in Fig.1 for E. coli, the antibacterial activities of chitosans against S. aureus also increased as the temperature increased (Fig. 2). In acidic pH values of 5.0 (Fig. 2A) and 6.0 (Fig. 2B), the activity for chitosans was gradually increased when chitosan MW increased. Again, the negative correlation between the activity against S. aureus and chitosan MW was observed at pH 7.0 (Fig. 2C), which was opposite from what was found in pH 5.0 (Fig. 2A) or pH 6.0 (Fig. 2B).

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As shown in Figures 1, at 37°C and 45°C in acidic pH condition (pH 5.0 and 6.0), the difference in the reduction count with the chitosan MW became smaller after reacting for 24 h. Therefore, we monitored the survival counts throughout the time course for E. coli and S. aureus after treatment with the six chitosan samples at 37°C and 45°C, as shown in Figures 3 and 4, respectively. At 37oC and pH 5.0 (Fig.3A), the survival counts were quite similar for E. coli after treatment with the six chitosan

samples for 4 h, whereas at pH 6.0 (Fig. 3B) using the same treatment time, the survival counts obtained with the two higher MW (300 and 156 kDa) chitosan treatments were significantly lower than those with the other four smaller MW chitosan treatments. In addition, the growth of E. coli without chitosan treatment was retarded at pH 5.0 (Fig. 3A), compared to that at pH 6.0 (Fig. 3B). It seemed that the acidic stress on E. coli might strengthen the antibacterial effects of chitosans. Accordingly, the difference in antibacterial effects in terms of chitosan MW became smaller at pH 5.0 (Fig. 3A). Similar growth pattern was observed for E. coli alone at pH 6.0 (Fig. 3B) and at pH 7.0 (Fig. 3C). At pH 7.0 the counts of E. coli in the presence of the three larger MW chitosans (300, 156, and 72.1 kDa) gradually increased with incubation time, which were similar to those obtained in the control. On the contrary, the E. coli counts generally decreased as the MW decreased for the three chitosans with lower MWs (29.2, 7.7, and 3.3 kDa) (Fig. 3C). At 45oC the growth of E. coli alone was greatly retarded at both pH 5.0 and pH 6.0 (Fig. 3D, 3E). Also, the E. coli cells died quickly in the presence of the chitosans at 45°C in acidic conditions (pH 5.0 and 6.0) (Fig. 3D, 3E). Similar survival patterns for E. coli in the presence of chitosans were observed at 45°C and pH 7.0 (Fig. 3F), compared with those at 37°C and pH 7.0 (Fig. 3C).

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Compared to E. coli (Fig. 3A), S. aureus was more acid-resistant that in the absence of chitosans it grows very well at 37oC and pH 5.0 (Fig. 4A). We found that the survival counts for S. aureus after treatments with chitosans at 37°C at both pH 5.0 and 6.0 decreased as the MWs of the chitosans increased (Fig. 4A, 4B). However, at 37°C and at neutral pH, similar to E. coli (Fig. 3C), S. aureus still grew in the presence of the three larger MW chitosans (300, 156, and 72.1 kDa); whereas its growth was inhibited by the three smaller MW chitosans and the survival count decreased as the MWs of the chitosans decreased (Fig. 4C). Similar survival patterns for S. aureus after treatment with chitosans at 45oC (Fig. 4D-F) and at 37oC (Fig. 4A-C) for each tested pH value were obtained. Several previous reports have noted that the antibacterial activities of chitosans

increase as the pH value decreases, where the presence of more–NH3+ residues favors binding with bacterial cells, thereby causing structural destabilization (Cruz-Romero et al., 2013; Georgantelis et al., 2007; Krajewska et al., 2011; No et al., 2002b; Tsai & Su, 1999). Increasing temperature obviously enhances the inhibitory activity of chitosan (Tsai & Su, 1999). The same results were also obtained in the present study, irrespective of chitosan MWs and the bacteria tested. However, we found equivocal results in terms of the correlation between the antibacterial activity and the MWs of chitosans. Some studies showed that the antibacterial activities of chitosans increased as the MW decreased, such as (Cao & Sun, 2009) who used chitosans with viscosities ranging from 10 to 1000 cp, Liu et al. (2006) who compared 11 chitosans of various MWs (50–550 kDa), and Ganan et al. (2009) who investigated chitosans of 120, 400, and 643 kDa. However, No et al. (2002b) found that the antibacterial activities of six larger chitosans (28–1671 kDa) were generally higher than those of six smaller MW chitosans (1–22 kDa). Qin et al. (2006) showed that the activities of the six enzymatically digested chitosans (1.4–130 kDa) increased with the MW. Lin et al. (2009) monitored the activities of enzymatically degraded chitosans (8.3–24.8 kDa) and demonstrated that the chitosan activity increased with the MW. In the present study, we found that the correlation between the antibacterial activity and the MW of chitosans was affected by pH. Thus, at an acidic pH (pH 5.0 and 6.0), the chitosan activity generally increased with MW, irrespective of the temperature and bacterial species tested. By contrast, at a neutral pH of 7.0, the chitosans with MW > 29.2 kDa exhibited little activity, for example, 72.1, 156, and 300 kDa, whereas the activities of the chitosans with lower MWs (29.2, 7.7, and 3.3 kDa) increased as the MW decreased.

3.2. pH Effect on solubility and ZP of chitosans The solubility and the available positive charge due to the presence of –NH3+ have been mentioned as important factors that influence the antibacterial activities of chitosans (Lin et al., 2009; Sekiguchi et al., 1994). The available positive charge on the surface of a chitosan can be represented by the ZP (Sadeghi et al., 2008). The correlation between the antibacterial activity and chitosan MW is affected by pH, as shown above, so we investigated the effects of pH on the solubility and ZP for chitosans with various MWs. The solubilities of chitosans with various MWs at pH 5.0–7.0 are shown in Table 1. Each chitosan (50 mg) was dissolved almost completely in 5 ml phosphate buffer at pH 5.0 and 6.0. However, except for the chitosans with MWs of 3.3 and 7.7 kDa, the solubilities of the other larger MW chitosans decreased greatly in neutral condition (pH 7.0). In particular, the 300-kDa chitosan did not dissolve at all and the solubility for chitosans with MWs of 156, and 72.1 kDa, was low at pH 7.0, whereas the 7.7- and 3.3-kDa chitosans retained almost complete solubility, and the 29.2-kDa chitosan retained about 50% solubility, compared with those at pH 5.0 and 6.0. Similarly, Lin, Chen and Peng (2008) showed that chitosans with an MW > 30 kDa cannot be used as antibacterial agents due to their poor solubility in aqueous solutions at neutral pH. Lin et al. (2009) demonstrated that enzymatically degraded chitosans (6.3–13.9 kDa) had good solubility, whereas the original chitosan (323.7 kDa) did not dissolve at all at pH 7.0. In general, chitosans should dissolve to obtain a conformational extension that binds with the bacterial cell surface to lyse bacterial cell (Tsai et al., 2006). Therefore, the low antibacterial activities of the chitosans with MWs of 300, 156, and 72.1 kDa at pH 7.0 may be due to their loss of solubility at pH 7.0.

Data are expressed as mean ± SD from triplicate experiments, and different superscripts (a-d) in the same column are significantly different (p< 0.05). * Maximum 50 mg chitosan was used for testing its solubility in 5 ml phosphate buffer with pH 5.0, 6.0, or 7.0. **:Not detectable.

Table 2 shows the ZPs of chitosans with various MWs at pH 5.0, 6.0, and 7.0. The ZP of each chitosan decreased gradually as the pH increased. In particular at pH 7.0 the ZP decreased as the chitosan MW increased. Negative ZPs (–5.34 to –12.03 mV) were determined for chitosans with MWs of 300, 156, and 72.1 kDa, whereas the chitosans with MWs<30 kDa (e.g., 29.2, 7.7, and 3.3 kDa) still retained their relative positive charges (1.3–5.53 mV). By contrast, at both pH 5.0 and 6.0, the ZP of each chitosan was positive, and its value decreased gradually as the chitosan MW decreased.

Table 2

Data are expressed as mean ± SD from triplicate experiments. Different superscripts (a-e) in the same column are significantly different (p< 0.05).

The pKa for large MW chitosan is around pH 6.2, and the pKa became 6.6 for chitosan with 5-14 sugar residues (Anthonsen & Smidsrød, 1995). Chitosan becomes polycationic at pH 5.0 and pH 6.0. As the pH becomes more acidic, more protonated–NH3+ residues become available as the MW of the chitosan increases.

Thus, higher ZPs were obtained (Table 2). In addition, all the chitosans exhibited good solubility in acidic conditions (Table 1) because they all became polycatonic. At pH 7.0, the larger chitosans (300, 156, and 72.1 kDa) deprotonated, and the chitosan molecules were aggregated (observed visually), probably due to inter-/intramolecular hydrogen bond formation. Therefore, the solubilities of these chitosans decreased greatly (Table 1). However, the smaller chitosans (29.2, 7.7, and 3.3 kDa) still had some protonated residues, and the chitosan molecules did not aggregate, probably due to their higher pKa values and less hydrogen bonding. These chitosans (especially those with MWs of 7.7 and 3.3 kDa) retained good solubility (Table 1). Similarly, Saïed and Aïder (2014) demonstrated that the ZP of chitosan (150 kDa) became negative at pH 7.0. Figure 5 plots the chitosan ZPs shown in Table 2 versus the reduction counts (designated as antibacterial activity) obtained by the chitosans at 37°C shown in Figures 1 and 2. The amount of positively charged–NH3+ residues has a critical effect on the bactericidal activity of chitosans (Cruz-Romero et al., 2013; Krajewska et al., 2011). Thus, we detected a positive correlation between ZP and the antibacterial activity against E. coli (Fig. 5A) and S. aureus (Fig. 5B) for chitosans with various MWs in this study, where the correlation coefficients (R2) were 0.815 and 0.911, respectively. (A)

(B)

25 20

y=3.633x - 12.01 R2=0.815

15 )Vm( laitn etop-at eZ

10 5 0 -5 -10 -15 0

2

4

6

Reduction count (Log CFU/ml)

8

10

25 20

y = 0.255x + 3.833 R 2 = 0.911

15 )Vm( laitn etop - at eZ

10 5 0 -5 -10 -15 0

2

4

6

8

10

Reduction count (Log CFU/ml)

Fig. 5. Chang et al.

4. Conclusions This systematic study shows clearly that the MWs of chitosans, temperature, and pH are important factors that affect the antibacterial activities of chitosans. The antibacterial activity of each individual chitosan increased as the pH decreased and as the temperature increased. Thus, the pH of the reaction mixture greatly affected the correlation between the antibacterial activity and the chitosan MW. At acidic pH (pH 5.0 and 6.0), the chitosan activity increased as the MW increased. By contrast, at pH 7.0, the chitosans with MWs> 29.2 kDa (i.e., 300, 156, and 72.1 kDa) greatly lost their activity, whereas the activities of the smaller chitosans (29.2, 7.7, and 3.3 kDa) increased as the MW decreased. These pH effects on the correlation between the antibacterial activity and chitosan MW may be explained by variations in the water solubility and ZP of chitosans with various MWs when the pH changed from acidic to neutral. The changes in water solubility and ZP increased as the MWs of the chitosans increased. Larger chitosans with MWs> 29.2 kD lost their water solubility almost completely, and they had negative ZPs. Thus, larger chitosans lose their activity at pH 7.0.

5. Acknowledgments Financial support from the Ministry of Science and Technology, Taiwan, R.O.C. (NSC101-2622-B-019-001-CC3, NSC102-2313-B-019-015) is gratefully acknowledged.

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Figure captions Figure 1. Reduction counts of E. coli BCRC 10675 treated with 150 ppm chitosans with various molecular weights in nutrient broth at pH 5.0 (A), 6.0 (B), and 7.0 (C), and at 4°C, 15°C, 37°C, and 45°C for 24 h. Reduction count (Log CFU/ml) = Final count (Log CFU/ml) without chitosan–Final count (Log CFU/ml) with chitosan All the data represent the mean (n=3). Different lowercase letters within a specific temperature differ significantly (p < 0.05).

Figure 2. Reduction counts of S. aureus BCRC 10780 treated with 150 ppm chitosans with various molecular weights in nutrient broth at pH 5.0 (A), 6.0 (B), and 7.0 (C), and at 4°C, 15°C, 37°C, and 45°C for 24 h. Reduction count (Log CFU/ml) = Final count (Log CFU/ml) without chitosan–Final count (Log CFU/ml) with chitosan All the data represent the mean (n=3). Different lowercase letters within a specific temperature differ significantly (p < 0.05).

Figure 3. Cell counts for E. coli BCRC 10675 in nutrient broth containing 150 ppm chitosans with various molecular weights at 37°C (A, B, and C) and 45°C (D, E, and F).

Chitosan MW:-▲,300 kDa; -■, 156 kDa; -▼, 72.1 kDa; -○, 29.2 kDa; -△, 7.7 kDa;

-□, 3.3 kDa; -●, control. All data are expressed as mean with SD (n=3) Figure 4. Cell counts for S. aureus BCRC 10780 in nutrient broth containing150 ppm chitosans with various molecular weights at 37°C (A, B, and C) and 45°C (D, E, and F). Chitosan MW:-▲, 300 kDa; -■, 156 kDa; -▼, 72.1 kDa; -○, 29.2 kDa; -△, 7.7 kDa; -□, 3.3 kDa; -●, control. All data are expressed as mean with SD (n=3)

Figure 5. Correlations between the zeta potentials and antibacterial activities of chitosans against E. coli BCRC 10675 (A) and S. aureus BCRC 10780 (B) at 37°C for 24 h. Antibacterial activity = Reduction count (Log CFU/ml) = Final count (Log CFU/ml) without chitosan–Final count (Log CFU/ml) with chitosan

Chitosan MW:-△,300 kDa; -□, 156 kDa; -▽, 72.1 kDa; -○, 29.2 kDa; -◇, 7.7 kDa;

-☆, 3.3 kDa, legend color filled with black, white, and gray represents pH 5.0, 6.0, and 7.0, respectively.

Table 1 Solubility of chitosans with various molecular weights in phosphate buffer MW

Solubility* (mg/ml)

Sample (kDa)

pH 5.0

pH 6.0

pH 7.0

300

9.999 ± 0.001a

9.969 ± 0.038a

ND**

156

9.999 ± 0.001a

9.986 ± 0.008a

0.118 ± 0.025d

72.1

10.000 ± 0.001a

9.990 ± 0.008a

0.543 ± 0.084c

29.2

9.995 ± 0.005a

9.989 ± 0.003a

4.998 ± 0.001b

7.1

9.998 ± 0.002a

9.989 ± 0.002a

9.990 ± 0.009a

3.3

9.997 ± 0.005a

9.992 ± 0.003a

9.994 ± 0.005a

Chitosan

Zeta potentials of chitosans with various molecular weights in NB with various pH values

Chitosan

Zeta potential (mV)

MW (kDa)

pH 5.0

pH 6.0

pH 7.0

300

20.33 ± 1.171a

16.23 ± 1.19a

-12.03 ± 1.01e

156

18.67 ± 1.00a

8.00 ± 0.71b

-10.98 ± 3.01e

72.1

17.50 ± 0.95a

7.06 ± 0.73b

-5.34 ± 3.44d

29.2

15.27 ± 0.90b

7.46 ± 0.36b

1.30 ± 0.74c

7.7

14.83 ± 0.59b

7.63 ± 0.19b

2.98 ± 0.62b

3.3

14.10 ± 0.26c

7.09 ± 0.14b

4.10 ± 0.54a