Collection, identification, and extraction of plant material
The leaves of Moringa oleifera were collected from matured healthy trees in a homestead garden at Oke—Ola in Ilaro, Ogun State, Nigeria. The leaves were removed from the branches, sorted, washed properly with sterile water to remove dirt and extraneous materials. The plant was identified by a botanist—Professor F.M. Ogbe of the Department of Biological Sciences, College of Basic and Applied Sciences, Samuel Adegboyega University, Ogwa, Edo State, Nigeria. An herbarium voucher specimen (UBH-M340) was deposited in the Herbarium of the Department of Plant Biology and Biotechnology, University of Benin, Nigeria.
The leaves of Moringa oleifera were air-dried for 12 days and then ground to a fine powder with a mortar and pestle. The large particles were removed, while the powder obtained was stored in a polythene bag before analysis.
For the aqueous extract, 45 g of the powdered leaves was soaked in 380 mL of distilled water and left to stand for 72 h to allow for maceration. The aqueous mixture was filtered using Whatman filter paper. For the ethanol extract, 45 g of the powdered leaves was soaked in 500 mL of ethanol and left to stand for 72 h to allow for maceration. The ethanol mixture was filtered using Whatman filter paper (Azwanida 2015; Nigussie et al., 2021).
Qualitative and quantitative phytochemical analyses of M. oleifera leaf extracts
The method of Akintelu and Amoo (2017) was used to test for alkaloids. Five grams of the powdered leaf was poured into a 250-mL beaker followed by the addition of 200 mL of 10% acetic acid in ethanol and left for 4 min and filtered. Concentrated ammonium hydroxide was added in drops to allow for complete precipitation. The precipitate was collected, rinsed with dilute ammonium hydroxide followed by filtration. The residue was dehydrated and measured.
$$\% \;{\text{Alkaloid}} = \frac{{w_{3} {-}w_{2} }}{{w_{1} }} \times 100$$
where w1 = weight of the sample, w2 = weight of filter paper, and w3 = weight of filter paper after drying.
The quantitative determination of flavonoids was done following outlined methods described by AOAC (2005) to test for flavonoids. A 0.5 g of the powdered leaf sample was placed inside a 250-ml titration flask, followed by 100 ml of 80% aqueous methanol. The mixture was thoroughly whirled for 4 h in a vortex machine. The entire mixture was filtered using a Whatman filter paper No. 42, and the entire procedure was repeated. The whole deposit was vaporized to dryness in a water bath and weighed
$${\text{Flavonoids }}\left( {{\text{mg}}/{1}00\;{\text{g}}} \right) = \frac{{{\text{weight}}\;{\text{ of}}\;{\text{ untreated}}\;{\text{ sample}}}}{{{\text{weight }}\;{\text{of}}\;{\text{ treated }}\;{\text{sample}}}} \times 100$$
The method of Akintelu and Amoo (2017) was used to test for saponin. The powder (2 g) of the sample was poured into a 250-mL beaker followed by 100 mL Isobutyl alcohol. The solution was mixed with a vortex machine for 5 h to make sure it is even and poured into a 100-mL beaker containing 20 ml of 40% saturated solution magnesium carbonate (MgCO3) followed by filtration to get a colorless solution. One milliliter of the solution was emptied in a 50-ml flask followed by 2 ml of 5% iron (III) chloride (FeCl3) solution, and distilled water was used to make up the volume required. The solution was left for 30 min in other to develop color. The absorbance was read against the blank at 380 nm.
Distilled water (50 ml) was added to 500 mg of powdered leaf sample in a beaker and allowed to stand on a mechanical shaker for 1 h to test for tannin. The sample was filtered using a Buchner funnel and Whatman No. 1 filter paper into a 50-mL volumetric flask. Distilled water was added to the desired volume. After that, 5 ml of the filtrate was poured into a test tube and mixed with 2 ml of 0.1 M FeCl3 in 0.1 M hydrogen chloride and 0.008 M potassium ferrocyanide. A spectrophotometer was used to read absorbance at 420 nm (Fapohunda et al. 2012).
Determination of proximate composition of M. oleifera leaf extracts
The determination of moisture, ash, and crude fiber contents of M. oleifera was determined by the difference in weight before and after drying divided by the weight before drying multiplied by 100%. Crude lipid content was determined by Soxhlet method. Crude protein was determined using the method of Kjeldahl flask. The sum of the percentages of the aforementioned parameters was subtracted from 100% to arrive at the carbohydrate content (AOAC 2005).
$$\frac{{{\text{weight}}\;{\text{before}}\;{\text{drying}}{-}{\text{weight}}\;{\text{ after}}\;{\text{ drying}}}}{{{\text{weight}}\;{\text{ before}}\;{\text{ drying}}}} \times 100$$
Gas chromatography–mass spectrometry (GC–MS) analysis of M. oleifera leaf extracts
Gas chromatography–mass spectrometry analysis of aqueous and ethanol extracts of Moringa. oleifera leaf extracts was done with Shimadzu Japan gas chromatography QP2010PLUS with a fused GC column (2010) and coated with polymethyl silicon (0.25 nm × 50 m) with the following conditions: temperature programming from 80 to 200o C held at 80o C for 1 min, rate 5 °C/min and at 200° C for 20 min, field ionization detector (FID) temperature 300 °C, injection temperature 220° C, nitrogen at a 1 ml/min flow rate, split ratio 1:75. The column length is 30 m with a diameter of 0.25 mm and a flow rate of 50 ml/min. The elute was emptied into a mass spectrometer with a detector voltage and sampling rate set at 1.5 kv and 0.2 s, respectively. The mass spectrum was connected to a computer-fed mass spectra data bank, Hermlez 233 M-Z centrifuge (Germany). The components of the extracts were preliminarily identified by corresponding peaks using Computer Wiley MS libraries and confirmed by comparison with peaks of the mass spectra in available literature (Balamurugan 2015).
Tested organisms
The test bacterial isolates (Staphylococcus aureus, Pseudomonas aeruginosa, Escherichia coli, and Salmonella species) were collected from the microbiology laboratory, Irrua Specialist Teaching Hospital, Irrua, Edo State, Nigeria. The organisms were isolated from stool samples of diarrhea patients who attended Irrua Specialist Teaching Hospital to seek medical treatment. The samples were inoculated into MacConkey agar (Becton Dickinson and Company, Cockeysville, MD, USA), blood agar and mannitol salt agar (both from HiMedia Laboratories, Mumbai, India) and incubated aerobically at 37 °C, for 24 h. After 24 h, plates without growth were incubated further for up to 48 h. The growth of microorganisms was identified by examining colony morphology followed by biochemical identification.
Antibacterial studies
Agar well diffusion method, as described by Erhabor (2017), was used. With the pure cultures of the test bacterial, antibacterial activity was performed to determine their respective tolerance to the extract. Sterilized agar plates were aseptically inoculated with a loopful of the test bacterial isolate. One milliliter (1 ml) of each inoculum was introduced into the petri dish. About 15 ml of Muller Hinton agar was poured and then swung to mix correctly and allowed to solidify. Five wells of 6 mm were bored with the aid of cock borer after solidification. The dry extracts were reconstituted by dissolving in 10% dimethyl sulfoxide (DMSO). In standardizing the isolates, a loopful of the stock culture of the organisms was inoculated into 5 ml sterile nutrient broth and incubated for 24 h. The broth culture of the organisms (0.2 ml) was inoculated into 20 ml of sterile nutrient broth and incubated for 3–5 h. The turbidity of the culture was compared with that of 0.5 Mac‐Farland to standardize the culture to 106 cfu/ml. Then, a volume of 0.2 ml of the reconstituted extract at the tested concentrations was dispensed into the wells. The commercial standard antibiotics discs were used as positive control. The plates were allowed to stand for 30 min for pre‐diffusion of the extract to occur and then incubated at 370C for 24 h. The efficiency of the extracts was carried out by measuring the diameter zone of inhibition around the well (mm). The mean of triplicate results was taken.
The following concentrations: 50 mg/ml, 25 mg/ml, 12.5 mg/ml and 6.25 mg/ml were used to determine the MIC of the leaf extracts. A sterile cotton wool swab was used to inoculate pure cultures on nutrient agar plates and allowed to dry, followed by boring wells (6 mm) with a sterilized cork borer. After that, 100 µL aliquots of the different concentrations of both leaf extracts were transferred into labeled wells. The plates were incubated at 370C for 24 h and, after that, observed for bacterial growth or not. The lowest of all concentrations that failed to give bacterial growth was recorded as the MIC. For the determination of MBC, the concentration that failed to give bacterial growth (MIC) was sub-cultured on the surface nutrient agar, followed by incubation at 370 C for 24 h. The lowest concentration that failed to grow after 24 h of incubation was taken as the MBC (Enerijiofi and Isola 2019).
Analysis data
Data were presented as mean ± SEM of the respective triplicate. One-way ANOVA was done to compare the means of the groups and Duncan's multiple range tests to analyze differences among different means. Differences at p < 0.05 were statistically significant. SPSS software was used for the analysis (Ogbeibu 2005).