Under salt stress, the morphological characters and physiological indexes of sensitive cultivars are more obvious than tolerant cultivars. When the sensitive plants were treated with H₂S, the ameliorative effects of H₂S will be easy to judge. Therefore, the sensitive cucumber (C. sativus L.) cultivar “Chunxiaqiuwang” was used in our experiments. Uniform seeds were sterilized in 10 % (v/v) sodium hypochlorite solution for 5 min followed by washing three times with distilled water. Sterilized seeds were germinated on moist filter paper in petri dishes at 25°C in the dark. After germination, seedlings were transplanted into perlite-filled plastic pots (240 mL, 65 × 45 × 70 mm, a seedling per pot) and watered with Hoagland’s solution. 10 mL Hoagland’s solution was applied for every other day after germination, which contained 5 mM KNO₃, 5 mM Ca(NO₃)₂, 1 mM NH₄H₂PO₄, 2 mM MgSO₄, 10 μM MnSO₄, 50 μM H₃BO₃, 0.7 μM ZnSO₄, 0.2 μM CuSO₄, 0.01 μM (NH₄)₆Mo₇O₂₄ and 70 μM Fe-EDTA-Na₂. The seedlings of cucumber were maintained in a controlled growth chamber with a light/dark regime of 14/10 h, relative humidity of 70%, temperature of 25°C and a PAR of 800 μmol.m^-2.s^-1. When the first true leaf emerged, uniform and healthy seedlings were selected and divided into three groups for the following treatments. (i) Hoagland’s nutrient solution (as Control); (ii) Hoagland’s nutrient solution + 200 mM NaCl (as NaCl); (iii) Hoagland’s nutrient solution + 200 mM NaCl + 5 or 10, or 15 or 20 μM NaHS (as NaCl + NaHS). In our preliminary experiment, the phenotypic characteristics of plants with treated by H₂S alone were no significant difference compared with the control group (data not shown). Thus there is no only H₂S treatment group in our study. NaHS is commonly used as an H₂S donor since it dissociates to water produce HS^- and Na^+, and then combination of HS^- with H⁺ produces H₂S (Lin et al., 2012). NaHS is responsible for induction of stress tolerance, but not Na₂S, Na₂SO₄, NaHSO₄, Na₂SO₃, NaHSO_3, or CH₃COONa (Deng et al., 2016). Each treatment was replicated three times under the same experimental conditions. The NaHS was purchased from Sigma (St Louis, MO, United States) and used as H₂S donor. The remaining treatment solution, which was not fully absorbed by plants, was removed and 10 mL fresh treatment solution was irrigated every other day at 9:00 AM during the whole culture process. After 7 days of treatment, the parameters of photosynthesis and chlorophyll fluorescence were determine using the first true leaf of seedlings with different treatments. And finally, at least 24 seedlings per treatment group were harvested and weighed separately to determine various physiological and biochemical parameters, while other samples were immediately frozen in liquid N₂ and then stored in a -80°C freezer until use.

Root length, seedling length (including aboveground and underground parts) and leaf area index were determined after 7 days of treatment. Leaf area index was measured using graph paper according to the method of Gupta et al. (2017). Plant roots were gently removed from the perlite, and then washed three times with deionized water to remove adhered perlite particles. After absorbing moisture from the root surface with a clean filter paper, the fresh weights of root and shoot were recorded, respectively. Shoot dry weight and root dry weight were, respectively, recorded after drying the same plants in an oven at 60 ° C until the weight became constant. Survival rate (%) was calculated as the number of survival seedlings / number of total seedlings. The rate of second leaf expansion (%) was calculated as the number of seedlings with appearance of the second leaf / number of total seedlings.

The first true leaf was chosen to determine the parameters of photosynthesis, chlorophyll fluorescence and chlorophyll Content. And the P_n and T_r were measured using an LI-6400 portable photosynthesis system (LI-COR, Lincoln, NE, United States). The leaf chlorophyll fluorescence parameters were measured using a portable chlorophyll fluorometer (OS-30p+, Opti-Science, Inc., Tyngsboro, MA, United States). The F_o emission was determined on dark-adapted leaves. The F_m was obtained during a subsequent saturating light pulse. A second saturating pulse of white light was imposed to determine the F_m′. The F_o′ was determined by illuminating the leaf with far-red light for 3 s. The fluorescence parameters were calculated according to Hu and Yu (2014): maximal quantum yield of PSII photochemistry, F_v/F_m = (F_m-F_o)/F_m; effective quantum-use efficiency of PSII in light-adapted state, F_v′/F_m′ = (F_m′-F_o′)/F_m′; quantum yield of PSII photochemistry, Φ_PSII = (F_m′-F_t)/F_m′; qP = (F_m′-F_t)/(F_m′-F_o′); and NPQ = (F_m-F_m′)/F_m′. The contents of Chlorophyll a (C_a), chlorophyll b (C_b), and C_x+_c were measured on fresh leaves as described by Lichtenthaler and Wellburn (1983).

The first true leaves were selected to measure the stomatal opening by scanning electron microscopy (S-3400N; Hitachi, Tokyo, Japan). The leaf epidermis and stomata were observed and photographed under 15 kv. The diameter of the stomata was measured by Motic Images Advanced 3.2 (Notic China Group Co., Ltd., China) software at 300× magnification.

Membrane integrity was evaluated in the leaves and roots by measuring electrolyte leakage. Fresh leaf or root samples (50 mg) were cut in small pieces followed by repeated washings with double distilled water. And then the samples were incubated at 28°C for 2 h in test tubes containing 10 mL of double distilled water for determining EC1 using a conductivity meter (Lei-Ci, DDSJ-308F, Shanghai, China). The same samples were again kept in water bath at 100°C for 20 min and the EC2 was recorded at room temperature. The electrolyte leakage was calculated as electrolyte leakage (%) = (EC1/EC2) × 100%. MDA content in the leaves or roots was measured by TBA reaction according to Shi et al. (2014) with modifications. Leaves or roots tissue (0.5 g) was homogenized in 10 mL of 0.1% (w/v) TCA, and the extract was centrifuged at 5, 000 × g for 10 min at 4°C. To measure MDA, 1.5 mL of the supernatant was added into 1.5 mL of 0.5% (w/v) TBA made in 5 % TCA. The mixture was heated at 100°C for 20 min and then quickly cooled in an ice bath. After centrifuging at 7, 888 × g for 10 min, the absorbance of the supernatant was measured at 450 nm, 532 nm, and 600 nm, respectively. The MDA content of leaves or roots (per 0.5 g, FW) was calculated using the formula: C (nmol. g^-1 FW) = 20 × [6.45 (OD₅₃₂ – OD₆₀₀) – 0.56 OD₄₅₀].

The production of O₂^•- was determined using the protocol described by Jiao et al. (2011) with some modifications. Samples (0.5 g) were homogenized in an ice bath with 1.2 mL of phosphate buffer (pH 7.8) and centrifuged at 5, 000 × g for 10 min at 4°C. The supernatant was reacted with 1 mL of hydroxylamine hydrochlorides for 1 h, then 1 mL of p-aminobenzene sulfonic acid and 1 ml of α-naphthylamine were added. The solution was kept at 25°C for 20 min. The optical density value of the solution was measured with a spectrophotometer at 530 nm using NaNO₂ as the standard curve and the corresponding calibration curves was Y = 552.16 X + 4.365 (R² = 0.9903) H₂O₂ was measured spectrophotometrically according to Jiang et al. (2012). Roots and leaves (0.2 g) were homogenized in an ice bath with 1 mL of 0.1% TCA and centrifuged at 12, 000 × g for 20 min at 4°C. The supernatant was used for measuring H₂O₂ content. The reaction mixture consisted of 0.5 mL of the extracted supernatant, 0.5 mL of 10 mM potassium phosphate buffer (pH 7.0) and 1 mL of 1 M potassium iodide. The reaction was developed for 1 h in darkness before the absorbance was measured at 390 nm. The blank was 0.1% TCA in the absence of root extract. The amount of H₂O₂ was calculated using a standard curve prepared with known concentrations of H₂O₂. The corresponding calibration curves was Y = 82.69 X – 1.11 (R² = 0.9959).

Endogenous H₂S content was determined by the formation of methylene blue from dimethyl-p-phenylenediamine in H₂SO₄ according to the method described previously (Zhang et al., 2008). Samples (0.5 g) were extracted in 1 mL phosphate buffer solution (50 mM, pH 6.8) containing 0.2 M ASA and 0.1 M EDTA. Subsequently, 1 M HCl (0.5 mL) was added to the homogenate in a test tube for releasing H₂S, and 0.5 mL of 1% (w/v) zinc acetate was used to absorb H₂S. After 30 min reaction, 0.3 mL 5 mM N,N-dimethyl-p-phenylenediamine dihydrochloride dissolved in 3.5 mM H₂SO₄ was injected into the trap. Then 0.3 mL of 50 mM ferric ammonium sulfate in 100 mM H₂SO₄ was injected into the trap. The amount of H₂S in zinc acetate traps was determined spectrophotometrically at 670 nm, after leaving the mixture for 15 min at room temperature. Solutions with different concentrations of Na₂S were prepared, treated in the same way as the assay samples and were used for the quantification of H₂S.

Samples (0.3 g) were ground with a mortar and pestle in liquid nitrogen and soluble proteins were extracted with 1.5 mL of cold extraction buffer containing 20 mM Tris–HCl (pH 8.0), 0.1% (w/v) DTT and 0.2% (w/v) sodium ascorbate. The homogenate was centrifuged at 13, 000 × g at 4°C for 15 min. The resulting supernatant was used for the determination of the activities of L/D-CD, CAS, and OAS-TL. Total L-CD activity was determined by the release of H₂S from L-cysteine as described in Riemenschneider et al. (2005) with minor modification. The assay contained in a total volume of 1 mL: 0.1 mL of 10 mM L-cysteine, 0.8 mL 100 mM Tris–HCl (containing 2.5 mM DTT, pH 9.0) and 0.1 mL of protein solution. After incubation for 60 min at 30° C, the reaction was terminated by adding 0.1 mL of 30 mM FeCl₃ dissolved in 1.2 N HCl and 0.1 mL of 20 mM N, N-dimethyl-p-phenylenedi-amine dihydrochloride dissolved in 7.2 N HCl. The formation of methylene blue was determined at 670 nm, and known concentrations of Na₂S were used in the calibration curve. D-CD activity was determined in the same way as L-CD activity except with D-cysteine instead of L-cysteine and the pH of the Tris–HCl buffer was 8.0. One unit of enzyme activity is defined as the amount of enzyme that produces 1.0 mmol/min H₂S under the stated assay conditions and expressed as U g^-1 FW. The total OAS-TL activity was determined following the method described Bloem et al. (2004). CAS activity was measured by the release of H₂S from L-cysteine in the same way as L-CD activity with the following modifications: the 1.0 mL volume of reaction mixture contained 0.1 mL of 10 mM L-cysteine, 0.7 mL 100 mM Tris–HCl (pH 9.0), 0.1 mL of 7.5 mM KCN and 0.1 mL of protein solution, as described in Meyer et al. (2003).

Dried plant material (200 mg) was subjected to acid digestion in 10 mL digestion mixture [HClO₄: HNO₃: H₂SO₄ (5: 1: 1, v/v)] on a hot plate (120°C) until the volume was reduced to 1 mL. After diluting with double distilled water, the Na⁺ and K⁺ contents were estimated using the flame photometer (Horneck and Hanson, 1998).

To study the effect of NaHS treatment on the expression of the SOS1, SKOR, and PM H⁺-ATPase genes in cucumber leaves and roots in response to salt stress, 18-day-old seedlings treated with either Hoagland’s nutrient solutions (control group), 200 mM NaCl, or 200 mM NaCl + 15.0 μM NaHS for a week were harvested. Total RNA of cucumber leaves or roots (100 mg) was isolated by grinding with mortar and pestle in liquid nitrogen to a fine powder and using TriZol Reagent (Invitrogen) in accordance with the manufacturer’s protocol. The total RNA samples were treated with RNAase-free DNase (TaKaRa Bio Inc., Dalian, China) to eliminate traces of DNA, followed by quantification using the NanoDrop 2000 (Thermo Fisher Scientific, Wilmington, DE, United States). Total RNA (2 μg) was reverse-transcribed using an oligo d (T) primer (50 μM, 1 μL) and M-MLV reverse transcriptase (200 U/μL, 1 μL) (BioTeke, Beijing, China). Real-time quantitative RT-PCR reactions were performed using an ABI 7000 (Applied Biosystems) with SYBR^®Premix ExTaq^TM (TaKaRa Bio Inc., China) and the cycling conditions of denaturation at 95°C for 5 min, followed by 40 cycles of denaturation at 95°C for 15 s, annealing at 60°C for 30 s, and extension at 72°C for 15 s. Using specific primers (Supporting information, Supplementary Table S1), the expression levels of the genes were presented as values relative to the corresponding control samples under the indicated conditions, with normalization of data to the geometric average of internal control gene GAPDH (a housekeeping genes). Three independent replicates were performed for each sample. The comparative threshold cycle (Ct) method was used to determine the relative amount of gene expression. Relative gene expression levels were calculated via the 2^-ΔΔCT method. The relative gene expression was determined as previously described by Livak and Schmittgen (2001).

The ASA content was determined using the DCIP method. Samples (0.50 g) were homogenized in 3.0 mL of 2 % oxalic acid supplemented with 0.50 mL of 30 % ZnSO₄ and 0.50 mL of 15% K₄Fe(CN)₆⋅3H₂O. The volume of the extract was adjusted to 10.0 mL with 1% oxalic acid, followed by centrifugation at 10, 000 × g for 10 min. Then 3 mL of the supernatant was mixed with 2.0 mL of dye solution (0.25% DCIP and 0.20% NaHCO₃) and 3 mL of dimethylbenzene. The absorbance was read at 500 nm, and the content of ASA was calculated using a standard curve. Total GSH was estimated using a kit (Jiancheng Bioengineering Institute, Nanjing, China). The absorbance of GSH was measured at 420 nm according to manufacturer’s instructions. CAT activity was measured according to the methods described by Velikova et al. (2000) using 10 mM potassium buffer (pH 7.00), with 100 μL of enzyme extract, and 33 mM H₂O₂. H₂O₂ was assayed from the decrease in absorbance in optical density at 240 nm, and the activity was calculated using the extinction coefficient of 40 mM cm^-1 of H₂O₂. GR activity was determined by measuring the absorbance change at 340 nm due to oxidation of NADPH (Foyer and Halliwell, 1976). SOD and POD activities were assayed as described by Chen et al. (2013).

Data were analyzed by one-way ANOVA using SPSS (version 21.0.0; IBM, Armonk, NY, United States). Different letters on the graphs or in the tables indicated that the mean values were statistically different at the P < 0.05 level.

The morphology, survival rate, and expansion rate of the second leaf were determined after cucumber seedlings exposed to 200 mM NaCl for 7 days. Under salt stress, most seedlings began to show leaf wilting, lodging and inhibition of the second leaves compared with control group (Supporting information, Supplementary Figure S1). In order to analyze the alleviation effect of NaHS on growth inhibition induced by NaCl in a dose-dependent manner, series of concentrations of NaHS were added to the nutrient solution. In cucumber seedlings, 15 or 20 μM NaHS significantly alleviated the inhibition of plant growth during salt stress (Supplementary Figure S1A). Salinity treatment significantly (P < 0.05) reduced the survival rate by 89 %, the rate of expansion of the second leaf by 62 % (Supplementary Figures S1B,C) and significantly (P < 0.05) affected general growth (Figure 1A). Compared with the control, the shoot fresh weight, root fresh weight, seedling fresh weight, seedling total length, shoot dry weight, root dry weight, leaf area and root length were significantly (P < 0.05) reduced by 66%, 58%, 65%, 43%, 50%, 53%, 90%, and 48% under salt stress, respectively (Figure 1B–I). However, all growth parameters showed different degree ameliorating effects with application of different concentrations of NaHS (Figure 1 and Supplementary Figure S1). For example, the combination NaCl+15 μM NaHS significantly (P < 0.05) reduced shoot fresh weight, root fresh weight, seedling fresh weight, seedling total length, shoot dry weight, root dry weight, leaf area and root length by 28%, 40%, 31%, 14%, 17%, 42%, 41%, and 15%, respectively, as compared to the control (Figure 1B–I). Thus, these results indicated that exogenous supplementation with 15 μM NaHS was the most ameliorating effect in boosting cucumber tolerance to NaCl stress.

In comparison with the control, a significant (P < 0.05) decline in the P_n (by 14.3%) and a sharp decline in the T_r (by 94%) were recorded in the leaves of salt-stressed cucumber plants (Table 1). The combination of NaCl and 15 μM NaHS showed a decline in the P_n (by 2.4%) and T_r (by 90.4%) compared with the control, respectively (Table 1). Nevertheless, 5 μM NaHS did not significantly affect the Tr under salt stress. In addition, 20 μM NaHS alleviated the declines of P_n, but did not cause a significant change of T_r under salt stress.

As shown in Table 1, salt stress significantly (P < 0.05) reduced F_v/F_m (by 6%), F_v/F_o (by 25%),Φ_PSII (by 7%) and qP (by 12%) compared with the control, while 15 μM NaHS treatment reduced these by 0.5%, 4.5%, 2% and 3.1%, respectively, compared with the control. In contrast, comparing with the control, the parameters of F_o and NPQ significantly (P < 0.05) increased by 39% and 78% under salt stress. As expected, 15 μM NaHS significantly reversed the increase in F_o (by 10.4%) and NPQ (by 34.2%) under salt stress (Table 1).

Salt stress resulted in a decrease in the contents of the photosynthetic pigments C_a, C_b, C_a+b and C_x+c compared to the non-treated plants. Compared with the control, salt stress reduced C_a, C_b, C_a+b and C_x+c content by 38%, 37%, 36%, and 35%, respectively, while 15 μM NaHS treatment reduced C_a, C_b, C_a+b, and C_x+c content by 11.2%, 8.7%, 7.8% and 15% (Table 1).

To determine the effects of H₂S on stomata in cucumber, the abaxial leaf surface was observed by a scanning electron microscope to analyze several stomatal indexes (Figure 2A–C). Salt stress induced stomatal closure and resulted in abnormal morphology of the epidermal and guard cells compared to the untreated control plants, whereas application of 15 μM NaHS significantly reversed these damages. Salt stress caused a noticeable reduction in stomatal opening rate by 67% (Figure 2D) and stomatal width by 74% (Figure 2E), and a significant increase in stomatal density by 41% (Figure 2F) and stomatal length by 36% (Figure 2G) compared with control plants. However, 15 μM NaHS treatment reduced stomatal opening rate by 30.8% (Figure 2D) and stomatal width by 52.8% (Figure 2E), and increased stomatal density by 12.6% (Figure 2F) and stomatal length by 12% (Figure 2G) compared with control plants. The diverse responses of the stomata indicated that there is co-ordination of the physiological (i.e., aperture) and morphological (i.e., stomatal density) changes induced by H₂S that control gas exchange and Tr when encountering unfavorable environments.

The exposure of the cucumber plants to salinity significantly (P < 0.05) increased electrolyte leakage by 1.1 fold in leaves (Figure 3A) and by 87.2% in roots (Figure 3B), and increased the MDA content by 45% in leaves (Figure 3C) and by 1.0 fold in roots (Figure 3D) compared with the control. However, treatment of salt-stressed plants with 15 μM NaHS resulted in an increase in electrolyte leakage by 27.6% in leaves and by 46% in roots (Figure 3A,B) and an increase in the MDA content by 21% in leaves and by 5.4% in roots as compared to the control (Figure 3C,D). These results indicated that NaHS clearly reduces cell membrane injury.

Salt stress treatment did not change the O₂^•- accumulation in cucumber leaves (Figure 3E), but it did increase by nearly 2.1 fold in roots as compared to control plants (Figure 3F). While different concentration of NaHS treatment increased the accumulation of O₂^•- in cucumber leaves and roots as compared to the control (Figure 3E,F). Salt stress caused a dramatic increase in H₂O₂ by 1.92 fold in leaves (Figure 3G) and 1.04 fold in roots (Figure 3H) compared with the control. Treatment with 15 μM NaHS increased the H₂O₂ by 21.7% in leaves (Figure 3G) and 19.7% in roots compared with the control (Figure 3H). These results indicated that exogenous application of NaHS suppresses ROS accumulation, thereby protecting cucumber plants from NaCl-induced oxidative stress.

In comparison with the control, a sharp decline in the content of endogenous H₂S was recorded in leaves (by 49%) (Figure 4A) and roots (by 45%) (Figure 4B) of salt-treated cucumber plants, whereas 15 μM NaHS treatment improved the content of endogenous H₂S by 9.9% in leaves (Figure 4A) and by 9.1% in roots (Figure 4B) compared with the control. Compared with the control, salt stress significantly (P < 0.05) decreased the activity of L-CD by 19% in leaves (Figure 4C) and 12% in roots (Figure 4D). The same change was found in the activity of D-CD after salt stress, which reduced by 13% in leaves (Figure 4E) and 33 % in roots compared with the control (Figure 4F), As expected, these changes were reverted by different concentrations of NaHS treatment. For example, treatment with 15 μM NaHS significantly (P < 0.05) improved the activities of L-CD (38.7% in roots) and D-CD (by 52.7% in leaves and 4.6 fold in roots) compared with the control (Figure 4C–F). These results indicated that NaHS treatment significantly enhanced the activities of L-CD and D-CD in cucumber seedlings under salt stress.

As for the activity of CAS, a completely opposite trend was found between leaves and roots. Salt stress significantly (P < 0.05) improved the activity of CAS in leaves by 19% (Figure 4G) and reduced it by 43% in roots (Figure 4H) compared with the control. Application of 5 or 10 μM NaHS enhanced the activity of CAS by 9.4% or 8.9% in leaves, in contrast, 15 or 20 μM NaHS reduced it by 31.7% or 22.8% compared with the control. In the roots, 5∼20 μM NaHS all reduced the activity of CAS in different degree compared with the control group (Figure 4G,H). In addition, the OAS-TL activity significantly (P < 0.05) increased in both leaves (by 65%) (Figure 4I) and roots (43%) (Figure 4J) under salt stress. Similarly, these effects were reverted by different concentrations of NaHS as compared to the plants receiving salinity treatment alone. For example, 15 μM NaHS treatment reduced the activity of OAS-TL by 42% in leaves and 2.4% in roots as compared to the control (Figure 4I,J).

In plants, maintaining a lower Na⁺/K⁺ ratio is critical to its ability to tolerate salt stress. Compared with the control, a sharp increase in the Na⁺ content was observed in leaves (by 2.13 fold) and roots (1.35 fold) under salt stress (Figure 5A,B). On the contrary, a significant decrease in the K⁺ content was recorded in leaves (by 30%) and roots (by 41%) under salt stress as compared to the control (Figure 5C,D). Accordingly, the Na⁺/K⁺ ratio significantly increased in leaves (by 3.4 fold) and roots (by 3 fold) of salt-treated plants than the control (Figure 5E,F). As expected, treatment with 15 μM NaHS improved the Na⁺ content in leaves by 1.9 fold and roots by 1.0 fold, and reduced the K⁺ content in leaves by 17.8% and roots by 28% compared with the control, indicating that 15 μM NaHS significantly reversed the increase of Na⁺ and the decrease of K⁺ under salt stress. In addition, the Na⁺/K⁺ ratio was significantly lower in the NaHS-treated plants when compared with the plants treated with salt alone (Figure 5E,F). Thus, H₂S eases excessive Na⁺ uptake and maintains the Na⁺/K⁺ homeostasis at the cellular level in cucumber plants under salt stress.

To further characterize the effect of H₂S on the modulation of Na⁺/K⁺ homeostasis in cucumber seedlings upon salt stress, the transcript levels of PM H⁺-ATPase, SOS1 and SKOR were analyzed. Real-time RT-PCR analysis showed that NaCl treatment resulted in the significant decreases in the transcripts levels of PM H⁺-ATPase by 66.9% (Figure 6A), SOS1 by 42.7% (Figure 6C) and SKOR by 72.4% (Figure 6E) in cucumber leaves compared with the controls, respectively. In contrast, the relative expression of PM H⁺-ATPase, SOS1 and SKOR were significantly increased by 72.6% (Figure 6B), 2.3 fold (Figure 6D) and 1.17 fold (Figure 6F) in the roots under NaCl stress compared with the control. The NaCl-induced increases in PM H⁺-ATPase, SOS1 and SKOR transcripts were significantly reversed by 15 μM NaHS treatments in the cucumber roots. As shown in Figure 6, compared with the control, treatment with 15 μM NaHS+NaCl reduced the relative expression of PM H⁺-ATPase, SOS1 and SKOR by 57.1%, 59.5% and 75.1% in leaves, respectively. On the contrary, treatment with 15 μM NaHS+NaCl improved the relative expression of PM H⁺-ATPase, SOS1 and SKOR by 2.1 fold, 2.59 fold and 3.21 fold in leaves, respectively, as compared to the control.

To evaluate the role of H₂S in mediating salt-induced antioxidant system, the levels of non-enzymatic antioxidants (ASA and GSH) and the activities of antioxidant enzymes (SOD, CAT, GR, and POD) were analyzed in leaves and roots of cucumber plants. As show in Table 2, compared with the control, the contents of ASA and GSH and the activity of GR were significantly increased by 98%, 73%, and 61% in the leaves, but they were decreased by 88%, 23%, and 2.25 fold, in the roots under NaCl treatment. However, application of exogenous NaHS ameliorated the effect of salt stress. For example, 15 μM NaHS treatment significantly increased the ASA content by 42.6% and the GR activity by 9.1%, and decreased GSH by 44% in the leaves as compared to the control. On the contrary, 15 μM NaHS treatment decreased the ASA content by 23.9% and the GR activity by 27.8%, and increased GSH by 48.7% in the roots as compared to the control. Compared with the control, SOD activity was increased under salt stress in the leaves by 23% and roots by 43%. Application of NaHS showed a synergistic effect resulting in decreased SOD activity in the leaves and roots under NaCl stress. The application 15 μM NaHS significantly decreased SOD activity in the leaves by 19% and roots by 15% as compared to NaCl stress alone. The NaCl treatment significantly decreased the activity of CAT in the leaves by 6 % and roots by 59% as compared to control, but increased the POD activity in the leaves by 25% and roots by 10%. Application of NaHS reversed these changes in the leaves and roots induced by salt stress.

Results showed that salinity stress inhibited plant growth performance as judged by decreased shoot fresh weight, root fresh weight, shoot dry weight, root dry weight, leaf area and root length compared with control, but this inhibition was significantly alleviated by H₂S donor NaHS (Figure 1). Similarly, the reversion of salt stress responses by H₂S has been observed in other plant species, like Medicago sativa (Lai et al., 2014), Oryza sativa (Mostofa et al., 2015b) and Triticum aestivum (Deng et al., 2016). However, it is well-known that H₂S must be within a rather narrow concentration range to act as a signal molecule and that high concentrations of H₂S induce a toxicity response. In our study, the capacity of H₂S to enhance salt tolerance firstly increased and then decreased with the concentration improvement. Ameliorative effects of phenotypes were seen from 5 to 15 μM NaHS, but plants exhibited slight toxicity under the 20 μM NaHS treatment. It may be that excessive high levels of H₂S might lead to the ROS over-production, which further leads to the inhibition of the mitochondrial electron transport chain (Mancardi et al., 2009). In our study, 20 μM NaHS treatment indeed caused the ROS over-production and cell membrane injury than 15 μM NaHS as judged by the parameters of electrolyte leakage, MDA, O₂^•- and H₂O₂ (Figure 3). Hence, the use of H₂S is a double-edged sword, and the maintenance of a suitable level of intracellular H₂S is key conferring salt tolerance in cucumber seedlings.

In this study, the F_v/F_m was significantly decreased under salt stress, and they were remarkably improved with addition of 15 μM NaHS (Table 1). Similarly, the results of Christou et al. (2013) showed that 100 mM NaCl stress significantly decreased the values of F_v/F_m, while 100 μM NaHS significantly reversed the decline of F_v/F_m. In addition, a phenomenon that H₂S prevented the reduction of photosynthetic pigments C_a, C_b, C_a+b, and C_x+c induced by salt stress was also observed in our study (Table 1). The results are consistent with these of Mostofa et al. (2015b), who reported that H₂S improved overall growth and biomass of salt-stressed rice plants which could be attributed to its role in protecting Chl_a, Chl_b and Cx+c from salt-induced damage. As the photosynthetic parameters P_n and T_r, our results showed that H₂S also alleviated the decrease in the photosynthetic parameters P_n and T_r. These results are consistent with these of Dawood et al. (2012), who reported that H₂S reduced the Aluminum-induced inhibition of P_n, G_s and T_r. Another study also demonstrated a similar result that NaHS treatment increased the P_n and G_s values in Spinacia oleracea seedlings under drought (Chen et al., 2016). Together these results suggest that H₂S plays a role in plant photosynthesis.

Earlier investigations demonstrated that hypoxia, salt and Cd stress induce notable increases in endogenous H₂S production in different plant species, such as pea, strawberry, alfalfa and Chinese cabbage (Cheng et al., 2013; Christou et al., 2013; Lai et al., 2014; Zhang et al., 2015). The activation of endogenous H₂S level after stress treatments indicated that H₂S might also be an important secondary messenger of stress sensing which in turn modulated plant physiological changes and downstream gene expressions (Christou et al., 2013). Surprisingly, a remarkable decline in endogenous H₂S production was observed in cucumber leaves and roots upon treatment with 200 mM NaCl for 7 days in our experiment. Further investigation is still required to detail the reason. Although different kinds of environmental stress factors caused differential dynamic changes in endogenous H₂S metabolism, exogenously applied NaHS, a well-known H₂S donor, all greatly enhanced H₂S concentration.

Generally, synthesize H₂S in plants is associated with the enzymes L-CD, D-CD, OAS-TL or CAS, often in response to environmental stress, leading to accumulation of endogenous H₂S and the acquisition of stress tolerance. The activities of L-CD in leaves and D-CD in both roots and leaves all showed a significantly decreased after salt stress treatment as compared to the control (Figure 4C–F). Lai et al. (2014) found that NaCl could induce L-CD in Medicago sativa roots, while the total activity of D-CD was not significantly altered. Meanwhile the expression of L-CD and D-CD at the transcripts level were strongly induced by drought stress in Arabidopsis thaliana (Jin et al., 2011). In our study, exogenously applied 15 μM NaHS significantly enhanced the activities of D-CD in both roots and leaves and L-CD in roots compared with the control or salt stress group (Figure 4D–F). However, Cheng et al. (2013) found that exogenously applied NaHS decreased the total activity of L-CD and D-CD in Pisum sativum L. seedlings under hypoxia stress. In addition, NO could induce the activities of to L-CD, OAS-TL and CAS regulate the endogenous H₂S biosynthesis in maize roots (Peng et al., 2016). In our study, salt-stress significantly improved the activity of CAS in leaves and OAS-TL in leaves and roots and significantly reduced the activity of CAS in roots (Figure 4G–J). This is also in line with a report of Cheng et al. (2013), and their results showed that exogenously applied NaHS enhanced the activity of CAS and OAS-TL in Pisum sativum L. seedlings under hypoxia stress (Cheng et al., 2013).

Maintenance of Na⁺ and K⁺ ion homeostasis is critically important for plants to survive under salt stress. Our study provided firm evidence that H₂S prevented Na⁺ uptake and improved K⁺ uptake in both leaves and roots, thereby restraining the salt-induced decrease of the Na⁺/K⁺ ratio (Figure 5). Similarly, Lai et al. (2014) observed that the biologically beneficial role of NaHS was due to its specific ability to retain K⁺, thus preventing the increase of the Na⁺/K⁺ ratio in stellar cells under salt stress. In addition, Christou et al. (2013) showed that H₂S regulates the expression of SOS pathway genes and kept the balance of Na⁺ and K⁺ in strawberry plants under salt stress. Deng et al. (2016) also found that H₂S significantly improved the salt tolerance of wheat seedlings by regulating the membrane-bound translocation proteins of the SOS1 pathways to increase Na⁺ extrusion and decrease Na⁺ uptake. Interestingly, salt stress induced the up-regulation of PM H⁺-ATPase, SOS1 and SKOR at the transcript levels in roots compared with the control, but a completely opposite trend in leaves. This is also in line with a report of Wang et al. (2013), and their results showed that the expression of CsSOS1 increased in roots and decreased in leaves with the concentration of NaCl increasing in cucumber. Perhaps the root irrigation treatment method in this study affected the expression of genes or some signal first in roots. In our study, H₂S alleviated the NaCl-induced increases in PM H⁺-ATPase expression in roots. It has been known that NaCl stress also up-regulates the expression of genes encoding PM H⁺-ATPase in roots (Shi et al., 2000). Moreover, it is well known that PM H⁺-ATPase can sustain an H⁺ gradient by promoting Na⁺ efflux and H⁺ influx to drive Na⁺/H⁺ antiport across the plasma membrane. It was also found that the administration of NaHS effectively prevented the NaCl-triggered K⁺ efflux in the mature zone of alfalfa seedling roots, which might be partially related to the SKOR channel (Lai et al., 2014). These studies suggested that salt-tolerant need plants have an ability to prevent transport of Na⁺ from roots to shoots and an improved ability to exclude Na⁺ from the roots (Takahashi et al., 2007a,b).

Plants suffering from NaCl toxicity often exhibit symptoms associated with oxidative stress and membrane lipid peroxidation, which can result in accumulation of ROS and MDA (Hossain et al., 2005). In this study, high salinity triggered overproduction of O₂^•- and H₂O₂, increased the electrolyte leakage, and caused accumulation of MDA in both cucumber leaves and roots (Figure 3). H₂S alleviated the salinity-induced oxidative damage, as evident by reduced levels of O₂^•-, H₂O₂, MDA and electrolyte leakage in both cucumber leaves and roots (Figure 3). This is in line with a report of Mostofa et al. (2015b), and their results showed that H₂S reduced the salt-induced increase in the levels of O₂^•-, H₂O₂ and MDA in rice. Similar results were also reported by Chen et al. (2016), who demonstrated that NaHS treatment counteracted the accumulation of ROS and increased the MDA content of Spinacia oleracea L. seedlings under drought. The balance between ROS production and antioxidant defense determines the extent of oxidative damage within the plant. In our study, NaHS treatment significantly alleviated the improvement of ASA and GSH contents and SOD, GR, and POD activities in leaves of cucumber under salt stress (Table 2). In cucumber roots, H₂S also successfully counteracted the decline in the content of ASA and GSH and the activities of CAT, GR and POD under salt stress (Table 2), indicating that H₂S may enhance the ability of cucumber roots to scavenge ROS so that the plant can maintain membrane integrity under salt stress. The similar result was reported by Wang et al. (2012), who founded that NaHS pretreatment significantly activated the antioxidant enzymes activities including SOD, CAT, POD, and APX and their expression at the transcripts level under 100 mM NaCl stress in Medicago sativa L., thus resulting in the alleviation of oxidative damage induced by NaCl.