1 Introduction

The growing demand for food, which has accompanied exponential growth in the global population, has propelled the need for food preservation and packaging technologies focused on sustainability. The growing transition from synthetic plastics to more biodegradable packaging materials has led to an interest in biodegradable polymers as packaging alternatives, with edible films and coatings garnering much attention [1,2,3]. Consumers are also committed to having healthier eating habits, involving the consumption of fresh or minimally processed foods, including highly perishable fruits and vegetables, which pose a challenge to preserving them fresh [4, 5]. The early onset of spoilage in fresh fruits and vegetables is a significant contributor to postharvest losses and food wastage. Edible films and coatings have shown great potential for ameliorating this problem as well as improving quality.

An edible film or coating is a thin layer of material applied to the surface of food commodities that can be consumed with the food, provide barrier properties, maintain freshness and prevent spoilage among others [6]. The words film and coating are used interchangeably, with the main differences being their method of manufacture and the mode of application on the commodity [7]. Edible films and coatings made from natural biodegradable biopolymers (polysaccharides, proteins and lipids), show promise for preserving fresh fruits and vegetables and will not cause harm to the environment [8]. One recent study used a composite film from industry brewing waste to delay spoilage and improve freshness of fresh strawberries [9]. The conditions during production, as well as the application method of the film/coating, are also important for obtaining a resultant good quality film. Polysaccharide–based edible films such as chitosan and starch have been utilised extensively due to their availability and excellent barrier properties, the latter being a key attribute for the preservation of not only fruits and vegetables [10] but also fish products [11]. Starch has been categorised as the polysaccharide of greatest importance for film formulations and starch–based films and coatings are very common due to the natural abundance and low cost of starch [12]. Although the best film–forming properties are exhibited by polysaccharides with linear and neutral polymer chains, starch and chitosan (also from a naturally abundant source) polymers, which are branched and cationic, respectively, also show good film–forming properties. The branching of polysaccharide chains and the presence of charges affect the film–forming properties, as both factors hinder the close association of polymer chains, which negatively impacts the strength of the films formed [2]. The shortcomings of these individual films have led to the production of composite films and the use of additives for the optimisation of film properties, which create an opportunity for additional research aimed at determining the best blends.

Apart from its relative abundance and good film–forming properties, chitosan, a linear β-1,4-D glucosamine obtained from the deacetylation of chitin is preferred for film production because of its inherent antimicrobial effects [7, 8]. The mechanism of action of the antimicrobial properties of chitosan has not been fully elucidated, but researchers believe that the amino functional group of chitosan plays a key role in its interaction with the negative charges of the bacterial cell membrane [13]. Apart from its antimicrobial properties, chitosan has limitations [14]. Chitosan films are incorporated with other components that exhibit antioxidant activity, such as metals (gold and silver), metal oxides (titanium oxide and zinc oxide) and plant extract essential oils, to improve antimicrobial properties [8, 15, 16]. Plant extracts are preferred because they contain natural, organic compounds that are generally regarded as safer and healthier [11]. Plectranthus caninus Roth and Plectranthus barbatus Andrews leaves are part of the Lamiaceae family and have medicinal and nutritional properties and are commonly used in folk medicine [17]. They are commonly referred to as Panadol plants in St. Lucia. Although Panadol is an established antibiotic, there is a paucity of studies on the incorporation of Panadol leaf extracts in ameliorating the antimicrobial properties of chitosan composite films.

The banana family of plants (Musaceae) is quite prevalent and considered a staple food, with bananas being the second most abundant fruit group in the world [18]. The regular banana variety seems to be more utilised than the other varieties, such as plantain. Plantains are a bit harder and less palatable in the green stage and are mainly consumed in the ripened stage. The underutilisation of plantains, ultimately leads to significant spoilage as they are highly perishable. Their starch availability served as the basis for their use as a polysaccharide based edible film in this study. The study objectives were to analyse the physicochemical and anti-microbiological properties of composite films produced by the casting method from two common polysaccharides (plantain starch and chitosan) by incorporating two concentrations (0.2% and 2%) of the Panadol leaf extracts from two plant species (Plectranthus barbatus Andrews and Plectranthus caninus Roth).

2 Materials and methods

2.1 Collection and preparation of plant material

Plectranthus barbatus Andrews and Plectranthus caninus Roth leaves were obtained from plants in the southwestern district of Choiseul, St. Lucia. The plants were washed with tap water, rinsed with distilled water and dried in a conventional oven at 50 °C for 24 h. The dried leaves were weighed, packaged in Ziploc® bags and stored at room temperature. Ethanolic extraction from the dried leaves was carried out using the methods described in Saleh et al. [19] and Lyekowa et al. [20] with modifications. The leaves were ground into a powder using a blender and weighed using an Ohaus top pan balance.

Fifty grams of powder from each leaf type were added to a 1000 mL beaker, and 450 mL of 99% ethanol was added (1:9 ratio). The beaker was then foiled, and the mixture was allowed to steepen in a dark cupboard for 3 days at room temperature followed by 2 days at 5 °C. The mixture was then vacuum-filtered through Whatman No. 1 filter paper. The residue was prepared for a second extraction, and the filtrate was placed in a 600 mL beaker and allowed to evaporate in the fume hood. The dried extract was weighed, and the percent yield was calculated. The leaf extract from Plectranthus barbatus Andrews was referred to as leaf extract 1 (LE1), and the leaf extract from Plectranthus caninus Roth was referred to as leaf extract 2 (LE2).

2.2 Plantain starch extraction

Mature green plantains pulp and peel were used for starch extraction. The peel was placed in 0.5% citric acid to prevent darkening. Starch extraction was performed following the method of Ogechukwu et al. [21] with some modifications. A total of 2.80 kg of pulp was steeped in 5.6 L of a 0.25% (w/v) sodium hydroxide (NaOH) solution at a 1:2 ratio of pulp to solution at a temperature of 4 °C in a chiller for 24 h. The NaOH solution was decanted, and the steeped plantain was washed 3 times with distilled water before being blended with 5.6 L distilled water for 2 min. The slurry was filtered twice through a 150-mesh screen and through a 4–layered muslin cloth. The filtrate was allowed to stand overnight to allow the starch to settle. The sediment was centrifuged using a Sorvall Superspeed centrifuge (model S/N A 3815; Thermo Scientific, Asheville, North Carolina, USA) at 3000 rpm for 15 min. The washing and centrifugation steps were repeated until a white starch layer was obtained. The starch was dried at 40 °C for 24 h, and the final weight recorded. The procedure was repeated for starch extraction from the peels (2.95 kg).

2.3 Total phenolic content, total flavonoid content and antioxidant activity of the leaf extracts

The total phenolic content of the leaf extracts was evaluated using the Folin–Ciocalteu method with gallic acid (Sigma Aldrich, St. Louis, Missouri, USA) as the standard [22]. Briefly, 0.1 mL of each dilution or film extraction solution was mixed with 7 mL of distilled water and 0.5 mL of Folin–Ciocalteu reagent. The mixture was incubated at room temperature for 8 min, after which 1.5 mL of sodium carbonate (7.5%) and 0.9 mL of distilled water were added. The mixture was stored in the dark for 2 h, after which the absorbance at 760 nm was measured using a UV–vis spectrophotometer (model S/N 2RIN326001; Thermo Fisher Scientific, Wisconsin, USA). The absorbances of the leaf extract samples were used to determine the concentration of total phenolics (mg/g gallic acid equivalents) in the samples using the standard curve equation.

The total flavonoid content of the extracts was determined using the aluminium chloride method with 95% quercetin hydrate (Acros Organics) used as the standard [22, 23]. A 100 µL of each sample, 4 mL of distilled water and 0.3 mL of 5% NaNO2 were added to a test tube. After 5 min, 0.3 mL of 10% AlCl3 was added to the mixture, and after 6 min, 2 mL of 1 N NaOH was added and this was followed by 3.3 mL of distilled water to bring the volume to 10 mL. The mixture was vortexed, and the absorbance was measured at 510 nm using a UV–vis spectrophotometer. For quantification, a quercetin standard curve was generated (0.1–5.0 mg/mL), and the mean sample (leaf extract) absorbance was compared to the curve to determine the flavonoid content (mg/g quercetin equivalent).

The antioxidant activity of the leaf extracts was determined using the radical scavenging activity of DPPH [22, 24]. The extract solution (0.2 mL) was mixed with 7.8 mL of DPPH in a sterile test tube, vortexed and stored at 37 °C for 30 min. The absorbance was measured at 517 nm with ethanol (99%) as the blank using a Thermo Scientific UV–vis spectrophotometer. The absorbance of the DPPH radical (0.1 mM) was also measured as a control with 0.2 mL of ethanol (99%) replacing the extract solution as described above. The antioxidant activity was calculated from equation (Eq. 1):

$$\% {\text{ DPPH inhibition }} = \, \left[ {\left( {{\text{A}}_{control} {-}{\text{ A}}_{sample} } \right)/{\text{A}}_{control} } \right] \times {1}00$$
(1)

where A is the absorbance at 517 nm.

2.4 Preparation and treatment of film–forming solutions

Film-forming solutions were prepared using extracted plantain starch and low-molecular weight chitosan (Sigma–Aldrich, St. Louis, Missouri, USA), following modified methods from Gao et al. [14] and Wang et al. [25]. To create the chitosan solution, 2.5 g of chitosan was dissolved in 100 mL of 1% acetic acid at 60 °C. After cooling to 40 °C, 1 mL of glycerol was added before the solution was poured into petri dishes and dried for 4 days at room temperature. For the chitosan-starch films, 100 mL of a 2.5% chitosan solution was mixed with 100 mL of a 5% starch solution at 90 °C. After cooling, 2 mL of glycerol was added, and the mixture was poured into petri dishes and dried. Additional solutions were prepared by incorporating 2% and 0.2% leaf extracts after the mixture had cooled.

2.5 Physicochemical analysis of unripe plantain pulp starch

The moisture, ash, protein and lipid contents of the unripe plantain pulp starch were determined using AOAC methods [26]. Additionally, moisture and ash percentages were also determined for the various films formed along with colour of the plantain pulp starch.

The moisture content was determined using the oven convection method. The samples were placed in a preheated Precision Theclo convection oven (model S/N 605061277; Virginia, USA) at 105 °C for 2 h, after which they were cooled in a desiccator for 30 min to ambient temperature and subsequently weighed. The moisture content of each sample was determined from equation (Eq. 2):

$${\text{Moisture \% }} = \,\left( {\left[ {M_{initial} \, - \,M_{dried]/} \,M_{initial} } \right]} \right)\, \, \times { 1}00$$
(2)

where M initial is the initial mass before drying and M dried is the mass after drying expressed in grams (g).

The ash content was measured gravimetrically using a box furnace. Weighed samples in crucibles were placed in a box furnace (model BF51794C-1; North Carolina, USA) and heated at 550 °C for 5 h. After heating, the crucibles were allowed to cool in a desiccator for 30 min. The contents of each crucible were weighed, and the weight of the residue was calculated. The ash content (% ash) was calculated from equation (Eq. 3):

$${\text{Ash }}\% \, = \, \left( {{\text{weight of residue}}/{\text{weight of starting material}}} \right) \times {1}00$$
(3)

The crude protein content of the samples was determined using the Kjeldahl method. A Gerhardt digestion block (model KBL20S) and distillation unit (model S/N VAP004667; Gerhardt Analytical Systems, Konigswinter, Germany) were used, and the nitrogen and subsequent protein percentages were calculated using the following equations (Eqs. 4 and 5):

$$\% {\text{ Nitrogen }} = \, \left( {{\text{mL standard }} - {\text{mL blank}}} \right) \, \times {\text{ Molarity of acid }} \times { 1}.{4}00{7})/{\text{weight of sample }}\left( {\text{g}} \right)$$
(4)
$$\% {\text{ Protein }} = \, \% {\text{ Nitrogen }} \times { 6}.{25}$$
(5)

The lipid content was determined gravimetrically by solvent extraction methods using the SER 148 solvent extraction unit (model S/N 344546) with petroleum ether as the solvent. The lipid content was calculated using equation (Eq. 6):

$$\% {\text{ Lipid }} = \,\left( {{\text{T}}2 - {\text{T}}1} \right)/{\text{SW}}\, \times {1}00$$
(6)

where T1 and T2 are the weights of the cup (g) before and after extraction, respectively, and SW is the sample weight (g).

The amylose content of the extracted starch was determined spectrophotometrically using the method of Elvis [27]. To create a standard curve, a stock solution of pure amylose (0.4 mg/mL) was diluted to produce a series of concentrations ranging from 0.004 to 0.02 mg/mL. Each dilution was treated with 0.2% iodine solution, and the absorbance of the resulting blue complex was measured at 620 nm using a UV–vis spectrophotometer after 20 min of incubation. For the starch sample, a 1 mg/mL stock solution was prepared, and 5 mL of this stock solution was used to create a similar dilution series. Each diluted sample was incubated with iodine solution for 20 min, and absorbance was measured at 620 nm. The absorbance values were compared to the standard curve to determine the amylose content of the starch sample, as calculated using equation (Eq. 7).

$${\text{Amylose content }} = \, ({\text{C}} \times {\text{V}}_{{1}} \times {1}00\% )/{\text{V}}_{{2}} \times {\text{M}} \times {1}000)$$
(7)

where C is the amylose content in mg from the standard curve, V1 is the volume of starch solution prepared and V2 is the volume of solution used.

The pasting properties of the extracted plantain starch were determined using AACC methods [28] with the Rapid Visco Analyser (model S/N 2031531; Newport Scientific, New South Wales, Australia).

The pH of the various film–forming solutions was measured using a Hanna pH meter (model pH 211; Hanna Instruments, Rhode Island, USA).

The carbohydrate content was determined using the sulfuric acid-UV method as outlined by Albalasmeh et al. [29]. A glucose standard curve was prepared by diluting a 100 mg/L glucose stock solution to concentrations of 10, 30, 50, and 70 mg/L. For analysis, 1 mL aliquots of each carbohydrate solution (both dilutions and stock) were mixed with 6 mL of concentrated sulfuric acid, vortexed, and then allowed to cool. The absorbance was measured at 315 nm using a UV–vis spectrophotometer. The absorbance of a 100 mg/L starch solution, treated similarly, was compared to the glucose standard curve to determine its carbohydrate content (mg glucose/L).

The water and oil absorption capacities were determined using methods employed by Sofi et al. [30], with slight adjustments. The weight of an empty 50 mL centrifuge tube was measured, and approximately 1 g of starch was weighed and added to the tube along with 10 mL of either water or oil. The mixture was vortexed for 15 s at 5-min intervals for a total of 20 min. The mixture was then centrifuged using a Sorvall Superspeed centrifuge (model S/N A 3815; Thermo Scientific, Asheville, North Carolina, USA) at 3000 rpm at 25 °C for 10 min, after which the supernatant was discarded. The weight of the centrifuge tube containing the residue sample was measured, and the gain in weight was used to calculate the oil or water absorption capacity from equation (Eq. 8):

$${\text{Oil}}/{\text{water absorption capacity }}\% \, = \, \left( {\left[ {{\text{M}}_{final} {-}{\text{ M}}_{initial} } \right] \, /{\text{ M}}_{initial} } \right){\text{ x 1}}00$$
(8)

where M initial is the mass of the empty centrifuge (g) and M final is the mass of centrifuge tube with residue (g).

2.6 Physicochemical assessment of the various film types

Colour readings were recorded using a Konica Minolta CR-410 chroma meter calibrated with a white standard tile. The CIELAB colour coordinates L*, a* and b* were measured at room temperature and readings were taken for the three different films on days 5 (after solidification) and 12.

The film thickness was measured accurately to the nearest 0.01 mm using a Fischer Scientific digital Vernier calliper (S/N 130195756;) according to the ASTM standard [31]. The measurements were taken at 5 different points along the film, including the centre, and three separate films were assessed. The average thickness for each film type was calculated to determine the density of the corresponding film. The thickness and radius of the films were used to calculate their volume from equation (Eq. 9):

$${\text{Volume }} = { 4}\Pi {\text{r}}^{{2}} \times {\text{ h}}$$
(9)

where r is the radius and h is the height represented by the thickness. The weights of the respective films were also measured, and the densities of the films were calculated using equation (Eq. 10):

$${\text{Density }} = {\text{ m}}/{\text{v}}$$
(10)

where m is the mass (g) and v is the volume (cm3).

The moisture content of the film compositions was determined by measuring the weight loss of films upon drying in an oven at 105 °C for 24 h following the ASTM standard procedure [32]. The ash content was measured gravimetrically using a box furnace as previously described.

The optical properties of the films were measured in both the ultraviolet and visible regions of the light spectrum according to the method of Park et al. [33]. A rectangular sample of each film (4 cm × 1 cm) was placed in a cuvette which was placed in a UV–vis spectrophotometer, and the absorbances at 300 nm and 600 nm were measured for each of the three film types using an empty cuvette as a blank. The opacity of each film type was calculated using the average absorbance at 600 nm and the average film thickness via equation (Eq. 11):

$${\text{Opacity }} = {\text{ Abs}}_{{{6}00}} /{\text{ x}}$$
(11)

where x is the average film thickness and Abs600 is the average absorbance at 600 nm.

The water solubility of the films was determined by the percentage of dry mass dissolved in water after 24 h of exposure using the method of Ogechukwu et al. [21]. The films (4 cm × 1 cm) were weighed using an analytical balance (model PW 254; Adam Equipment Incorporated, Connecticut, USA) to determine the initial dry weight (Wi). The strips were placed in 50 mL of distilled water in a petri dish with agitation for 24 h and then removed and dried in a Precision Thelco Laboratory oven at 105 °C for 6 h. The strips were weighed to determine the final dry weight (Wf) and the water solubility was calculated from equation (Eq. 12):

$${\text{Water solubility }}\left( \% \right) \, = \, \left[ {\left( {{\text{W}}_{{\text{i}}} {-}{\text{ W}}_{{\text{f}}} } \right)/{\text{W}}_{{\text{i}}} \times { 1}00} \right]$$
(12)

where Wi is the initial dry weight of the film and Wf is the final dry weight of the film.

The swelling degree of the films was determined by the difference in weights method as described by Ogechukwu et al. [21]. Film strips (4 cm × 1 cm) were weighed (Wi) and placed in petri-dishes containing 30 mL of distilled water. The strips were removed after 24 h, the excess water was gently removed from the surface with filter paper, and the films were weighed (Wf). The swelling degree of each of the three films was calculated using equation (Eq. 13):

$${\text{SD }}\left( \% \right) \, = \, \left[ {\left( {{\text{W}}_{{\text{f}}} {-}{\text{ W}}_{{\text{i}}} } \right)/{\text{W}}_{{\text{i}}} \times { 1}00} \right]$$
(13)

where Wi is the initial weight of the film and Wf is the final weight of the film.

The water vapour transmission rate (WVTR) of the various films was determined gravimetrically using the ASTM method [34] with some modifications. Cylindrical test vials (18 mm × 70 mm) were filled with 25 mL of distilled water, and the test films were mounted on the open ends of the vials. Parafilm was used along the circumference of the opening to ensure that the mounted films were airtight. Duplicate samples were set up for each film type, and the initial weight of each test vial was measured using an analytical balance. The vials were placed in a fume hood for 12 h a day, and weight loss was measured daily for 1 week. A graph of weight loss (g) against time (days) was plotted to determine the WVTR (expressed as the slope of the graph).

2.7 Microbiological evaluation of the composite films

2.7.1 Preparation of culture media

The bacterial strains Escherichia coli (E. coli) ATCC 35218, Staphylococcus aureus (S. aureus) ATCC 12600, Shigella sonnei (S. sonnei) ATCC 25931 and Shigella flexneri (S. flexneri) ATCC 12022 were cultured through passage 3. The E. coli and S. aureus strains cultured on Oxoid Mueller Hinton Agar (MHA) (CM0337, Thermo Fischer Scientific, Massachusetts, USA) plates were obtained from the laboratory (passage 1) and the Shigella strains were cultured from culti–loops on Oxoid Nutrient Agar (NA) (CM0003) plates.

2.8 Differentiating bacteria: Gram staining

The slides for Gram staining were prepared by heat fixation. The fixed sample was stained with crystal violet solution for 1 min and gently washed with deionised water. Subsequently, the slide was flooded with iodine solution for 1 min, and the wash was repeated with deionised water. The slides were decolourised with ethanol (95%) for 15 s, rinsed, and counterstained with safranin for 1 min before being blot-dried with bibulous paper. The samples were viewed under a microscope (40X and 100X) with oil immersion to determine whether the fixed bacteria were Gram–positive/negative.

2.9 Identifying bacteria: selective media streaking

Various selective media were used for further identification of the various cultured bacteria. S. aureus ATCC 12600 was streaked on Oxoid Mannitol Salt Agar (MSA) (CM0085), Oxoid Baird Parker Agar (CM0275), E. coli ATCC 35218 on Violet Red Bile Agar (VRBA) (M049-500G, Hi-Media, India), Oxoid Eosin-Methylene Blue (EMB) (CM0069) agar, S. sonnei ATCC 25931 and S. flexneri ATCC 12022 on Oxoid MacConkey agar (CM0007) and Oxoid XLD agar (CM0469). The various agar plates were prepared according to the manufacturer’s instructions, and these were allowed to solidify and stored at 4 °C. The refrigerated plates were acclimatised to room temperature before streaking. After streaking, the plates were placed inverted in a 35 °C incubator for the required incubation times, as specified by the agar type. VRBA, XLD, EMB and MacConkey agars were incubated for 24 h, whereas Baird–Parker Agar and MSA were incubated for 48 h.

2.10 Previous contamination test

A previous contamination test was carried out with slight modifications as previously described by Lozano-Navarro et al. [35] to test the ability of the films to prevent microbes from contacting the surface during handling. A 1 cm2 film sample from each film type was added to each sterilised vial containing sterilised distilled water (5 mL). After 24 h, 1 mL of each sample was added to a sterile petri dish, and previously prepared Oxoid Plate Count Agar (PCA) (CM0325) was added to the petri dish. The petri dishes were labelled and placed inverted in a 32 °C incubator for 48 h.

2.11 Culturing bacteria: testing for nutrient broth growth

To facilitate antimicrobial testing, the bacterial cultures from the agar plates were cultured in Oxoid Nutrient Broth (CM0001) to allow swab plating. The nutrient broth (10 mL portions) was distributed in test tubes and autoclaved at 121 °C for 15 min. Colonies of E. coli ATCC 53218 (3 colonies), S. sonnei ATCC 25931 (4 colonies) and S. aureus ATCC 12600 (4 colonies) were transferred to tubes containing nutrient broth and placed in a shaking incubator at 37 °C and 100 rpm. After 3 h, 0.1 mL of sample from each tube was transferred to tubes containing fresh nutrient broth and placed in a shaking incubator for another 3 h under the same conditions as previously described. A sterile swab was dipped into the E. coli nutrient broth culture mixture, pressed against the wall of the tube to remove excess liquid and used to swab the entire surface of the NA plate. This process was repeated on an MHA plate, and the other bacterial strains (S. sonnei) were swabbed on both types of nutrient plates, as was the case for S. aureus on the MHA. The plates were inverted in a 35 °C incubator and checked for growth (18–24 h).

2.12 Antimicrobial testing: Disk diffusion method

The disk diffusion method was carried out using circular film samples (1 cm in diameter) from the various film types [35, 36]. MHA plates were prepared for inoculation and the bacterial species (E. coli, S. aureus and S. sonnei) were cultured in nutrient broth as previously described. The liquid cultures were used to inoculate the entire surface of MHA plates (duplicate for each bacteria) by swabbing with sterile swabs. After allowing the cultures to dry, one circular sample from each film type was placed on the surface of each inoculated MHA plate, ensuring that the samples were adequately spaced out. Circular filter paper (1 cm in diameter) dipped in 10% alcohol, water and ampicillin was also placed on the surface of the inoculated plates as a control. The plates were inverted in a 35 °C incubator overnight, and the antibacterial activity recorded by observing the presence or absence of an inhibition zone around the circular samples [8].

2.13 Antimicrobial testing: Plate count agar (PCA) method

Various samples were prepared to investigate the antimicrobial effects of the leaf extracts by examining growth via PCA. PCA and nutrient broth cultures of E. coli, S. aureus and S. sonnei were prepared as previously described. The bacterial cultures (200 µL) and 4800 µL of leaf extract (0.2%) were pipetted into sterile petri dishes in a biosafety cabinet. PCA was poured into the petri dishes with careful shaking to ensure proper mixing of the samples. This process was repeated with 4800 µL of peptone water, ethanol extract (10%) or ampicillin.

2.14 Statistical analysis

The data are reported as the average ± standard deviation of triplicate analyses unless otherwise stated. The differences among films with regard to physicochemical parameters were analysed using single factor ANOVA, where p ≤ 0.05 represented statistical significance, using Minitab software (version 21.4.10).

3 Results and discussion

3.1 Yield of leaf and starch extracts

Both plantain starch extraction and leaf extraction produced favourable yields, although these extraction methods may be further optimised for film formulation. The overall extraction yields for the plantain peel and pulp seemed slightly lower at 1.20% and 9.76% (Table 1), respectively, compared to the previously reported results of plantain pulp yields ranging from 6.67–15.00% [27], and 1.86 g/kg on a dry mass basis [20]. One study reported a yield of 6% for another starch source from male banana pulp [37]. Extraction yields of 12.0 g/kg and 97.6 g/kg were recorded for plantain peel and pulp, respectively. The optimisation of starch extraction by the wet milling process allowed a favourable extraction yield. However, researchers have reported high extraction yields of 16.6 to 48.5% for plantain peels [38], while others have reported lower yields (4.50 to 5.59%) [39] than what was obtained in this research. This low extraction yield could have resulted from the failure to optimise the extraction by the use of an alkali, which has been reported to increase the starch extraction yield [40].

Table 1 Physicochemical properties of unripe plantain starch
Full size table

Most extraction processes involve a compromise between the extraction yield, purity and integrity of the extract, as conditions for improving one may inhibit the other. For example, improving the starch extraction yield by increasing the alkaline concentration may result in reduced purity [40]. A significant amount of protein was produced from starch extraction, which resulted in an increase in the loss of starch in the protein fraction during extraction [41]. Temperature and alkaline conditions affect the chemical composition and purity of the starch obtained, which may impact the film properties. Other factors that may have led to the reduced yield include the use of simple separation techniques, such as scraping off the protein layer and filtering through the muslin cloth. Some starch would have been lost with both of these techniques. There was also starch loss in the discarded fibrous waste and the decanted water (resuspended fine starch). Ethanolic leaf extraction produced similar results and good yields of 8.20 and 8.44% for the 2 types of Panadol leaves utilised (Table 2) suggesting comparable extraction efficiencies. The purity and stability of the leaf extracts were maintained by the high-quality ethanol used and the temperature conditions. It appeared that the use of the dried leaf samples produced more stable extracts and the concentration of polyphenols in the extract was also optimised by the use of ethanol [42]. In summary, the similar and good yields obtained for Plectranthus barbatus (LE1) and Plectranthus caninus (LE2) using ethanolic extraction likely stemmed from the chemical similarity of the plants, the efficacy of ethanol as a solvent, and the consistency in extraction methods and leaf quality.

Table 2 Percentage yield, total phenolic content, total flavonoid content and DDPH antioxidant activity of the Panadol leaf extracts
Full size table

3.2 Physicochemical properties of unripe plantain pulp starch

3.2.1 Proximate analysis

The proximate analysis, as shown in Table 1, revealed a starch purity of 85.9%, and this also represented the carbohydrate content determined by the difference method [26]. The proximate analysis of the extracted starch correlated with the results reported in the literature [43]. The moisture content of the extracted starch was slightly greater than the reported values of 11.56 to 13.15% for plantain starch cultivars [35], 11.7% for green banana starch [37] and 11.20% for native plantain starch [43] which indicated that the starch could have undergone further drying. Unripe plantain pulp starch exhibited a low ash content, indicating minimal mineral impurities and low protein and crude fat contents. The small variations in these values compared to the reported results could be attributed to factors such as the specific variety of plantain, growing conditions, and processing methods [10].

3.3 Amylose content

The amylose content of the starch sample was calculated as 41.8 ± 0.41% (Table 1), and was higher than that typically reported (13.87–38.79%) for plantain starch but fell within the range reported for banana starches (21.91 to 42.07%) [43, 44], indicating that the plantain variety utilised may be more closely related to bananas. Growth conditions such as soil type, climate and geographic location also play a role in the amylose content of crops [10, 18], but little research has been conducted on Caribbean plantain varieties for comparison. This high amylose content was advantageous because it provided the starch with good film–forming properties as films made from amylose are stronger than those made from amylopectin [12]. This strength was exhibited with the films produced.

3.4 Pasting properties

Table 1 highlights the pasting properties of the extracted plantain starch. Some of the parameters corresponded with results reported in the literature for plantain starch, but others showed deviations [43]. The peak viscosity as illustrated in Fig. 1 indicates the maximum thickness of the paste. For unripe starch, this peak is often lower than that of ripe starch due to less extensive swelling. A high peak and final viscosity were recorded, indicating good gelling properties. The peak time and pasting temperature recorded were lower than the values reported for plantain starch [43]. The pasting properties of starch depend on its ability to form a gel or paste, which is influenced by the ability of the starch granules to hydrate and swell [12]. The hydration and swelling power of the starch granules are further influenced by several factors, including the amylose/amylopectin ratio, granule size and extent of hydrogen bonding between the granules [7, 10]. The swelling power of starch will ultimately influence its pasting properties. High-quality starch has a high swelling potential and low solubility [27]. The extracted plantain starch exhibited good swelling power (12.94%), which was greater than the swelling powers of 9.48–10.76% [27], and 10.28% [39] reported for plantain starch. The swelling power of starch is influenced by the amylose/amylopectin ratio which is positively correlated with the amylopectin content [27, 45]. Amylose inhibits water binding, thus reducing the swelling power and viscosity at elevated temperatures. This inhibition occurs as a result of amylose complexing with the phospholipids in the starch granules. On the other hand, a high amylopectin content results in high viscosity and swelling power at low temperatures [45]. This theory supported the results obtained for the extracted plantain starch, which exhibited a high swelling power and viscosity at a lower temperature than what has been reported for plantain starch (low pasting temperature of 50.15 °C), which indicated a higher amylopectin content than what was previously reported (lower amylose). The presence of holes and channels in the starch structure has also been reported to increase hydration, thus increasing the swelling power and viscosity by inhibiting the amylose effect [46]. This may account for the results obtained for extracted starch with a high amylose content. The size of the starch granules becomes insignificant for swelling and viscosity values, when there is a major difference in the chemistry of the starches [44].

Fig. 1
figure 1

Pasting curve of plantain pulp starch

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The lower pasting temperature (50.15 °C) and peak time (4.40 min) recorded indicated that the viscosity of the starch started increasing at a lower temperature, reaching its peak viscosity in a shorter time frame. Pasting temperature and peak time ranges of plantain starch of 81.37–87.13 °C and 4.72–7.00 min, respectively have been reported [43]. The high setback viscosity and final viscosity influence the properties of the final paste and product [43]. A high setback viscosity is indicative of the formation of a cohesive paste for the extracted starch. This was important for the production of films with good texture.

3.5 Total phenolic content, total flavonoid content and antioxidant activity of the leaf extracts

The phenolic content (TPC) expressed as gallic acid equivalents (mg/g) was 86.6 and 69.6 mg/mL, and the flavonoid content (TFC) was 91.2 and 74.4 mg/mL quercetin for Plectranthus barbatus Andrews (LE1) and Plectranthus caninus Roth (LE2), respectively (Table 2). The DPPH inhibition of LE1 and LE2 antioxidant activity was 74.60% and 64.77%, respectively. The use of dried leaves as well as the use of ethanol as the extractant ensured the maintenance of phenolic and antioxidant activity, which was supported by the high DPPH antioxidant activity, phenolic content and flavonoid content (Table 2). Ethanol was reported as the preferred solvent for extraction of Plectranthus compared to aqueous extracts as a result of the high antioxidant activity of the ethanolic extracts as previously noted [42]. The total phenolic content was significantly higher in LE1 compared to LE2 which may suggest that Plectranthus barbatus Andrews has a richer profile of phenolic compounds associated with potential health benefits. The total flavonoid content also differed with LE1 showing greater levels compared to LE2 (Table 2). Flavonoids are known for their antioxidant properties suggesting that LE1 may offer greater protection against oxidative stress. The DPPH antioxidant activity results indicated that LE1 exhibited higher inhibition compared to LE2. It was reported by Cordeiro et al. [47] that flavonoids, cinnamic acid derivatives, steroids and ellagic acid were present in both aqueous and organic extracts of Plectranthus barbatus Andrews, which supports our findings and the potential of Plectranthus barbatus Andrews as a source of natural antioxidants.

3.6 pH of the film–components

The pH of the various solutions utilised in film formation was measured as highlighted in Table 3. The pH of the film–forming solution plays an important role in the antimicrobial properties of the film. Microbial growth is inhibited in highly acidic environments. Compared with starch solutions, which are alkaline in pH, chitosan film solutions exhibited an acidic pH (Table 3), which is believed to be a contributing factor to the antimicrobial properties of chitosan films. The weak alkaline nature of the starch solution did not outweigh that of acidic chitosan, as the pH of the composite mixture of chitosan and starch was also acidic, which probably helped confer antimicrobial properties to the composite films. The acidic nature of the acetic acid solution and leaf extracts as shown in Table 3 also helped contribute to the acidic pH of the final film solution mixture.

Table 3 pH of various film-forming solutions
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3.7 Colour parameters of the unripe plantain pulp starch and the various film formulations

The purity of the extracted plantain pulp starch was reflected by its colour parameters, with an L* value of 100, indicating maximum whiteness, and there was relatively no significant change (p > 0.05) in colour parameters from day 0 to 15 (Table 4). White starches contain lower levels of proteins and pigments, which make them suitable for industrial applications particularly in the food and pharmaceutical sectors [37].

Table 4 Colour of plantain pulp starch and film formulations
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The L* a* b* colour parameters did not change significantly (p > 0.05) with age between days 0 and 12 for the 2.5% chitosan-only film, but the parameters differed significantly (p < 0.05) for the composite films both with and without the leaf additive, with the b* values showing the most deviation as shown in Table 4. Over time, certain coloured compounds present in the natural leaf extracts may degrade or oxidize, leading to a reduction in the intensity of colour, thus increasing the lightness (L* value). Additionally, drying of the film with aging could have caused an increased transparency or changes in the porosity of the film with time which may have affected how light is reflected and potentially could lead to an increase in lightness. Nearly all the film types had positive a* values, which corresponded to redness, and with time, a reduction in a* values (which represents the red-green spectrum) was observed which could have been caused by anthocyanins or other flavonoids present in the leaf extract which may have degraded. Additionally, the interaction between the leaf extract and the starch/chitosan matrix might have altered the colour balance [18, 48]. For example, changes in pH or chemical environment in the film matrix over time could affect the stability of pigments. Oxidative reactions over time could lead to the loss of red pigments or changes in their structure, causing a shift towards a greener or less intense colour. There was a shift towards increased yellowness (increased b* value) for films with higher leaf extracts after 12 days. The increased yellowing of the films with aging (change from blue to yellow) by the increasing b* value, may be partly due to the presence of phenolic and antioxidant compounds (usually of colour) in the leaf extracts being degraded by the presence of light and oxygen [35] as well as changes in the film matrix.

3.8 Physicochemical parameters of the various film components

3.8.1 Functional properties

The different types of films formed are illustrated in Fig. 2 and some physicochemical properties of the different types of films formed are highlighted in Table 5. There was a significant difference (p < 0.05) in thickness for the different film types comprising chitosan (2.5%), chitosan (2.5%)/plantain starch (5%) and chitosan (2.5%)/plantain starch (5%) with leaf extracts, but there was no significant difference (p˃0.05) for the two different films with leaf extracts and the two concentrations of leaf extracts.

Fig. 2
figure 2

Different types of films synthesised: 1 chitosan (2.5%), 2 chitosan (2.5%) and starch (5%), 3 chitosan-starch with 0.2% leaf extract and 4 chitosan-starch with 2% leaf extract 1 (CH), 2 (CH + PS), 3 (CH + PS + 0.2%LE), 4 (CH + PS + 2.0%LE)

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Table 5 Physicochemical parameters of film components and film formulations
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The addition of starch paste increased the film density. Plantain starch, when incorporated into the chitosan matrix, probably contributed to additional molecular weight. Starch molecules, which are typically larger and bulkier than individual chitosan molecules, can add to the overall mass and density of the film through bonding interactions making the film structure more compact and thus increasing the film thickness and density [48]. There was no significant difference in density with the two types and concentrations of Panadol leaf extracts.

It appeared that the films were not dried completely as the moisture content ranged from 17.1 to 25.5% for the different film types. Despite drying the sample starch to a constant mass, there was still some residual moisture present. Plantain starch is highly hydrophilic and has a significant water-binding capacity. When added to the film formulation, it tends to absorb and retain more moisture, which increases the overall moisture content of the film and some moisture is essential for functionality and it plays an essential role in the ability of starch films to confer good sealing properties [49]. This residual water represented the water from the monolayer, which was tightly bound and required harsher drying conditions for its removal [50]. The moisture content lost during drying of the starch sample represented the free and loosely bound water particles. The moisture content decreased with decreasing concentrations of leaf extract and vice-versa. Panadol leaf extracts contain various compounds that can influence the moisture content of the film. It was hypothesized that some of these compounds may affect the film's ability to retain or release moisture. This could be attributed to the reduced interaction of the films with water, as there was an increase in the interaction with the hydrophilic components of the extracts; this effect was less pronounced as the number of hydrophilic groups significantly increased with increasing leaf extract concentration.

The ash content of the films did increase when compared to that of the native starch as several chemical compounds were added to the film formulation. While chitosan has a relatively low ash content, plantain starch may have contributed to the ash content due to impurities or minerals present in the starch, but these would vary depending on the source and processing conditions. The variations in the ash levels in the film particularly with the leaf extracts could be as a result of the presence of minerals or other ash-forming substances in the extract based on the species used and the processing conditions.

The optical properties detailed in Table 5 analyze light absorption in the UV (300 nm) and visible regions (600 nm) of the spectrum. Absorbance serves as an indirect measure of light transmission, being inversely related to transmittance [51]. Higher absorbance indicates greater light absorption which can be attributed to the presence of specific compounds in the film. Conversely, opacity measures the extent to which light is blocked or absorbed by the film with greater opacity values indicating a stronger light-blocking capacity. Among the samples, the 2.5% chitosan film exhibited the lowest opacity, allowing more light to pass through compared to the other formulations. The inclusion of plantain starch reduced absorbance at 300 nm and increased opacity, suggesting that it affected the film's light absorption properties. Furthermore, the addition of leaf extract 1 (LE1) at 0.2% further reduced absorbance at 300 nm and enhanced opacity relative to the formulation containing only plantain starch. This indicated that LE1 positively influenced the film’s light absorption and opacity. Increasing the concentration of LE1 to 2% increased both absorbance at 300 nm and opacity. This suggested that the higher concentration of extract significantly affected the film's ability to absorb light at this wavelength and further increased its opacity [51]. The leaf extract 2 (LE2) at 0.2% had a similar effect on absorbance as LE1 at the same concentration but resulted in slightly lower opacity compared to LE1. Increasing the concentration of LE2 to 2% also increased both absorbance and opacity, though the changes were not as pronounced as those seen with LE1 at the same concentration. Overall, the presence and concentration of different leaf extracts significantly influenced the absorbance, with LE1 having the most substantial impact. The addition of plantain starch and leaf extracts increased opacity, with higher concentrations of leaf extracts leading to more opaque films. This suggested that the extracts contributed to the film's light-blocking properties similar to recent studies by Nxumalo et al. [48]. Moreover, the leaf extracts modified the film's properties, with higher concentrations resulting in greater opacity and changes in absorbance. Notably, LE1 appeared to have a more pronounced effect on both parameters compared to LE2. Film thickness is also a contributing factor to film barrier properties. Film thickness is influenced not only by the chemical composition of the film–forming components, but also by the type of gelatinisation of the starch. Cold gelatinisation (using alkali) produces thinner films as this type of gelatinisation involves hydrolysis of starch components [10].

3.9 Water solubility of the various film formulations

The water solubilities of the different types of films (Fig. 3a) showed some interesting characteristics. Generally, films with lower water solubility are preferred for their water resistance and stability which make them more durable. Although pure chitosan will not dissolve in pure water, pure chitosan films are very susceptible to water and are degraded in water (highly soluble) [52]. This phenomenon was observed when chitosan films were added to water during this study. The addition of starch (although also hydrophilic) to the chitosan films decreased the solubility, as shown in Fig. 3a. Both chitosan and starch individual films were reported to be susceptible to water degradation [35, 52], but they exhibited synergistic resistance to water when combined. This synergy may have resulted from increased interactions between the starch and chitosan polymers, resulting in reduced interactions with the water molecules. Like water molecules, carbohydrate molecules contain many hydroxyl groups that can form hydrogen bonding with chitosan.

Fig. 3
figure 3

Functional properties of the various film samples measured as: (a) water solubility, (b) water swelling and (c) water vapour transmission rate (WVTR). Error bars represent the standard deviation of mean values of three replicates. Values designated by different letters are significantly different (p < 0.05)

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The addition of leaf extracts yielded some interesting results as the solubility of the films at low concentrations (0.2%) greatly increased but significantly decreased at higher concentrations (2.0%). This behaviour, can attributed to the hydrophilic nature of the leaf extracts and their ability to engage in intramolecular and intermolecular interactions in solution. At low extract concentrations, fewer intramolecular interactions occurred among the chitosan and starch molecules, allowing more free hydrophilic extract molecules to interact with water, thereby increasing film solubility. Conversely, at higher concentrations of leaf extract, the interactions between starch and chitosan were disrupted, allowing intramolecular interactions between the leaf extracts and the polymer molecules. This resulted in fewer free extract molecules available for interaction with water, which decreased the hydrophilicity and solubility of the films. Additionally, the increase in insoluble polyphenols and flavonoids at higher extract concentrations further contributed to the decrease in film solubility [51]. Although the leaf extract conferred enhanced microbial properties, too high a concentration can negatively impact the physiochemical properties of the films. Similar phenomena have been reported in other studies with extracts, such as tea polyphenols [14]. Nxumalo et al. [48] recently reported enhanced swelling degree and water solubility in chitosan-based films infused with plant extracts due to improved hydrophilicity. However, high water solubility, can lead to weaker films that disintegrate easily, compromising their barrier properties.

CH-Chitosan.

PS-Plantain starch.

LE1- The leaf extract from Plectranthus barbatus Andrews was referred to as leaf extract 1.

LE2-The leaf the extract from Plectranthus caninus Roth was referred to as leaf extract 2.

3.10 Swelling degree of the various film formulations

The high swelling degree of the chitosan film (2.5%) also supported the lack of water resistance [39]. The addition of starch significantly decreased the swelling degree of the films (Fig. 3b). Starch does not undergo swelling in water at ambient temperature, as its granules swell only at high temperature, resulting in gelatinisation, as previously mentioned. The addition of the leaf extracts caused a slight increase in swelling degree as a result of their hydrophilic properties [15, 42, 48] and the more open structure resulting from their addition to the film–forming solutions.

3.11 Water vapour transmission rate of the various film formulations

The water vapour transmission rate (WVTR) was determined via the weight loss per day, with slight modifications. Moreover, there was no significant difference (p < 0.05) among the WVTRs of the various film types (Fig. 3c). The films reduced water vapor movement but did not provide significant barrier properties. Their water vapor transmission rate (WVTR) was lower than that of an uncovered vial, but not as low as that of a vial covered with foil. The foil offered superior water vapor barrier properties compared to the films. The hydrophilic nature of the films (all polysaccharides) resulted in their poor ability to prevent water loss. The presence of plasticisers such as glycerol used in the film preparation in this study may have also contributed to increased movement of water across films, as it promoted a less rigid and open structure (reduced intermolecular bonding of polymer chains) [7, 12].

3.12 Microbiological assessment

The results of the previous contamination tests showed no colonies on PCA for most of the films, except for one sample with 2 mould colonies (CH + PS + LE1 (2%) film). Based on the standards, none of the films were contaminated, as plates with ˃ 10 CFU were considered to be contaminated by fungi. This indicated that the films possessed some resistance to environmental microbes and handling and did not need sterilisation before antimicrobial testing.

The disk diffusion method did not produce the anticipated results. There was some inhibition of bacterial growth under the applied disks, but there was no diffusion of antimicrobial activity; thus, zones of inhibition on the outer edges of the disks were not observed. However, it should be noted that the clearing zone under the disks was more prominent for the films with extracts than for the others.

The PCA method was utilized to assess the ability of various film-forming components to inhibit inoculum formation. Bacterial growth was observed on all plates except those containing leaf extracts, indicating that the leaf extracts had antimicrobial properties [7, 35]. On plates showing growth, the number of colonies was too numerous to count (TNTC). Film coatings containing the leaf extracts were expected to exhibit antimicrobial effects on the surfaces of the coated products, which was the intended outcome. The PCA analysis confirmed the antimicrobial properties of the leaf extracts, as no bacterial growth was detected for the various strains inoculated on the PCA plates. Both gram-positive bacteria (S. aureus) and gram-negative bacteria (E. coli and S. sonnei) were inhibited by the leaf extracts. Peptone-enriched PCA served as a positive control, while PCA with alcohol was used to demonstrate that the solvent in the extract did not contribute to the inhibition. However, mould growth on one PCA plate suggested reduced inhibition of mould or potential contamination, as only one plate exhibited this issue.

4 Conclusions

The study successfully extracted plantain starch from both the peel and pulp using alkaline extraction and optimisation techniques. While the plantain pulp yielded high purity and better starch yields than the peel, it is important to note that variations in extraction efficiency can occur based on the specific conditions used and the inherent properties of the raw materials. The extracted starch displayed promising physicochemical properties, including high swelling capacity and relatively low water solubility, which are advantageous for various applications. Notably, the starch also exhibited favourable pasting characteristics, such as a low pasting temperature and peak time along with high peak viscosity, breakdown viscosity and final viscosity. These attributes suggest potential for effective film formation, however the practical implications of these properties in real-world applications require further exploration. The resulting starch-chitosan composite films exhibited improved physicochemical properties, such as increased opacity, density, moisture content and decreased solubility compared to films made with chitosan alone. Moreover, there was no significant difference in the water vapour transmission rate of the various films. It should be emphasized that while there were observable changes in properties like opacity, density, moisture content and swelling index, the significance of these changes may vary depending on environmental conditions and film applications scenarios. The inclusion of the leaf extracts positively influenced the film’s properties but these were dependent on the concentration applied. The higher phenolic and flavonoid contents along with enhanced DPPH inhibition of Plectranthus barbatus Andrews (LE1) compared to Plectranthus caninus Roth (LE2) suggests a need for further investigation into specific bioactive compounds in LE1. At a concentration of 0.2%, LE1 increased film solubility compared to the starch-chitosan film, however increasing the concentration to 2% resulted in a significant decrease in solubility. A similar trend was observed with LE2, indicating a complex relationship between concentration and film properties. The antimicrobial activity of the film–forming solution was evidenced through PCA method results, which indicated inhibition of both gram-positive and gram-negative bacteria by the leaf extracts. However, the disk diffusion method did not yield significant results, highlighting a limitation in the assessment of the antimicrobial efficacy. In conclusion, the study demonstrates the potential benefits of incorporating the Panadol leaf extracts of Plectranthus barbatus Andrews and Plectranthus caninus Roth in improving composite film properties containing 2.5% chitosan and 5% plantain starch. Future research directions could include exploring the use of essential oils from the leaf extracts to minimize colour changes in films, determining minimum inhibitory concentrations for the extracts, testing for yeast and mould inhibition, and optimizing chitosan-starch ratios and concentrations for enhanced film properties. Additionally, employing techniques like scanning electron microscopy and X-ray diffraction to analyse starch morphology, including granule size, and pore availability, could provide valuable insights into the swelling behaviour and gelatinization characteristics of the starch, ultimately influencing its functionality.