The leaves of M. oleifera were harvested from Silanwali, district Sargodha, Punjab, Pakistan at four different stages /periods (15^th October/ early stage, 15^th November/mid stage, 15^th December/penultimate stage and 15^th January/late stage) with the gap of 1 month during the year 2017. Leaves were washed with water to remove any dusty particles, then shade-dried for 1 week and finally pulverized into a fine powder. The powdered sample was used for different experiments and analyses.

Moringa leaf powder was weighed (20 g) and mixed with 200 ml of 80% methanol in a conical flask; shaking was carried out by using an orbital shaker for 8 h at 200 rpm. After shaking, the extracts were separated from the residue by filtering through Whatman filter paper No. 1. The residue was extracted again in a similar fashion, and both the filtrates recovered were combined. The extra solvent was distilled off under reduced pressure/vacuum using a rotary evaporator machine until a solvent-free extract was obtained. The extract was kept at −4°C for further studies.

Total phenolic contents (TPC) were analyzed by the Folin-Ciocalteu reagent method (Zahoor et al., 2016). A weighed amount of extract was placed in a test tube and mixed with Folin-Ciocalteu reagent (0.5 ml) and distilled water (7.5 ml). The mixture was kept at room temperature (22°C) for 5 min, and then 2 ml of 20% Na₂CO₃ was added. After that, the mixture was further incubated for 20 min at around 40°C and then absorbance was noted by a spectrophotometer at a wavelength of 700 nm. TPC was calculated using a gallic acid standard calibration curve.

For TFC determination, the weighed amount of aqueous extract (0.5 ml) was mixed with 0.3 ml of 5% NaNO₂ in a test tube. After 5 min, 0.3 ml of 10% AlCl₃ was added and the mixture incubated for 6 min, followed by the addition of 2 ml of 1.0 M NaOH. The absorbance of the final reaction mixture was measured at 510 nm using a spectrophotometer, and the amount of TF was calculated as catechin equivalents using a standard calibration curve (Aiyegoro and Okoh., 2010; Babbar et al., 2011).

For the preparation of Bradford reagent, 100 mg of Coomassie brilliant blue G250 was dissolved in 50 ml of ethanol. Then 100 ml of 85% phosphoric acid was added till the complete dissolution of the dye and filtered through the Whatman No.1 filter paper. Briefly, 1 ml of different concentrations of stock solutions like 20, 40, 60, 80, and 100 mg were taken and then in each of the test tubes, 5 ml of Bradford reagent was added. Similarly, 1 ml of each sample was taken in a labeled test tube and 5 ml of the Bradford reagent was added and the mixture incubated for 5 min. A blank, containing only water and 5 ml of Bradford reagent, was also prepared. The absorbance was measured at 595 nm against a blank solution and the amount of protein was calculated (Stoscheck, 1990).

Five Gram of each sample was taken into porcelain crucible and heated on flame until turned black. This black ash was then placed into the muffle furnace for about 5 h until it turned into white or gray ash. The furnace temperature was slowly increased from room temperature to 500°C, and the percentage of ash content was determined gravimetrically (Turkekul et al., 2004; Farhoosh and Riazi., 2007).

The sample was ashed and then placed in Eppendorf tube for further analysis. For acidic digestion of ash, two drops of HNO₃ and 3-4 drops of acidic water were added to the ash. The acidic water was prepared by adding a few drops of HCL to 200 ml of distilled water. This solution was then filtered into a volumetric flask and further diluted. Finally, the analysis was made by using an atomic absorption spectrophotometer using the respective minerals/metals’ hollow-cathode lamps. The mineral content was determined by preparing and analyzing a series of standard solutions of the minerals through the construction of calibration curves (Varadharaju et al., 2001).

Different standard amino acid solutions (0.1–1.0 ml) were pipetted into the respective marked test tubes and further diluted with distilled water. One mL of ninhydrin reagent was added to all the test tubes. All the test tubes were placed in a water bath for 15 min. Similarly, test samples were prepared/treated. After cooling the sample and standard test tubes, 1 ml of ethanol was added to each test tube and absorbance was recorded at 570 nm for each solution. The amount of amino acids were calculated following the standard calibration curve (Yemm et al., 1955).

The determination of vitamin C was carried out spectrophotometrically using 2,4-dinitrophenylhydrazine reagent after pre-treatment of samples with trichloroacetic acid (TCA). The absorbance of the sample and standard solution was recorded at 540 nm (Tantray et al., 2017).

A well-blended sample (5–25 g) containing 10–500 mg total carotenoids was taken in a conical flask. The carotene extraction was performed using a mixture of petroleum ether and distilled water via liquid–liquid partitioning. The absorbance of the test solution recovered along with the standard solutions was measured at 452 nm. The total carotenoid content was estimated using a standard curve, and the results were expressed as mg/100 g of sample (Mustapha and Babura., 2009).

The data were expressed as mean values ± SD for triplicate measurements. A 2-way ANOVA was performed using SPSS version 22.0.0.0 software.

The plantation of M. oleifera is considered suitable in regions where temperature ranges from 25–35°C. The soil should be loamy with a moderate acidic to alkaline pH. Most of the soils in Pakistan are calcareous in nature with a pH of 8. The pH of the soil in Silanwali is also reported to be 7.9–10.4. At this pH, the availability of P, K, Zn, Cu, Fe, Mn, and B is high, which is useful for the health and growth of most plants (Aslam et al., 2005).

The percentage yield of phenolic bioactive extracted from M. oleifera leaves using 80% methanol ranged from 14.21 to 9.69% in relation to different maturity stages (Figure 1). Extraction yield of early, mid, penultimate, and late stages are 14.21, 13.66, 10.99, and 9.69%. There was no literature available regarding the effect of maturity stages on the extract yield of M. oleifera leaves. However in the studies conducted by Gull et al., 2012 on guava fruits, they found that the fully ripened fruits from the three selected regions exhibited the highest methanolic extract yields (18.92–24.91%), while those picked at the un-ripe stage offered the lowest (12.05–13.23%).

The total phenolic contents of the 80% methanol extract of M. oleifera leaves at different maturity stages are shown in Table 1. The total phenolic contents in M. oleifera leaves were found to vary from 95.26 to 38.22 mg GAE/g. The TPC in early, mid, penultimate and late stage samples was 95.26 ± 0.89, 60.36 ± 1.14, 46.85 ± 1.55, and 38.22 ± 1.61 mg/g, respectively. The highest contents of phenolics were recorded in young leaves, and the lowest contents were present in late stage leaves. In line with current results, Jahan et al., 2015 also reported that the TPC of the tender leaves (high chlorophyll) and mature leaves (low chlorophyll) were observed at 35.51 and 30.83 mg/g GAE, respectively.

The total flavonoid content of an 80% methanolic extract of M. oleifera leaves at four different maturity stages ranged from 12.26 to 30.07 mg CE/g. Values of TFC in early to late stages ranged from 12.26 ± 0.47, 16.84 ± 0.72, 25.49 ± 1.41 and 30.07 ± 1.09 mg CE/g, respectively. In line with TPC results, Jahan et al., 2015 described in their work that the mature leaves (high chlorophyll) were found to have high flavonoids (98.67 mg CE/g) as compared with tender leaves (low chlorophyll), which had lower flavonoids (32.74 mg CE/g).

Phenolics and flavonoids of M. oleifera methanolic leaf extract were individually identified by using HPLC with a UV-visible detector. The signals were detected at 280 nm and the identification of unknown compounds was made by comparing the retention times with those of pure standards. The HPLC results revealed the presence of six phenolics and flavonoids such as quercetin, gallic acid, benzoic acid, sinapic acid, ferulic acid, and p-coumaric acid (Table 2). The major flavonoid was quercetin with a concentration of 21.64 ppm, followed by benzoic acid, ferulic acid, sinapic acid, gallic acid, and p-coumaric acid with concentrations such as 13.03, 8.85, 3.39, 2.88, and 1.59 ppm, respectively. The HPLC chromatogram of M. oleifera leaves is shown in Figure 2.

The values of ash content in M. oleifera leaves were found at 10.36% at first stage, 11.14% at 2nd stage, 13.20% at 3rd stage and 15.9% at the late stage (Table 3). The lowest ash content was found in the early stage (10.36%) and the highest amount of ash was found in the late stage (15.9%). However, the studies conducted by Agamou et al., 2015 demonstrated that young and mature leaves had almost similar total ash contents, such as 8.90 ± 0.45 and 8.97 ± 0.50 g per 100 g, respectively. Likewise, they also observed no difference in the ash content of M. oleifera leaves collected from different localities.

The concentration of proteins in M. oleifera leaves at different maturity stages ranged from 50.3 to 56 mg/100 g. The protein concentrations in the early to late stage were 50.3, 52.1, 54.5, and 56 mg/100g, respectively. Maximum protein content was found in mature leaves, and minimum content was observed in tender leaves. Moyo et al. (2011) presented in studies that M. oleifera leaf powder had a considerable amount of proteins, such as 29 g/100. The protein content in moringa leaves was also affected (11.98%) by the stages of maturity of the plant.

In the current study, among the amino acids analyzed, valine content ranged from 90.87 to 197.66 μg/g (Table 4). The maximum content was noted at late stage (197.66 ± 0.18 μg/g) and minimum content was found in early stage (90.87 ± 0.10 μg/g). Alanine concentration was found to be in the range of 53.07 to 114.3 μg/g. The maximum content was exhibited in late stage samples (114.3 ± 0.41 μg/g) and minimum in early stage sample (53.07 ± 0.08 μg/g). Leucine content was found to be in the range of 55.21 to 114.2 μg/g. The maximum content was exhibited by late stage (114.2 ± 0.32 μg/g) and minimum content was observed in early stage (55.21 ± 0.03 μg/g). Phenylalanine levels were found to be in the range of 48.65 to 104.5 μg/g. The maximum content was shown in late stage sample (104.5 ± 0.06 μg/g) and minimum in early stage sample (48.65 ± 0.26 μg/g). The amino acid peak area of standards and samples was calculated to determinate the amino acid concentration.

β- Carotene concentration was found to be in the range of 89 to 112 mg/100 g (Table 5). The maximum content was shown in the late stage (112 ± 1.40) and the minimum content was found in the early stage (89 ± 0.28). The maximum content was shown in the late stage (112 ± 1.40) and the minimum content was found in the early stage (89 ± 0.28). Our results are contrary to the studies reported by Moran-Palacio et al. (2014), wherein mature leaves had low carotene content (122.46 ± 1.38) and tender leaves had more carotene content (92.77 ± 1.19). In another study, the concentration of carotenoids was appreciably higher in young leaves than in mature leaves and was also affected by the maturity stages. Other than carotenoids, ascorbic acid (271 mg/100 g) was also present in moringa leaves (Shih et al., 2011).

The concentration of vitamin C in M. oleifera leaves at different maturity stages ranged from 143.14 to 164.81 mg/100 g. Highest levels were observed in late stage and minimum in early maturity stage samples. The results agree with the literature, such as fresh M. oleifera samples showed vitamin C content of 62.66–143.58 mg/100 g for tender leaves, 51.22–150.15 mg/100 g for matured leaves. Kalappurayil and Joseph, 2016, demonstrated that the contents of vitamin C in moringa leaves were not affected by maturity stages, but a minor effect was observed by the locality. Whereas the young and mature moringa leaves had almost the same amount of vitamin C, such as 50.22 and 50.78 mg/100g, respectively. Fat-soluble vitamins such as vitamin-A (pre-cursor of beta-carotene) and vitamin-C were also present in M. oleifera (Mbikay., 2012). In another study conducted by Phullakhandam and Failla., 2007, the amounts of lutein and β-carotene were 418 and 272 mg/kg, respectively, in the fresh leaves of moringa and 472 and 166 mg/kg in dried powder.

The composition of different minerals such as copper, nickel, manganese, iron, and manganese was also observed in the current work (Table 6) and found in confirmation with the literature (Castillo-López et al., 2017). The level of copper in the samples of M. oleifera leaves at different maturity stages ranged between 0.595 and 2.08 mg/100 g. The maximum Cu content was found in the late stage (2.08 ± 0.09 mg/100 g), and the minimum copper content was found in the early stage (0.595 ± 0.05 mg/100 g). The levels of nickel in the samples of M. oleifera leaves at different maturity stages ranged from 13.0 and 17.75 mg/100 g. The maximum Ni content was found to be in the penultimate stage (17.75 ± 0.09 mg/100 g), and the minimum Ni content was found to be in the early stage (13.0 ± 0.38 mg/100 g). The level of iron in the samples of M. oleifera leaves at different maturity stages ranged between 21.96 and 58.68 mg/100 g. The maximum Fe content was found to be in the late stage (58.68 ± 0.39 mg/100 g), and the minimum Fe content was observed in the early stage (21.96 ± 0.85 mg/100 g). The level of cadmium in the samples of M. oleifera leaves at different maturity stages ranged from 0.532 and 1.236 mg/100 g. The maximum Cd content was found to be in the late stage (1.236 ± 0.11 mg/100 g), and the minimum Cd content was found to be in the early stage (0.532 ± 0.04 mg/100 g). The level of manganese in the samples of M. oleifera leaves at different maturity stages ranged between 5.56 and 13.84 mg/100 g. The maximum Mn content was found to be in the late stage (13.84 ± 0.37 mg/100 g), and the minimum Mn content was found to be in the early stage (5.56 ± 0.48 mg/100 g). The moringa leaves are also rich in minerals such as potassium, zinc, magnesium, iron, and copper (Kasolo et al., 2010).