In Vitro Antimicrobial and Antioxidant Activity of the Essential Oil of Lemon Grass

Essential oils are biosynthetically produced combinations of plant secretions that are predominantly made of terpenes or terpenoids. They are stored in structures or canals in the plant. Essential oils have auditory and hysterical properties and differ from one another. They can be made of (in order of possible predominance) phenylpropanoids, alcohols, or even ketones. The two highest classes are monoterpenes and sesquiterpenes. Due to this, monoterpenes (C10) and sesquiterpenes (C15) are some of the highest constituents, representing over three-fourths of the essential oil. Many essential oils have either monoterpenes and/or sesquiterpenes, and all of these constituents can define the bioactivity and odor of the oil [1]. Oxygenated and aromatic derivates of terpenoids such as (in order of predominance) alcohols, phenols, aldehydes, and ketones are often empirically proven to possess the most antimicrobial and antioxidant properties of these essential oil constituents as a whole. Terpenes derived from essential oils can have major variances or lack of variances due to numerous factors including plant genetics, geographic location of where the plant was grown, time of harvest, time of processing, and method of extraction [2]. Essential oils are amphipathic and have a relatively small MB. These two things help essential oils to gain rapid permeability and provide a wide spectrum of phytotherapeutic properties. Essential oils are lipophilic, which provide rapid permeability. Essential oils have been a focal point in recent applied research due to issues with multilayer drug resistant pathogenic microorganisms. Essential oils have shown great properties to resolve inflammation and provide analgesia. Essential oils also have great antioxidant properties which have been shown to help with issues of oxidative stress. In relation to these properties, essential oils have also shown great potential as a natural food preservative and attractive ingredient to help maintain the safety and quality of cosmetics [3].

Antimicrobial activity of essential oils

Research in the past focused a lot on the antimicrobial properties of essential oils against a wide range of microorganisms such as fungi, bacteria, viruses, and yeast. Traditional antibiotics function by inhibiting a specific biochemical target, such as the synthesis of a cell wall or the inability to translate proteins. In contrast, essential oils target multiple sites and pathways, which significantly minimizes the development of antibiotic resistance. The cell membrane of the microorganism is the main target [4-6]. The hydrophobic nature of essential oil constituents allows them to penetrate the membrane phospholipid bilayer which causes a disruption in the membrane and makes it more permeable, leading to leakage of the ions, ATP and other cellular contents, and the membrane potential. This, in turn, dissipates the membrane potential, disrupting the cellular respiration and the microorganisms ultimately die. Biochemical structures such as the monoterpene such as thymol and carvacrol have been shown to cause a general destabilization of the outer membrane of Gram-negative bacteria, and some phenolic compounds, such as eugenol, also have been shown to inhibit the activity of ATPases [7]. The Gram-positive bacteria are more susceptible to the essential oils as compared to the Gram-negative bacteria because the Gram-negative bacteria have the outer membrane which is composed of a lipopolysaccharide that acts as a barrier to the hydrophobic compounds. However, some essential oils that are rich in compounds such as aldehydes, for example, citral which is from lemongrass, and cinnamaldehde which is from cinnamon, still can have an appreciable activity against the Gram-negative microorganisms such as Escherichia coli and also Pseudomonas aeruginosa. The range of the minimum inhibitory concentrations of essential oils and the ranges are from 0.01% to 1.0 % (v/v) have an appreciable activity against a range of microorganisms, although the ranges are highly dependent on the microorganism and the composition of the oil [8]. Due to the global rise in antibiotic resistance, essential oils are being researched more frequently as stand-alone antimicrobial agents or as synergistic adjuvants that restore the effectiveness of traditional antibiotics. This provides a positive alternative for addressing resistant strains in clinical, veterinary, and food preservation situations [9].

Antioxidant Activity of Essential Oils

Oxidative stress occurs when reactive oxygen species (ROS) exceed the action of the antioxidant defense. Chronic diseases like cancer, heart disease, neurodegeneration, and aging all show signs of oxidative stress. While synthetic antioxidants can aid in the reduction of oxidative stress, like BHT, and BHA, the hepatotoxic potential and carcinogenic nature of these additives can outweigh the benefits [10]. Therefore, the use of extracted plant antioxidants is enticing. The core of the antioxidant potential inherent in plant oils is the ability of phenolic terpenes and monoterpenes to reduce the reactivity of free radicals, use oxidative metal ions, and break the cross-linking of the oxidative chain. The main actions of phenolic groups are to donate a hydrogen atom (THAT), or a single electron (SET) to lessen the reactivity of the free radical, which then forms a resonance-stabilized phenoxyl radical. Radical scavenging activities in screening for the antioxidant power are often characterized by DPPH (2,2-Diphenyl-1-Picrylhydrazyl), the qualitative reduction of ABTS (2,2′-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)) and the measurement of FRAP (Ferric Reducing Ability of Plasma) and total phenolic contents using the Folin-Ciocalteu reagent. Plant oils high in thymol, carvacrol, eugenol, and citral show a high degree of reactivity with free radicals to the extent that they are often comparable to ascorbic acid and Trolox [11]. Oregano (carvacrol and thymol) and clove (eugenol) oils, for example, show DPPH IC50 results of as low as 5-50 µg/mL. Although essential oils can show great potential for antioxidant activity, the volatility and the metabolic conversion and the bioavailability in the human body as well as the in vitro antioxidant activities all play important roles for determining the practical utility of the essential oils.
 The addition of antioxidant essential oils in food, cosmetics, and pharmaceuticals is a natural method to antioxidant lipid peroxidation, increase shelf life, and reduce the effects of oxidative damage in the body [12].

Previous studies on lemon grass essential oil

Lemongrass (Cymbopogon citratus) is a perennial grass that has been historically used in folk medicine to treat various diseases such as digestion, fever, high blood pressure, and infection. Lemongrass is naturally present in tropical and subtropical regions around the world. Numerous studies have shown that lemongrass essential oils have been effective at suppressing the growth of different pathogens [13]. Lemongrass essential oil has an average citral content of 65% to 85% and can contain myrcene, citronellal, geraniol, limonene, and β-caryophyllene. Lemongrass oil has shown an inhibitory zone that ranges from 15 to 40 mm against pathogens such as Staphylococcus aureus, Escherichia coli, Salmonella typhimurium, and Candida albicans, and the minimum inhibitory concentration (MIC) values primarily ranged from 0.125% to 0.5% (v/v). The majority of the antimicrobial activity for lemongrass essential oil comes from citral. Citral is a derivation of the essential oil obtained from lemongrass and is primarily an aldehyde. This mean that citral has an -CHO functional group that can interact and bind to Gram -(–) bacteria outer membrane proteins to enhance the oil's antimicrobial efficacy among other Gram -(–) bacterial strains [14]. Furthermore, Lemongrass essential oil has shown a moderate to high level of activity in the ABTS and DPPH radical scavenging activity assays. Studies have shown that the lemongrass essential oil with the highest citral percentage has shown a DPPH IC50 of 20 - 150 µg/mL depending on the oil's geographical source. The total gallic acid phenolic content achieved 50 mg/g of oil. Lemongrass oil is usually judged weaker than clove and oregano oil, but is considered stronger than most culinary herbs. Some studies examining combinative effects have found that lemongrass oil together with standard antibiotics like amphotericin B and ciprofloxacin have demonstration synergy against resistant bacteria. While these were promising in vitro findings, there are some issues affecting lemongrass’ therapeutic value. These issues include volatility, organoleptic effects, and organ toxicity at high concentrations. Still, lemongrass oil is being widely used to preserve food, to enhance the shelf life of food products like meat, fish and fruits and to provide calming and immune stimulating effects when used in aromatherapy. it should also be used in topical formulations for the treatment of skin that has been infected with fungi, as well. Carefully executed research in the future should be directed to the development and implementation of such formulations combined with potent preservatives to enhance the self-life while maintaining the calming effects of lemongrass oil lipid-bilayer [15].

MATERIAL & METHODS

Collection and authentication of plant material

Leaves of lemon grass (Cymbopogon citratus) were harvested during the flowering period of March to April. The harvesting was done early in the morning (7:00 to 8:00 A.M) to optimize the concentration of the essential oil and the minimal loss of the volatile compounds. Careful selection was done in order to choose leaves that were expanded fully and that were safe from diseases and insects. The raw material was thoroughly washed with running tap water and then with distilled water to remove surface contaminants, debris, and dust. After the washing, the leaves were dried in the shade and ambient temperature (25 ± 2°C) for NFT 7 to 10 days to achieve constant weight and the leaves were turned periodically to avoid the growth of fungi. The dried up leaves were ground in to a coarse powder with the help of an electric grinder (mesh 1-2 mm).

Extraction of essential oil

Following the recommendations of the European Pharmacopoeia, the Clevenger type apparatus was improved so that the extraction of Lemon grass Essential Oil by Hydrodistillation could be performed. For a single round of extraction, 100 grams of dry, ground Lemon grass leaves and 1 liter of distilled water was added into the 2 liter round-bottom flask. The flask was placed on a temperature-controllable heating mantle, and distilled for four hours after the first drop of condensate was observed. For the prevention of thermal degradation of compounds that were labile to heat, the distillation rate was set to 2-3 mL of distillate per minute. The distillate was separated into two phases. The upper phase was the essential oil, and the lower phase was the aqueous hydrosol. The essential oil was separated carefully from the aqueous hydrosol, and the aqueous phase was dried by the addition of an anhydrous sodium sulfate (Na2SO4). The dried aqueous phase was further dried by the use of a membrane filtration that reinforced the PTFE and had a filtering pore size of 0.22 micrometers. The oil was transferred to an amber glass vial so that the vial was flushed with nitrogen gas and preserved from oxidation. The vial was kept at 4° C until further analysis occurred. The extraction yield was calculated as (v/w) concerning the dry plant material. The average yield from three repeated extractions was reported as [e.g., 0.8–1.2% (v/w)].

ANTIMICROBIAL ASSAYS

Test microorganisms

The antimicrobial effect of lemongrass essential oil was studied using the reference microbial strains. The bacterial strains included Staphylococcus aureus (ATCC 25923), Bacillus subtilis (ATCC 6633), Enterococcus faecalis (ATCC 29212), Escherichia coli (ATCC 25922), Pseudomonas aeruginosa (ATCC 27853), and Salmonella typhimurium (ATCC 14028). The yeast Candida albicans (ATCC 10231) and the filamentous mold Aspergillus niger (ATCC 16888) were the two fungal strains included. All of the microbial strains came from [source, e.g. the American Type Culture Collection or a national microbial culture repository]. The fungal stock was kept on the slants of Sabouraud dextrose slants and the bacterial stock was kept on the slants of nutrient agar at 4°C. Just before each experiment, the bacterial strains were transferred to the Mueller Hinton agar (MHA) after which they were incubated at 37°C for 18-24 hrs. The fungal strains were transferred to the potato dextrose agar (PDA) and incubated at 28°C for 48-72 hrs. so as to obtain fresh, viable colonies.

Disc Diffusion Method

The disc diffusion assay was conducted based on the recommendations of the Clinical and Laboratory Standards Institute (CLSI) (M02-A11). An aqueous solution of each tested microorganism was prepared in sterile saline (0.85% NaCl) and adjusted to a turbidity of the 0.5 McFarland standard (approximately 1–2 × 10? CFU/mL and 1–5 × 10? CFU/mL for bacteria and fungi, respectively). A sterile cotton swab was dipped into the test microorganism suspension. To remove excess fluid, the swab was rotated against the tube wall, and the swab was used to evenly streak the surface of the agar plates, either Mueller Hinton agar (for bacteria) or potato dextrose agar (for fungi), in three directions to create a confluent lawn. To prepare substance impregnated discs, sterile Whatman No. 1 filter paper discs (6.0 mm) were respectively impregnated with 10 µL of lemongrass essential oil, either neat, or with concentrations of 5%, 10%, 20% and 50% (v/v) in 1% Tween 80. Control discs were prepared by impregnating the discs with 10 µL of sterile distilled water with 1% Tween 80 (negative control), and standard antibiotic discs (positive control). The antibiotics were used as follows: gentamicin (10 µg) for bacteria and nystatin (100 µg) for fungi. Using sterile forceps, the discs nestled in the agar plates and were subsequently incubated in the appropriate growth conditions (bacteria: 37 °C for 24 h; fungi: 28 °C for 48–72 h). Once the incubation was complete, the diameter of each inhibition zone (including the diameter of the disc) was measured with the aid of a digital Vernier calliper. All tests were performed in triplicate and expressed as mean ± SD.

Minimum Inhibitory Concentration (MIC)

Lemongrass essential oil’s antimicrobial potential and MIC against various pathogens were assayed using the broth microdilution method as described in the CLSI guidelines (M07-A9 for bacteria and M27-A3 for fungi). Essential oil was a prepared two-fold serial dilution (0.024% v/v) in 96-well microtiter plates. Essential oil was first diluted in 1% Tween 80 for improved solubility in rhe medium that was subsequently used to dilute the oil. In the case of bacteria, Mueller Hinton broth was used, and for fungi, Shake RPMI 1640 was used, to further complete the dilution to a total of 5% (v/v). In each well, 100 µL of the diluted essential oil was paired with 100 µL of a microbial suspension of 1 × 10? CFU/mL for bacteria or 1 × 10? CFU/mL for fungi, resulting in a total volume of 200 µL in each well. Internal control wells were used to determine the presence or absence of microbial growth (oil void) and absence of bacteria (sterile broth). Plates were incubated under the same conditions as described for the disc diffusion method. Following the incubation period, the degree of bacterial growth was assessed visually, and measured in terms of 600 nm using a microplate reader. The MIC was the lowest concentration of essential oil resulting in complete inhibition of bacterial growth. In case of oil void wells, 10 µL of each were subcultured to solid media, and the MBC/MFC was the lowest concentration resulting in absence of colony growth. This was repeated three times.

ANTIOXIDANT ASSAYS

DPPH Radical Scavenging Assay

Lemon grass essential oil was studied for its ability to scavenge the DPPH (2,2-diphenyl-1-picrylhydrazyl) radical using the almost unchanged technique pioneered by Brand-Williams et al. Prior to use, a DPPH solution at 0.1 mM was prepared in methanol and stored in the dark at 4 °C for up to 24 hours. Using methanol, essential oil samples were diluted to 25, 50, 100, 200, 400, and 800 µg/mL. In test tubes, 2 mL of the prepared DPPH solution was combined with 2 mL of the oil-diluted samples. The mixture was placed in a vortex mixer and incubated in the dark at room temperature (25 °C) for 30 minutes. The samples were compared to the blank (methanol) and the absorbance of the negative (without DPPH) was measured using the UV-Vis spectrophotometer at 517 nm. The same concentration of ascorbic acid was used as a positive control. In addition, the negative control was made with only methanol and DPPH in the absence of the oil sample. The DPPH scavenging activity was measured with the equation Scavenging activity (%) = (A_control − A_sample) / A_control] × 100, where the DPPH absorbance sample was denoted A_sample and the DPPH absorbance control was denoted A_control. The linear regression analysis yielded the IC50, the concentration required to scavenge 50% of DPPH radicals. Mean graphs of the collected data were plotted and a standard deviation was placed. All tests were conducted in triplicate for the same purpose.

ABTS Radical Scavenging Assay

The protocol of Re et al. was followed for the ABTS radical scavenging assay. This involved generating ABTS radical cation (ABTS•?) by reacting a 7 mM ABTS stock solution and a 2.45 mM potassium persulfate (final concentration) at a 1:1 ratio. The mixture was kept covered in a dark room at room temperature for 12–16 hrs. ABTS•? cation was diluted in ethanol to have an absorbance of 0.700 ± 0.020 at 734 nm. The concentrations of lemon grass essential oil were made to be 5, 10, 20, 40, 80, and 160 µg/mL in ethanol. 20 µL of each of the oil dilutions were mixed with 980 µL of ABTS•? diluted solution. The absorbance was measured at 734 nm after incubating for 6 minutes in the dark. Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid) was used as a positive control. ABTS radical scavenging was calculated as follows = Scavenging activity (%) = [(A_control – A_sample) / A_control] × 100. Also, as for the DPPH assay, IC50 was calculated. Each concentration was performed in triplicate.

Total Phenolic Content (TPC)

The phenolic constituents of lemongrass oil are described by the Folin-Ciocalteu method. A calibration curve was constructed by making an order of dilutions of the syringaldazine standard (i.e. 0, 20, 40, 60, 80, and 100 mg/L) in methanol. Methanolic diluted phenolic compounds and the Folin-Ciocalteu reagent are also mixed in a 1:5 ratio, in which 2.5mL of the diluent and 0.5mL of the mixed phenolics is taken in a test tube. The mixture is inverted 5-10 times and left to settle for 3 minutes, whereafter sodium carbonate (2.0 mL of 7.5% w/m) is added. The mixture is left covered in a dark for 60 minutes, the dark color developed is due to the resultant phenolic compound. The absorbance of this compound is measured against a blank for the standard at 765 nm, and mg GAE/g oil is expressed as the final result of the calculated phenolic constituents where determined GAE is the phenolic standard, and mg GAE/g oil is the calculated phenolic standard, and mg GAE/g oil is expressed when the tube is left to settle without the addition of the sample to ensure optimum condition. All the estimates were taken thrice and the final expressed results of the calculated phenolic constituents where determined are expressed as standard GAE.

Statistical Analysis

All experiments were conducted in triplicate, and the results were expressed as mean ± standard deviation (SD). Statistical analysis was performed using GraphPad Prism version 9.0 (GraphPad Software, San Diego, CA, USA) and IBM SPSS Statistics version 26.0. For antimicrobial assays, differences in mean inhibition zone diameters among different concentrations of essential oil and standard antibiotics were analyzed using one-way analysis of variance (ANOVA) followed by Tukey's post hoc test for multiple comparisons. 

 RESULTS AND DISCUSSION

Chemical composition of lemongrass essential oil

Hydro–distilling dried leaves of Cymbopogon citratus produced a light-yellow oil with a lemony scent having an average extraction yield of 1.05% (v/w). The oil was composed of 98.4% of 28 constituents identified through GC–MS analysis. Citral and its isomers, geranial (47.2%) and neral (34.5%), accounted for 81.7% of the total oil. Other constituents included 4.6% β-myrcene, 2.8% limonene, 2.1% geraniol, 1.9% citronellal, and 1.3% β-caryophyllene. Based on the literature, the presence of citral forms the signature lemongrass essential oil, so its presence is chiefly responsible for the oil’s bioactivity.

Antimicrobial Activity

The antimicrobial activity of lemongrass essential oil was evaluated against six bacterial and two fungal strains using disc diffusion and broth microdilution methods. The results are summarized in Table 1.

Fig: 1  Illustrates the Antimicrobial Activity of a substance (likely the lemongrass essential oil previously discussed) against three different microorganisms: Candida albicans, Staphylococcus aureus, and Pseudomonas aeruginosa.

Table 1. Antimicrobial activity of lemongrass essential oil (mean ± SD, n=3)

Lemongrass essential oil was found to show concentration dependent antimicrobial activity toward all microorganism groups analyzed in Table 1. The inhibition zone varied from 6.5 mm to 42.3 mm for 5% oil against P. aeruginosa and neat oil against C. albicans, respectively. For Gram-positive bacteria, inhibition zones were larger than those for Gram-negative bacteria. For example, at 50% oil, the inhibition zones for S. aureus and B. subtilis were 31.4 mm and 29.6 mm, respectively, compared to 20.5 mm for P. aeruginosa. The larger inhibition zones and the differential susceptibility can also be attributed to the Gram-negative bacteria’s lipopolysaccharide outer membrane, which hinders the penetration of some hydrophobic components found in most of the essential oils. The oil was shown to have good activity against E. coli and S. typhimurium as well with E. coli and S. typhimurium showing MIC of 0.5% (v/v) which is higher than the observation in the literature of MIC of 0.5-1% (Tyagi & Malik 2011). The MIC values of lemongrass oil was recorded to be 0.125% (v/v) for C. albicans, S. aureus, and B. subtilis, and were found to be 0.25% (v/v) for E. coli, S. typhimurium, E. faecalis, A. niger, and the most resistant organism of the bacterium studied was P. aeruginosa with MIC of 0.25% (v/v). The MBC/MFC values was equal to or were twice the MIC values which indicate the lemongrass oil to be a bactericidal/fungicidal activity rather than a bacteriostatic/fungistatic active. The oil was significantly more active when tested against C. albicans (zone 42.3 mm, MIC 0.125%) when compared with the filamentous mold A. niger. Yeasts are shown to be more susceptible to citral rich oils that inhibit the synthesis of ergosterol in the fungal membranes (Silva et al., 2016). While compared to the positive controls, undiluted lemongrass oil showed inhibition zones greater than gentamicin against S. aureus (38.7 mm vs 28.3 mm) and B. subtilis (36.2 mm vs 26.7 mm) and was similar to nystatin against C. albicans (42.3 mm vs 31.6 mm) when the inhibition was compared to the positive controls. Therefore, it could be concluded that lemongrass essential oil is a good choice for a natural source of topical antimicrobial interventions for drug resistant Candida and Staphylococcus strains.

Antioxidant activity

The antioxidant potential of lemongrass essential oil was evaluated using DPPH radical scavenging, ABTS radical scavenging, and total phenolic content (TPC) assays. Figure 1 illustrates the DPPH radical scavenging activity of the oil in comparison to ascorbic acid.

Figure 2 DPPH radical scavenging activity of lemongrass essential oil and ascorbic acid (positive control)