Safflower seeds (cultivars TSF1 and SM) procured from Tanduru ARS (Rangareddy District India) were sown in earthen pots containing red soil and farmyard manure (3:1). The pots were kept under natural photoperiod in the botanic garden, treated with water once a day and maintained for 19 days. Nineteen day-old plants were subjected to salt stress by adding 0 (control), 1.0%, 1.5% and 2.0% NaCl solutions to the potted plants. For each treatment, twenty pots, each with three plants were maintained; control and treated pots were maintained daily with tap water. Care was taken to avoid leaching out of soluteion by adding water slightly less than field capacity. After 12 days of stress imposition, the leaves and roots were separated and stored in liquid nitrogen at −80˚C until further use.

The plants were carefully uprooted from pots and washed immediately with running tap water. Plant growth was determined by measuring the length of the root and shoot. The dry weight (DW) was measured after drying the uprooted plants at 80˚C for 48 hrs.

The leaf samples were washed with deionised water, blotted dry and fresh weights (FW) recorded. The leaves were immersed in distilled water for 4hrs, blotted dry and the turgid weight (TW) was taken. The leaves were kept at 80˚C in a hot air oven for 48 hrs and dry weights (DW) were recorded. RWC was calculated according to Turner, [21] using the formula RWC = FW − DW/TW − DW × 100.

The reducing sugars were estimated by Nelson’s method [22] , as modified by Somogyi [23] . 1 ml of alcoholic extract was taken into a test tube. To this 1.0 ml of a freshly prepared mixture of 25 parts of reagent “A” and 1 part of reagent “B” (25:1) was added and mixed with the solutions. The test tubes were incubated in a boiling water bath for 20 min and cooled under a running tap water. 1 ml of arsenomolybdate reagent was added. The colour developed very rapidly. The mixture was diluted to 10.0 ml after 15 min and the absorbance was measured in a UV-Visible Spectrophotometer (Shimadzu-1601) at 500 nm. Reagent blanks were used to adjust absorbance to zero. Reducing sugar content was estimated by using a standard curve prepared with dextrose.

Free proline content of control and NaCl treated samples was determined by following Bates [24] method. Fresh plant material (0.5 g) was homogenized in a pre-chilled mortar with a pestle using 3% aqueous sulfosalicylic acid. The homogenate was filtered through four layered muslin cloth and the filtrate was collected. The extraction was repeated twice, all the filtrates were pooled and made up to known volume. 2 ml of the filtrate was taken into a test tube and 2 ml of acid ninhydrin, 2 ml of glacial acetic acid were added. The tubes were incubated at 100˚C for 1 h in boiling water bath. After incubation, they were transferred to an ice bath to terminate the reaction. 4 ml of toluene was added to the contents of the tubes and mixed thoroughly using a cyclomixer (CM101, REMI India) for 15 sec. Chromophore containing toluene was aspirated from the aqueous phase. Then the absorbance was measured in a UV-spectrophotometer at 520 nm against toluene. Proline was measured from the standard curve prepared with authentic proline and its amount was calculated on dry weight basis.

The measurement of total AsA was carried out as described by Cakmak and Marschner [25] . Fresh leaf samples (500 mg) were extracted with 5 ml of 5% metaphosphoric acid and centrifuged at 15,000 × g for 15 min. The reaction mixture contained 0.2 ml aliquot, 0.5 ml 150 mM phosphate buffer with pH 7.4 having 5 mM EDTA, 0.1 ml 10 mM DTT and 0.1 ml 0.5% (w/v) N-ethylmaleimide to remove excess DTT. 10% TCA, 44% orthophosphoric acid, 2.1 bipyridine in 70% ethanol (0.4 ml each) and 0.2 ml 3% FeCl3 were added to that mixture for the development of colour. The mixtures were then incubated in a water bath at 40˚C for 40 min and the colour developed was read at 525 nm. AsA was used as a standard and the concentration of AsA was expressed as µmol∙g⁻¹ dry weight.

QACs were extracted and estimated according to Grieve and Grattan, [26] . Dried finely ground plant material (0.5 gm) was mechanically shaken with 20 ml of deionized water for 24 hrs at 25˚C. The samples were then filtered and the filtrates were diluted (1:1) with 2N H2SO4. Aliquots (0.5 ml) were taken into centrifuge tubes and cooled in ice water for 1 hr. Cold KI-I2 reagent (0.2 ml) was added and then reactants were gently stirred. The tubes were stored at 4˚C for 16 hrs and then centrifuged at 10,000 rpm for 15 min at 0˚C. The supernatant was carefully aspirated with a fine tipped glass tube. The periodide crystals were dissolved in 9.0 ml of 1, 2-dichloroe- thane and mixed vigorously. After 2 hrs the absorbance was measured at 365 nm by using a spectrophotometer. Reference standards of glycine betaine (50 - 200 mg/ml) were prepared in 1 N H₂SO₄. The amount of glycine betaine was calculated using following formula:

Glycine betaine content = (A525 − 0.0121)/0.035 × 1.5 × 25/0.5.

In situ accumulation of and H₂O₂ was detected by histochemical staining with NBT and DAB according to Romero-Puertas [27] , with minor modifications. For detection the leaves of control and salt-stressed plants were excised and immersed in a 0.1% solution of NBT in 10 mM phosphate buffer (pH 7.8). The leaves immersed were illuminated for 1 - 2 h until the appearance of dark spots, specific to blue formazan precipitates. For localization of H₂O₂ excised leaves were immersed in 10 mM phosphate buffer (pH 3.8) solution containing 0.1% DAB and then incubated at room temperature for 8 h until brown spots, appear by the reaction of DAB with H₂O₂. The leaves were then bleached in warm ethanol to visualize the blue and brown spots.

Superoxide anion content was determined using the method described by Doke [28] . The leaves were cut into pieces and placed in the test tubes filled with 7 ml of a mixture containing 50 mM of NaN₃ and incubated in dark for 5 min. Then 2 ml of the solution was taken from the tubes and heated for 15 min at 85˚C. The samples were cooled on ice for 5 min and the absorbance was measured at 580 nm.

The estimation was done according to Singh [29] . Fresh leaves (0.1 g) were extracted in ice bath with 5 ml 0.1% (w/v) TCA. The homogenate was centrifuged at 12,000 × g for 15 min and the supernatant (0.5 ml) was added to 0.5 ml of 10 mM potassium phosphate buffer (pH 7.0) and 1 ml of 1 M KI. H₂O₂ concentrations of the samples were calculated and expressed in µmol∙g⁻¹ dry weight by using H₂O₂ as a standard.

MDA the product of Lipid peroxidation was determined by estimating thiobarbituric acid reactive substances (TBARS) as described by Hodges et al. [30] . 500 mg of leaf tissue was ground to a fine paste with 15 ml of ethanol: water (95:5 v/v and 0.1% butylated hydroxyl toluene) the supernatant was collected after centrifugation 3000 × g for 10 min. Later the supernatant (1 ml) was added to 20% TCA (1 ml) along with 0.65% thiobarbituric acid (TBA) in a clean glass tube and mixed vigorously. Another 1 ml of supernatant was treated with 1 ml of 20% TCA and mixed vigorously. The solution was heated at 95˚C for 30 min in a water bath, cooled immediately by ice then centrifuged at 3000 × g for 10 min. Absorbance at 532 nm was recorded for MDA. In addition, the absorbance at 440 nm (carbohydrates) and 600 nm (phenylpropanoid pigments) was also recorded to avoid overestimation of MDA.

[(Abs532 + TBA) − (Abs600 + TBA) − (Abs532 − TBA) − (Abs600 − TBA)] = A.

[(Abs440 + TBA − Abs600 + TBA) 0.0571] = B.

[MDA equivalents (μmol∙ml⁻¹) = (A − B/157000)103] = C

MDA equivalents (μmol∙g⁻¹) = (C × 15 × 1/0.5).

Cell membrane stability measured according to the method of Premchandra et al., [31] . 30 leaf discs of 0.75 cm diameter were cut from healthy plants of both cultivars using a leaf punch and were placed into a 100 ml conical flask and washed thrice with deionized distilled water. Leaf discs were submerged in 30 ml of 1%, 1.5% and 2% NaCl solutions for 24 hrs at 100˚C. Discs submerged in distilled water served as controls. Both control and salt induced discs were dipped in 30 ml of deionized distilled water for 24 hrs at 100˚C, the flasks were warmed to 25˚C shaken thoroughly and electrical conductivity was measured using an electrical conductivity meter (C₁ and T₁). Following the conductivity measurements, the leaf tissues were killed by autoclaving for 15 min, at 15 lbs/cm³, cooled to 25˚C and electrical conductivity was measured (C₂ and T₂). Cell membrane stability of the leaf tissues was calculated as per the membrane percentage injury from the following equation:

Percentage membrane injury = 1 − (1 − T₁/T₂)/(1 − C₁/C₂) × 100.

Fresh leaf tissue was homogenized in 50 mM Tris-HCl (pH 7.5) buffer containing 40 mM phenyl methyl sulfonyl fluoride (PMSF), and 2% (w/v) polyvinyl poly pyrrolidone (PVPP) with the addition of 2 mM AsA for the APX assay. The extract was centrifuged at 15,000 × g for 20 min at 4˚C and the resultant supernatant was used for all the enzyme assays. The amount of protein was calculated according to Lowry et al. [32] . All enzymatic activities were measured spectrophotometrically at 25˚C.

CAT (EC 1.11.1.6) activity was measured according to Beers and Sizer [33] , with minor modifications. The decrease of H₂O₂ was monitored by reading the absorbance at 240 nm at the moment of H₂O₂ addition and 1 minute later. The difference in absorbance (DA240) was divided by the H₂O₂ molar extinction coefficient (36 M⁻¹∙cm⁻¹) and the enzyme activity expressed as mmol of H₂O₂ min⁻¹∙mg⁻¹ protein.

APX (EC 1.11.1.1) activity was determined as described by the method of Nakano and Asada [34] . The oxidation of As A in the reaction mixture was measured using the rate of decrease in absorbance at 290 nm and was calculated using an extinction coefficient of 2.8 m M⁻¹∙cm⁻¹.

GPX (EC 1.11.1.7) Guaiacol peroxidase (GPX) activity was determined in terms of oxidation of guaiacol by H₂O₂ [35] as measured by the increase in absorbance at 420 nm (E = 26.6 mM⁻¹∙cm⁻¹).

SOD (EC1.15.1.1) activity was assayed using the method described by Urbanek et al., [36] . One unit of SOD activity was defined as the amount of enzyme required to cause 50% inhibition of the NBT photo reduction rate.

Data was analyzed by using the SPSS (Statistical Package for the Social Sciences) version 16.0. Data presented here are mean values and standard deviation (±SD). One-way ANOVA was carried out using Post hoc multiple comparisons from Duncan’s test at a significance level of p < 0.05.

Root and shoot growth of two cultivars differed greatly to varied levels of salinity Figure 1(a). The percent decrease in root and shoot growth were 28.40 and 33.51 at 2% NaCl level in cultivar TSF1 and cultivar SM, respectively. At 1.5% NaCl stress, a slight decrease was noticed in both cultivars when compared to control. Reduction in shoot growth was also in proportionate to NaCl stress levels. Comparatively, cultivar TSF1 maintained better root and shoot growth over cultivar SM even at high NaCl stress level.

NaCl salinity levels restricted the biomass of both cultivars Figure 1(b). The reduction in biomass was more significant in cultivar SM than cultivar TSF1 in root and shoot at high salt stress levels, but insignificant reduction was observed in both cultivars at moderate NaCl stress.

Salinity effects relative water content of leaf in two cultivars; but the effect was more pronounced at high salinity. There was a gradual decline in leaf water content Figure 1(c). The percent decrease were 13, 15 and 20.4 in cultivar TSF1 ;while they were 15, 34 and 42 in cultivar SM at 1%, 1.5%, and 2% NaCl respectively, when compared with the control. This implies the cultivar TSF1 maintained more tissue water content even at high salinity level than SM.

The increase in sugars content was observed in root and leaf tissues in both cultivars at all stress levels. The percent increase was relatively more in cultivar TSF1 than in cultivar SM Figure 1(d). The degree of increase in soluble sugar content was dependent on the severity of stress in both cultivars, but with greater accumulation of soluble sugar content in cultivar TSF1 than in cultivar SM.

There was a proportionate increase of free proline content with stress intensity was ob-served in both cultivars. The percent increase was 72.7 and 51.1 in cultivar TSF1 and in cultivar SM at high NaCl stress. Maintenance of better proline accumulation supports the tolerance potential of cultivar TSF1 Figure 2(a).

Salinity affected AsA content at all regimes of NaCl stress but significantly in cultivar TSF1 than cultivar SM at 2% NaCl stress Figure 2(b).

Accumulation of glycine betaine gradually increased with increasing stress level and the percent increase was more in cultivar TSF1 than in cultivar SM Figure 2(c).

Salinity induced and H₂O₂ content was observed in both cultivars at all levels of

NaCl stress. However, a smaller increase in and H₂O₂ content was observed in cultivar TSF1at high salinity level conferring its efficient defensive mechanism than in cultivar SM (Figure 3(a), Figure 3(b)).

Malondialdehyde content Figure 3(c) of both cultivars was elevated due to salinity and the magnitude of elevation was concentration dependent in both the cultivars. However, the percent increase in MDA content was 31 and 67 in cultivar TSF1and in cultivar SM, respectively.

The data was shown in terms of percent membrane injury. The per cent membrane in jury was relatively less in cultivar TSF1 than in cultivar SM indicating better membrane

stability in that cultivar TSF1 Figure 3(d).

CAT activity was more in cultivar TSF1 than in cultivar SM at higher salinity gradient as depicted in Figure 4(a). Similarly, APX and GPX activities also increased with the gradual increase of stress intensity and this was more pronounced in cultivar TSF1 than in cultivar SM (Figure 4(b), Figure 4(c)). Salinity caused a significant increase in the total SOD enzyme activity with increased salinity stress in both cultivars, however percent increase was more in cultivar TSF1 than in cultivar SM (Figure 4(d), Figure 5).