The essential oil extracted with superheated steam extraction from Cinnamomum cassia from Pakistan has potential antimicrobial and antioxidant applications

Abstract

Pharmaceutical, food, beverage, cosmetics, and other industries face severe challenges from bacterial infections and oxidative degradation of active molecules. Their prevention has led to a large use of synthetic preservatives that, however, impact negatively on human health. Natural antioxidants and antimicrobial agents are therefore actively sought, and essential oils (EOs) from different plants are among the natural products with such potentialities. In this context, aimed at finding the method of choice for producing an essential oil (EO) from a Cinnamomum cassia bark from Pakistan with such characteristics, the yield and the antimicrobial and antioxidant properties of the oil extracted by superheated steam extraction (SHSE) were cmpared with those obtained using the standard hydro- and steam distillation techniques. More than 99% of the oil components, including predominant sesquiterpene hydrocarbons, were identified by gas chromatography-mass spectrometry. The EO obtained by SHSE exhibited the highest yield (2.07%) and the most potent antimicrobial effects against Escherichia coli, Pasteurella multocida, Staphylococcus aureus, Bacillus subtilis, Alternaria alternate, Fusarium solani, Aspergillus flavus, and Aspergillus niger. The antimicrobial ativity was determined by the resazurin microtiter-plate, the disc diffusion, the micro-dilution broth susceptibility, and the biofilm assays. In addition, the SHSE EO displayed the most potent effects (93.60%) in the DPPH free radical scavenging assay, as well as the highest (86.66%) hydrogen peroxide free radical scavenging activity and the most powerful antioxidant/reducing properties (60.97 mg of gallic acid/100 g oil). Based on these results, the SHSE technique is strongly recommended over the two traditional distillation methods for the extraction of the EO from the bark of C. cassia.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-026-55841-x.

Introduction

Pharmaceutical, food, beverage, cosmetics, and other industries face severe challenges from bacterial infections and oxidative degradation of active molecules. For example, ingredient oxidation and the action of spoilage bacteria, e.g., Bacillus subtilis, Clostridium botulinum, Escherichia coli, Campylobacter jejuni, Staphylococcus aureus, Listeria monocytogenes, contribute to shortening the shelf life of perishable foods by causing discoloration, off-smell, gas production, bad flavor and mold production¹. Moreover, ingestion of food contaminated by some pathogenic bacteria, viruses, or parasites may cause foodborne illness², which is of great concern for public health.

Other severe infections are caused by biofilms formed by organized microbial communities enclosed in an extracellular polymeric matrix, mainly composed of polysaccharides. The bacterial biofilms have the ability to adhere to both biotic and abiotic surfaces³, causing, for example, dental infections and periodontitis, lung infections caused by cystic fibrosis, implant-associated infections, chronic rhinosinusitis, contamination of contact lenses, catheters and other devices in intensive care units, and human gastrointestinal tract infections^4,5. The biofilm reduces drug penetration through cell membranes, eventually resulting in the development of antibiotic resistance and unsuccessful antimicrobial treatments⁴. Moreover, biofilm formation reduces food quality and can cause illness in consumers. In another field, careful management of the growth of oral microorganisms in biofilms, including aciduric bacteria, is critical to prevent the onset and progression of dental caries and periodontal diseases⁶. Synthetic preservatives are commonly used to combat food pathogens, although they may have long-term negative effects on human health⁷. Similarly, synthetic agents used to inhibit biofilm formation can be toxic. The alternative overuse of bactericidal antibiotics in humans, animals, and plants is the main factor in the development of antimicrobial resistance, which is one of the global public health and development threats⁸.

In another context, food waste has emerged as a promising feedstock for renewable energy production, including its conversion into hydro-treated vegetable oils and biogas as sustainable fuels by anaerobic digestion using advanced monitoring techniques, including artificial intelligence. In addition, apart from its usefulness for fuel production, food waste also contains valuable bioactive compounds such as EOs, which are often overlooked and pose a challenge as inhibitory compounds^9,10. In this regard, pre-treatment of food waste to remove inhibitory constituents can improve the efficiency of anaerobic digestion.

On the other hand, free radical oxidative reactions may occur in different materials and biological systems¹¹. Especially unsaturated molecules easily undergo radical reactions with reactive oxygen species leading to autoxidation, with formation of degradation molecules and even toxic compounds, altered material products, etc.¹². Synthetic antioxidants, such as tertiary butyl hydroquinone (TBHQ), butylated hydroxy anisole (BHA), and butylated hydroxytoluene (BHT), which are commonly used to prevent molecular oxidative deterioration, have recently been reported to be dangerous to human health¹³.

There is, therefore, a growing interest in developing powerful natural antioxidant and antimicrobial agents as green substitutes for antibiotics and synthetic preservatives¹².

Essential oils (EOs) are considered among the most promising natural, nontoxic antimicrobial and antioxidant agents¹⁴, as demonstrated by the EOs from Rosmarinus officinalis L., Origanum vulgare L., Cinnamomum zeylanicum Blume, and Ocimum basilicum L.¹⁵. In fact, besides their pleasant aromatic features, which are highly valued in the production of perfumes, cosmetics, food flavors and fragrances¹⁶, EOs often exhibit potent antibacterial, anticancer, anti-inflammatory, and antioxidant properties¹⁷.

Essential oils (EOs) are extracted from various parts of a plant by different methods. It has been reported that chemical composition and biological activities of EO depend on the part of plants, extraction method, total extraction time, the season, the composition of soil, and environmental conditions^18,19.

The species Cinnamomum cassia (L.) Presl (family Lauraceae), commonly known as Chinese cassia or Chinese cinnamon, is an evergreen tree native to southern China and nowadays widely cultivated there and other Southern and Southeastern Asian countries including Bangladesh, Pakistan, Seychelles, and Madagascar^20,21. It has also been introduced into South America and Hawaii²². The roots, which have camphor as the main volatile component, are less valued than the bark, which contains high amount of trans-cinnamaldehyde (see below). Since ancient times the aromatic dried bark, both powdered or in a “stick” form, has widely been used as a flavoring agent in different cookeries worldwide, especially in Asian food recipes. The buds are also used as a spice, especially in India. The plant is also utilized in traditional medicines to treat dyspepsia, peptic ulcer, diabetes, and ischemic brain injury^23,24. Moreover, the cinnamon bark is included in the Chinese Pharmacopoeia as a major medicinal substance and it has significant economic importance, due to the wide uses in pharmaceutical preparations, seasonings, cosmetics, foods, drinks, commodity essences and chemical industries. Scientific studies have confirmed that C. cassia is a medicinal plant with a wide range of pharmacological effects, including antitumor, anti-inflammatory, analgesic, anti-diabetic and anti-obesity, antibacterial and antiviral properties, as well as cardiovascular protective, cytoprotective, neuroprotective, immunoregulatory and anti-tyrosinase activities²⁴. On the other hand, the EO from the bark, the so-called cinnamon cassia oil, displays antioxidant, antiallergy, anticancer, inhibitory tyrosinase activity and antibacterial effects against several microbial strains, including Staphylococcus aureus, Aspergillus niger, Escherichia coli, Pseudomonas aeruginosa, and Klebsiella pneumoniae²⁵. The oil has been used to treat stomach pain and to prevent diabetes, tumor, hypertension, and other diseases²⁵. Phenylpropanoids are the main bioactive components of C. cassia. In this context, it appears to be of great interest that a C. cassia EO and its main component, trans-cinnamaldehyde, displayed in vitro the ability to resensitize microbial strains to drugs and increased the efficacy of different antibiotics against drug-resistant bacteria²⁶.

HD and SD have been the traditional techniques used to extract cinnamon cassia oil, although both processes have limited extraction efficiency²⁷. In fact, the yield varied from 1.18 to 1.5% with HD, while it ranged from 0.9 to 1.8% using HD (Table 1).

Table 1.

Extraction yields reported in the literature for the cinnamon cassia oil produced using HD and SD techniques.

Technique used	Yield	Reference
Hydrodistillation	1.5%	22
Hydrodistillation	1.3%	28
Hydrodistillation	1.18%	29
Hydrodistillation	1.3%	30
Steam distillation	1.8%	31
Steam distillation	1.44%	32
Steam distillation	1.65%	33
Hydro distillation	0.3%	34
Steam distillation	0.9%	35

A modified improved version of the steam distillation is superheated steam extraction (SHSE). In this process, an EO is extracted quickly from a plant by a steam heated at a temperature (100–180 °C) higher than the vaporization point at a given pressure. High extraction efficiency and capacity, high thermal conductivity, and low oxidizing conditions are beneficial features of the SHSE technique. The high temperature of the steam facilitates the breakage of the plant cell walls and thus the extraction of volatiles³⁶. Furthermore, the polarity and dielectric constant of a superheated steam can be modulated by changing the temperature, allowing the extraction of compounds with different polarity.

The SHSE technique has been rarely used so far in the domain of EO extraction. Examples of its applications are the extraction of the EOs from Boswellia serrata oleo gum resin³⁷, and from thyme and black pepper³⁶. On the contrary, to the best of our knowledge, SHSE has not yet been applied to the extraction of EO from the bark of C. cassia.

Aimed at finding the method of choice for producing an essential oil (EO) from a Cinnamomum cassia bark from Pakistan with potential antimicrobial and antioxidant applications, first the yields of the oils obtained with the HD, SD and SHSE extraction techniques were compared. The EOs were qualitatively and quantitatively analyzed by gas chromatography–flame ionization detector (GC-FID) and gas chromatography–mass spectrometry (GC-MS).

Subsequently, the antimicrobial and the biofilm inhibitory potential of the EOs against fungi and bacteria causing food-borne disease and producing biofilms was determined with the disc diffusion method, the microdilution broth assay, and the resazurin microtiter plate assay. Finally, the antioxidant properties of the three oils were compared with different antioxidant/antiradical assays.

In summary, to the best of our knowledge, this is the first study reporting the chemical and biological investigation of the essential oil isolated from a bark sample of C. cassia trees grown in Pakistan. Furthermore, this is the first application of the SHSE technique for extracting cinnamon cassia oil.

Methods and materials

All the chemical reagents, solvents (reagent or HPLC grade) and nutrient broths were purchased from Merck/Sigma-Aldrich (Milan, Italy).

Collection of plant material

Fresh C. cassia bark (3 Kg) was removed from adult trees grown at Pattoki, in the Punjab province of Pakistan (31°1′ N, 73°51′ E) in September 2022. The plant material was authenticated by Fahim Arshad, Associate Professor, Department of Botany, University of Okara, Pakistan and voucher specimen with the code PTK-04 has been deposited at the herbarium, Department of Botany, University of Okara, Pakistan. The bark was washed with deionized distilled water, air-dried in the shade at room temperature until constant weight, finely ground with a blade mill, and stored in glass containers at 4 °C until analysis.

The collection and use of plant materials complied with relevant institutional, national, and international guidelines and legislation. Cinnamomum cassia is not listed as an endangered or protected species under the IUCN Red List or the Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES).

Isolation of the essential oil

The HD, SD and SHSE EOs were isolated from the same sample of powdered C. cassia bark (100 g in each experiment) using the three techniques described in 2.2.1–2.2.3. Moisture was removed from each sample by anhydrous Na₂SO₄; afterwards, each oil was filtered through a microfilter and stored at 4 °C in amber glass vials until analysis.

The percentage yield of each EO was determined with the following equation:

1

Three experiments were repeated for each oil for reproducible results³⁸, and the mean yield ± SD was calculated.

Hydrodistillation (HD)

A batch of C. cassia bark was soaked in water (3 L) for 30 min in a 5 L round bottom flask in a standard Clevenger apparatus. The mixture was then heated using a heating mantle. The resulting vapors of water and oil were condensed in a tube from the bottom of which the aqueous phase returned to the flask (Supplementary Material, Fig. S1). At the end of the process (3 h) the two phases were removed from the tube and separated in a separating funnel³⁹.

Steam distillation (SD)

The SD process was carried out in the standard manner³⁸. A steam produced in an external 5 L round bottom flask heated using a heating mantle flowed for 3 h through a batch of C. cassia bark placed in a 2 L flask that was connected to a condenser cooled with a jacket of circulating ice water (Supplementary file, Material S2). Condensed steam and EO vapors were separated using a separating funnel³⁸.

Superheated steam extraction (SHSE)

An equipment made specifically (SHSD-001, PAMICO Technologies, Faisalabad, Pakistan) for the SHSE was used. It consisted of a stainless-steel extraction vessel containing the biomass through which flowed a superheated steam (SHS) produced in an electrical steam generator. The temperature of the superheated steam was monitored using thermocouples mounted at the inlet and outlet of the stainless-steel vessel. The system also included a condenser and an EO collection tank (Supplementary Material, Fig. S3), The EO was extracted for 60 min using superheated steam at 150 °C, and then separated from the hydrosol using a separating funnel³⁸.

GLC-FID analysis

Each EO isolated in the three experiments described in 2,2.1.−2.2.3. was analyzed using a Shimadzu GC-2010 gas-chromatograph equipped with a flame ionization detector (FID) and a fused silica DB-5 capillary column (5% phenylmethylpolysiloxane, 50 m × 0.32 mm i.d., film thickness 0.25 μm), Rtx-5 GC Capillary Column, 30 m, 0.53 mm ID, 3.00 μm perchased from Restek. The initial oven temperature (60 °C) was gradually increased to 240 °C at a rate of 5 °C/min, then held at 240 °C for 1 min. The injector and detector temperatures were set at 230 °C and 250 °C, respectively; helium was used as the carrier gas, with a linear velocity of 1.5 mL/min. 1.0 µL of each EO solution in n-hexane (5 mg/mL) was injected into the injection port in a split mode (split ratio: 15 : 1). The linear retention index (LRI_calcd) of each EO constituent was calculated relative to a homologous series of standard n-alkanes (Merck/Sigma-Aldrich, Milan, Italy, No. CE 203–777-6), from n-octane (C₈) to n-tetracosane (C₂₄), that were injected after each oil under the same experimental conditions⁴⁰.

Each EO component was quantified, and the relative concentration was expressed as % over all components. The procedure included the calculation of the FID response factor (RF) of the oil components whose standards were available. RF overall compensated for the irreproducibility of manual injections into the gas-chromatograph. To this aim, a known amount of an internal standard (n-undecane) was added to each EO solution, so that the ratio of the areas of an analyte and the internal standard remained constant from one run to another, irrespective of small differences of injection volumes (and hence the peak areas). The area of the internal standard thus became the value to which the area of each oil component was referred. The area (A) of a peak in the chromatogram was proportional to the molarity (M) times the volume (V) and a constant of proportionality (k), A = k × M × V. Since V was the same for the internal standard and any EO component, the ratio A_c/A_is was given by the following equation:

2

where A_c = peak area of a component, M_c = molarity of a component, M_is = molarity of the internal standard, A_is = peak area of the internal standard. The constant k_c/k_is was defined as the response factor (RF_c) of an EO component and was calculated from the Eq. (2) using the Eq. (3):

3

To calculate the concentration (Mi) of an oil i component, a solution containing known amounts (Mc) of a standard of the i component and n-undecane was injected into the GC and the response factor, RFc, of the i component was calculated from the integrated peak areas, according to the Eq. (6). Subsequently, a separate solution of an EO containing an unknown amount of the i component and a known amount of n-undecane was injected. The % of the i component was calculated using the following equations:.

corrected peak area = integrated peak area of the i component/RF_c of the i component.

The average peak areas from three independent chromatographic runs were considered. For a component with an unavailable standard, the RF_c was set at 1.0.

GC–MS analysis

The GC–MS analysis of each EO was performed on a Shimadzu gas chromatograph GC-2010 system connected to a QP-2010 plus mass detector [Shimadzu (Asia Pacific) Pte Ltd., Singapore]. Gas-chromatographic conditions were the same as described in Sect. 2.3. Mass spectra: electron impact (EI⁺) mode; electron energy, 70 eV; filament emission current, 100 mA; ion source, 175 °C; transfer line, 240 °C. Scanning was performed at a rate of 1 scan sec^− 1 from 40 to 500 a.m.u. The mass spectra were acquired using the TurboMass software, version 5.4 (Perkin Elmer Inc, Shelton, CT, USA)³⁹.

Identification of the EOs components

Each EO component (Table 2) was identified by comparing the calculated linear retention index (LRI_calcd)⁴⁰ with the values (LRI_lit) in the literature⁴¹ and comparing the mass spectrum of each peak with the spectra contained in the NIST 08 and Adams⁴¹ and databases. Moreover, the identity of most oil components was confirmed by GC coelution with authentic standards purchased from Merck/Sigma-Aldrich (Milan, Italy) or from BenchChem (Pasadena, CA, United States).

Table 2.

Chemical composition of the EOs isolated from C. cassia dried bark from Pakistan using different distillation techniques.

No.	Component	Rita	LRIcalcb LRIlitc	HD EO % ± SD	SD EO % ± SD	SHSE EO % ± SD
1	α- Pinened,*	2.357	938 935	0.03 ± 0.00	0.06 ± 0.01	0.08 ± 0.01
2	Myrcened,*	2.516	991 983	0.09 ± 0.00	0.11 ± 0.02	0.14 ± 0.02
3	Limonened,*	2.799	1031 1032	0.39 ± 0.01	0.53 ± 0.04	1.08 ± 0.01
4	γ-Terpinened,*	2.969	1055 1050	0.04 ± 0.00	0.05 ± 0.00	0.12 ± 0.01
5	Linaloole,*	3.203	1098 1100	0.09 ± 0.01	0.23 ± 0.02	–
6	Fenchonee,*	3.271	1090 1088	0.14 ± 0.02	0.34 ± 0.03	0.48 ± 0.03
7	Citrale,*	3.601	1240 1241	–	0.04 ± 0.00	–
8	Hydrocinnamaldehydef,*	3.9	1160 1162	2.62 ± 0.15	1.28 ± 0.12	–
9	Borneole,*	3.999	1165 1165	1.59 ± 0.11	0.09 ± 0.01	–
10	Terpinen-4-ole,*	4.08	1177 1178	1.61 ± 0.10	1.12 ± 0.09	–
11	Anisolef,*	4.248	917 915	2.68 ± 0.12	1.36 ± 0.11	–
12	Estragolef,*	4.253	1202 1196	0.87 ± 0.06	1.98 ± 0.09	2.33 ± 0.16
13	trans-Cinnamaldehydef,*	4.595	1266 1266	14.10 ± 1.12	16.64 ± 1.17	18.51 ± 1.14
14	2-Methyl-3-phenylpropanalf,*	4.864	1244 1244	0.69 ± 0.03	1.01 ± 0.11	1.34 ± 0.11
15	α-Citrale,*	5.14	1271 1270	0.25 ± 0.02	0.14 ± 0.00	0.13 ± 0.01
16	Cinnamyl alcoholf,*	5.896	1259 1277	0.14 ± 0.06	0.25 ± 0.01	0.30 ± 0.05
17	δ-Elemeneg	6.26	1339 1339	–	0.04 ± 0.00	0.07 ± 0.01
18	Ylangeneg	6.349	1373 1375	0.12 ± 0.03	0.27 ± 0.02	0.55 ± 0.03
19	Cyclosativeneg,#	6.88	1368 1368	–	–	0.27 ± 0.04
20	β-Elemeneg,*	7.242	1375 1387	0.55 ± 0.02	0.77 ± 0.06	1.32 ± 0.13
21	Eugenolf,*	7.378	1351 1356	4.66 ± 0.21	5.97 ± 0.27	6.02 ± 0.26
22	Sativeneg,#	7.439	1394 1392	1.87 ± 0.07	2.09 ± 0.04	2.45 ± 0.06
23	Isosativeneg	7.702	1410 1418	0.98 ± 0.10	1.35 ± 0.23	2.04 ± 0.18
24	Caryophylleneg,*	7.877	1418 1418	1.79 ± 0.14	2.61 ± 0.17	2.86 ± 0.11
25	α-Bergamoteneg,#	7.942	1434 1436	1.02 ± 0.13	1.19 ± 0.16	1.42 ± 0.15
26	Isocaryophylleneg,#	8.077	1413 1408	1.91 ± 0.16	1.07 ± 0.13	0.92 ± 0.10
27	Cinnamyl acetatef,*	8.148	1409 1440	2.11 ± 0.13	1.44 ± 0.06	0.53 ± 0.03
28	1,5,9,9-Tetramethyl-(1Z,4Z,7Z)-cycloundecatrieneg	8.484	1519 1519	–	2.16 ± 0.43	2.91 ± 0.42
29	Aromadendreneg,#	8.618	1439 1440	–	0.42 ± 0.02	0.43 ± 0.02
30	Coumarinf,*	8.801	1429 1432	3.10 ± 0.17	0.64 ± 0.04	–
31	Copaeneg,#	8.916	1376 1377	5.79 ± 1.16	7.67 ± 1.18	8.04 ± 1.19
32	α-Amorpheneg	9.494	1485 1484	8.58 ± 1.19	11.24 ± 2.23	12.02 ± 2.11
33	γ-Muuroleneg,#	9.68	1477 1477	0.29 ± 0.01	0.37 ± 0.04b	0.56 ± 0.02
34	δ-Cadineneg,#	9.827	1524 1524	9.42 ± 1.44	11.48 ± 1.56	12.58 ± 1.72
35	α-Calacoreneg,#	10.339	1548 1542	–	1.91 ± 0.35	2.23 ± 0.43
36	Germacrene-4-olh	11.054	1574 1574	–	2.04 ± 0.11	1.30 ± 0.07
37	Isoaromadendrene epoxideh	11.17	1612 1612	–	–	0.53 ± 0.01
38	Caryophyllene alcoholh,#	11.173	1568 1572	0.96 ± 0.01	1.31 ± 0.04	1.99 ± 0.06
39	β-Germacreneg,#	11.298	1560 1556	1.54 ± 0.24	–	–
40	Globulolh,*	11.575	1576 1582		3.09 ± 0.14	1.51 ± 0.03
41	Humulolh	11.7	1618 1618	2.27 ± 0.08	1.13 ± 0.05	1.00 ± 0.02
42	Cubenolh,#	11.72	1642 1641	–	–	0.41 ± 0.00
43	Isolongifoleneg,#	11.821	1387 1402	0.38 ± 0.06	0.59 ± 0.04	0.86 ± 0.12
44	τ-Cadinolh	11.827	1640 1642	2.53 ± 0.03	–	–
45	α-Cedreneg,*	12.206	1419 1412	3.79 ± 0.14	2.88 ± 0.57	2.61 ± 0.64
46	δ-Cadinolh	12.398	1636 1632	–	–	1.81 ± 0.02
47	α-Cadinolh	12.552	1653 1654	10.5 ± 0.15	5.66 ± 0.09	2.94 ± 0.05
48	α-Bisabololh,*	12.71	1701 1697	–	–	0.45 ± 0.02
49	Cadala-1(10),3,8-trieneg	12.817	1563 1562	–	–	0.74 ± 0.06
50	Cadaleneg	13.155	1674 1670	3.65 ± 0.16	2.35 ± 0.68	0.71 ± 0.08
51	β-Bisabololh,#	13.249	1683 1672	4.52 ± 0.03	2.65 ± 0.02	1.05 ± 0.03
52	Acorenoneh	13.28	1685 1685	–	–	0.35 ± 0.01
53	Longiverbenoneh	13.31	1649 1649	1.74 ± 0.08	–	–
Chemical families
Monoterpene hydrocarbons (%)				0.55	0.75	1.42
Oxygenated monoterpenoids (%)				3.68	1.96	0.61
Sesquiterpene hydrocarbons (%)				41.68	50.46	55.59
Oxygenated sesquiterpenoids (%)				22.52	15.88	13.34
Phenylpropanoids and minor aromatic compounds (%)				30.97	30.57	29.03
Total identified compounds (%)				99.40	99.62	99.99

^a RT= retention time (min); ^b LRI_calc = calculated linear retention index⁴⁰, ^c LRI_li30t = linear retention index taken from the literature^41,49; ^d monoterpene hydrocarbon; ^e oxygenated monoterpenoids; ^f phenylpropanoids and other aromatic compounds; ^g sesquiterpene hydrocarbons; ^h oxygenated sesquiterpenoids; %, percentage; SD, standard deviation. Both values were calculated as the means of three separate GC analysis. * Commercially available from Merck-Sigma-Aldrich. # Commercially available from BenchChem.

Antiradical and antioxidant activities

DPPH free radical scavenging activity

The radical scavenging activity of each EO was evaluated with the 2,2-diphenyl-1-picrylhydrazyl radical (DPPH.) assay, following the protocol reported in the literature with small modifications⁴². Gallic acid was used as a positive control. A methanol solution of DPPH. (0.3 mM, 1.0 mL) was separately mixed with each EO (2.5 mL of a solution of 100 mg/mL in MeOH) or gallic acid (100 ppm) in EtOH and the mixture was shaken vigorously while being incubated in the dark for 20 min at 22 ◦C until a stable absorbance value (Abs_sample) at 517 nm was reached. The absorbance was measured against the blank using a Lambda 25 UV/Visible spectrophotometer N.3903 (Perkin Elmer Inc, Shelton, CT, USA). The lower the absorbance of the reaction mixture, the greater the free radical (DPPH.) scavenging (FRS) activity. Methanol (1.0 mL) plus each EO solution (2.5 mL) or gallic acid, in the absence of DPPH, was used as the blank. A methanol DPPH solution (0.3 mM, 1.0 mL) plus MeOH (2.5 mL) was used as the control to measure the Abs_control at 517 nm.

The per cent free radical scavenging activity (FRS%) was calculated using the Eq. (4):

4

Each experiment was carried out in triplicate.

Potassium ferricyanide reducing antioxidant power assay (PFRAP)

A protocol of the literature⁴³ reporting the potassium ferricyanide reducing antioxidant power (PFRAP) assay was followed with minor modifications to determine the total antioxidant activity of each EO. The activity was expressed as gallic acid equivalent, i.e., mg GAE per 100 g EO, using a calibration curve. To determine the standard curve, gallic acid (5 mg) was dissolved in H₂O (25 mL) to give a stock solution with a concentration of 200 mg/L, from which nine dilutions of gallic acid were prepared with concentrations of 100, 75, 50, 25,15,10, 7, 5, 2.5 mg/L, respectively. 1 mL of each EO or gallic acid solution was mixed with 2.3 mL of 0.2 M phosphate buffer (pH 6.6) and 2.5 mL of 1% (w/v) K₃Fe (CN)₆, and the resulting mixture was incubated for 20 min in a water bath heated to 37 °C. Subsequently, a 10% (w/v) trichloroacetic acid solution (2.5 mL) was added and the mixture was centrifuged at 1000 rpm for 10-min. Then, 2.5 mL of the resulting supernatant layer was mixed with 0.1% (w/v) FeCl₃ (0.5 mL) and diluted with distilled water (2.5 mL). After standing for 10 min, the absorbance of the mixture was measured at 700 nm against the blank using the UV-Vis spectrophotometer cited above. Each experiment was performed in triplicate.

Hydrogen peroxide scavenging activity

The hydrogen peroxide scavenging activity of each EO was measured spectrophotometrically by using a protocol of the literature⁴⁴. A H₂O₂ solution (2 mM) was prepared in phosphate buffered (0.17 M, pH 7.4) distilled water, and each EO (1 mL) was mixed separately with this solution (1 mL). After an incubation of 10 min at 22 °C, the absorbance (Abs_sample) of the mixture at 230 nm was measured with the UV-Visible spectrophotometer against a blank solution containing the phosphate buffer without H₂O₂. The buffered aqueous solution of H₂O₂ without the sample was used as the control. Ascorbic acid (100 ppm) was used as the positive standard. The lower the absorbance of the sample solution, the greater the hydrogen peroxide scavenging activity. The percentage of hydrogen peroxide scavenging activity of each EO or ascorbic acid was calculated using the Eq. (5):

5

Where Abs_control is the absorbance of the control and Abs_sample is the absorbance in the presence of the tested sample. Each experiment was performed in triplicate.

Antimicrobial activity

Disc diffusion method

The antifungal and antibacterial activity of the EOs was determined with a slightly modified disc diffusion method described in the literature⁴⁵, using the bacteria Bacillus subtilis, Staphylococcus aureus, Pastrulla multocida, and Escherichia coli, and the fungi Fusarium solani, Aspergillus niger, Alternaria alternate, and Aspergillus flavus, respectively. The strains were obtained from the Department of Microbiology, University of Agriculture Faisalabad, Pakistan, where the strains were identified and their homogeneity was assessed. The bacterial strains were grown on nutrient agar (NA) plates at 37 °C for 24 h, whereas the fungal strains were grown on potato dextrose agar (PDA) plates at 30 °C for 48 h. Subsequently, a suspension (100 µL) of a bacterium (10⁸ CFU/mL) or a fungus (10⁴ spores/mL) was added to liquid NA or PDA (20 mL), respectively, and the mixture was transferred onto 6 mm-sized paper disks. After 10 min of agar solidification at 22 °C, 15 µL of each EO solution (1 mg/mL DMSO) was placed on each disk. After an incubation period (24 h at 37 °C for bacteria and 40 h at 30 °C for fungi), the antimicrobial activity against each strain was evaluated by measuring the width of the inhibition zones using a digital Vernier caliper. Amoxicillin and fluconazole (25 µg/disc) were used as the positive controls to compare the sensitivity of bacteria and fungi, respectively, while disks without the tested samples were used as a negative control. Separate experiments indicated that DMSO (dimethyl sulfoxide) was not toxic to microorganisms at the concentration used.

MIC determination using the resazurin microtiter-plate assay

The minimum inhibitory concentration (MIC) of each EO against each bacterial strain was evaluated using the 7-hydroxy-3H-phenoxazin-3-one 10-oxide (resazurin) microtiter-plate assay⁴⁶. A homogeneous resazurin solution was prepared by dissolving a commercial tablet (270 mg) thoroughly in sterilized water (40 mL), using a vortex mixer. Polymeric microtiter 96 well plates were sterilized and labelled under aseptic conditions. A 10% (v/v) stock solution of each EO, prepared by diluting the oil (100 µL) with DMSO (900 µL), was placed in the first row of a plate, while Mueller Hinton (M-H) Broth (50 µL) was added to all other wells. Each EO was serially diluted with the M-H Broth from 10 mg/mL to 0.046 mg/mL. After dilution, each well was left with the tested sample (50 µL), the resazurin indicator solution (10 µL), and a bacterial suspension (10 mL, 5 × 10⁶ cfu/mL). A cling film was wrapped loosely around each well to keep microbes hydrated. In each plate a column of wells contained serial dilutions of amoxicillin used as the positive control, a column contained the solutions without the tested sample as a negative control, and a column contained all the solutions, bacterium-free and added with the nutrient broth (10 µL). The plates were prepared in triplicate and incubated at 37 °C for 18–24 h. The change of color from purple to pink or colorless was then evaluated visually, noticing the lowest concentration that caused such a variation. The average concentration ± SD from three plates was calculated and this value was considered the MIC of the sample against the tested bacterial strain.

Antifungal activity determined using the broth microdilution susceptibility assay

The minimum inhibitory concentration (MIC) of each EO against each fungal strain was determined using the broth microdilution susceptibility assay recommended by the National Committee for Clinical Laboratory Standards (NCCLS), making minor changes to a procedure described in the literature⁴⁷. A fungal suspension incubated overnight was diluted with a sterilized Sabourand Dextrose Broth (SDB) medium. A 10% (v/v) stock solution of each EO was prepared by diluting the oil (100 µL) with DMSO (900 µL). Sterilized SDB (100 µL) was added to each well of a sterilized 96-well flat-shaped microtiter plate. The stock solution (100 µL) of tested EO was then placed in the first well of the plate and was two-fold serially diluted with the SDB medium to obtain a series of concentrations in the range of 0.01–32.0 mg/mL). Subsequently, a fungal suspension was added to wells with the final concentration adjusted to about 5 × 10⁵ cfu/ml. Fluconazole was used as a reference antifungal sample; the SDB medium containing the fungal suspension was used as a positive control and DMSO was used as a negative control. Plates were prepared in triplicate and then incubated for 48 h at 30 °C. The cloudiness of the broth or the separation of a cell layer at the bottom of a well indicated that fungal growth had occurred. The lowest concentration of the tested sample that stopped fungal growth was recorded, and the mean ± SD, calculated from three independent experiments, was considered as the MIC of the sample against the tested fungal strain.

Biofilm formation inhibition assay

The inhibitory activity of each EO on the biofilm formation by two Gram-positive bacteria (Bacillus subtilis, Staphylococcus aureus) and two Gram-negative bacteria (Pastrulla multocida, Escherichia coli), was determined in triplicate using a slightly modified crystal violet (CV) binding method⁴⁸. Nutrient broth (100 µL), an EO (100 µL), and a bacterial culture (10 µL, 5 × 10⁵ cfu/ml) were added to each well of a sterilized 96-well flat-shaped microtiter plate. After having been incubated at 37 °C for 18 h under static conditions to allow for bacterial growth and biofilm maturation, the suspension was removed from each well, which was rinsed twice with phosphate buffer saline (PBS, 220 µL) to remove unattached bacterial cells and medium components. The formed biofilm was treated with MeOH (99%) for 15 min; subsequently, each well was stained with a 7% (w/v) aqueous solution of CV (200 µl) and incubated at room temperature for 10 min. Then, the wells were rinsed thoroughly with deionized water to remove unbound dye, and air dried for 5 min. The bound CV dye was dissolved in 33% aqueous acetic acid (125 µl) added to each well, and the plate was incubated at 22 °C for 15 min. Subsequently, the optical density (OD) of each well was measured at 630 nm against the blank (33% aqueous acetic acid) using a microplate ELISA reader (BIO-RAD, Hercules, CA, USA). The extent of biofilm inhibition was calculated with respect to the amount of biofilm grown in the absence of the tested sample, which was considered as a negative control. Rifampicin (10 mg/ml) was used as a positive control. The % biofilm inhibition in each experiment was calculated using the Eq. (6):

6

Statistical analysis

Each assay was performed in triplicate, and the mean value ± standard deviation (SD) was calculated. Statistical analysis of the data was performed by Analysis of Variance (ANOVA) using STATISTICA 5.5 (Stat Soft Inc., Tulsa, OK, USA) software. Results were considered statistically significant at p < 0.05.

Results and discussion

EO extraction yield from C. cassia bark

The EOs isolated from the powdered bark of C. cassia from Pakistan using the HD, SD, and SHSE techniques had a pale-yellow color and a pleasant aroma. The percent EO yields (%, w/w) were calculated using the Eq. (1) (Sect. 2.2). The isolation methods affected the yields of the oils significantly. In fact, the highest yield (2.07%) was obtained with the SHSE process, while the lowest one was observed using the HD method (1.11%) and the SD technique gave an intermediate value (1.51%). These results were comparable with those previously obtained by other authors for cinnamon cassia oil using HD and SD methods (Table 1 in Sect. 1).

SHSE involves controlled heating to produce SHS; the shorter extraction time (one hour) and the higher extraction efficiency compared to HD and SD compensate for the initial energy consumption. In contrast, hydrodistillation and steam distillation typically involve longer extraction times (three hours) and constant heating, which usually lead to higher costs. The improved extraction yield using SHSE suggests that it may be applied on a large-scale for EO extraction. However, although SHSE shows promising laboratory-scale performance, a pilot-scale techno-economic analysis should be performed in future works to better demonstrate its industrial applicability.

Chemical composition of the EOs

A total of 53 compounds with percentages ˃ 0.03% were identified in the three EOs distilled from dried bark of C. cassia and analyzed on a DB-5 (methylpolysiloxane phase with 5% phenyl groups) capillary column. They accounted for the 99.40, 99.62, and 99.99%, respectively, of the HD, SD, and SHSE EOs, respectively. The components of each oil, with the corresponding percentages, are listed in Table 2, according to the increasing retention time (RT, min).

The main components of the three EOs extracted from C. cassia bark from Pakistan are listed in Table 3, while their chemical structures are depicted in Fig. 1. Only the relative stereochemistry is shown.

Table 3.

Main components occurring in the EOs extracted from C. cassia bark from Pakistan.

Compound	EO
	HD % ± SD	SD % ± SD	SHSE % ± SD
trans-Cinnamaldehyde (1)	14.10 ± 1.12	16.64 ± 1.17	18.51 ± 1.14
α-Cadinol (2)	10.5 ± 0.15	5.66 ± 0.09	2.94 ± 0.05
δ-Cadinene (3)	9.42 ± 1.44	11.48 ± 1.56	12.58 ± 1.72
α-Amorphene (4)	8.58 ± 1.19	11.24 ± 2.23	12.02 ± 2.11
Copaene (5)	10.77 ± 1.24	11.34 ± 0.02	8.14 ± 0.24
Eugenol (6)	4.66 ± 0.21	5.97 ± 0.27	6.02 ± 0.26
Β-Bisabolol (7)	4.52 ± 0.03	2.65 ± 0.02	1.05 ± 0.03
α-Cedrene (8)	3.79 ± 0.14	2.88 ± 0.57	2.61 ± 0.64

Fig. 1.

Chemical structures of the main components of the EOs from the bark of C. cassia from Pakistan.

The GC-FID and GC-MS analyses (Table 2) indicated that the HD, SD, SHSE EOs consisted mainly of sesquiterpene hydrocarbons (41.68, 50.46, and 55.58%, respectively), followed by phenylpropanoids and minor aromatic compounds (30.97, 30.57, and 29.03, respectively), and oxygenated sesquiterpenoids (22.52, 15.88, and 13.34%, respectively). Instead, monoterpenoids only occurred in minor amounts. Noteworthy, only eight compounds occurred with percentages ˃ 1.5% in the three oils (Table 3 and Fig. 1). The profiles of the three oils were similar qualitatively, but the quantitative compositions were significantly different. The amounts of monoterpene and sesquiterpene hydrocarbons increased significantly from the HD to the SHSE oils with the concurrent marked decrease of the oxygenated terpenoids, while the amount of phenylpropanoids and aromatic compounds remained almost the same in the three EOs. This finding was likely due to alcohols’ dehydration and esters hydrolysis caused by the overheated steam used in the SHSE. trans-Cinnamaldehyde (1) predominated in all the three oils with percentages increasing from 14.1% in the HD EO to 16.64% in the SD EO and 18.5% in the SHSE EO (Table 3). Aldehyde 1 was accompanied by copaene (5) (10.77%), α-cadinol (2) (10.50%) and δ-cadinene (3) (9.42%) as the most abundant components of the HD EO, while δ-cadinene (3) (11.48%), copaene (5) (10.77%) and amorphene (4) (11.24%), and δ-cadinene (3) (12.58%), α-amorphene (4) (12.02%) and copaene (5) (8.14%) were the other main components of the SD EO and the SHSE EO, respectively. These results are consistent with the literature, in which cinnamaldehyde is consistently reported as the main component of the EOs extracted from different bark specimens of C. cassia (L.) Presl and related cinnamon species, such as C. zeylanicum Blume. However, the amount of 1 and other C. cassia oil components varied with the authors and the distillation method. For example, 42.37%²⁵ and 75.66%³³ yields of trans-cinnamaldehyde were obtained by SD of dried bark of C. cassia collected in China, while yields of 24.41 and 28.44% were attained by SD of fresh bark and fresh leaves, respectively³³. (E)−2’-Methoxycinnamaldehyde (6.08%), cinnamaldehyde (3.47%) and dihydrocinnamaldehyde (2.78%) were the other main components of the SD EO isolated from the bark³³. Other studies indicated that the composition of the HD EO from a C. cassia bark also varied significantly with the growth stage of the plant and the type of bark. The content of trans-cinnamaldehyde varied from 33.95 to 76.4% with the age of the trees, reaching the maximum amount in the stem bark of six-years-old trees⁵⁰. In another study, the content of trans-cinnamaldehyde varied from 66.28 to 77.21% in the HD EOs isolated from seven whole barks of 11-16yearold cinnamon trees grown in different locations in two provinces of southern China²⁰. Similarly, the percentages of other major components of the oil, e.g., cinnamyl acetate (0.09–49.63%), (E)−2’-methoxycinnamaldehyde (0.09–6.69%), and copaene (1.09–14.3%) showed high fluctuations, reaching the maximum value at different age of the plant⁵⁰. In addition, the oils contained variable amounts of cis-cinnamaldehyde (0.72–2.10%), guaiene (4.27–7.62%), copaene (0.58–3.86%), (E)−2’-methoxycinnamaldehyde (0.98–2.59%), muurolene (0.47–1.83%), and calacorene (0.57–1.28%). Instead, the content of eugenol, which is highly valued commercially, was very low in all bark samples (0.0–0.21%)²⁰.

Antiradical/antioxidant activity

There is general consensus that there is no single method suitable for determining the potential antioxidant/antiradical activity of a sample. Therefore, the antiradical/antioxidant potential of the HD, SD, and SHSE EOs from C. cassia bark from Pakistan was determined by three common methods, i.e., the DPPH assay, the PFRAP assay and the hydrogen peroxide scavenging assay⁵¹.

DPPH.(1,1-diphenyl-2-picrylhydrazyl radical) is a stable free radical with a free electron delocalized throughout the entire molecule, which results in a deep violet color in MeOH with a maximum absorption at around 520 nm. A stable non-radical form of DPPH is produced when DPPH. is quenched by a substrate functioning as a H. donor, while the color of the solution simultaneously changes from violet to pale yellow. The spectrophotometrically measured percent free radical scavenging activity (% FRSA) of HD, SD and SHSE EOs are reported in Table 4. The EO isolated using the SHSE technique exhibited the highest FRSA value (93.60%), followed by the SD (92.87%) and the HD (91.47%) oils. Notably, the antiradical activity of the SHSE EO surpassed that shown by 100 ppm of gallic acid (81.03%) as well as the activities of sage and oregano EOs⁵². Moreover, the values are consistent with the radical scavenging activity of other C. cassia EOs reported in the literature⁵³.

Table 4.

DPPH free radical scavenging, total antioxidant/FRAP, and hydrogen peroxide scavenging activities of EOs extracted from C. cassia bark from Pakistan using different distillation methods^a.

Sample	DPPH FRSA (%)	Potassium ferricyanide reducing/antioxidant power (PFRAP)b	Hydrogen peroxide scavenging activity (%)
HD EO	91.47 ± 0.20	47.37 ± 0.71	80.0 ± 0.40
SD EO	92.87 ± 0.23	54.92 ± 0.80	83.33 ± 0.50
SHSE EO	93.60 ± 0.31	60.97 ± 0.91	86.66 ± 0.70
Ascorbic acidc (100 ppm)		–	95.33 ± 0.90
Gallic acidd (100 ppm)	81.03 ± 0.1	–	–

^aActivity values are the means ± SD from three separate experiments; ^bexpressed as mg GAE per 100 g EO; ^creference antioxidant agent; ^dreference antiradical agent.

In the PFRAP (potassium ferricyanide reducing/antioxidant power) assay, which is a non-radical single electron tranfer (SET)-based method, the reducing power of an antioxidant EO is determined using a colorimetric reaction in which the complex K₃[Fe^III(CN)₆] is reduced to K₄[Fe^II(CN)₆] in acidic conditions (pH 3.6). The subsequent reaction with added Fe^IIICl₃ produces the dark blue complex Fe^III₄[Fe^II(CN)₆]₃. The Prussian blue absorbance is then measured spectrophotometrically at 700 nm and the reducing power of each tested sample is expressed as mg gallic acid equivalent (GAE) per 100 g EO. The values determined for the EOs extracted from the C. cassia bark from Pakistan (Table 4) clearly indicated that the distillation method greatly affected the results of the FRAP assay. In fact, the highest reducing power was observed for the SHSE EO (60.97 mg GAE/100 g) followed by the SD (54.92 mg GAE/100 g) and the HD EO (47.37 mg GAE/100 g) oils.

Finally, the data of the hydrogen peroxide scavenging activity of the three EOs well agreed with the results of the other two assays (Table 4): the highest activity was shown by the SHSE EO (86.66%) while the weakest effects were shown by the HD EO (80.0%).

The antioxidant/antiradical activities were consistently different between the three EOs. Such differences were likely due to different qualitative and quantitative chemical compositions of the oils (Table 2), although it is challenging to establish a relationship between the effects and the structures of the EOs constituents⁵⁴. In addition, synergistic effects of different components may affect the activity. However, cinnamaldehyde and eugenol have been considered potent natural antioxidant/radical scavenger agents^52,55, and it is interesting to note that the activity of the EOs from the bark of C. cassia from Pakistan (Table 4) increased with the increasing concentration of these two compounds in the oils, from the HD to the SHSE EO (Table 2). The antioxidant activity of these compounds is likely due to the high oxidation potential, which enables them to work as reductants, hydrogen donors, and oxygen quenchers⁵⁵.

Antimicrobial activity

The antibacterial activities of the HD, SD, and SHSE EOs against two Gram-positive bacteria, Staphylococcus aureus and Bacillus subtilis, and two Gram-negative bacteria, Escherichia coli and Pastrulla multocida, were evaluated by measuring the zones of inhibition (ZOIs) using the disc diffusion method⁴⁵, the minimum inhibitory concentration (MIC) using the resazurin microtiter plate assay⁴⁶, and the inhibition of bacterial biofilm formation using the crystal violet (CV) binding method⁵⁶.

The antifungal activity of the three EOs against Fusarium solani, Aspergillus niger, Alternaria alternate, and Aspergillus flavus strains was estimated by measuring the corresponding MICs using the broth microdilution susceptibility assay⁴⁷ and measuring the zones of fungal growth inhibition (ZOIs) using a disc diffusion method⁴⁵.

Antibacterial activity (disc diffusion method and MIC)

The results of the MIC and the ZOI assays are reported in Table 5. The three EOs from C. cassia bark exhibited high antibacterial activity against all the strains, with MIC values in the range of 0.04–2.50 µg/mL and ZOI diameters, for 25 µg EO/disc, ranging from 16.50 to 27.12 mm, in accordance with the literature⁵³. In comparison, the MIC values of the positive control amoxicillin were in the range of 0.01–0.31 µg/mL while 25 µg/disc of this antibiotic produced ZOI diameters ranging from 31.00 to 39.86 mm. All the three EOs, as well as amoxicillin, exhibited the highest antibacterial activity against Pastrulla multocida. The activity decreased in the order P. multocida > B. subtilis > S. aureus > E. coli, and from the SHSE to the HD EO. Noticeably, the effects of the SHSE EO were of the same order of magnitude as those of the well-known antibiotic amoxicillin.

Table 5.

Antibacterial activity of EOs isolated from C. cassia bark from Pakistan using different distillation techniques.^§.

Bacterial strain	HD EO	SD EO	SHSE EO	A amoxicillin†
Minimum inhibitory concentration (MIC, µg/mL)
Staphylococcus aureus	0.63 ± 0.02a	0.31 ± 0.01b	0.16 ± 0.01c	0.078 ± 0.01d
Escherichia coli	2.50 ± 0.02a	1.25 ± 0.01b	0.63 ± 0.00c	0.31 ± 0.00d
Bacillus subtilis	0.31 ± 0.01a	0.16 ± 0.02b	0.08 ± 0.01c	0.04 ± 0.01d
Pastrulla multocida	0.16 ± 0.01a	0.08 ± 0.02b	0.04 ± 0.00c	0.01 ± 0.00d
Diameter (mm) of the zone of inhibition
Staphylococcus aureus	18.00 ± 0.21d	19.00 ± 0.17c	21.00 ± 0.19b	31.00 ± 0.15a
Escherichia coli	16.50 ± 0.12d	19.51 ± 0.12c	20.02 ± 0.08b	38.06 ± 0.32a
Bacillus subtilis	20.10 ± 0.15d	21.12 ± 0.12c	23.19 ± 0.15b	34.02 ± 0.16a
Pastrulla multocida	23.72 ± 0.23d	26.44 ± 0.32c	27.12 ± 0.18b	39.86 ± 0.32a

^§Activity values are expressed as the mean ± SD from three separate determinations; ^† used as a standard antibiotic. ^{a, b,c, d}Different letters represent a significant difference (p < 0.05) between the three EOs.

Antibiofilm activity

The percent inhibitory activities of bacerial biofilm formation by the three EOs extracted from C.cassia bark from Pakistan are shown in Table 6.

Table 6.

Antibiofilm activity of the EOs isolated from C.cassia bark from Pakistan using different distillation techniques^§.

Microbial strain	HD EO#	SD EO#	SHSE EO# rifampicin*
Biofilm inhibition activity (%)
Staphylococcus aureus	20.07 ± 0.09a	29.66 ± 0.08b	33.27 ± 0.12c 35.44 ± 0.10d
Escherichia coli	30.56 ± 0.012a	31.28 ± 0.09b	39.05 ± 0.09c 43.12 ± 0.2d
Bacillus subtilis	25.78 ± 0.04a	34.46 ± 0.06b	38.06 ± 0.08c 42.53 ± 0.14d
Pastrulla multocida	42.64 ± 0.0a	45.24 ± 0.06b	51.52 ± 0.06c 56.76 ± 0.18d

§The activity values are expressed as the mean ± SD from three separate determinations; #100 µL; *(10 mg/ml), used as a reference standard. a, b,cDifferent letters represent a significant difference (p < 0.05) between the three EOs.

The order of activity of the three oils was, consistently, SHSE ˃ SD ˃ HD oils, as determined in the other two antibacterial assays described above, while P. multocida was the most sensitive of the four tested bacteria to the oils. Noticeably the effects of 100 µL of the SHSE EO against the four strains were compable with those of the very well-known bactericidal antibiotic rifampicin.

The studies of antibacterial and antibiofilm properties of different cinnamon EOs have been mainly focused on C. zeylanicum and C. cassia oils^5,53,57. In general, the mechanism of action of an essential oil depends on its chemical composition, and the antimicrobial activity is not attributable to a single mechanism⁵⁸. Although the mechanisms underlying the antibacterial effects of cinnamon EOs have not yet been fully elucidated, the main active constituents have been identified in cinnamaldehyde^5,57,59,60 and eugenol and, to a minor extent, in terpenoid constituents, such as β-caryophyllene, citral, linalool, limonene. In fact, due to the highly electronegative unsaturated aldehyde group, cinnamaldehyde readily reacts with biological molecules such as proteins and nucleic acids, thus interfering with biological processes. On the other hand, eugenol may form a hydrogen bond with enzymes’s active centers⁵⁹; moreover, it may denature proteins and change the permeability of cell membranes⁶⁰. On the other hand, due to their hydrophobicity, terpenoids can penetrate the bacterial cell wall and disrupt the lipid structure⁶¹, ultimately resulting in protein dysfunction, cell lysis, cytoplasmic leakage, and death of bacteria⁶².

Antifungal activity (disc diffusion method and MIC)

The results of the antifungal activity tests conducted on the HD, SD, and SHSE EOs from the Pakistani sample of C. cassia bark are shown in Table 7. The MIC and ZOI values (for 25 µg EO/disc), were in the range of 0.04–5 µg/mL and 11.64–19.54 mm, respectively, while the values of the positive control fluconazole were in the range of 0.02–0.63 µg/mL and 33.18–38.24 mm, respectively. The EOs exhibited the highest activity against Alternaria alternate while Aspergillus niger was the least sensitive of the four fungi. As observed for the antibacterial effects, the SHDS EO showed the highest activity against all four strains, followed by the SD and the HD oils. Noteworthy, the MICs of the SHDS EO were of the same order of magnitude as those of fluconazole.

Table 7.

Antifungal activity of C.cassia bark EOs isolated by different techniques^§.

Fungal strain	HD EO	SD EO	SHSE EO	Fluconazole†
Minimum inhibitory concentration (MIC, µg/mL)
Fusarium solani	1.25 ± 0.04a	0.63 ± 0.03b	0.31 ± 0.05c	0.16 ± 0.04d
Aspergillus niger	5.00 ± 0.02a	2.50 ± 0.01b	1.25 ± 0.00c	0.63 ± 0.01d
Alternaria alternate	0.31 ± 0.01a	0.16 ± 0.05b	0.04 ± 0.05c	0.02 ± 0.03d
Aspergillus flavus	1.25 ± 0.05a	0.65 ± 0.05b	0.32 ± 0.06c	0.32 ± 0.03d
Diameter of the zone of inhibition (mm) Fluconazole†, #
Fusarium solani	13.26 ± 0.08d	14.82 ± 0.04c	16.34 ± 0.08b	34.74 ± 0.48a
Aspergillus niger	11.64 ± 0.12d	13.22 ± 0.08c	14.78 ± 0.12b	33.18 ± 0.32a
Alternaria alternate	16.24 ± 0.05d	18.22 ± 0.12c	19.54 ± 0.08b	38.24 ± 0.16a
Aspergillus flavus	12.46 ± 0.17d	14.42 ± 0.07c	15.98 ± 0.09b	34.38 ± 0.12a

^§The activity values were expressed as the mean ± SD from three separate experiments; ^† used as a standard antifungal agent; ^#25 mg/disc. ^{a, b,c, d} Different letters represent a significant difference (p < 0.05) between the three EOs.

These results are consistent with previous studies concerning the antifungal activity of C. cassia EOs against different fungal strains^53,63,64. Cinnamaldehyde has been identified as the major active fungitoxic component of the cinnamon bark oil^64–66. Remarkably, this compound was effective in vitro against fluconazole-resistant Candida species that cause respiratory tract mycoses and other important clinical infections⁶⁵. Inhibition of ATPases, cell wall biosynthesis, and alteration of membrane structure and integrity are considered the main antifungal action mechanisms of cinnamaldehyde⁶⁵. This aldehyde is thought to inhibit the activity of β−1,3-glucan and chitin synthases, two enzymes involved in the production of essential components of fungal cell walls.

Conclusions

The present work contributes to reducing the lack of information regarding the influence of extraction methods on the chemical composition and biological activities of the essential oil from C. cassia bark, in particular the one of trees grown in Pakistan. The results clearly showed that the extraction techniques significantly influenced the yield, the chemical profile, and the antioxidant, antimicrobial, and antibiofilm properties of the EO. Notably, SHSE was found to be the most effective extraction method compared to the traditional HD and SD techniques, affording an EO with the highest yield and the best biological activities. The improved performance of SHSE can be attributed to its higher efficiency in mass transfer and its ability to more effectively extract volatile bioactive constituents from disrupted cells in the biomass. This research suggests that SHSE is an environmentally friendly and efficient method of extracting valuable EOs. However, to improve the efficiency of the process and facilitate future large-scale applications of the EO, the optimization of different SHS extraction parameters such as extraction temperature, extraction time, steam flow rate, and biomass particle size must be addressed in future studies.

The remarkable antioxidant and antimicrobial properties of the SHSE oil suggest its potential application in the pharmaceutical and cosmetic fields. Although such applications have not been our priority in this work, assessing the absence of toxicity of the oil, including its cytotoxicity, will be a necessary step in our future study to strengthen the validity of the current results.

In conclusion, this research provides a solid foundation for the use of SHSE as an efficient extraction method of EOs and opens new opportunities for the sustainable development and applications of cinnamon cassia oil.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

TBHQ

Tertiary butyl hydroquinone

BHA

Butylated hydroxyanisole

BHT

Butylated hydroxytoluene

EOs

Essential oils

HD

Hydrodistillation

SD

Steam distillation

SHSE

Superheated steam extraction

GC-FID

Gas chromatography–flame ionization detector

GC-MS

Gas chromatography–mass spectrometry

PFRAP

Potassium ferricyanide reducing antioxidant power

DPPH

2,2-diphenyl-1-picrylhydrazyl radical

MIC

Minimum inhibitory concentration

OD

Optical density

Author contributions

Conceptualization, M.A.A.; validation, A.H., K.D.I. and Z.H.J.; formal analysis, H.I.M.A. and G.V.; investigation, Q.U.A and H.I.M.A.; resources, Z.H.J., J.R., C.A., K.D.I. and K.Y.A.; writing—original draft preparation, M.A.A and Q.U.A.; writing—revision and editing, M.A.A, A.H., H.I.M.A, and G.V.; supervision, M.A.A. and H.I.M.A.; project administration, M.A.A.; funding acquisition, M.A.A., C.A., J. R., and Z.H.J. All authors have read and agreed to the published version of the manuscript.

Funding

This work is a part of research project funded (Grant No. 20-15988/NRPU/R&D/HEC/2021) by the Higher Education Commission (HEC), Islamabad, Pakistan and, the Universidad Técnica Particular de Loja (UTPL) for supporting open access publication.

Data availability

The data will be available from corresponding authors upon a reasonable request.

Declarations

Competing interests

The authors declare no competing interests.