Determination of antibacterial activity by agar well diffusion method

The crude extract (1.5 g each) was dissolved in 15 ml of dimethyl sulfoxide (DMSO) (for organic extracts) and distilled water (for aqueous extracts) using a vortex mixer to make a stock solution of 100 mg/ml. The bacterial strains tested included sensitive and resistant strains of S. aureus, E. coli, K. pneumonie, P. aeruginosa, S. pyogenes, and S. typhi. The species were selected because they have been reported to be the most prevalent bacteria that cause skin infections and life-threatening multi-drug resistant organisms in skin infections [42]. S. typhi is also known for causing cutaneous skin and sternal wound infections [22, 35]. The organisms were obtained from the Department of Pharmacology College of Veterinary Medicine, Animal Resources, and Biosecurity, Makerere University. The organisms were isolated on a nutrient broth, and diluted with 20 ml of the sterile nutrient growth media. The dilutions formed the bacterial stock solutions that were used in the agar-well diffusion assays. Ciprofloxacin (CIP), a standard antibacterial drug was used for control experiments [7].

Muller Hinton agar plates were inoculated with test bacteria strains separately by spreading the cultured organism on the surface of solidified agar to obtain a uniform inoculum. Four wells (6 mm in diameter) were punched in the agar. The plant extract (100 μL of 10 mg/ml each) was dispensed into each well. For the control experiment, ciprofloxacin disks (10 μg) were dispensed into the well. The plates were incubated at 37 °C for 24 h. The experiment was performed in triplicate and the diameter of zones of inhibition was measured using a ruler. The results were recorded in millimeters as the average zone of inhibition. Extracts with a zone of inhibition greater than the diameter of a well were considered active [7, 41].

The lowest concentration of the extracts which inhibited the growth of tested bacteria was measured by the MIC using the broth micro-dilution method. Five test tubes were arranged in a rack and labeled 1 to 5. Each test tube was filled with 1.0 ml of DMSO, 1.0 ml of the plant was having a concentration of 100 mgml⁻¹ was added to the first test tube. Serial twofold dilutions were made from the second test tube to the 5th test tube to make a concentration of 50.00, 25.00, 12.50, 6.25, and 3.12 mg/ml. Six wells were punched in the agar and labeled according to the dilution order. The diluted extract (100 μl each) was dispensed into each respective well. All experiments were performed in triplicate. The plates were incubated at 37 °C for 24 h. The activity was determined visually by the presence or absence of colonies. MICs were determined as the lowest concentrations of extracts showing clear wells [5].

In determining the MBCs, five test tubes were arranged in a rack and labeled 1 to 5. Each test tube was filled with 1.0 ml of growth media followed by 1.0 ml of the extract having a concentration of 100 mgml⁻¹. Serial twofold dilutions were made from the first test tube to the 5 consecutive test tubes to make a concentration of 50.00, 25.00, 12.50, 6.25, and 3.12 mg/ml. The test bacteria (10 μl) was added to each test tube (1-5) and then incubated at 37 °C for 24 h. The agar plate was divided into five partitions and labeled according to the dilution order. The cultured mixture (10 μL) was smeared on each respective partition. All experiments were performed in triplicate. The plates were again incubated at 37 °C for 24 h. Partitions without growth were observed visually and MBCs were recorded as the lowest concentration of the extract that killed the tested bacteria [5].

Acute dermal toxicity assay was carried out using OECD guideline 402 as described by Banerjee et al. [6]. A total of 14 young adult healthy Wistar albino rats weighing 80 to 120 g were divided into 2 groups (treated group/transdermal patch and control/non-treated group). The treated group consisted of 2 animals and non-treated/control group consisted of 1 animal. One day before the acute dermal toxicity started, the backs of rats were clipped with an electric clipper. Each rat was caged individually and left undisturbed for 24 h. The exposed skin was cleaned with non-irritating distilled water. On the test day, the extracts were dissolved in distilled water and applied evenly to the exposed skin at a dose of 8000 and 10,000 mg/kg body weight and covered with a semi-occlusive dressing. Distilled water (3 ml/kg body weight) was topically applied to the exposed skin of the control rats. The rats were then returned to their cages. The animals were observed twice daily for 14 days for signs of irritation, general behavior, and possible mortality. Body weight measurement, food, and water consumptions were taken daily for 14 days. On the 15th day, all animals in the vehicle control and treated groups were killed, organs were carefully taken out and weighed. Histopathological examination of animals was performed at the termination of the study on day 15. The aqueous extracts were selected because water is the common solvent used by traditional herbalist to prepare herbal drugs. Blood for clinical chemistry was placed in vacuum blood collection tubes devoid of anticoagulant (serum tube) and allowed to clot at room temperature. Blood samples were centrifuged at 3000 rpm for 10 min after collection and then the serum was separated. Serum biochemistry parameters including creatinine (CREJ), alanine aminotransferase (ALT), and aspartate aminotransferase (AST) were analyzed by COBAS 6000 analyzer machine. The serum of the experimental rats was compared with those of control rats.

The antioxidant activity of the crude extracts was determined according to the method of Himesh et al. [16] with slight modifications. Each crude extract (11 mg) was dissolved in 100 ml of methanol for organic extract and water for the aqueous extract to make 110 μg/ml stock solution. Ascorbic acid was used as a standard and was prepared in the same way as the extracts using distilled water. DPPH solution of concentration 0.5 mM and 0.1 mM was prepared and kept in darkness for 45 min at room temperature. The scavenging activity of S. princeae was measured using 0.5 mM whereas 0.1 mM of DPPH was used to measure the scavenging activity of P. caespitosus, P. febrifugum, and E. tomentosa. The sample solution (2 ml) was pipetted and mixed with DPPH (2 ml) solution in a cuvette. The mixture was kept in darkness for 15 min to stabilize. The absorbance of the mixture was measured at 517 nm using Shimadzu UV-VIS double beam spectrophotometer against a blank. The percentage inhibition of radicals was calculated using the following formula;

Control Absorbance is the absorbance of DPPH only and Sample Absorbance is the absorbance of sample mixed with DPPH. The stock solutions were serially diluted five times and the antioxidant activity of the diluted solutions was determined. The concentration of the extract (antioxidant) required to decrease the initial DPPH concentration by 50% (IC₅₀) was calculated using the Logit regression analysis. A lower IC₅₀ value corresponded to a larger scavenging power. All experiments were conducted in triplicate and values expressed as mean ± standard deviation (SD).

The sun protection factor was determined using a modified method reported by Dutra et al. [12]. Each crude extract (0.1 g) was dissolved in 50 ml of ethanol to make a solution of concentration 2 mg/ml without ultra-sonication. The absorption data of each sample was measured using Shimadzu UV-VIS double beam spectrophotometer against ethanol as a blank. The absorption data were obtained for every 5 nm interval between the range of 290 to 320 nm, and four determinations were made at each point and the sun protection factor was determined using the Mansur equation.

Where CF is the correction factor (=10), EE is the erythemal effect spectrum, I is the solar intensity, and Abs is the absorbance.