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Discover Food
Research
A preliminary investigation oftheproperties ofplantain
starch‑chitosan composite films containing Panadol leaf extracts
ChadLafeuillee1· RohanieMaharaj1
Received: 21 August 2024 / Accepted: 2 January 2025
© The Author(s) 2025 OPEN
Abstract
The study explores composite polysaccharide lms made from plantain pulp starch and chitosan, incorporating extracts
from Panadol leaves of Plectranthus barbatus and Plectranthus caninus to improve physicochemical and antimicrobial
properties. Plantain pulp starch was extracted using 25% NaOH and lms were created via solvent casting by combining
equal volumes of 5% starch and 2.5% chitosan. Phytochemical screening of the ethanolic leaf extracts employed spec-
troscopic methods. Evaluations included antioxidant capacity, total phenolic and avonoid contents, water solubility,
swelling indices, water vapour transmission rates and optical properties. Antimicrobial activity was tested using the disk
diusion method and plate count agar. Antioxidant activities showed % DPPH inhibition of 74.60 ± 0.05 and 64.77 ± 0.07
for Plectranthus barbatus and Plectranthus caninus, with phenolic contents of 86.56 ± 0.03 and 69.59 ± 0.04mg/g gallic
acid equivalents, and avonoid contents of 91.25 ± 0.005 and 74.49 ± 0.003mg/g quercetin equivalents respectively. The
composite lms exhibited increased opacity, density and moisture content alongside decreased swelling indices. Water
solubility varied by component with no signicant dierence in water vapour transmission rates among the lms. Both
gram-positive and gram-negative bacteria were inhibited by the leaf extracts. The starch-chitosan composite lms with
leaf extracts demonstrated enhanced physicochemical and antimicrobial properties making them suitable for sustain-
able food packaging.
Keywords Plantain starch· Chitosan· Edible lms· Panadol leaf· Antimicrobial
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 lms and coatings garnering much attention [1–3]. 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 signicant contributor to postharvest losses and food wastage. Edible lms and coatings have shown
great potential for ameliorating this problem as well as improving quality.
An edible lm 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 lm and
* Rohanie Maharaj, rohanie.maharaj@sta.uwi.edu | 1Department ofChemical Engineering, The University oftheWest Indies, St. Augustine
Circular Road, St.Augustine, TrinidadandTobago.
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coating are used interchangeably, with the main dierences being their method of manufacture and the mode of applica-
tion on the commodity [7]. Edible lms 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 lm 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 lm/coating, are also
important for obtaining a resultant good quality lm. Polysaccharide–based edible lms 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 sh products [11]. Starch has been categorised as the
polysaccharide of greatest importance for lm formulations and starch–based lms and coatings are very common due
to the natural abundance and low cost of starch [12]. Although the best lm–forming properties are exhibited by polysac-
charides 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 lm–forming properties. The branching of polysaccharide
chains and the presence of charges aect the lm–forming properties, as both factors hinder the close association of
polymer chains, which negatively impacts the strength of the lms formed [2]. The shortcomings of these individual
lms have led to the production of composite lms and the use of additives for the optimisation of lm properties, which
create an opportunity for additional research aimed at determining the best blends.
Apart from its relative abundance and good lm–forming properties, chitosan, a linear β-1,4-D glucosamine obtained
from the deacetylation of chitin is preferred for lm production because of its inherent antimicrobial eects [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 lms 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 nutri-
tional 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 lms.
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 signicant spoilage as they are highly perishable. Their starch
availability served as the basis for their use as a polysaccharide based edible lm in this study. The study objectives were
to analyse the physicochemical and anti-microbiological properties of composite lms 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 andmethods
2.1 Collection andpreparation ofplant 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 conven-
tional oven at 50°C for 24h. 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 etal. [19] and Lyekowa
etal. [20] with modications. 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 1000mL beaker, and 450mL 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 3days at room
temperature followed by 2days at 5°C. The mixture was then vacuum-ltered through Whatman No. 1 lter paper. The
residue was prepared for a second extraction, and the ltrate was placed in a 600mL beaker and allowed to evaporate in
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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 etal. [21] with some modications.
A total of 2.80kg 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 24h. The NaOH solution was decanted, and the steeped plantain was
washed 3 times with distilled water before being blended with 5.6L distilled water for 2min. The slurry was ltered twice
through a 150-mesh screen and through a 4–layered muslin cloth. The ltrate 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
Scientic, Asheville, North Carolina, USA) at 3000rpm for 15min. The washing and centrifugation steps were repeated
until a white starch layer was obtained. The starch was dried at 40°C for 24h, and the nal weight recorded. The proce-
dure was repeated for starch extraction from the peels (2.95kg).
2.3 Total phenolic content, total flavonoid content andantioxidant activity oftheleaf 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]. Briey, 0.1mL of each dilution or lm extraction solution was mixed
with 7mL of distilled water and 0.5mL of Folin–Ciocalteu reagent. The mixture was incubated at room temperature for
8min, after which 1.5mL of sodium carbonate (7.5%) and 0.9mL of distilled water were added. The mixture was stored
in the dark for 2h, after which the absorbance at 760nm was measured using a UV–vis spectrophotometer (model
S/N 2RIN326001; Thermo Fisher Scientic, 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 avonoid 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, 4mL of distilled water and 0.3mL of 5%
NaNO2 were added to a test tube. After 5min, 0.3mL of 10% AlCl3 was added to the mixture, and after 6min, 2mL of
1 N NaOH was added and this was followed by 3.3mL of distilled water to bring the volume to 10mL. The mixture was
vortexed, and the absorbance was measured at 510nm using a UV–vis spectrophotometer. For quantication, a quercetin
standard curve was generated (0.1–5.0mg/mL), and the mean sample (leaf extract) absorbance was compared to the
curve to determine the avonoid 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 (0.2mL) was mixed with 7.8mL of DPPH in a sterile test tube, vortexed and stored at 37°C for 30min. The
absorbance was measured at 517nm with ethanol (99%) as the blank using a Thermo Scientic UV–vis spectrophotom-
eter. The absorbance of the DPPH radical (0.1mM) was also measured as a control with 0.2mL of ethanol (99%) replacing
the extract solution as described above. The antioxidant activity was calculated from equation (Eq.1):
where A is the absorbance at 517nm.
2.4 Preparation andtreatment offilm–forming solutions
Film-forming solutions were prepared using extracted plantain starch and low-molecular weight chitosan (Sigma–Aldrich,
St. Louis, Missouri, USA), following modied methods from Gao etal. [14] and Wang etal. [25]. To create the chitosan
solution, 2.5g of chitosan was dissolved in 100mL of 1% acetic acid at 60°C. After cooling to 40°C, 1mL of glycerol was
added before the solution was poured into petri dishes and dried for 4days at room temperature. For the chitosan-starch
lms, 100mL of a 2.5% chitosan solution was mixed with 100mL of a 5% starch solution at 90°C. After cooling, 2mL
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.
(1)
%
DPPH inhibition =
[(
A
control
−A
sample)
∕A
control ]
×
100
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2.5 Physicochemical analysis ofunripe 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 lms 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 2h, after which they were cooled
in a desiccator for 30min to ambient temperature and subsequently weighed. The moisture content of each sample
was determined from equation (Eq.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 5h. After heating, the crucibles were
allowed to cool in a desiccator for 30min. 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):
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):
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):
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 (0.4mg/mL) was diluted to produce a series of concentra-
tions ranging from 0.004 to 0.02mg/mL. Each dilution was treated with 0.2% iodine solution, and the absorbance of
the resulting blue complex was measured at 620nm using a UV–vis spectrophotometer after 20min of incubation. For
the starch sample, a 1mg/mL stock solution was prepared, and 5mL of this stock solution was used to create a similar
dilution series. Each diluted sample was incubated with iodine solution for 20min, and absorbance was measured at
620nm. The absorbance values were compared to the standard curve to determine the amylose content of the starch
sample, as calculated using equation (Eq.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 Scientic, New South Wales, Australia).
The pH of the various lm–forming solutions was measured using a Hanna pH meter (model pH 211; Hanna Instru-
ments, Rhode Island, USA).
The carbohydrate content was determined using the sulfuric acid-UV method as outlined by Albalasmeh etal. [29]. A
glucose standard curve was prepared by diluting a 100mg/L glucose stock solution to concentrations of 10, 30, 50, and
70mg/L. For analysis, 1mL aliquots of each carbohydrate solution (both dilutions and stock) were mixed with 6mL of
concentrated sulfuric acid, vortexed, and then allowed to cool. The absorbance was measured at 315nm using a UV–vis
(2)
Moisture %
=
([
M
initial
−M
dried]∕
M
initial ])
×
100
(3)
Ash %=(weight of residue∕weight of starting material)×100
(4)
%Nitrogen =(mL standard −mL blank)×Molarity of acid ×1.4007)∕weight of sample (g)
(5)
%Protein =%Nitrogen ×6.25
(6)
%Lipid =(T2 −T1)∕SW ×100
(7)
Amylose content =(C×V1×100%)∕V2×M×1000)
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spectrophotometer. The absorbance of a 100mg/L starch solution, treated similarly, was compared to the glucose stand-
ard curve to determine its carbohydrate content (mg glucose/L).
The water and oil absorption capacities were determined using methods employed by So etal. [30], with slight adjust-
ments. The weight of an empty 50mL centrifuge tube was measured, and approximately 1g of starch was weighed and
added to the tube along with 10mL of either water or oil. The mixture was vortexed for 15s at 5-min intervals for a total
of 20min. The mixture was then centrifuged using a Sorvall Superspeed centrifuge (model S/N A 3815; Thermo Scientic,
Asheville, North Carolina, USA) at 3000rpm at 25°C for 10min, 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):
where M initial is the mass of the empty centrifuge (g) and M nal is the mass of centrifuge tube with residue (g).
2.6 Physicochemical assessment ofthevarious 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 dif-
ferent lms on days 5 (after solidication) and 12.
The lm thickness was measured accurately to the nearest 0.01mm using a Fischer Scientic digital Vernier calliper
(S/N 130195756;) according to the ASTM standard [31]. The measurements were taken at 5 dierent points along the lm,
including the centre, and three separate lms were assessed. The average thickness for each lm type was calculated
to determine the density of the corresponding lm. The thickness and radius of the lms were used to calculate their
volume from equation (Eq.9):
where r is the radius and h is the height represented by the thickness. The weights of the respective lms were also
measured, and the densities of the lms were calculated using equation (Eq.10):
where m is the mass (g) and v is the volume (cm3).
The moisture content of the lm compositions was determined by measuring the weight loss of lms upon drying in
an oven at 105°C for 24h following the ASTM standard procedure [32]. The ash content was measured gravimetrically
using a box furnace as previously described.
The optical properties of the lms were measured in both the ultraviolet and visible regions of the light spectrum
according to the method of Park etal. [33]. A rectangular sample of each lm (4cm × 1cm) was placed in a cuvette which
was placed in a UV–vis spectrophotometer, and the absorbances at 300nm and 600nm were measured for each of
the three lm types using an empty cuvette as a blank. The opacity of each lm type was calculated using the average
absorbance at 600nm and the average lm thickness via equation (Eq.11):
where x is the average lm thickness and Abs600 is the average absorbance at 600nm.
The water solubility of the lms was determined by the percentage of dry mass dissolved in water after 24h of
exposure using the method of Ogechukwu etal. [21]. The lms (4cm × 1cm) 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 50mL of distilled water in a petri dish with agitation for 24h and then removed and dried in a Precision
Thelco Laboratory oven at 105°C for 6h. The strips were weighed to determine the nal dry weight (Wf) and the water
solubility was calculated from equation (Eq.12):
where Wi is the initial dry weight of the lm and Wf is the nal dry weight of the lm.
(8)
Oil
∕water absorption capacity %=
([
M
final
−M
initial ]
∕M
initial )x 100
(9)
Volume =4Πr2×h
(10)
Density =m∕v
(11)
Opacity =Abs600∕x
(12)
Water solubility
(%)=
[(
W
i
−W
f)
∕W
i
×100
]
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The swelling degree of the lms was determined by the dierence in weights method as described by Ogechukwu
etal. [21]. Film strips (4cm × 1cm) were weighed (Wi) and placed in petri-dishes containing 30mL of distilled water. The
strips were removed after 24h, the excess water was gently removed from the surface with lter paper, and the lms
were weighed (Wf). The swelling degree of each of the three lms was calculated using equation (Eq.13):
where Wi is the initial weight of the lm and Wf is the nal weight of the lm.
The water vapour transmission rate (WVTR) of the various lms was determined gravimetrically using the ASTM
method [34] with some modications. Cylindrical test vials (18mm × 70mm) were lled with 25mL of distilled water, and
the test lms were mounted on the open ends of the vials. Paralm was used along the circumference of the opening to
ensure that the mounted lms were airtight. Duplicate samples were set up for each lm type, and the initial weight of
each test vial was measured using an analytical balance. The vials were placed in a fume hood for 12h a day, and weight
loss was measured daily for 1week. 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 ofthecomposite films
2.7.1 Preparation ofculture media
The bacterial strains Escherichia coli (E. coli) ATCC 35218, Staphylococcus aureus (S. aureus) ATCC 12600, Shigella sonnei
(S. sonnei) ATCC 25931 and Shigella exneri (S. exneri) ATCC 12022 were cultured through passage 3. The E. coli and S.
aureus strains cultured on Oxoid Mueller Hinton Agar (MHA) (CM0337, Thermo Fischer Scientic, 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 xation. The xed sample was stained with crystal violet solution
for 1min and gently washed with deionised water. Subsequently, the slide was ooded with iodine solution for 1min,
and the wash was repeated with deionised water. The slides were decolourised with ethanol (95%) for 15s, rinsed, and
counterstained with safranin for 1min before being blot-dried with bibulous paper. The samples were viewed under a
microscope (40X and 100X) with oil immersion to determine whether the xed bacteria were Gram–positive/negative.
2.9 Identifying bacteria: selective media streaking
Various selective media were used for further identication 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. exneri 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 specied by the agar type. VRBA, XLD, EMB and Mac-
Conkey agars were incubated for 24h, whereas Baird–Parker Agar and MSA were incubated for 48h.
2.10 Previous contamination test
A previous contamination test was carried out with slight modications as previously described by Lozano-Navarro etal.
[35] to test the ability of the lms to prevent microbes from contacting the surface during handling. A 1 cm2 lm sample
from each lm type was added to each sterilised vial containing sterilised distilled water (5mL). After 24h, 1mL 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 48h.
(13)
SD
(%)=
[(
W
f
−W
i)
∕W
i
×100
]
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2.11 Culturing bacteria: testing fornutrient 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 (10mL portions) was distributed in test tubes and autoclaved at
121°C for 15min. 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 100rpm.
After 3h, 0.1mL of sample from each tube was transferred to tubes containing fresh nutrient broth and placed in a
shaking incubator for another 3h 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 (18–24h).
2.12 Antimicrobial testing: Disk diffusion method
The disk diusion method was carried out using circular lm samples (1cm in diameter) from the various lm 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 (dupli-
cate for each bacteria) by swabbing with sterile swabs. After allowing the cultures to dry, one circular sample from each
lm type was placed on the surface of each inoculated MHA plate, ensuring that the samples were adequately spaced
out. Circular lter paper (1cm 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 eects 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 (200µL) and 4800µL 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
4800µL 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 dierences
among lms with regard to physicochemical parameters were analysed using single factor ANOVA, where p ≤ 0.05 rep-
resented statistical signicance, using Minitab software (version 21.4.10).
3 Results anddiscussion
3.1 Yield ofleaf andstarch extracts
Both plantain starch extraction and leaf extraction produced favourable yields, although these extraction methods may be
further optimised for lm formulation. The overall extraction yields for the plantain peel and pulp seemed slightly lower at
1.20% and 9.76% (Table1), respectively, compared to the previously reported results of plantain pulp yields ranging from
6.67–15.00% [27], and 1.86g/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 12.0g/kg and 97.6g/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].
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Most extraction processes involve a compromise between the extraction yield, purity and integrity of the extract, as condi-
tions 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 signicant 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
aect the chemical composition and purity of the starch obtained, which may impact the lm properties. Other factors that
may have led to the reduced yield include the use of simple separation techniques, such as scraping o the protein layer and
ltering through the muslin cloth. Some starch would have been lost with both of these techniques. There was also starch loss
in the discarded brous waste and the decanted water (resuspended ne starch). Ethanolic leaf extraction produced similar
results and good yields of 8.20 and 8.44% for the 2 types of Panadol leaves utilised (Table2) suggesting comparable extraction
eciencies. The purity and stability of the leaf extracts were maintained by the high-quality ethanol used and the tempera-
ture 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 ecacy of ethanol as a solvent, and the consistency in extraction methods and leaf quality.
3.2 Physicochemical properties ofunripe plantain pulp starch
3.2.1 Proximate analysis
The proximate analysis, as shown in Table1, revealed a starch purity of 85.9%, and this also represented the carbohydrate
content determined by the dierence 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
Table 1 Physicochemical
properties of unripe plantain
starch
Values are means of triplicate samples ± SD
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
Parameter Value
Starch extraction yield (%)
% Yield (Peel)
% Yield (Pulp) 1.20 ± 0.05
9.76 ± 1.07
Proximate analysis
Moisture (%) 13.90 ± 0.07
Ash (%) 0.06 ± 0.00
Lipid (%) 0.10 ± 0.06
Protein (%) 0.08 ± 0.00
Carbohydrate (%) 85.90 ± 0.01
Amylose and glucose content
Amylose (%)
Carbohydrate (g glucose/L) 41.80 ± 0.41
0.23 ± 0.00
Pasting properties
Peak viscosity (cP)
Trough viscosity (cP)
Breakdown viscosity (cP)
Setback viscosity (cP)
Final viscosity (cP)
Pasting temperature (ºC)
Peak Time (min)
4245 ± 40
3474 ± 32
771 ± 12
1264 ± 55
4738 ± 45
50.15 ± 1.2
4.40 ± 0.4
Functional properties
Solubility index (%) 19.80 ± 0.58
Swelling power (g/g) 12.94 ± 0.01
Water absorption capacity (%) 106.00 ± 6.08
Oil absorption capacity (%) 176.50 ± 3.90
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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 specic 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% (Table1), 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 lm–forming properties as lms made from amylose are stronger than those made from
amylopectin [12]. This strength was exhibited with the lms produced.
3.4 Pasting properties
Table1 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 nal 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 inuenced by the ability of the starch granules
to hydrate and swell [12]. The hydration and swelling power of the starch granules are further inuenced 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 inuence its pasting properties. High-quality starch has a high swell-
ing 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 inuenced 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 eect [46]. This may account for the results obtained for extracted starch with a high amylose
content. The size of the starch granules becomes insignicant for swelling and viscosity values, when there is a major
dierence in the chemistry of the starches [44].
The lower pasting temperature (50.15°C) and peak time (4.40min) 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
Table 2 Percentage yield,
total phenolic content, total
avonoid content and DDPH
antioxidant activity of the
Panadol leaf extracts
Values are means of triplicate samples ± SD
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
% Yield Total phenolic content
(mg/g) Total avonoid content
(mg/mL) DDPH antioxidant
activity (% inhibi-
tion)
LE1 LE2 LE1 LE2 LE1 LE2 LE1 LE2
8.20
± 0.54 8.40
± 0.44 86.60
± 0.34 69.60
± 0.39 91.25
± 0.34 74.50
± 0.34 74.60
± 0.65 64.77
± 0.90
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peak time ranges of plantain starch of 81.37–87.13°C and 4.72–7.00min, respectively have been reported [43]. The high
setback viscosity and nal viscosity inuence the properties of the nal 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 lms
with good texture.
3.5 Total phenolic content, total flavonoid content andantioxidant activity oftheleaf extracts
The phenolic content (TPC) expressed as gallic acid equivalents (mg/g) was 86.6 and 69.6mg/mL, and the avonoid
content (TFC) was 91.2 and 74.4mg/mL quercetin for Plectranthus barbatus Andrews (LE1) and Plectranthus caninus Roth
(LE2), respectively (Table2). The DPPH inhibition of LE1 and LE2 antioxidant activity was 74.60% and 64.77%, respec-
tively. 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 avonoid content
(Table2). 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
signicantly higher in LE1 compared to LE2 which may suggest that Plectranthus barbatus Andrews has a richer prole of
phenolic compounds associated with potential health benets. The total avonoid content also diered with LE1 showing
greater levels compared to LE2 (Table2). Flavonoids are known for their antioxidant properties suggesting that LE1 may
oer greater protection against oxidative stress. The DPPH antioxidant activity results indicated that LE1 exhibited higher
inhibition compared to LE2. It was reported by Cordeiro etal. [47] that avonoids, cinnamic acid derivatives, steroids and
ellagic acid were present in both aqueous and organic extracts of Plectranthus barbatus Andrews, which supports our
ndings and the potential of Plectranthus barbatus Andrews as a source of natural antioxidants.
3.6 pH ofthefilm–components
The pH of the various solutions utilised in lm formation was measured as highlighted in Table3. The pH of the lm–form-
ing solution plays an important role in the antimicrobial properties of the lm. Microbial growth is inhibited in highly
acidic environments. Compared with starch solutions, which are alkaline in pH, chitosan lm solutions exhibited an
acidic pH (Table3), which is believed to be a contributing factor to the antimicrobial properties of chitosan lms. 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 lms.
The acidic nature of the acetic acid solution and leaf extracts as shown in Table3 also helped contribute to the acidic
pH of the nal lm solution mixture.
Fig. 1 Pasting curve of plan-
tain pulp starch
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3.7 Colour parameters oftheunripe plantain pulp starch andthevarious film formulations
The purity of the extracted plantain pulp starch was reected by its colour parameters, with an L* value of 100, indicat-
ing maximum whiteness, and there was relatively no signicant change (p > 0.05) in colour parameters from day 0 to 15
(Table4). White starches contain lower levels of proteins and pigments, which make them suitable for industrial applica-
tions particularly in the food and pharmaceutical sectors [37].
The L* a* b* colour parameters did not change signicantly (p > 0.05) with age between days 0 and 12 for the 2.5%
chitosan-only lm, but the parameters diered signicantly (p < 0.05) for the composite lms both with and without the
leaf additive, with the b* values showing the most deviation as shown in Table4. 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 increas-
ing the lightness (L* value). Additionally, drying of the lm with aging could have caused an increased transparency or
changes in the porosity of the lm with time which may have aected how light is reected and potentially could lead
to an increase in lightness. Nearly all the lm 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 avonoids 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 lm matrix over time could aect the stability of pigments. Oxidative reac-
tions 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 lms with higher leaf extracts
after 12days. The increased yellowing of the lms 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 lm matrix.
3.8 Physicochemical parameters ofthevarious film components
3.8.1 Functional properties
The dierent types of lms formed are illustrated in Fig.2 and some physicochemical properties of the dierent types
of lms formed are highlighted in Table5. There was a signicant dierence (p < 0.05) in thickness for the dierent lm
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 signicant dierence (p˃0.05) for the two dierent lms with leaf extracts and the two
concentrations of leaf extracts.
The addition of starch paste increased the lm density. Plantain starch, when incorporated into the chitosan matrix,
probably contributed to additional molecular weight. Starch molecules, which are typically larger and bulkier than indi-
vidual chitosan molecules, can add to the overall mass and density of the lm through bonding interactions making the
lm structure more compact and thus increasing the lm thickness and density [48]. There was no signicant dierence
in density with the two types and concentrations of Panadol leaf extracts.
It appeared that the lms were not dried completely as the moisture content ranged from 17.1 to 25.5% for the dif-
ferent lm types. Despite drying the sample starch to a constant mass, there was still some residual moisture present.
Table 3 pH of various lm-
forming solutions
Values are means of triplicate samples ± SD
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
Parameter pH Value
pH of lm components
Starch (5%)
Chitosan (2.5%)
Starch-Chitosan
Acetic acid (1%)
LE1 (10% stock)
LE2 (10% stock)
8.89 ± 0.01
4.53 ± 0.02
4.68 ± 0.07
2.65 ± 0.01
4.62 ± 0.03
4.54 ± 0.02
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Plantain starch is highly hydrophilic and has a signicant water-binding capacity. When added to the lm formulation, it
tends to absorb and retain more moisture, which increases the overall moisture content of the lm and some moisture
is essential for functionality and it plays an essential role in the ability of starch lms to confer good sealing properties
[49]. This residual water represented the water from the monolayer, which was tightly bound and required harsher dry-
ing 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 inuence the moisture content of the lm. It was
hypothesized that some of these compounds may aect the lm’s ability to retain or release moisture. This could be
attributed to the reduced interaction of the lms with water, as there was an increase in the interaction with the hydro-
philic components of the extracts; this eect was less pronounced as the number of hydrophilic groups signicantly
increased with increasing leaf extract concentration.
The ash content of the lms did increase when compared to that of the native starch as several chemical compounds
were added to the lm 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
Table 4 Colour of plantain
pulp starch and lm
formulations
Values are means of triplicate samples ± SD
* Dierent superscript letters in a column indicate values are signicantly dierent (p < 0.05)
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
Plantain pulp starch L*a*b*
Day 0
Day 15 100.01 ± 1.29a
100.01 ± 0.22a0.83 ± 0.02a
0.90 ± 0.01a2.06 ± 0.04a
2.69 ± 0.07a
Film formulations L* a* b*
CH-2.5%: Day 0 38.78 ± 0.04a0.80 ± 0.02a−1.53 ± 0.03a
CH-2.5%: Day 12 38.82 ± 0.14a0.75 ± 0.05a−1.42 ± 0.16a
CH-2.5%/PS-5%: Day 0 39.00 ± 0.08a0.94 ± 0.01b−2.09 ± 0.02b
CH-2.5%/PS-5%: Day 12 48.47 ± 2.43b0.61 ± 0.06c−1.49 ± 0.05a
CH-2.5%/PS-5%/LE1-0.2%: Day 0 35.98 ± 0.16c0.64 ± 0.02c−1.09 ± 0.03c
CH-2.5%/PS-5%/LE1-0.2%: Day 12 44.76 ± 0.98d0.43 ± 0.11d0.15 ± 0.06d
CH-2.5%/PS-5%/LE1-2%: Day 0 36.40 ± 0.05c0.55 ± 0.01d1.70 ± 0.07e
CH-2.5%/PS-5%/LE1-2%: Day 12 45.37 ± 0.66d−0.29 ± 0.02e4.93 ± 0.16f
CH-2.5%/PS-5%/LE2-0.2%: Day 0 36.19 ± 0.33c0.68 ± 0.07c−1.25 ± 0.04a
CH-2.5%/PS-5%/LE2-0.2%: Day 12 47.68 ± 3.08b0.40 ± 0.04d0.18 ± 0.08d
CH-2.5%/PS-5%/LE2-2%: Day 0 37.21 ± 0.04c0.04 ± 0.01f2.08 ± 0.07e
CH-2.5%/PS-5%/LE2-2%: Day 12 39.80 ± 0.91a−0.19 ± 0.01e2.56 ± 0.46f
Fig. 2 Dierent types of
lms 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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and processing conditions. The variations in the ash levels in the lm 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 Table5 analyze light absorption in the UV (300nm) and visible regions (600nm) 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 specic compounds
in the lm. Conversely, opacity measures the extent to which light is blocked or absorbed by the lm with greater opacity
values indicating a stronger light-blocking capacity. Among the samples, the 2.5% chitosan lm exhibited the lowest
opacity, allowing more light to pass through compared to the other formulations. The inclusion of plantain starch reduced
absorbance at 300nm and increased opacity, suggesting that it aected the lm’s light absorption properties. Further-
more, the addition of leaf extract 1 (LE1) at 0.2% further reduced absorbance at 300nm and enhanced opacity relative to
the formulation containing only plantain starch. This indicated that LE1 positively inuenced the lm’s light absorption
and opacity. Increasing the concentration of LE1 to 2% increased both absorbance at 300nm and opacity. This suggested
that the higher concentration of extract signicantly aected the lm’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 eect 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 dierent leaf extracts signicantly inuenced 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 lms. This suggested that the extracts contributed to
the lm’s light-blocking properties similar to recent studies by Nxumalo etal. [48]. Moreover, the leaf extracts modied
the lm’s properties, with higher concentrations resulting in greater opacity and changes in absorbance. Notably, LE1
appeared to have a more pronounced eect on both parameters compared to LE2. Film thickness is also a contributing
factor to lm barrier properties. Film thickness is inuenced not only by the chemical composition of the lm–forming
components, but also by the type of gelatinisation of the starch. Cold gelatinisation (using alkali) produces thinner lms
as this type of gelatinisation involves hydrolysis of starch components [10].
3.9 Water solubility ofthevarious film formulations
The water solubilities of the dierent types of lms (Fig.3a) showed some interesting characteristics. Generally, lms 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 lms are very susceptible to water and are degraded in water
(highly soluble) [52]. This phenomenon was observed when chitosan lms were added to water during this study. The
addition of starch (although also hydrophilic) to the chitosan lms decreased the solubility, as shown in Fig.3a. Both
chitosan and starch individual lms 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
Table 5 Physicochemical parameters of lm components and lm formulations
Values are means of triplicate samples ± SD
* Dierent superscript letters in a column indicate values are signicantly dierent (p < 0.05)
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
Film formulations Thickness (mm) Density (g/cm−3) Moisture (%) Ash (%) Abs 300nm Opacity (Abs600/mm)
CH-2.5% 0.04 ± 0.01a2.08 ± 0.11a20.54 ± 0.24a2.37 ± 0.02a0.568 ± 0.03a1.35 ± 0.01a
CH-2.5%/PS-5% 0.02 ± 0.00b3.85 ± 0.70b25.21 ± 1.21b3.36 ± 0.03b0.313 ± 0.01b1.97 ± 0.09b
CH-2.5%/PS-5%/LE1-0.2% 0.03 ± 0.00c3.36 ± 0.53c15.92 ± 0.45c3.33 ± 0.03b0.356 ± 0.02b2.58 ± 0.16c
CH-2.5%/PS-5%/LE1-2% 0.03 ± 0.01c3.22 ± 0.15c27.54 ± 2.91b1.20 ± 0.01c0.839 ± 0.02c4.03 ± 0.09d
CH-2.5%/PS-5%/LE2-0.2% 0.03 ± 0.01c3.32 ± 0.51c17.07 ± 0.26c0.67 ± 0.01d0.354 ± 0.03b2.12 ± 0.11c
CH-2.5%/PS-5%/LE2-2% 0.03 ± 0.01c3.40 ± 0.07c24.45 ± 2.16b1.22 ± 0.01c0.645 ± 0.03d3.01 ± 0.13e
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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.
The addition of leaf extracts yielded some interesting results as the solubility of the lms at low concentrations
(0.2%) greatly increased but signicantly 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 lm solubility.
Conversely, at higher concentrations of leaf extract, the interactions between starch and chitosan were disrupted, allow-
ing 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 lms. Additionally,
the increase in insoluble polyphenols and avonoids at higher extract concentrations further contributed to the decrease
in lm solubility [51]. Although the leaf extract conferred enhanced microbial properties, too high a concentration can
negatively impact the physiochemical properties of the lms. Similar phenomena have been reported in other studies
with extracts, such as tea polyphenols [14]. Nxumalo etal. [48] recently reported enhanced swelling degree and water
solubility in chitosan-based lms infused with plant extracts due to improved hydrophilicity. However, high water solu-
bility, can lead to weaker lms 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 ofthevarious film formulations
The high swelling degree of the chitosan lm (2.5%) also supported the lack of water resistance [39]. The addition of starch
signicantly decreased the swelling degree of the lms (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 lm–forming solutions.
3.11 Water vapour transmission rate ofthevarious film formulations
The water vapour transmission rate (WVTR) was determined via the weight loss per day, with slight modications. Moreo-
ver, there was no signicant dierence (p < 0.05) among the WVTRs of the various lm types (Fig.3c). The lms reduced
water vapor movement but did not provide signicant 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 oered superior water
vapor barrier properties compared to the lms. The hydrophilic nature of the lms (all polysaccharides) resulted in their
poor ability to prevent water loss. The presence of plasticisers such as glycerol used in the lm preparation in this study
may have also contributed to increased movement of water across lms, 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 lms, except for one sample
with 2 mould colonies (CH + PS + LE1 (2%) lm). Based on the standards, none of the lms were contaminated, as plates
with ˃ 10CFU were considered to be contaminated by fungi. This indicated that the lms possessed some resistance to
environmental microbes and handling and did not need sterilisation before antimicrobial testing.
The disk diusion method did not produce the anticipated results. There was some inhibition of bacterial growth
under the applied disks, but there was no diusion 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 lms with extracts than for the others.
The PCA method was utilized to assess the ability of various film-forming components to inhibit inoculum forma-
tion. 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
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Fig. 3 Functional properties
of the various lm samples
measured as: (a) water solubil-
ity, (b) water swelling and (c)
water vapour transmission
rate (WVTR). Error bars rep-
resent the standard devia-
tion of mean values of three
replicates. Values designated
by dierent letters are signi-
cantly dierent (p < 0.05)
a
b
c
d
c
d
0
5
10
15
20
25
30
Water solubility (%)
(a)
a
b
bbb
b
0
500
1000
1500
2000
2500
Water swelling (%)
(b)
a
a
a
a
b
c
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
WVTR (g/day)
(c)
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(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 optimisa-
tion 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 applica-
tions 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.
Acknowledgements We are grateful to the laboratory sta of the Food Science and Technology unit of the Chemical Engineering department
(UWI) for their assistance in the completion of this study.
Author contributions R.Maharaj and C.Lafeuillee contributed to the conceptualisation of the work. C. Lafeuillee constructed the methodol-
ogy and performed the formal analysis, investigation and data analysis. R. Maharaj performed the data analysis, wrote, reviewed, and edited
the manuscript.
Funding This research received no external funding.
Data availability The datasets generated during and/or analysed during the current study are available from the corresponding author upon
reasonable request.
Vol.:(0123456789)
Discover Food (2025) 5:5 | https://doi.org/10.1007/s44187-025-00270-4
Research
Declarations
Ethics approval and consent to participate The study does not involve research on human participants and/animals. The collection of the
plants used in the study complies with local or national guidelines with no need for further armation. Plectranthus barbatusAndrew- was
collected in St. Lucia. The plant material was identied by Chad Lafeuillee, and a voucher specimen was deposited at the University of the West
Indies (UWI) herbarium with ID TRIN number 51587. Plectranthus caninus Roth—was collected in St. Lucia. The plant material was identied
by Chad Lafeuillee, and a voucher specimen was deposited at the University of the West Indies (UWI) herbarium with ID TRIN number 51588.
Competing interests The authors declare no competing interests.
Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which
permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to
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do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party
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material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds
the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http:// creat iveco
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