Nutritional value, antioxidant and antidiabetic properties of nettles (Laportea alatipes and Obetia tenax)

Nettles are commonly consumed in South Africa, Europe and Asia and are used in traditional medicine to treat a variety of ailments. In this study, the nutritional value of the leaves of nettles (Laportea alatipes and Obetia tenax) was evaluated and compared, when cooked and uncooked. The results showed a decrease in the concentrations of crude protein, vitamin A, vitamin E and metals after cooking of nettles. Although cooking reduced the concentrations of essential elements in nettles, their contribution to the diet remained adequate. L. alatipes presented with reduced levels of Cd (from 1.86 to 0.810 mg kg−1) and Pb (from 2.87 to 1.88 mg kg−1) after cooking. Similarly, Cd (from 2.97 to 0.780 mg kg−1) and Pb (from 2.21 to 0.795 mg kg−1) levels in O. tenax decreased after cooking, demonstrating the significance of cooking. The antioxidant activity of the nettles was determined using the 2,2-diphenyl-l-picrylhydrazyl (DPPH) free radical and ferric reducing antioxidant power (FRAP) assays. The methanol extract of Obetia tenax showed high ferric reducing power whilst the radical scavenging activity was due to the presence of the bioactive molecule, β-carotene, in the plants which exhibited higher DPPH radical scavenging ability relative to test samples and standards. The in vitro antidiabetic activity of the extracts and compounds from the nettles was better than or comparable to that of the known standard, acarbose, which underscores the prospective antidiabetic properties of nettles. Overall, our study provides scientific validation for the ethno-medicinal use of nettles and supports their consumption, which highlights their potential as nutraceuticals.


Nutrient analysis
All solvents and reagents were of analytical grade and were purchased from Merck (Darmstadt, Germany) and Sigma Aldrich (St Louis, USA). Nutrient analysis was performed as described in our previous study 1 . The moisture and ash content were established by well-known methods 2,3 . Moisture content was determined by drying fresh sample (approximately 1.00 g weighed in a crucible) in the oven at 105 °C, to constant mass. Thereafter, the crucible with the dried sample was removed from the oven and placed in the desiccator to cool. The percentage moisture was calculated by the following equation: where W1 is the weight of the crucible, W2 is the weight of the crucible with the fresh sample and W3 is the weight of the crucible with the dried sample. 3 The ash content was determined by igniting the dried sample in a muffle furnace at 600 ℃ for 12 h. Fat content was determined by the soxhlet method using n-hexane. Briefly, approximately 5 g of dried sample was placed into an extraction thimble and 100 mL of n-hexane was poured into a 200 mL round bottom flask. The soxhlet apparatus was then assembled on a heating mantle set at 60 °C and the extraction process was continued for 6 h. The solvent was removed using a rotary evaporator. The resulting oil was quantified gravimetrically and the percentage oil was calculated using the following equation: where M1 is the weight of the empty vessel, M2 is the weight of the vessel with the fat residue and E is the sample weight.
Protein content was determined by Kjeldahl distillation method, multiplying nitrogen by 6.25 4 . Total carbohydrate was obtained by difference (subtracting the protein, ash, and fat content from the total dry mass of the sample). The vitamin C content was determined by the iodometric method 5 . Dried samples (approximately 20 g) were placed in a mortar and 20 mL doubly distilled water was added. The mixture was stirred for 20 min and filtered by suction using a Buchner funnel. The filtrate was placed into a 50 mL volumetric flask and made up to the mark with doubly distilled water. A 25 mL aliquot of the leaf solution was placed into a conical flask then 5 mL of 0.15 M potassium iodide, 0.1 M hydrochloric acid and 1 mL of 1% starch solution indicator was added. This was titrated against a potassium iodate solution (0.009 M). The endpoint was observed by the permanent blue-black color due to the starch-iodine complex.
The vitamin A content (β-carotene) was determined using previously described methods 6 .
Briefly, approximately 0.5 g of dried sample was mixed with 4 mL cold ethanol and extracted with 8 mL hexane followed by vortex mixing for 2 min then centrifuged at 1000 rpm for 15 4 min. β-carotene was determined spectrophotometrically (Biochrom Libra S11, Cambridge, England) at 450 nm.
Vitamin E content was determined by previously described methods 7  at 536 nm using a UV spectrophotometer (Biochrom Libra S11, Cambridge, England).
Absolute ethanol served as a blank. A calibration curve of α-tocopherol (prepared in absolute ethanol) at various concentrations was used to determine the vitamin E content in the leaves.

Phytochemical analysis
Leaves of L. alatipes (196 g), and O. tenax (292 g) were air-dried, ground and extracted with hexane, dichloromethane (DCM), and methanol (MeOH), in turn, by continuous shaking on an orbital shaker for 48 h. The aqueous MeOH extract was partitioned with DCM followed by ethyl acetate (EtOAc). All mixtures were filtered thereafter the filtrate (extract) was concentrated to dryness and stored in the fridge until analyzed. The extracts were spotted on TLC plates (Merck silica gel 60, 20 x 20 cm F254 aluminum sheets) and were subjected to column chromatography using Merck Kieselgel 60 silica gel (0.063-0.200 mm, 70-230 mesh ASTM). 5 The hexane and DCM extract of L. alatipes leaves were combined due to similar TLC profiles.
The combined extract (12.23 g) was loaded onto the column and separated using a hexane: EtOAc step gradient starting from 100% hexane that was stepped by 5% to 100% EtOAc.

Elemental analysis
All glassware and polyethylene bottles were prewashed with 10% HNO3 and repeatedly rinsed with doubly distilled water. Digestion of the plant samples followed the same procedure as described in our previous study 1 where AC is the absorbance of the control (DPPH without the sample) and AS is the absorbance of the sample. 8

Ferric reducing power
The reduction of Fe 3+ to Fe 2+ by L. alatipes, O. tenax and compounds was measured by the previously described FRAP method 9 . In brief, 1 mL of extract or compound of various concentrations (10 -500 μg mL -1 ) was mixed with 2.5 mL of 0.2 M phosphate buffer (pH 6.6) and 2.5 mL of 1% (w/v) potassium ferricyanide followed by incubation at 50 °C for 20 min.
After incubation, 2.5 mL of 10% trichloroacetic acid (TCA) was added into the mixture followed by vortex mixing for 5 min. The mixture was centrifuged at 5000 rpm for 10 min.
About 2.5 mL of the upper layer was mixed with 2.5 mL of doubly distilled water and 0.5 mL of 0.1% ferric chloride then left to stand for 10 min. The absorbance of the samples and blank was measured at 700 nm using a UV spectrophotometer (Biochrom Libra S11, Cambridge, England). A mixture of reagents and MeOH without the sample served as a blank. Ascorbic acid and α-tocopherol were used as positive controls. Experiments were conducted in triplicate.

Alpha amylase
The α-amylase inhibition assay was established by a method previously described with some modifications 10 . Plant extracts or isolated compounds at various concentrations (200 μL) were added to sodium phosphate buffer (20 mM, pH 6.9, 250 μL) containing NaCl (0.006 M) and porcine pancreatic amylase (0.5 mg mL -1 , 250 μL) and incubated at 25 °C for 10 min. To the reaction mixture 500 μL of 1% starch solution (in 20 mM sodium phosphate buffer with 0.006 M NaCl, pH 6.9) was added and further incubated at 25 °C for 10 min. To terminate the reaction 1mL of 3,5-dinitrosalicylic acid (DNS) was added and the mixture was placed in a boiling water bath for 10 min then cooled to room temperature. The mixture was then diluted with 10 mL distilled water and the absorbance was measured at 540 nm using a UV spectrophotometer (Biochrom Libra S11, Cambridge, England). A solution containing the reagents and enzyme 9 without the sample was used as the control. Acarbose was used as a positive control. All experiments were conducted in triplicate. The IC50 values were estimated from the doseresponse curves.

Alpha glucosidase
The α-glucosidase inhibition assay was established by a method previously described with some modifications 10 . Experiments were conducted in triplicate. Plant extracts or compounds at various concentrations (200 μL) were added to 100 μL of 0.5 mg mL -1 of α-glucosidase (in 0.1 M phosphate buffer, pH 6.9) and incubated at 25 °C for 10 min. Thereafter, 50 μL of 5 mM p-nitrophenyl-α-D-glucopyranoside (in 0.1 M phosphate buffer, pH 6.9) was added and incubated further at 25 °C for 5 min. The absorbance was measured at 405 nm using a UV spectrophotometer (Biochrom Libra S11, Cambridge, England). A solution containing the reagents and enzyme without the sample was used as the control. Acarbose was used as a positive control. The IC50 values were estimated from the dose-response curves.
The percentage inhibition was calculated as follows: where AC is the absorbance of the control and AS is the absorbance of the sample.

Statistical analysis
Experimental data are expressed as mean (standard deviation). Statistical analyses were performed using the Statistical Package for the Social Sciences (PASW Statistics, Version 25, IBM Corporation, Cornell, New York). Significant differences between means were established by Tukey's Post-hoc tests (p < 0.05). Statistical analyses for antioxidant and antidiabetic activity were done using GraphPad Prism 5.0 (GraphPad Software Inc., San Diego, CA). One-way Anova was performed on the data followed by Dunnett's test (p < 0.05).