Transcription profile analysis identifies marker genes to distinguish salt shock and salt stress after stepwise acclimation in Arabidopsis thaliana and Zea mays

Many physiological and molecular responses to salt stress have been investigated after a salt shock. However, salt shock rarely happens in agricultural practice. In the field, salts accumulate gradually due to poor agricultural management. Thus in salinity research, it is more reasonable to investigate plant reaction after stepwise acclimation to salt stress. Previous studies demonstrate that salt shock induces Phase 0, a short-term effect that shows transient water loss and rapid turgor decrease; salt stress after stepwise acclimation avoids Phase 0 effects and induces Phase 1. During Phase 1, plants show maintenance of turgor. In this study, salt shock and stepwise acclimation to salt stress were separated at physiological and transcriptional levels. Four major experiments were conducted: 1) leaf turgor changes were monitored in real time after salt application to separate Phase 0 and Phase 1 effects at the physiological level, 2) RNA-sequence analysis was conducted in Arabidopsis thaliana L. to identify potential marker genes that are involved in plant water relations to distinguish Phase 0 and Phase 1 at transcript level, 3) these selected marker gene candidates were identified in Arabidopsis at different Phase 0 and Phase 1 time points via qRT-PCR, 4) these candidates were further evaluated in Zea mays L. (a model plant for applied research in plant physiology and an important crop plant) via qRT-PCR. In future salinity research, marker genes that are both applicable in Arabidopsis and maize have the potential to differentiate salt shock and stepwise acclimation to salt stress.


Introduction
Salinity is considered a major threat to crop yield worldwide because most crops are sensitive to salt stress (Munns and Tester, 2008). Several practical strategies have been developed to alleviate damages caused by salt stress on crops, including conventional plant breeding (Schubert et al., 2009) and transgene technology (Roy et al., 2014). However, many approaches have not successfully increased crop yield or salt resistance. One reason for this failure might be that most physiological and molecular responses to salt stress are investigated after salt shock where plants are suddenly exposed to high levels of salt. However, salt shock does not usually occur in agricultural practice (Shavrukov, 2013). In the field, salts accumulate gradually because of high-salt irrigation water and poor agricultural management that allows ground water to rise to soil surface (Rengasamy, 2006). Therefore, it is more conducive to investigate salt stress in a way that allows plants to stepwise acclimate to salt treatment like in the field.
According to Munns (1993) and Munns and Tester (2008), plants suffer salt stress in two phases: one is an early, also called osmotic phase (Phase 1) that physiologically inhibits growth of young leaves; the other is a late ion-toxicity phase (Phase 2) that accelerates senescence in mature leaves. In addition, Schubert (2011) suggested to add a transient phase (Phase 0) before Phase 1 under salt shock situations. During Phase 0, plants show transient decreases in turgor and growth rate in the first few minutes and hours after the imposition of salt. These changes are transient and recover after some time (Thiel et al., 1988;Munns, 1993). The extent of recovery depends on the levels of applied salts, the sensitivity of plants, and the salt acclimation capacity of plants (Munns and Tester, 2008). It is important to avoid Phase 0 effects in salinity research because its transient character is not useful in plant breeding (Schubert, 2011). During Phase 1, ion accumulation surrounding the root medium reduces the water availability to plants and plants suffer osmotic stress (Munns and Tester, 2008;Vetterlein et al., 2004). Plant growth is inhibited during this phase and the growth inhibition showed in this phase is similar to that induced by other osmotica such as PEG, mannitol, and KCl (Munns, 1993). Thus, this effect is not salt-specific. During this phase, carbohydrate supply is sufficient (Schubert, 2011) and turgor is maintained because of osmotic adjustment (De Costa et al., 2007;Thiel et al., 1988). However, Sümer et al. (2004) found that Na + also contributes to the decreased growth in maize during Phase 1. During Phase 2, plants develop chlorosis or even necrosis resulting from ion toxicity (Munns and Tester, 2008).
Turgor is the driving force of plant cell expansion and elongation (Cosgrove, 2005). In earlier research, turgor was calculated by measuring the total water potential and osmotic potential separately via psychrometer and osmometer, respectively. In those methods, tissues under test are destroyed, which inevitably impacts the following research. Recently, a novel method conducted by ZIM YARA water sensors has been developed which can measure turgor pressure in real time without damaging the plants (Zimmermann et al., 2008). The maximum value of recorded patch pressure (P m ) occurs around noon and is a potential parameter to determine whether plants are suffering a water deficit. Under water stress, the P m value increases. This parameter has been successfully used in grapevines (Westhoff et al., 2009), wheat (Bramley et al., 2013), and Arabidopsis (Ache et al., 2010) under experimental and field conditions.
In the early stages of salt stress (Phase 0 and Phase 1), the plants' first response is to alleviate salt stress-induced water deficiency by reducing leaf water loss (Deinlein et al., 2014). Leaf transpiration and leaf hydraulic conductance are two key processes that mediate leaf water loss (Defraeye et al., 2014;Kosma et al., 2009;Pou et al., 2013). When plants experience water stress, they reduce the leaf water loss mainly by decreasing stomatal transpiration through closing stomata (Defraeye et al., 2014). The rest of transpired leaf water loss is determined by cuticular transpiration . The cuticle is a hydrophobic layer on epidermal cells, comprised of two major components (cutin and cuticular waxes). It has been demonstrated that water deficit increases the amount of cuticular wax per unit area and leaf cuticle thickness in Arabidopsis plants to enhance their resistance (Lü et al., 2012). In addition, the efficiency of water transport within leaves is reduced under water stress conditions (Pou et al., 2013), and it is closely related to the transcript abundance of aquaporins in sunflowers (Nardini et al., 2005), walnuts (Cochard et al., 2007), and grapevines (Pou et al., 2013).
According to Munns and Tester (2008) and Schubert (2011), turgor pressure decreases dramatically after salt shock (Phase 0) and is maintained after stepwise acclimation to salt stress (Phase 1). Our first goal of this study was to distinguish these two types of salt stress at physiological level. For this purpose, turgor pressure changes were measured in a non-invasive online-monitoring way via ZIM YARA water sensor in Arabidopsis thaliana (a model plant for genetics and molecular biology) and Zea mays (an important crop plant). In a second step, to separate Phase 0 and Phase 1 at transcriptional level, RNA-sequence analysis was conducted in Arabidopsis to select marker gene candidates that are highly and specifically affected by each type of salt stress. Third, only genes involved in the regulation of plant water relations were selected as marker genes in Arabidopsis to avoid genes that are related to Phase 2 (ion toxicity) getting involved. To test whether the marker candidates allow for differentiation between Phase 0 and Phase 1, they were first evaluated in Arabidopsis by means of qRT-PCR at different time points of Phase 0 and Phase 1, and then validated in maize plants via qRT-PCR. We hypothesized that there are marker genes which can be used to differentiate Phase 0 (salt shock) from Phase 1 (stepwise acclimation to salt stress).
The relative turgor pressure was monitored by ZIM YARA water sensors which were clamped to leaves via two magnets. The clamp pressure (P clamp ) exerted by magnets is constant during the measurements and can be adjusted by screwing the movable magnet. The leaf turgor pressure (P c ) is opposed to clamp pressure in the leaf patch. The output of sensors (patch pressure, P p ) is a power function of the turgor pressure P c (Zimmermann et al., 2008): Among these, a and b are constants. F a is a leaf-specific attenuation factor. Equation (1) shows that the recorded P p pressure is inversely correlated with leaf turgor pressure P c , which means that P p is large when P c is small and vice versa.
Sensors were attached to fully expanded rosette leaves of Arabidopsis and the 4th leaves of maize to measure leaf turgor pressure in real time. Moreover, sensors were applied to the leaves 2 days prior to the treatments for accommodation. Due to individual compressibility of leaves and pressure applied through sensors, the initial P p value varied among leaves.

c) Quantification of cation and anion concentrations
Five-week-old Arabidopsis thaliana plants were harvested after 2 days of 90 mM NaCl treatment (stepwise acclimated, 15 mM per day). Plant shoots and roots were harvested and dried at 80°C separately. In order to determine Na + , Ca 2+ , K + , and Mg 2+ concentrations, samples were prepared by dry-ashing (550°C) overnight and cations were measured with an atomic absorption spectrophotometer (SpectrAA220 FS, Varian, Mulgrave, Victoria, Australia). d) RNA sequence RNA sequence was performed by Max Planck Institute for Heart and Lung (Bad Nauheim, Germany). The total RNA was isolated from rosette leaves according to manufacturer protocols of RNeasy Plant Mini Kit (Qiagen). Genomic DNA was removed using RNase-Free DNase Set (Qiagen) to avoid contamination. Four milligrams of total RNA were used as input for Truseq Stranded mRNA Library preparation following the low sample protocol (Illumina). Sequencing was performed on the NextSeq500 instrument (Illumina) using v2 chemistry, resulting on W. Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245 average in 30 Mb reads per library with 1 × 75 bp single end setup. The resulting raw reads (Table S1) were assessed for quality, adapter content and duplication rates with FastQC (FastQC: a quality control tool for high throughput sequence data. Available online at: http://www. bioinformatics.babraham.ac.uk/projects/fastqc). Reaper version 13-100 was employed to trim reads after a quality drop below a mean of Q20 in a window of 10 nucleotides. Only reads between 30 and 150 nucleotides were cleared for further analyses. Trimmed and filtered reads were aligned versus the Ensembl Arabidopsis genome version TAIR10 (version 34) using STAR 2.4.0a with the parameter "-out-FilterMismatchNoverLmax 0.1" to increase the maximum ratio of mismatches to mapped length to 10%. The number of reads aligning to genes was counted with featureCounts 1.4.5-p1 tool from the Subread package. Only reads mapping at least partially inside exons were admitted and aggregated per gene. Reads overlapping multiple genes or aligning to multiple regions were excluded. Differentially expressed genes were identified using DESeq2 version 1.62. Only genes with |log 2 FoldChange| ≥ 1, false discovery rate (FDR) ≤ 0.05, and a minimum combined mean of five reads were deemed to be significantly differentially transcribed. The Ensemble annotation was enriched with UniProt data (release 06.06.2014) based on Ensembl gene identifiers (Activities at the Universal Protein Resource (UniProt)). According to the regularized log transformation (R-Log transformation) in DESeq2, the replicates in each group work well (Supplemental Fig. S1). The RNA-sequence data can be found in the NCBI Gene Expression Omnibus (GEO) database under accession number GSE113138. It is currently private; please email us to get the secure token. e) qRT-PCR Total RNA was extracted using RNeasy Plant Mini Kit (Qiagen), and was reversely transcribed with RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific) according to the manufacturer protocols. Realtime PCR was performed in a StepOnePlus system (Applied Biosystems). Relative quantification of gene expression in mRNA level was calculated using equation by 2 -△Ct . △Ct is the target gene Ct subtracted by the geometric mean of the endogenous reference genes Ct. At-actin2 and Zm-GaPDh were considered as endogenous reference genes in Arabidopsis thaliana and Zea mays, respectively. Primers used for qRT-PCR are listed in Table S2.

Diurnal changes of leaf turgor pressure in phase 0
The relative leaf turgor changes were measured in real-time by ZIM YARA water sensors in a non-invasive way. After 2 weeks, the leaf patches beneath the sensors were brighter than the surroundings but had no signs of necrosis both in Arabidopsis thaliana and Zea mays A. Phenotypes of Arabidopsis plants exposed to 90 mM NaCl for 2 days. White bar = 2 cm. B/C. Quantification of shoot fresh weight (B), and shoot dry weight (C) of Arabidopsis plants treated with 90 mM NaCl for 2 days. Significant differences are indicated by asterisks (*, **, *** significant differences of shoot fresh weight during Phase 1 in comparison to the control with P < 5%, 1%, 0.1%, respectively; ttest). According to t-test, no significant difference of shoot dry weight was observed during Phase 1 in comparison to the control. D. Cation concentrations (Na + , Ca 2+ , K + , and Mg 2+ ) in Arabidopsis plants shoots and roots were quantified after 2 days treatment with 90 mM NaCl. Significant differences among Na + , Ca 2+ , K + , and Mg 2+ concentrations in shoots and roots are indicated by different letters (P < 5%; one-way ANOVA and Post-hoc analysis using Tukey's Honestly Significant Difference Method). Significant differences of Na + , Ca 2+ , K + , or Mg 2+ concentrations in shoots or roots after salt treatment in comparison to Na + , Ca 2+ , K + , or Mg 2+ concentrations in shoots or roots under control conditions are indicated by asterisks (*P < 5%, **P < 1%, ***P < 0.1%, ttest). Data show means of four replicates. Error bars represent the standard errors (SE). W. Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245 (Supplemental Fig. S2). Thus, YARA water sensors are suitable for the measurements of turgor pressure in Arabidopsis and maize.
To induce Phase 0 in Arabidopsis plants, salt shocks induced by 105 mM, 125 mM, and 200 mM NaCl were conducted. When 200 mM NaCl were added to Arabidopsis plants, P p increased rapidly within minutes but could not recover from this shock (Fig. 1A). Similarly, P p could not recover from 125 mM NaCl salt shock in Arabidopsis either (Supplemental Fig. S3). When the strength of treatment was reduced to 105 mM NaCl, P p value showed a rapid rise, just like after the 200 mM and 105 mM NaCl shocks, and reached its maximum value after 1 h (Fig. 1B). After the peak, P p recovered within one day and the maximum value of P p (P m ) increased a bit in the following days (Fig. 1B). When maize plants were treated with 200 mM NaCl salt shock, their P p increased rapidly within minutes and peaked after 2 h (Fig. 1C). The P p and P m values of maize plants both returned to the original levels 1 day after (Fig. 1C).

Diurnal changes of leaf turgor pressure in phase 1
When NaCl was added gradually, P p showed a different response to salt stress. NaCl was added to nutrient solution of Arabidopsis in daily increments of 25 mM, and reached a final concentration of 100 mM 4 days later. The P m value of Arabidopsis continued to rise with increasing NaCl, even after the addition was stopped ( Fig. 2A). Actually, the turgor should be maintained in this period. If the daily application was decreased from 25 mM to 15 mM NaCl and the addition period was increased from 4 days to 6 days, P m showed a slight increase after each 15 mM NaCl application, and fully recovered within 1 day (Fig. 2B). The P m value remained mostly stable in the first 6 days and only had a slight increase when NaCl addition was stopped in Arabidopsis (Fig. 2B). When maize plants were stepwise acclimated to 100 mM NaCl (25 mM NaCl per day), the P m value decreased slightly when 25 mM NaCl were added daily, and was maintained when NaCl application was stopped (Fig. 2C).
The growth of Arabidopsis was inhibited after 90 mM NaCl treatment (stepwise adapted, as described above) (Fig. 3A). Under salt stress, shoot fresh weight decreased which was 70.8% relative to the control (Fig. 3B). However, shoot dry weight showed no significant change (Fig. 3C). The Na + concentrations in shoots and roots increased after salt treatment, whereas other cation (Ca 2+ , K + , and Mg 2+ ) concentrations decreased (Fig. 3D). However, no toxicity and deficiency symptoms were observed in these plants.

Differentially transcribed genes in phase 0 and phase 1 for Arabidopsis thaliana
Nine samples from Arabidopsis thaliana were harvested and used for RNA-sequence analysis with three biological replicates for each treatment. One group was harvested under control conditions; another group was harvested 1 h after 105 mM NaCl treatment when P p reached its peak value; and the last group was harvested 2 days after reaching 90 mM NaCl when the P m value remained stable using the stepwise acclimation as described above. Based on the selection conditions (see "Materials and methods"), 213 differentially transcribed genes (DTGs) were identified in Phase 0, 2505 DTGs were observed in Phase 1, and 303 DTGs were affected both in Phase 0 and Phase 1 (Fig. 4).

Responses of DTGs involved in the inhibition of leaf water loss
To alleviate the salt stress-induced water deficiency, plants have developed several strategies to reduce leaf water loss such as reducing leaf transpiration and leaf hydraulic conductance (Defraeye et al., 2014;Kosma et al., 2009;Pou et al., 2013).

DTGs involved in the regulation of leaf hydraulic conductance
Leaf hydraulic resistance contributes 30% or more to the whole plant hydraulic resistance (Pou et al., 2013). Water transport in leaves includes an apoplastic and a cell-to-cell pathway (Pou et al., 2013). The cell-to-cell pathway is mainly triggered by water channels (aquaporins). Transcriptional changes in several water channels were observed in Phase 0 and Phase 1 ( Table 2). Two aquaporin genes were down-influenced in Phase 0; aquaporin PIP2-7, located in the plasma membrane had 2.2-fold lower transcript abundance compared to controls; aquaporin TIP2-1, located in tonoplast membrane had 4.0-fold lower transcript levels than under control conditions. One aquaporin gene, PIP1D, was down-influenced both in Phase 0 and Phase 1, and the transcript levels were 2.1 and 2.0-fold lower than controls, respectively. Nevertheless, two aquaporin genes (PIP2A and TIP1-3) were up-influenced in Phase 1. The transcription level of PIP2A was 2.0-fold higher than that in controls, and TIP1-3 was 6.0-fold higher.

Candidate marker genes for phase 0 and phase 1 in Arabidopsis thaliana
In Phase 0, 16 highly induced or inhibited DTGs (|log 2 FoldChange (Phase 0/Control)| ≥ 2) were identified and these are listed in Table 3.  Table 1 Genes related to stomatal movement differentially transcribed in Phase 0 and Phase 1. Pink indicates genes involved in ABA pathway; yellow indicates genes affecting ion channels (Ca 2+ and K + ) localized in plasma membrane and tonoplast; green indicates genes influencing the dynamic changes of apoplast and tonoplast in guard cells (e.g. volumetric elastic modulus and vacuole fusion); purple indicates genes affecting low molecular weight compounds (e.g. inositol hexakisphosphate and sphingosine-1-phosphate); blue indicates genes correlated with H + -ATPase.
(continued on next page) W. Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245 The transcript levels of AT4G28140, ERF053, and AHK5 were 14.6, 6.9, and 6.2-fold higher than controls, and they are all involved in the regulation of stomatal movement. TIP2-1, a tonoplast water channel, was 4.0-fold lower compared to controls in transcriptional level. In addition, these four genes showed no significant transcript changes in Phase 1. In Phase 1, 33 DTGs (|log 2 FoldChange (Phase 1/Control)| ≥ 5) were observed (Table 4). Among these, UF3GT, LDOX, TT8, GSTF12, SWEET15, and AT5G62360 were selected as marker genes based on the transcriptional changes. The transcriptional levels of UF3GT, LDOX, TT8, and GSTF12 were 77, 70, 64, 37-fold higher in Phase 1 compared to the control. These four genes are involved in anthocyanin metabolism. The transcript level of SWEET15 was 64-fold higher than the control and AT5G62360 showed 141-fold decrease on transcript abundance compared to controls. SWEET15 and AT5G62360 are involved in sucrose metabolism.
To evaluate the selected Phase 0 marker gene candidates, qRT-PCR was conducted in samples harvested after 1 h, 2 h, and 3 h of 105 mM  Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245 NaCl shock. The qRT-PCR results show that the transcriptional level of AT4G28140, AHK5, and TIP2-1 had no significant differences with the elongated treated time (Fig. 5A). The transcription of ERF053 increased after 1 h, 2 h and 3 h of 105 mM NaCl shock, but the fold changes were getting smaller with the prolonged treatment time (Fig. 5A).
To evaluate the Phase 1 marker gene candidates, Arabidopsis samples harvested after 1 day, 2 days, and 3 days of 90 mM NaCl treatment (stepwise acclimated, 15 mM NaCl per day) were analyzed using qRT-PCR. The absolute value of fold change at transcriptional level of all these Phase 1 candidates (except GSTF12) increased with the elongated treating time (Fig. 5B). The absolute value of log 2 FoldChange ≥1 was used as cutoff to evaluate significant differences in gene transcription. All these Phase 0 and Phase 1 marker candidates were beyond the cutoff at transcriptional level after 1 h, 2 h, and 3 h of 105 mM NaCl shock or 1 day, 2 days, and 3 days of 90 mM NaCl treatment (Fig. 5), indicating high reliability of the suggested marker gene candidates.

Marker genes tested in Zea mays
To evaluate the applicability of the selected marker genes, qRT-PCR was conducted in an important crop plant, Zea mays. The Phase 0 samples were harvested after 1 h, 2 h, and 3 h of 200 mM NaCl shock. Likewise, the Phase 1 samples were harvested after 1 day, 2 days, and 3 days of 100 mM NaCl treatment (4 days acclimation: 25 mM NaCl added daily). Using the amino acid sequences of the suggested marker genes (Arabidopsis thaliana) as templates, a BLAST was conducted in Zea mays database (B73 RefGen_v4). Analogs with the highest similarity were selected as the marker gene candidates in Zea mays, and details of these analogs are presented in Table S3. The relative transcript levels ≥2 or ≤0.5 were used as cutoff to evaluate the significant up-regulated or down-regulated genes, respectively.
In Phase 0, the transcript levels of all the Phase 0 candidates in maize were not in the range ≥2 or ≤0.5 after 1 h of 200 mM NaCl shock (Fig. 6A). After 2 h of 200 mM NaCl shock, the transcript levels of GRMZM2G141638, GRMZM2G125023, and GRMZM2G027098 exceeded the cutoff (Fig. 6A). The transcript levels of GRMZM2G141638, GRMZM2G025579, and GRMZM2G125023 were in the range ≥2 or ≤0.5 after 3 h of 200 mM NaCl shock (Fig. 6A). These results indicate that GRMZM2G141638 and GRMZM2G125023 whose transcript levels exceeded the cutoff after 2 h and 3 h of 200 mM NaCl shock are suitable Phase 0 marker genes for maize.

Turgor decrease in phase 0 and maintenance in phase 1
According to Fig. 1A and Supplemental Fig. S3, Arabidopsis cannot endure 200 mM and 125 mM NaCl shocks. First, the P p value could not recover after the peak which indicates that Arabidopsis had lost the ability to adjust turgor. Second, the diurnal change of P p was abolished in the following days. Generally, turgor decreased during the day because of transpiration and was restored at night. As P p is inversely correlated with turgor, P p increased during the day and recovered at night under normal conditions. The irregular diurnal changes of P p in Arabidopsis indicate that plants cannot recover from salt shocks induced by 200 mM and 125 mM NaCl. In contrast, maize plants can endure 200 mM NaCl shock and Arabidopsis plants can recover from the 105 mM NaCl shock. Under proper salt shock treatments, the P p value showed transient increase in the first few minutes and recovered within 1 day both in Arabidopsis and maize plants ( Fig. 1B and C). This indicates that turgor pressure decreased rapidly at first, and then recovered both in Arabidopsis and maize during Phase 0. This is consistent with the results obtained in barley and maize root tissue (Pritchard et al., 1996;Thiel et al., 1988).
During Phase 1 (salt stress after stepwise acclimation), Arabidopsis could not tolerate 100 mM NaCl treatment (25 mM increments per day) because the P m value kept increasing before and after NaCl addition was stopped ( Fig. 2A). This indicates that turgor decreased after 100 mM NaCl treatment. In contrast, turgor in Arabidopsis almost remained constant during and after the addition of 90 mM NaCl (15 mM increments per day) (Fig. 2B). Moreover, turgor in maize increased during NaCl adaptation (100 mM; 25 mM increments per day) and was maintained after stopping addition (Fig. 2C). The maintained turgor observed in Arabidopsis and maize under proper salt stress is in agreement with studies conducted in barley leaves with different levels of NaCl treatments (Thiel et al., 1988), maize root tissues under osmotic stress (Pritchard et al., 1996), and maize leaves with stepwise-acclimated salt stress (De Costa et al., 2007). These studies demonstrate that the decreased osmotic potential contributes to the turgor recovery. Moreover, the different responses of turgor to NaCl application between Arabidopsis and maize may be attributed to their different growth rate. According to Hanway (1963), maize plants with four or six fully emerged leaves are in the stage 1 or 1.5, respectively. In this study, maize plants developed five leaves, when the ZIM YARA water sensors were first applied. So the tested 4th leaves were still growing, undergoing cell expansion. This result agrees with the increased turgor pressure observed in expanding tissues by Thiel et al. (1988) and Pritchard et al. (1996).
In conclusion, the turgor measurements conducted in Arabidopsis and maize indicate that these two plant species have different capacity of salt acclimation. Moreover, salts need to be carefully applied under experimental conditions to induce salt shock or stepwise acclimation to salt stress. As shown in (Figs. 1), 125 mM and 200 mM NaCl shocks were too strong for Arabidopsis plants to survive. However, the NaCl concentration used in many Arabidopsis salinity researches exceeded 125 mM. To induce stepwise acclimation to salt stress, the daily increment of NaCl differs between Arabidopsis and maize. For Arabidopsis, the daily increment is 15 mM NaCl; and for maize, it is 25 mM NaCl. However, this is not always the case, and it may differ for different Arabidopsis ecotypes and maize cultivars.

Growth response to salt stress
After 2 days of 90 mM NaCl treatment (stepwise acclimated as described in Materials and Methods), the shoot fresh weight of Arabidopsis plants decreased (Fig. 3B), which indicates that the water uptake and transport were inhibited during this period. However, shoot dry weight had no significant difference after salt treatment (Fig. 3C). This indicates that the photosynthesis was not inhibited and the carbohydrate supply was sufficient in this period. The Na + concentration in shoots was around 20 mg/g on dry matter basis after salt treatment (Fig. 3D). As far as we know, there is no literature demonstrating the Na + toxic level for Arabidopsis thaliana. Furthermore, Arabidopsis plants did not show any toxicity and nutrient deficiency symptoms during this period. The turgor pressure of Arabidopsis plants was maintained after 2 days of 90 mM NaCl treatment (Fig. 2B). Thus, we demonstrate that Arabidopsis plants were still in Phase 1 rather than Phase 2 after 2 days of 90 mM NaCl treatment. However, we cannot exclude that genes related to ion toxicity (e.g. genes involved in Na + exclusion and Na + compartmentation; Phase 2) may also have responded in Phase 1. Therefore, the following analyses based on RNA-W. Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245 sequence were mainly focusing on the genes related to plant water relations (e.g. water transport and transpiration) to avoid the genes related to Phase 2 getting involved.

The inhibition of leaf water loss
The genes involved in the regulation of stomatal closure were mostly affected in Phase 1 (Table 1). Plants regulate stomatal movement in different ways. Among those, ABA (abscisic acid)-induced stomatal closure is the most functional strategy and has been well studied (Kim et al., 2010). One-third of the genes listed in Table 1 regulate stomatal movement through the ABA pathway, including ABI1, HAI-1, RD20, PP2CA, AKS1, ABI2, RZPF34, BGLU18, RBOHF, and BHLH128. Apart from the ABA pathway, plants can mediate stomatal movement via ion channels especially through Ca 2+ channels (e.g. PCaP1, ArathEULS3, GLR2, PHOT1, and CRY2). Moreover, EXPA1 and SYP22 take part in stomatal opening through influencing the volumetric elastic modulus and vacuole fusion, respectively. ABCC4, APK1B, and ELF3 showed in Table 1 are involved in the regulation of H + -ATPase activity which is a key regulator of stomatal opening. TOD1 and ABCC5 participate in stomatal movement by influencing the inositol hexakisphosphate and sphingosine-1-phosphate metabolic processes, respectively. Table 2 shows that genes related to wax biosynthesis were mostly influenced in Phase 1 at transcriptional level especially the eceriferum family genes (e.g. CER1, CER3, CER4, CER8, and CER60). CER1 and CER3 both encode fatty acid hydroxylase superfamily proteins. Overexpressing CER1 in Arabidopsis increased the odd-carbon-numbered alkanes and reduced soil water deficit susceptibility (Bourdenx et al., 2011). CER4 encodes an alcohol-forming fatty acyl-CoA reductase. Arabidopsis cer4 mutants displayed a decreased primary alcohols and wax esters in stem, indicating that CER4 was involved in cuticular wax biosynthesis (Rowland et al., 2006). CER8 encodes an acyl-CoA synthetase; the Arabidopsis cer8 mutants reduced the accumulation of wax in the stem and leaf, indicating that CER8 was involved in cuticular wax and cutin biosynthesis . CER60 encodes a member of the 3-ketoacyl-CoA synthase family which was involved in the biosynthesis of very long chain fatty acids (Trenkamp et al., 2004). The aquaporin activity showed contrasting transcription patterns between Phase 0 and Phase 1 ( Table 2). The salinity-induced short-term down-influenced and long-term up-influenced aquaporins are consistent with the findings in Arabidopsis (Jang et al., 2004) and rice (Kawasaki et al., 2001). Moreover, Hill et al. (2004) suggested that the initially down-regulated aquaporins may contribute to decreased water loss in the early stages of salt stress, and the later up-influenced aquaporins may play a role in the subsequent water uptake to maintain water homeostasis.  Table 3 Differentially transcribed genes with statistically significant changes in Phase 0. False discovery rate (FDR) ≤ 0.05 and the maximum value of |log 2 FoldChange (Phase 0/Control)| ≥ 2 were used as cut-off to evaluate significant differences in transcription. △ indicates marker gene candidates related to stomatal movement. ▲ indicates marker gene candidates related to water transportation.  Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245

Marker genes for phase 0 and phase 1
According to the RNA-sequence analysis in Arabidopsis plants, AT4G28140, ERF053, AHK5, and TIP2-1 were suggested as marker genes for Phase 0. First, they were all differentially and highly influenced at transcriptional level in Phase 0 and had no fundamental changes in Phase 1 (Table 3). Second, they are all involved in the inhibition of leaf water loss according to previous studies. AT4G28140 and ERF053 belong to the ERF/AP2 transcription factor family. Overexpressing AtERF053 in Arabidopsis leads to more sensitive stomatal behavior and greater drought tolerance (Cheng et al., 2012). The role of AT4G28140 in stomatal movement requires further investigation. AHK5 encodes a member of the histidine kinase family and mediates stomatal closure in an ABA-independent way (Desikan et al., 2008). TIP2-1 encodes a tonoplast water channel. Leitão et al. (2014) demonstrated that the grapevine TIP2-1 might function as a volume regulator to mediate vacuolar size and shape in response to water deficiency.
The highly and differentially influenced UF3GT, LDOX, TT8, GSTF12, SWEET15, and AT5G62360 were selected as markers for Phase 1 (Table 4). According to the previous studies, UF3GT, LDOX, TT8, and GSTF12 are involved in anthocyanin modification, biosynthesis, and transportation, respectively. Chalker-Scott (2002) demonstrated that anthocyanin is functioning as an osmolyte under salinity and drought stress to fight against osmotic stress. The UDP-glucosyltransferase (UGT) and leucoanthocyanidin dioxygenase (LDOX) proteins are key enzymes mediating biosynthesis of anthocyanin. UF3GT (also called UGT79B1) is a homologous UDP-glucosyltransferase like UGT79B2. Over-expressing UGT79B2 in Arabidopsis leads to anthocyanin accumulation and enhances resistance to drought and salt stresses (Li et al., 2017). Yeast two-and three-hybrid assays indicate that TT8, TT2, and TTG1 work together as a ternary complex to regulate proanthocyanin and anthocyanin biosynthesis (Kitamura et al., 2004). GSTF12 (also called TT19) is a carrier to transport anthocyanins from cytosol to tonoplast (Kitamura et al., 2004). SWEET15 and AT5G62360 are involved in sucrose transport and synthesis, respectively. Sucrose can act as an osmolyte to reduce the intracellular osmotic potential and prevent damage to proteins and cellular structures from salt stress (Deinlein et al., 2014). SWEET15 encodes a member of sucrose uniporter, involved in the transportation of sucrose from source to sink. It was demonstrated that AtSWEET15 is induced under osmotic stress, and alleviates damages by mediating the senescence process (Seo et al., 2011). AT5G62360 encodes a plant invertase/pectin methylesterase inhibitor, and its transcription was inhibited during Phase 1 (Table 4). This is supported by the findings of Koh et al. (2008), who demonstrated that ABA enhanced invertase activity by down-regulating its inhibitor in water-stressed plants.
These Phase 0 and Phase 1 marker gene candidates were further tested in Arabidopsis samples harvested at different time points of Phase 0 (1 h, 2 h, and 3 h of 105 mM NaCl shock) and Phase 1 (1 day, 2 days, and 3 days of 90 mM NaCl treatment) via qRT-PCR. All these marker candidates showed significant differences at transcriptional level in different Phase 0 and Phase 1 time points (Fig. 5), indicating that these 10 suggested candidates are suitable marker genes for the differentiation between Phase 0 and Phase 1 in Arabidopsis plants.
To evaluate the reliability of the marker genes, they were further tested in Zea mays (a salt-resistant maize cultivar Pioneer 3906 was used). The qRT-PCR results indicate that analogs of ERF053 and TIP2-1 are potential marker genes for Phase 0; likewise, UF3GT, GSTF12, and SWEET 15 are marker genes for Phase 1 in maize (Fig. 6). In conclusion, genes or analogs of ERF053, TIP2-1, UF3GT, GSTF12, and SWEET 15 that are affected in Arabidopsis and maize, can serve as potential Table 4 Differentially transcribed genes with statistically significant changes in Phase 1. False discovery rate (FDR) ≤ 0.05 and the maximum value of |log 2 FoldChange (Phase 1/Control)| ≥ 5 were used as cut-off to evaluate significant differences in transcription. △ indicates marker gene candidates related to anthocyanin biosynthesis and transport. ▲ indicates marker gene candidates related to sucrose synthesis and transport.  Zhao, et al. Plant Physiology and Biochemistry 143 (2019) 232-245 markers to differentiate between Phase 0 and Phase 1 in future salinity research. The transcriptional changes of marker genes in maize were not as strong as those observed in Arabidopsis under salt stress. This may be attributed to the different salt sensitivity of Arabidopsis and maize plants. The ecotype of Arabidopsis thaliana used in this study is Columbia-0, which is a relative salt-sensitive ecotype. The maize cultivar Pioneer 3906 used in this study is salt-resistant, which has strong Na + -exclusion ability. It was developed by crossing Pioneer 165 and Pioneer 605 inbred lines, which are efficient in Na + exclusion from the root and shoot, respectively (Schubert et al., 2009). As shown in Fig. 1 C, maize plants can endure 200 mM NaCl shock and the P m value recovered to the original level on the following day. However, Arabidopsis plants cannot live through 200 mM NaCl shock (Fig. 1 A). Though Arabidopsis can endure 105 mM NaCl shock, the P m value in Arabidopsis plants cannot completely recover on the next day (Fig. 1 B). These results indicate that Arabidopsis plants are more sensitive to salt stress than maize plants. The different capacity of salt acclimation between Arabidopsis and maize further validates the reliability of marker genes.

Conclusions
As salt accumulates gradually in the field, it is more conducive to study salt stress in a stepwise-acclimated way rather than salt shock. Usually plants show wilting symptoms in salt-shock situations, but there are exceptions such as in Arabidopsis and maize under the experimental conditions described in this study. In this case, it is difficult to differentiate these two types of salt stresses in terms of phenotype. In this study, two potential parameters are provided. At the physiological level, our results show that turgor decreased rapidly in a salt-shock situation, but was maintained under stepwise-acclimated salt stress. According to the turgor measurements, we suggest that salts need to be carefully applied both in salt shock and stepwise-acclimated salt stress Fig. 5. Based on the RNA-sequence analysis, the selected Phase 0 and Phase 1 marker gene candidates were further tested via qRT-PCR. Five-week-old Arabidopsis plants were treated with 105 mM NaCl shock and harvested after 1 h, 2 h, and 3 h treatments to test the Phase 0 marker gene candidates (A). Five-week-old Arabidopsis plants were stepwise acclimated to 90 mM NaCl (15 mM per day) and harvested after 1 day, 2 days, and 3 days treatments under 90 mM NaCl to test Phase 1 marker gene candidates (B). The y-axis values are the log 2 FoldChange of transcript level after salt treatments in comparison to the control. Error bars represent standard errors (SE). Significant differences of each gene's relative transcription after 2 h and 3 h of 105 mM NaCl shock or 2 days and 3 days of 90 mM NaCl treatment in comparison to the transcript level after 1 h of 105 mM NaCl or 1 day of 90 mM NaCl treatment are indicated by asterisks (*P < 5%, **P < 1%, ***P < 0.1%, ****P < 0.01%, t-test). |log 2 FoldChange| ≥ 1 was used as cutoff to evaluate differentially transcribed genes. Red lines indicate cutoff points.
conditions, otherwise plants cannot recover from these stresses. At the transcriptional level, genes or analogs of ERF053, TIP2-1, UF3GT, GSTF12, and SWEET 15 are suggested as marker genes to separate these two types of salt stress in Arabidopsis and maize. These marker genes were first selected in Arabidopsis by means of RNA-sequence, then evaluated in Arabidopsis at different time points via qRT-PCR, and finally validated in maize plants via qRT-PCR. These experiments confirmed the reliability of the selected marker genes and we expect that these marker genes are applicable to other plants. Fig. 6. Test marker gene candidates in Zea mays. Two-week-old Pioneer 3906 plants were treated with 200 mM NaCl shock and 100 mM NaCl by stepwise application (25 mM NaCl per day), respectively. The Phase 0 samples were harvested after 1 h, 2 h, and 3 h of 200 mM NaCl shock. The Phase 1 samples were harvested after 1 day, 2 days, and 3 days of 100 mM NaCl treatment. The graphs (y-axis) indicate the x-fold transcription of the genes during Phase 0 (A) and Phase 1 (B) as compared to the control (0 mM NaCl). Data show means of four biological replicates. Error bars represent the standard errors (SE). Significant differences are indicated by asterisks (*, **, ***, **** significant differences of each gene's relative transcription during Phase 0 and Phase 1 in comparison to the control with P < 5%, 1%, 0.1%, 0.01%, respectively, t-test). The relative transcript level ≥2 or ≤0.5 were used as cutoff to evaluate significant up-regulated or down-regulated genes, respectively. Red lines indicate cutoff points. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)