Methyl cellosolve

The assay used was modified according to the method described by Li et al. [53 (link)]. 4-MU oleate was used as substrate for pancreatic lipase activity. Briefly, 50 μL of lipase (50 U/mL) dissolved in phosphate buffer (200 mM, pH 7.4) were added to 50 μL of different concentrations of extracts and standard (Orlistat) in a 24-well microtiter plate and allowed to react for 10 min. After that, 100 μL of 1 mM 4-MU solution (dissolved in methyl cellosolve) was added and incubated at 25 °C for 30 min. The reaction was stopped by adding 100 μL sodium citrate solution (0.1 M, pH 4.2). The amount of 4-methylumbelliferone released was measured using a fluorescence reader at an excitation wavelength of 355 nm and an emission wavelength of 460 nm. Experiments were done in triplicates. Orlistat was used as a positive control. Percentage lipase inhibition was calculated as shown below: $\mathrm{Lipase}\mathrm{Inhibition}\mathrm{Rate}(\%)=1-\left(\frac{F_{\mathit{test}}-F_{\mathit{test}\mathit{blank}}}{F_{\mathit{control}}-F_{\mathit{control}\mathit{blank}}}\right)\times 100$
where F_test and F_{test blank} are the fluorescent values of test samples with and without the substrate 4-MU oleate, respectively. F_control and F_{control blank} are the fluorescent values of control with and without the substrate 4-MU oleate, respectively. The results were expressed in terms of IC₅₀ representing the concentration of test extracts required to cause the enzyme inhibition by 50%.

Two separate experiments were conducted to investigate the physiological mechanisms of improved germination performance and stress tolerance after osmopriming sorghum seed with PEG. Six primed or unprimed sorghum seeds were planted in each pot. Pots were fertilized immediately after planting by spraying 60 ml Hoagland solution [18 ]. The aforementioned SMTs were applied immediately after planting. Plants were thinned to three per pot after emergence. Each SMT comprised three replicate pots. Each pot represented an experimental unit, and each of the three seedlings in a pot is a sampling unit. There were a total of 120 pots arranged in a randomized block design. Relative water content (RWC), chlorophyll, root viability, antioxidant system, lipid peroxidation, O²⁻ content, plasma membrane stability, and osmotic adjustment were determined at 12 and 24 DAP.
Relative water content (RWC) of leaves and roots of sorghum were determined using a formula of RWC (%) = [(FW–DW)/(TW–DW)] × 100, where FW, DW, and TW are fresh weight, dry weight, and turgid weight, respectively. DW was determined when sample weight stabilized in an oven at 65°C. TW was measured 24 h after the saturation of plant samples in deionized water at 4°C [19 ].
Chlorophyll a and b content was determined with the procedure as described by Arnon [20 (link)]. Fresh leaves were cut into 0.5 cm fragments and extracted for 24 h using 80% acetone at -10°C. The resulting extract was centrifuged and the absorbance of the supernatant was measured at 645 and 663 nm using a spectrophotometer (UV-2401, Shimadzu Corporation, Japan).
Root viability was determined by measuring the activity of dehydrogenase using the 2,3,5-triphenyl tetrazolium chloride (TTC) reduction method. Fresh root material (0.2 g) was sampled from the root base, middle root, and root tip. Root material was then cleaned with distilled water and incubated in a 10 ml solvent mixture containing 5 ml 0.4% v/v TTC and 5 ml 0.06 mol∙L^-1 phosphate buffer (pH 7.0) in darkness at 37°C for 3 h. The reaction was terminated by adding 2 ml of 1 mol L^-1 sulfuric acid in the tubes. Samples were centrifuged and absorbance of supernatant was measured at 485 nm using a spectrophotometer. Root viability was expressed as the quantity of TTC reduced per gram of root dry mass per h (μg g^-1 h^-1).
APX in plant leaves was determined according the procedure described by Nakano and Asada[21 ] by measuring the decrease in absorbance of the oxidized ascorbate at 290 nm. A 1 ml reaction mixture contained 50 mM potassium phosphate buffer (pH 7), 10 μl enzyme extracts, 0.1 mM H₂O₂, and 0.5 mM ascorbate was used. The reaction was initiated by adding H₂O₂.
SOD activity was determined following the method of Giannopolitis and Ries [22 (link)], with minor modifications. Fresh leaf material (0.2 g) was used for measuring SOD activity. A 3 ml reaction solution containing 50 μM nitroblue tetrazolium (NBT), 1.3 μM riboflavin, 13 mM methionine, 75 nM EDTA, 50 mM phosphate buffer (pH 7.8), and 30 μl of enzyme extract. The test tubes were irradiated under 15 fluorescent lamps at 78 μmol m^-1 s^-1 for 15 min. The absorbance of the irradiated solution was 560 nm with a spectrophotometer. The amount of enzyme required to cause 50% photoreduction of NBT was regarded as one unit of SOD activity.
CAT and POD activities in plant leaves were determined using the method developed by Bradford [23 (link)], with slight modifications. A 3 ml CAT reaction solution containing 100 μl enzyme extract, 5.9 mM H₂O₂, and 50 mM phosphate buffer (pH 7.0) was used. The biochemical reaction was initiated by adding the enzyme extract. Changes in absorbance of the reaction solution at 240 nm were read for every 20 s to determine CAT activity. Similarly, a 3 ml POD reaction solution contained 20 mM guaiacol, 50 mM phosphate buffer (pH 5), and 40 mM H₂O₂. Changes in absorbance were read at 470 nm for every 20 s to determine POD activity. One unit of CAT or POD activity was defined as an absorbance change of 0.01 units per min.
Malondialdehyde (MDA) content was measured according to a modification of the method used by Noreen et al.[24 ]. Fresh leaf (1.0 g) was homogenized in 3 ml 1.0 w/v trichloroacetic acid (TCA) at 4°C and centrifuged at 12000 g for 10 min. A 0.5 ml of supernatant was transferred to 3 ml 0.5 v/v thiobarbituric acid (TBA) in 20% TCA. The resulting mixture was incubated in boiling water for 50 min. After cooling in an ice water bath, the mixture was centrifuged at 12000 g for 15 min. The absorbance of supernatant was read at 532 and 600 nm with a spectrophotometer.
The determination of O²⁻ content was conducted by using a modification of the procedure described by Doke [25 ]. The O²⁻ content was determined based on its ability to reduce NBT. Fresh leaf tissues (0.5 g) were excised and immersed in 10 mM potassium phosphate buffer (pH 7.8), containing 0.05% nitro blue tetrazolium and 10 mM NaN₃. The sample was incubated for 1 h at room temperature. Following incubation, 2 ml of this reaction solution was heated at 85°C for 15 min and cooled in an ice bath. Optical density of solution was determined at 560 nm for 15 min using a spectrophotometer. The O²⁻ content was expressed as the increase in absorbance per unit dry weight.
Relative electrolyte leakage was measured to determine the membrane permeability, according to the method by Blum and Ebercon [26 ]. Sorghum leaf materials were sampled and excised to 5 mm segments. Leaf tissues were rinsed with distilled water and immersed in a test tube containing 6 ml distilled water for 12 h at 18°C. The relative electrolyte leakage of solution was measured (E₁) using a conductivity meter (Model DDS, Shanghai Leici Instrument Inc., Shanghai, China). Samples were subsequently autoclaved for 10 min at 120°C. After cooling to 25°C, the second relative electrolyte leakage was measured (E₂). The conductivity of deionized water was also measured (E₀). The relative electrolyte leakage was determined with the formula of electrolyte leakage (%) = (E₁ –E₀)/(E₂ –E₀) × 100.
The free amino acid pool in plant leaves was determined according to a minor modification of the procedure used by Moore and Stein[27 (link)]. Plant leaf material (0.5 g) was sampled and homogenized with 10 ml of 80% boiling ethanol. The homogenate was centrifuged at 5000 g for 10 min. This extraction was repeated four times and the supernatants were combined and transferred to new tubes. The ethanol extract was evaporated in a fume hood and the residue was dissolved in 5 ml 0.2 M citrate buffer (pH 5.0). A 2 ml aliquot of the sample was mixed with 1 ml of ninhydrin reagent in methyl cellosolve and 0.2 M acetate buffer. The samples were boiled for 20 min and cooled at room temperature. Absorbance was then read at 570 nm with a spectrophotometer.
Proline content in leaf samples was determined following the method of Bates et al. [28 ]. Fresh leaf material (0.5 g) was homogenized in 10 ml 3% sulphosalicylic acid and centrifuged at 1200 g for 10 min. A 2 ml supernatant was mixed with 2 ml acid ninhydrin reagent and 2 ml glacial acetic acid. The sample was subsequently incubated at 100°C for 60 min. The sample materials were cooled in an ice bath prior to adding 4 ml toluene to each sample. The toluene layer was read at 520 nm with a spectrophotometer.
Soluble and reducing sugars in plant leaves were determined following a modification of the methods used by Dubois et al.[29 ] and Van Handel [30 (link)], with minor modifications. Plant leaves (0.25 g) were placed in a boiling water bath for 1 h. Total soluble sugar content was subsequently analyzed with the phenol-sulfuric method after hydrolysis of starch using perchloric acid. Sucrose content was determined using the anthrone method. Reducing sugar content was calculated as the difference between total soluble sugar and sucrose.

The pancreatic lipase inhibition assay was conducted following the protocol described in the literature [7 (link)] using 50 µL (1 mg/mL in 70% ethano) of the extracts mixed with 50 µL (50 U/mL) of lipase enzyme in methyl cellosolve at room temperature. The mixture was preincubated for 10 min at room temperature before adding 100 µL of 1 mM 4-Methylumbelliferyl (4-MU) in methyl cellosolve and then further incubated for 30 min at room temperature. Finally, 100 µL of 0.1 M, pH 4.2 of sodium citrate solution was used to stop the reaction. The fluorescence of the tested samples was measured using a microplate reader at wavelengths 355 nm and 460 nm. The experiment was performed in triplicates and the percentage lipase inhibition capacity was calculated as follows: $$\mathrm{Lipase} \mathrm{Inhibition} (\%)=[1-\left(\frac{Ftest-Ftest blank}{ Fcontrol-Fcontrol Blank}\right)]\times 100,$$
where Ftest and Ftest blank represent the fluorescent readings for the test samples with and without the substrate 4-Methylumbelliferyl (4-MU) oleate, respectively, while Fcontrol and Fcontrol blank were the fluorescent readings of control with and without the substrate 4-MU oleate, respectively.

The pancreatic lipase inhibition assay was conducted following the protocol described in the literature [7 (link)] using 50 µL (1 mg/mL in 70% ethano) of the extracts mixed with 50 µL (50 U/mL) of lipase enzyme in methyl cellosolve at room temperature. The mixture was preincubated for 10 min at room temperature before adding 100 µL of 1 mM 4-Methylumbelliferyl (4-MU) in methyl cellosolve and then further incubated for 30 min at room temperature. Finally, 100 µL of 0.1 M, pH 4.2 of sodium citrate solution was used to stop the reaction. The fluorescence of the tested samples was measured using a microplate reader at wavelengths 355 nm and 460 nm. The experiment was performed in triplicates and the percentage lipase inhibition capacity was calculated as follows: $$\mathrm{Lipase} \mathrm{Inhibition} (\%)=[1-\left(\frac{Ftest-Ftest blank}{ Fcontrol-Fcontrol Blank}\right)]\times 100,$$
where Ftest and Ftest blank represent the fluorescent readings for the test samples with and without the substrate 4-Methylumbelliferyl (4-MU) oleate, respectively, while Fcontrol and Fcontrol blank were the fluorescent readings of control with and without the substrate 4-MU oleate, respectively.