Development and application of active films for food packaging using antibacterial peptide of Bacillus licheniformis Me1


  • V. Nithya,

    1. Department of Food Microbiology, CSIR-Central Food Technological Research Institute, Mysore, India
    Current affiliation:
    1. Division of Energy and Bioengineering, Dongseo University, Busan, Korea
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  • P.S.K. Murthy,

    1. Department of Food Packaging Technology, CSIR-Central Food Technological Research Institute, Mysore, India
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  • P.M. Halami

    Corresponding author
    • Department of Food Microbiology, CSIR-Central Food Technological Research Institute, Mysore, India
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Prakash M. Halami, Department of Food Microbiology, CSIR-Central Food Technological Research Institute, Mysore-570 020, India. E-mail:



An attempt was made to evaluate the effectiveness of partially purified antibacterial peptide (ppABP) produced by Bacillus licheniformis Me1 for food preservation by means of active packaging.

Methods and Results

The active packaging films containing ppABP were developed using two different packing materials [low-density polyethylene (LDPE) and cellulose films] by two different methods: soaking and spread coating. The activated films showed antibacterial activity against pathogens. The release study of ppABP from coated film showed that the LDPE films liberated ppABP as soon as it comes in contact with water, while gradual release of coated ppABP was observed in case of cellulose films. The activated films showed residual activity in different simulating conditions, such as pH of food and storage temperatures. The activated films demonstrated its biopreservative efficacy in controlling the growth of pathogens in cheese and paneer.


The ppABP-activated films were found to be effective for biopreservation. The ppABP from active films got diffused into the food matrix and reduced the growth rate and maximum growth population of the target micro-organism.

Significance and impact of the Study

Both types of ppABP-activated films can be used as a packaging material to control spoilage and pathogenic organisms in food, thereby extending the shelf life of foods.


In response to the changes in market trends and increasing demand of consumers for high quality, safe and extended shelf life of food products, active packaging is creating a niche in the market and becoming increasingly significant. Active packing has been defined as ‘a type of packaging in which the package, the product and the environment interact to extend shelf life or improve safety and convenience or sensory properties, while maintaining the quality and freshness of the product’ (Vermeiren et al. 1999).

Antimicrobial packaging is a promising and innovative form of active packaging. One of the major concerns of the food industry is the spoilage of foods and food poisoning by microbial contamination. Most of the contamination of the foods occurs mainly on the surface due to postprocessing and handling (Perez-Perez et al. 2006). The delay or prevention of spoilage of foods has been done either by dipping and spraying foods with antimicrobials or by packing the foods with antimicrobial packages. The former approach is less efficient as these compounds may get neutralized on contact or get diffused rapidly into the food matrix or may get diluted to below effective concentration (Hoffman et al. 2001; Quintavalla and Vicini 2002), whereas the antimicrobial packaging is an efficient technology and helps in reducing the risk of pathogens development, as well as extending shelf life and maintaining food quality and safety (Han 2000; Mauriello et al. 2005). Furthermore, the packaging films with antimicrobial agents confer residual activity during transport, storage and distribution (Quintavalla and Vicini 2002).

Studies of new food-grade bacteriocins as biopreservatives and development of suitable systems for activation of plastic films with bacteriocins for food packaging are important issues in food biotechnology; both are meant for implementing and improving the effective hurdle technologies for the better preservation of food products. The use of bacteriocins or other natural antimicrobials in packaging films to control food spoilage and pathogenic organisms has increased significantly as it involves lesser risk for the consumers (Perez-Perez et al. 2006). Among bacteriocins, nisin produced by Lactococcus lactis has been the subject of extensive study and use, either as direct application in food or indirect use in antimicrobial packages for food biopreservation (Natrajan and Sheldon 2000; Hoffman et al. 2001; Lee et al. 2004; Mauriello et al. 2005). Nisin is ‘generally recognized as safe’ (GRAS) by FDA for application in food industry and agriculture (U.S. Food and Drug Administration 1988). However, the application of nisin as a biopreservative is restricted in certain conditions, such as use in alkaline foods due to its very low activity at a neutral or an alkaline pH (Schillinger et al. 1996), in foods requiring high processing temperature due to its lower activity in higher temperatures and in foods where Gram-negative bacteria and wide range of bacterial population are to be kept in check. Some strains of Listeria monocytogenes demonstrating increased tolerance or resistance to nisin have also been reported (Mazzotta et al. 2000). Like lactic acid bacteria, some of the Bacillus spp. such as B. subtilis, B. licheniformis and B. coagulans are also found safe for use in food and agricultural industry (Sharp et al. 1989; PR Newswire 2009) and are known to produce several antimicrobial compounds that show inhibitory activity against broad range of food-borne pathogens (Stein 2005; Abriouel et al. 2010). Studies on new food-grade bacteriocins from Bacillus as biopreservatives and the development of active packaging system with such bacteriocins may offer an alternative to nisin. Moreover, the use of Bacillus bacteriocins in packaging materials is not yet reported.

Thus, keeping in view the potential application of antimicrobial compounds and antimicrobial packages, the biopreservative efficacy of ppABP-activated packaging materials was evaluated. To determine this, active films were developed by incorporating ppABP of the culture B. licheniformis Me1 to packaging materials. Further, the effectiveness of ppABP-activated films for inhibiting the growth of common food-borne pathogens was evaluated. The release of ppABP from the activated film and the efficacy of the developed films in inhibiting the growth of L. monocytogenes Scott A during the storage of dairy products were also verified.

Materials and methods

Packaging material and bacteriological media

The packaging films [low-density polyethylene (LDPE) and cellulose] used for developing active films were obtained from the Department of Food Packaging, CFTRI, Mysore, India. Brain–heart infusion (BHI) medium used for the growth of the bacterial strains was procured from HiMedia, Mumbai, India.

Bacterial strain and culture conditions

The ppABP used for activating the packaging material is obtained from a native isolate B. licheniformis Me1. The culture is isolated from milk and is able to produce a proteinaceous antibacterial peptide, which exhibits broad spectrum of inhibitory activity and is stable over a wide range of temperature and pH (Nithya and Halami 2012). The pathogens used in this study included Micrococcus luteus ATCC 9341, L. monocytogenes Scott A, Staphylococcus aureus FRI 722, Salmonella typhimurium MTCC 1251 and Bacillus cereus F 4433. The cultures used in this study were maintained at −20°C in BHI broth, containing 20% (v/v) glycerol. All the cultures were propagated aerobically twice in BHI broth before use.

Preparation of antimicrobial agent

For the production of antibacterial peptide, the culture B. licheniformis Me1 was grown in a modified media (pH 8 ± 0·2) consisting of corn steep liquor (2%), yeast extract (0·5%) and NaCl (0·25%) for 24 h at 37°C and with agitation speed of 150 cycles min−1. After incubation, the cell-free supernatant (CFS) was collected by centrifugation (10 000 g for 15 min), and protein was precipitated from CFS by slow addition of ammonium sulfate to a 65% saturation. After 12 h of incubation at 4°C, the precipitated protein was collected by centrifugation (10 000 g for 10 min), and the pellet was then resuspended in 0·1 mol l−1 phosphate-buffered saline (pH 7) and further extracted with n-Butanol. Butanol was evaporated using Rotary vacuum evaporator (BUCHI India Pvt. Ltd., Mumbai, India.) at 14 lb and 55°C, and the dried residue was dissolved in 1/100th volume of sterile distilled water. The resulting partially purified ABP (ppABP) was stored at −20°C, until further use. The antibacterial activity of the ppABP against the food-borne pathogens was determined by agar spot assay (Todorov 2008). The minimum inhibitory activity/residual activity were determined by serial twofold dilution method against the indicator organism M. luteus ATCC 9341 (Nithya and Halami 2012). Activity was defined as the reciprocal of the dilution after the last serial dilution giving a zone of inhibition and expressed as activity unit (AU) per millilitre.

Preparation of antimicrobial packaging films


A solution of ppABP was prepared at a final concentration of 800 and 6400 AU ml−1. Samples of LDPE and cellulose films of size 2 × 2 cm were soaked in ppABP solution of the above concentrations for different incubation periods (1 and 5 h for LDPE and 1, 3 and 5 h for cellulose films). After soaking, the films were air-dried, and then, their antibacterial activity was assayed against the indicator organisms by bioactive assay.

Coating by spreading

The ppABP was prepared at a concentration of 6400 AU ml−1 in sterile distilled water. LDPE and cellulose films (size 30 × 3 cm) were spread-coated with ppABP uniformly with the help of a spreader (triangle shaped) dipped in bacteriocin solution. After spreading, the films were exposed to warm air to dry the ppABP solution and promote a homogenous distribution of the ppABP onto the surface of the films. Once dried, the treated films were assayed for antibacterial activity against the indicator organisms by bioactive assay.

Antibacterial activity

The above treated LDPE and cellulose films were assayed for antimicrobial activity against food-borne pathogens, such as M. luteus ATCC 9341, L. monocytogenes Scott A, Staph. aureus FRI 722, B. cereus F 4433 and Salm. typhimurium MTCC 1251 as described elsewhere (Mauriello et al. 2004). Briefly, samples of the treated films (2 × 2 cm) were placed onto the surface of BHI soft (0·8%) agar plates seeded with 104 CFU ml−1 of 16 ± 2 h grown indicator organism. The treated face of the film was in contact with the agar. The untreated films were also assayed and served as negative controls. After incubating the plates at 37°C for 18 ± 2 h, the antagonistic activity was evaluated by observing a clear zone of growth inhibition in correspondence with the active film.

Adsorption and release of ppABP from spread-coated active films

For checking the adsorption rate of ppABP by the films, drops of 20 μl of 6400 AU ml−1 ppABP were spotted on the surface of untreated LDPE and cellulose films and then removed after 1, 5, 10, 15, 30, 45 and 60 min (Mauriello et al. 2004). The film was then placed on BHI agar plates seeded with M. luteus ATCC 9341 for checking the antibacterial activity, as described above.

The study of release of ppABP from active LDPE and cellulose films was performed as described previously (Mauriello et al. 2004). Briefly, 20 μl of sterile deionized water was spotted onto the surface of active films with ppABP and then incubated in a humid chamber. The water was removed at 1 h interval for 5 h, and the antibacterial activity of the collected water samples against M. luteus ATCC 9341 was checked by agar well diffusion assay (Motta and Brandelli 2002).

To check the residual activity of the films after release of ppABP, the release experiment was also conducted by another method. Briefly, the active films (2 × 2 cm) with ppABP were placed in 2 ml of sterile deionized water and kept under gentle stirring (100 cycles min−1) at 28 ± 2°C. The antibacterial activity of the solution in which the film was dipped was checked against M. luteus ATCC 9341 at 1, 2, 4, 8, 24 and 48 h of incubation using agar well diffusion assay. The residual activity of the treated films was also checked by placing the film after treatment on the agar plates seeded with indicator organism.

Similar experimental procedure was followed to determine the release of ppABP from the active films under stimulating conditions, such as temperature and pH. The activated films were placed in 2 ml of sterile distilled water in different test tubes and then incubated at different temperatures (4 and 37°C). At different incubation times (1, 3, 8, 12 and 24 h), samples of 50 μl of water from each incubation temperatures were taken and assayed for antimicrobial activity against M. luteus ATCC 9341 by agar well diffusion assay. Simultaneously, the activity of the temperature-treated ppABP-activated films removed at different incubation period was also evaluated as described previously.

For determining the effect of pH on the release of ppABP from the activated films, the films were placed in 5 ml of PBS solutions of pH 3, 7 and 9 and incubated at room temperature (25 ± 2°C) for 6 h. After incubation, the pH of the PBS solutions with pH 3 and 9 was neutralized to pH 7 using sterile 1 N of NaOH or HCl, respectively, before testing its activity. Also, the films were dried and assayed for antimicrobial activity against the indicator strain, M. luteus ATCC 9341.

Application of active films as packing material

Paneer was prepared as follows: to 1000 ml of hot milk, 5 ml of lemon juice was added in to separate the curd from the whey. Then, the curd was drained, pressed out in cheese cloth to remove excess water and subsequently kept for moulding. After moulding, the paneer was cut into pieces of 1 g and divided into three sets. The paneer pieces were surface inoculated by dipping paneer pieces in a diluted suspension of overnight grown L. monocytogenes Scott A. The paneer pieces from the first and second set were wrapped individually with LDPE and cellulose films spread-coated with ppABP solution of concentration 1600 AU ml−1. The third set of paneer pieces was wrapped individually by untreated film of both types separately and served as the negative control. All the wrapped paneer pieces were stored in sterile containers at 4 ± 2°C. Individual samples were removed at an interval of 3 days for a period of 12–15 days, homogenized in 9 ml of saline and then appropriate dilutions were placed on Listeria Oxford (LO) agar. The plates were then incubated at 37°C for 24 h, and CFU of the pathogen per gram of the paneer sample was determined.

For the preservation studies of cheese using active packages, the cheese was purchased from local market and made into pieces (1 g each). The procedure for inoculation of cheese with indicator strain, packaging and detecting the effect of active packages on the viability of the inoculated pathogen was carried out as described for paneer. The results of both paneer and cheese biopreservation studies are based on three independent experiments.


Antibacterial activity of activated films

The packaging films LDPE and cellulose films were activated with ppABP of B. licheniformis Me1 by two different methods: soaking and spread coating. Both types of activated films showed antibacterial activity against indicator organisms M. luteus ATCC 9341, L. monocytogenes Scott A, Staph. aureus FRI 722, B. cereus F 4433 and Salm. typhimurium MTCC 1251. Some of the images showing inhibitory activity of the ppABP-activated LDPE and cellulose films against tested pathogens are shown in Fig. 1. The zone of inhibition was not confined to the film area of both types of film. The soaked activated films showed regular zone of inhibition along the periphery of the film as compared to spread-coated active films, suggesting even diffusion of the ppABP from the film into the agar. The activated films prepared with two different concentrations of ppABP (800 and 6400 AU ml−1) exhibited difference in the intensity of the inhibitory activity against the indicator strains. There was no marked difference in the intensity of the antimicrobial activity among the activated LDPE films against L. monocytogenes Scott A prepared by soaking at different incubation times (1 and 5 h). However, the activated cellulose films showed increase in zone of inhibition with increase in incubation time. In all the cases, the untreated films did not show any activity against the indicator strains. Interestingly, even after 3 months of incorporation, the ppABP-activated films (both LDPE and cellulose) kept at 4°C and room temperature displayed a clear and stable antilisterial activity. The activated films showed antibacterial activity when checked after rubbing also.

Figure 1.

Antibacterial activity of ppABP-activated low-density polyethylene films (i) and cellulose films (ii) against food-borne pathogens. Inhibition zone of soak-activated (I) and spread-activated (II) films with 6400 AU ml−1 of ppABP (a), 800 AU ml−1 of ppABP (b) and untreated films (c) against pathogens (1) Listeria monocytogenes Scott A, (2) Staphylococcus aureus FRI 722, (3) Bacillus cereus F 4433 (4) Salmonella Typhimurium MTCC 1251 and (5) M. luteus ATCC 9341.

Adsorption of ppABP to films with increasing contact time

To assess whether the binding of ppABP was affected by the time of incubation with antibacterial substance, aliquots (20 μl) of ppABP solution (6400 AU ml−1) were spotted on the surface of the LDPE and cellulose films for different contact times (1, 5, 10, 15, 30, 45 and 60 min), and then, the antimicrobial activity of the film was tested.

As shown in Fig. 2(i), in LDPE film, the observed antibacterial activity, in correspondence to the spot area of ppABP solution, was almost the same for all of the contact time except a slight increase at 60 min. While in case of cellulose film, an increasing trend in the diameter of the inhibition zone against the indicator organism was observed in correspondence to the ppABP-spotted region as the contact time of the ppABP solution with the film increased (Fig. 2(ii)).

Figure 2.

Antibacterial activity of low-density polyethylene (LDPE) and cellulose film spotted with ppABP for different contact time. Inhibition zone corresponding to 1, 5, 10, 15, 30, 45 and 60 min of contact time with ppABP (6400 AU ml−1) in case of LDPE (i) and cellulose (ii) films.

Release of ppABP from the activated film

The activated LDPE films were subjected to ppABP release in water at different incubation times (every 1 h till 5 h). The water spots of 20 μl collected after different incubation times showed the same intensity of antibacterial activity against M. luteus ATCC 9341 in agar well diffusion assay. There was no back-absorption of the ppABP from the water to the film until 5 h, as no marked difference in zone of inhibition of the collected water samples was noted. Although the release of the ppABP was confined to the area where the water drops were placed, the activated films, after being assayed for the ppABP release, showed inhibitory activity. This may be attributed to the fact that the ppABP present in the outer circumference of the water drop placed on the film might be getting diffused and inhibiting the pathogens in the area of the spot of released ppABP. To further confirm the release of ppABP from LDPE, the film was soaked in water under stirring condition, and the antibacterial activity of 20 μl of water collected at every 1 h interval for 5 h was checked. The water samples collected showed maximum activity from the initial 1 h onwards. There was no marked difference in the zone of inhibition for any of the other water samples collected till 5 h (Fig. 3(i)(b)). After the release assay, when the film was checked for antibacterial activity, it failed to show any zone of inhibition against M. luteus ATCC 9341 (Fig. 3(i)(a)), indicating complete release of ppABP into water.

Figure 3.

Antibacterial activity of ppABP-activated films placed in sterile water under stirring conditions. Release of ppABP from low-density polyethylene (i) and cellulose (ii) films; (a) the activity of the film after treatment and (b) the inhibitory activity of sterile water in which treated film was soaked for different incubation times.

The activated cellulose film showed contradictory results as compared to LDPE films in the ppABP release assay. Initially, the water drops did not show any zone of inhibition against M. luteus ATCC 9341 until 2 h. After 2 h of incubation, a zone of inhibition of very small diameter was observed, which slightly increased with further increase in incubation time of the water drops (Fig. 3(ii)(b)). The activated film, after being assayed for ppABP release, displayed antibacterial activity, indicating that the ppABP is still retained in the film (Fig. 3(ii)(a)).

Release of ppABP from the activated films under simulated conditions

Release of ppABP at different temperature

The activated LDPE film showed a loss in activity within 1 h of incubation at both the temperatures (4 and 37°C) (Table 1). However, a regain in activity was observed from 8 h onwards in the films, which were incubated at 4°C.

Table 1. Release of the ppABP from active films under different temperature
Incubation time (h)Residual activity of films at different Temperature (°C)
Low-density polyethyleneCellulose
  1. −, absence of activity; +, less activity; ++, maximum activity.


In the case of activated cellulose film, there was no marked difference in the release of ppABP at both the storage temperatures (Table 1). Even after 24 h of incubation in sterile distilled water, the activated cellulose film retained antibacterial activity, indicating its potential use in packaging of long-term-stored food products. Furthermore, there was a gradual increase in the inhibitory activity of the water, indicating controlled release of ppABP from the film and no dependency on the temperature difference.

Release of ppABP at different pH

The activated LDPE and cellulose films were kept under different pH (3, 7 and 9) for 6 h, and the films were then checked for antibacterial activity. In the case of LDPE, the activated film kept at the low pH (3) showed activity with less intensity, whereas at pH 9, the film did not exhibit any antibacterial activity. On the other hand, the cellulose film retained its activity at all the pH conditions. However, a decrease in intensity of the activity of the cellulose film kept at pH 9 was observed as compared to pH 3 and 7.

Inhibition of L. monocytogenes Scott A in dairy products

After preparation, samples of paneer pieces inoculated with L. monocytogenes Scott A were packed with active films (cellulose and LDPE) coated with ppABP and stored in sterile container at 4 ± 2°C. The effect of the activated films on the reduction in Listeria population was observed just within 24 h of incubation of paneer samples. The number of viable cells of L. monocytogenes Scott A in paneer samples packed with activated LDPE films showed a reduction of around 1-log CFU g−1 (Fig. 4a(i)). On the other hand, until 8 days, a slow and steady decline in the count of Listeria and a bacteriostatic effect was observed in paneer samples packed with activated cellulose film (Fig. 4a(ii)), and further incubation did not decrease the cells. However, after 12th day, there was an increase in the number of viable cells. This can be attributed to slow release of ppABP from the cellulose film as compared to the LDPE film. Moreover, the paneer samples covered with active LDPE films started spoiling by 8th day of incubation, while the cellulose-packed paneer samples were stable up to 12 days.

Figure 4.

The viable count of Listeria monocytogenes Scott A in paneer (a) and cheese samples (b) packed with either ppABP-activated low-density polyethylene (i) or cellulose (ii) films during incubation period; treated package (1600 AU ml−1) (●) and untreated package (♦). Each point is the mean ± SEM of three independent experiments.

A similar observation was noted for cheese samples packed with activated films. However, the effect of the ppABP was higher as compared to that of paneer samples. In the treated samples, a reduction of 2-log CFU of the inoculated pathogen per gram of the cheese samples was observed in both types of packaging (LDPE- and cellulose-packed cheese samples). In case of LDPE packed cheese, a bacteriostatic effect on pathogen was observed (Fig. 4b(i)), whereas in cellulose-packed cheese, the reduction was continuous and a bactericidal action was observed (Fig. 4b(ii)). A continuous increase in the number of viable cells of L. monocytogenes Scott A was noticed in the cheese samples packed with untreated films in both cases.


Foods are complex ecosystems with a range of microbial compositions, which may vary from (commercially) sterile foods to raw or fermented foods. In commercially sterile foods, postprocess contaminants may easily proliferate. Increasing interest of consumers for the use of biopreserved foods has sparked the use of bacteriocins, especially from food-grade micro-organisms because of their better adoptability in food systems (Galvez et al. 2007). The use of bacteriocins to assure microbial food safety is a novel approach and an alternative to chemical preservatives. Apart from direct application of bacteriocins by spraying and dipping, ex situ-produced bacteriocins can also be applied in the form of immobilized preparations, in which the partially purified bacteriocin or the concentrated cultured broth is bound to a carrier (Chen and Hoover 2003; Galvez et al. 2007). In the last few years, this method of application of bacteriocins has received considerable interest, because the carrier acts as a reservoir and the concentrated bacteriocin molecules diffuses slowly into the food matrix ensuring a gradient-dependent continuous supply of bacteriocin. Moreover, the localized application of bacteriocin molecules on the food surface requires much lower amounts of bacteriocin, decreasing the processing costs.

In the present study, the activated films (LDPE and cellulose) with ppABP from B. licheniformis Me1 showed a zone of inhibition that did not confine to the film area, indicating that the ppABP diffused from the films into the medium. Furthermore, the ppABP retained its activity in both methods of activation (soaking and spread coating). The soaking procedure proved to be more effective. Presumably, the regular inhibition zone along the periphery of the soak-activated films might be due to the homogeneous distribution of the bacteriocin on the surface of the films. However, Mauriello et al. (2004) observed irregular inhibition zone and heterogeneous distribution of the bacteriocin for soaked polythene-oriented polyamide films.

The interactions of the antimicrobial agents with the film matrix have a crucial effect on the antimicrobial activity of the active films (Papadokostaki et al. 1997; Han 2000). Lakamraju et al. (1996) reported that hydrophilic surface adsorbs a higher amount of nisin than hydrophobic one. The LDPE film shows hydrophobic properties and thus rejects the hydrophilic antimicrobial formulations to a greater extent than other films (Natrajan and Sheldon 2000). On the other hand, the cellulose film being hydrophilic polymer matrices absorbs higher amounts of ppABP. Similarly, in our experiments, the cellulose film exhibited a marked difference in the adsorption kinetics than LDPE indicating a higher binding ability for ppABP. Adsorption studies carried out by spotting the ppABP on the surface of the films demonstrated that even a quick contact of the bacteriocin with the surface of the film conferred activation, similar to the observation made by Mauriello et al. (2004). This observation lead to the conclusion that the ppABP adsorb or absorb to the surface of the films and not migrate from the cut margins into the film in the activity assay.

The diffusion rate of the antimicrobial agents and its concentration in the film must be sufficient to remain effective throughout the shelf life of the product (Cooksey 2000). Polymer structure affects the release of active compounds (Papadokostaki et al. 1997). Hydrophilic nature of the cellulose film creates greater retention of the ppABP by binding. As a consequence of this, in our study also the release rate of the ppABP from the cellulose films was lower and inhibition zones were smaller in the beginning of the incubation period, as compared to activated LDPE films. These results indicate that the release of ppABP from the activated cellulose film was time-dependent and in a controlled manner. This implies that the activated cellulose films are suitable for packing of solid foods as the coated ppABP will release slowly onto the food surface and inhibit the growth of surface spoilage and pathogenic bacteria during an extended period of transport and storage phase of food distribution (Perez-Perez et al. 2006).

Processed foods have different pH values and are exposed to different temperature profiles during handling, storage and distribution. The pH of a product affects the growth rate of target micro-organisms and changes the degree of ionization of the most active chemicals, as well as the activity of the antimicrobial agents (Han 2000). Several researchers have found that the increased storage temperature can accelerate the migration of the active agents in the film and deteriorate the protective action of antimicrobial films, due to high diffusion rates in the polymer (Wong et al. 1996). Furthermore, the storage temperature may also affect the activity of antimicrobial packages. Thus, release of the antibacterial peptide from films in simulating food conditions should be evaluated to determine the efficacy of the films in controlling pathogens in food systems. The result of the release studies at different temperatures indicates that lower temperatures allow the back-adsorption of the ppABP to the LDPE films, and thus, the film showed activity after 8 h of incubation at 4°C. The reason for this back-absorption behaviour remains unexplained as the mechanism for ppABP binding to the plastic film is not known. Some authors have also demonstrated a loss in activity of the antimicrobial LDPE packages at 25°C (Dawson et al. 2003; Mauriello et al. 2005). The cellulose film retained its activity at both the incubation temperatures and pH treatments. This might be probably due to the chemical nature of the cellulose films (Papadokostaki et al. 1997). This suggests the potential use of cellulose films as a packaging material for the control of spoilage microbes in acidic to alkaline foods, incubated at either lower or higher temperatures and when long-term storage is desired.

Most of the research work in antimicrobial packaging has been focused primarily on the development of various methods and model systems, whereas little attention has been paid to their preservation efficacy in actual foods (Han 2000). With increasing use of minimally processed food with no chemical preservatives and development of active films with bacteriocins as packaging material, research is essential to identify the types of food that can benefit most from antimicrobial packaging materials. Reports are available demonstrating the biopreservation of meat samples using food packaging materials containing bacteriocins (Dawson et al. 2003; Lee et al. 2004; Mauriello et al. 2004). However, reports on the preservation of dairy products using antimicrobial packaging materials are rare, especially the application of active packages with bacteriocins from Bacillus. The active packaging films (LDPE and cellulose) that were used to coat dairy products such as paneer and cheese demonstrated the control of inoculated pathogen Listeria. The released ppABP from the active films into the tested food samples reduced the growth rate and maximum growth population and delayed the lag period of the target micro-organism. Similar observations were reported in raw milk, pasteurized milk and ultrahigh temperature milk (Lee et al. 2004; Mauriello et al. 2005). An increase in L. monocytogenes Scott A viable counts was noted after certain days of storage in all the samples, which may be due to the particular mechanism of action of bacteriocins that can inhibit as many cells as molecules available in the medium (Moll et al. 1999). However, increasing the concentration of the bacteriocin in the coating solution may improve the preservative performance of the bacteriocin-coated films in storage of dairy products, as well as other food products. Furthermore, addition of hurdle molecules such as EDTA, lysozyme, citric acid, lactic acid, lauric acid into the coating solution may improve the antimicrobial performance of bacteriocin-activated films as reported in other studies (Natrajan and Sheldon 2000).


In conclusion, the cellulose film was found to be more efficient as a carrier of the ppABP produced by B. licheniformis Me1 and thus can be exploited for use in food packaging industries. However, further studies with respect to physiochemical properties of the film material after incorporation of ppABP are required. The activated films showed residual activity in different simulating conditions, such as pH of food and storage temperatures. Also, the films retained their activity during long-term storage at different temperatures. Moreover, both types of active films (LDPE and cellulose) with ppABP showed potential reduction in the population of tested bacteria in dairy products (cheese and paneer), which signifies the use of the ppABP from B. licheniformis Me1 in packaging material to control spoilage and pathogenic organisms in food. All these desirable properties of the activated film with ppABP of B. licheniformis Me1 make them practical for food industrial applications and prove that antimicrobial substances from Bacillus can also be used for developing antimicrobial food packaging materials.


The authors wish to acknowledge The Director, CFTRI, Mysore and Head, Food Microbiology Department, CFTRI for providing the facilities. The authors thank Mr. Binod Prasad for helping in the manuscript preparation. NV acknowledges CSIR for fellowship.