Phosphoproteomic Strategy for Profiling Osmotic Stress Signaling in Arabidopsis

High salinity, low temperature, and drought cause osmotic stresses, which is a major environmental factor that affects plant productivity^1,2. Protein phosphorylation is one of the most significant post-translational modifications mediating signal perception and transduction in plant response to osmotic stress^3,4,5. SNF1-related protein kinase 2s (SnRK2s) are involved in the osmotic stress signaling⁶. Nine of ten members of the SnRK2 family show significant activation in response to osmotic stress^7,8. The snrk2.1/2/3/4/5/6/7/8/9/10 decuple (snrk2-dec) mutant having mutations in all ten SnRK2 displayed hypersensitivity to osmotic stress. In snrk2-dec mutant, the osmotic stress-induced accumulation of inositol 1,4,5-trisphosphate (IP₃), abscisic acid (ABA) biosynthesis, and gene expressions are strongly reduced, highlighting the vital role of SnRK2s in osmotic stress responses⁶. However, it is still unclear how SnRK2s kinases regulate these biological processes. Profiling the phosphoproteomic changes in response to osmotic stress is an efficient way to bridge this gap and to delineate the osmotic stress-triggered defense mechanisms in plants.

Mass spectrometry (MS) is a powerful technique for mapping plant phosphoproteome⁹. Characterization of plant phosphoproteomics, however, remain a challenge due to the dynamic range of plant proteome and the complexity of plant lysate⁴. To overcome these challenges, we developed a universal plant phosphoproteomic workflow, which eliminates unwanted interferences such as from photosynthetic pigments and secondary metabolites, and enabling the deep coverage of plant phosphoproteome¹⁰. Several phosphopeptide enrichment methods such as immobilized metal ion affinity chromatography (IMAC) and metal oxide chromatography (MOC) have been developed for enriching phosphopeptides prior to MS analysis^{11,12,13,14,15,16}. Acidic non-phosphopeptides co-purifying with phosphopeptides are the major interferences for phosphopeptide detection. Previously, we standardized the pH value and organic acid concentration of IMAC loading buffer to eliminate the binding of non-phosphopeptides, to obtain more than 90% enrichment specificity bypassing the pre-fractionation step¹¹.

Sample loss in the multi-step process of phosphopeptide enrichment and fractionation hampers the sensitivity of phosphopeptide identification and the depth of phosphoproteomic coverage. Stop-and-go-extraction tips (stage tips) are pipette tips that contain small disks to cap the end of the tip, which can be incorporated with chromatography for peptide fractionation and cleaning¹⁷. Sample loss during the stage tip procedure can be minimized by avoiding sample transfer between the tubes. We have successfully implemented stage tip in Ga³⁺-IMAC and Fe³⁺-IMAC to separate low abundant multiple phosphorylated peptides from singly phosphorylated peptides, which improved the depth of human phosphoproteome¹⁵. In addition, the use of high pH reversed-phase (Hp-RP) stage tip has demonstrated the wider coverage of human membrane proteome compared to that of strong cation exchange (SCX) and strong anion exchange (SAX) chromatography¹⁸. Therefore, integrating IMAC and Hp-RP stage tip techniques can increase plant phosphoproteome coverage with simplicity, high specificity, and high throughput. We have demonstrated that this strategy identified more than 20,000 phosphorylation sites from Arabidopsis seedlings, representing an enhanced depth of plant phosphoproteome¹⁹.

Here, we report a stage tip-based phosphoproteomic protocol for phosphoproteomic profiling in Arabidopsis. This workflow was applied to study the phosphoproteomic perturbation of wild-type and snrk2-dec mutant seedlings in response to osmotic stress. The phosphoproteomic analysis revealed the phosphorylation sites implicated in kinase activation and early osmotic stress signaling. Comparative analysis of wild-type and snrk2-dec mutant phosphoproteome data leaded the discovery of a Raf-like kinase (RAF)-SnRK2 kinase cascade which plays a key role in osmore stress signaling in high plants.

1. Sample preparation

1. Harvest the control and stress treated seedlings (1 g) in an aluminum foil and flash freeze the samples in liquid nitrogen.
   NOTE: Higher protein concentration is usually observed from two-week-old seedlings than that from mature plants. One gram of seedlings generates approximately 10 mg of protein lysate, which is enough for the MS analysis. All centrifugation steps take place at 16,000 x g in step 1.
2. Grind frozen seedlings into a fine powder using a mortar and pestle filled with liquid nitrogen.
3. Add 1 mL of lysis buffer ((6 M guanidine-HCl in 100 mM Tris-HCl, pH 8.5) with 10 mM Tris (2-carboxyethyl)phosphine hydrochloride (TCEP), 40 mM 2-chloroacetamide (CAA), protease inhibitor and phosphatase inhibitor cocktails) to the mortar and mix with the aid of pestle.
4. Transfer the plant lysate to a 1.5 mL tube and heat at 95 °C for 5 min.
5. Place the tube on ice for 10 min. Sonicate the tube on ice for 10 s, pause for 10 s, and repeat thrice.
6. Centrifuge the tube for 20 min and aliquot 150 µL of plant lysate into a 1.5 mL tube.
7. Add 600 µL of 100% methanol into the tube. Vortex and spin down the tube.
8. Add 150 µL of 100% chloroform into the tube. Vortex and spin down the tube.
9. Add 450 µL of ddH₂O into the tube. Vortex and centrifuge the tube for 3 min.
10. Discard the upper aqueous layer. Add 600 µL of 100% methanol into the tube. Centrifuge the tube for 3 min and then discard the solution.
11. Wash protein pellets with 600 µL of 100% methanol. Discard the solution and air dry the protein pellet.
12. Resuspend protein pellets into 600 µL of digestion buffer (12 mM sodium deoxycholate (SDC)/12 mM sodium lauroyl sarcosinate (SLS) in 100 mM Tris-HCl, pH 8.5). Sonicate the tube until the suspension is homogenized.
13. Measure the protein concentration using a BCA kit and adjust the concentration to 4 µg/µL with the digestion buffer. Transfer 100 µL of the lysate (400 µg proteins) into a new tube.
14. Add 292 µL of 50 mM triethylammonium bicarbonate (TEAB) and 8 µL of Lys-C (2.5 Unit/mL) to the tube. Incubate the tube at 37 °C for 3 h.
15. Add 100 µL of 50 mM TEAB with 8 µg trypsin to the tube. Incubate the tube at 37 °C for 12 h.
16. Add 25 µL of 10% trifluoroacetic acid (TFA) to the tube. Vortex and centrifuge the tube for 20 min at 4 °C.
17. Transfer the supernatant into a conditioned desalting column for desalting. Dry the eluate using a vacuum centrifuge concentrator.
    NOTE: Activate the resin with 1 mL of methanol followed by 1 mL of 0.1% TFA in 80% acetonitrile (ACN). Wash out the organic solvent with at least 3 mL of 0.1% TFA in 5% ACN. Load acidified peptide samples prepared in step 1.16 into the column and then wash the column with at least 3 mL of 0.1% TFA in 5% ACN. More washes may be needed for samples with high salt concentrations. Elute the phosphopeptides with 1 mL of 0.1% TFA in 80% ACN.

2. Tandem Mass Tag (TMT) labeling

1. Resuspend the dried peptides into 100 µL of 200 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 8.5.
2. Dissolve the TMT reagent of each channel (0.8 mg) in 40 µL of anhydrous ACN. Vortex the TMT reagent tube for 5 min and spin it down.
3. Transfer 40 µL of TMT to the sample tube and incubate for 1 h at room temperature.
   NOTE: In this experiment, the channel 126 to 128 were labeled with three biological replicates of plants without mannitol treatment, and the channel 129 to 131 were labeled with three biological replicates of plants with mannitol treatment.
4. Add 8 µL of 5% hydroxylamine and incubate for 15 min.
5. Mix 6 samples in a 5 mL tube and add 14 µL of 10% TFA.
6. Add 3,876 µL of 0.1% TFA to the tube. Vortex and spin down.
7. Transfer the solution to a conditioned desalting column for desalting. Dry the eluate using an evaporator.

3. Preparation of IMAC stage tip

1. Use a 16 G blunt-ended needle to penetrate a polypropylene frit disk.
   NOTE: Hold the cutter perpendicular to the surface of the disk and roll the cutter a couple of times to make sure that the disk is completely excised. Place the disk in a Petri dish for storage. Place the needle into a 200 µL pipette tip and push the frit into the tip using a plunger.
2. Press the frit gently into the tip using a plunger to cap the end of tip.
   1. Place the frit disk in tips with the same pressure, which provides better reproducibility. The reproducibility of stage tips production is important for constant back pressure, which affects the timing of the centrifuge steps and reproducibility between technical replicates.
   2. If the stage tips show high back pressure during conditioning step, discard the tips to avoid potential tip clogging when while loading samples. It could be that the disk might have been pressed too hard into position.
3. Invert Ni-NTA spin column and place on a 1.5 mL tube.
4. Use a plunger to press the frit of Ni-NTA beads gently and push the beads into the tube.
   NOTE: Ensure to push the frit gently to avoid the potential bead loss or adsorption on the spin column.

4. Preparation of Hp-RP stage tip

1. Prepare Hp-RP stage tip as described for IMAC stage tip using a C8 disk.
   NOTE: Do not apply large force to the disk because it may result in a densely packed frit and increase the back pressure of the stage tip. All centrifugation steps take place at 1,000 x g for 5 min in step 4.
2. Suspend 1 mg of C18 beads in 100 µL 100% methanol and pass the beads solution through the tip.
3. Add 20 µL of buffer 8 (80% ACN/20% 200 mM NH₄HCO₂, pH 10.0) to the stage tip and pass buffer 8 through the tip by centrifugation.
4. Add 20 µL of Buffer A (200 mM NH₄HCO₂) to the stage tip and pass buffer A through the tip by centrifugation.
   NOTE: Balance the centrifuge during use and ensure that the Hp-RP stage tip is not completely dried.

5. Preparation of spin adaptor

1. Use sharp end tweezers to puncture a hole at the center of the lid of a 1.5 mL tube.
2. Insert the IMAC stage tip or Hp-RP stage tip into the hole of the spin adaptor.
   NOTE: Ensure that the stage tip is not close to the bottom of spin adaptor.

6. Phosphopeptide enrichment using IMAC stage tip

1. Suspend the 10 mg Ni-NTA beads with 400 µL of loading buffer (6% acetic acid (AA), pH 3.0) and load all of the beads solution into per tip. Pass the solution through the tip by centrifugation.
   NOTE: All centrifugation steps take place at 200 x g for 3 min in step 6 besides step 6.8.
2. Load 100 µL of 50 mM ethylenediaminetetraacetic acid (EDTA) to strip the nickel ions from IMAC stage tip and pass the solution through the tip by centrifugation.
3. Load 100 µL of loading buffer to the stage tip and pass the solution through the tip by centrifugation.
4. Load 100 µL of 50 mM FeCl₃ in 6% AA to the stage tip and pass the solution through the tip by centrifugation. Fe³⁺ ions will chelate to IMAC stage tip.
5. Load 100 µL of loading buffer to condition the stage tip and pass the solution through the tip by centrifugation.
6. Load 100 µL of loading buffer with sample peptides prepared in step 2.7 to the stage tip and pass the solution through the tip by centrifugation.
   NOTE: Take a new tube as a spin adaptor to collect the flow-through of sample and store the flow-through in the freezer for future analysis.
7. Load 100 µL of washing buffer (4.5% AA and 25% ACN) to the stage tip and pass the solution through the tip by centrifugation.
8. Perform the second wash with loading buffer. Load 100 µL of loading buffer to the stage tip and pass the solution through the tip using 1000 × g centrifuge for 3 min.
   NOTE: Ensure the washing buffer is replaced by loading buffer in the stage tip to eliminate the loss of phosphopeptides during elution step. Equilibrate the IMAC stage tip prior to phosphopeptide elution to ensure the concentration of ACN is below 5%.
9. Trim the IMAC stage tip at the front with a scissors and place the trimmed IMAC stage tip inside the Hp-RP tip. Ensure that the two layers of stage tips do not touch the lid of the centrifuge.

7. Phosphopeptide fractionation using Hp-RP C18 stage tip

1. Add 100 µL of elution buffer (200 mM ammonium phosphate (NH₄H₂PO₄)) to the trimmed IMAC stage tip and pass the solution through the two layers of stage tips by centrifugation.
   NOTE: If the solution does not pass through the stage tip, increase the speed of spin down. All centrifugation steps take place at 1,000 x g for 5 min in step 7. All the Hp-RP buffers are listed in Table 1.
2. Discard the IMAC stage tip using a tweezers and add 20 µL of Buffer A to the Hp-RP stage tip and pass the buffer through the tip by centrifugation.
   NOTE: Ensure all the elution buffer pass through the IMAC stage tip before discarding it.
3. Add 20 µL of Buffer 1 to elute fraction 1 and collect the eluate in a new tube by centrifugation. Repeat the process with Buffers 2, 3, 4, 5, 6, 7, and 8 to collect each fraction in a new tube. Dry the final eluate of each fraction using a vacuum concentrator.
   NOTE: Prepare fresh fractionation buffers. Take 8 new tubes as the spin adaptor of each fraction. Collect the eluate of each fraction in a different spin adaptor. Phosphopeptides are not stable under basic conditions, so dry the eluates by a concentrator or acidified by 10% TFA immediately.

8. LC-MS/MS analysis and data analysis

1. Add 5 µL of 0.1% formic acid (FA) and analyze the sample by a mass spectrometer.
2. Run a 90 min gradient with 6-30% buffer B (80% ACN and 0.1% FA) for each fraction.
3. Fragment the top 10 labeled peptides using high collision energy dissociation (HCD). Complete a database search to identify phosphorylation sites.
4. Load the raw files into the MaxQuant software, name the experiments, and set fractions and PTM.
   NOTE: The tutorial of MaxQuant software can be found at https://www.maxquant.org/.
5. Select reporter ion MS2 in the type of LC-MS/MS run and 6plex TMT as isobaric labels. Enable filter by PIF function and set the min PIF as 0.75.
6. Select Acetyl (Protein N-term), Oxidation (M), and Phospho (STY) to the panel of variable modifications. Select Carbamidomethyl (C) as fixed modifications.
7. Set digestion mode as specific, select Trypsin/P as digestion enzyme, and two missed cleavage.
8. Set PSM false discovery rate (FDR) and protein FDR as 0.01. Set the minimum score for modified peptides as 40.
9. Add Arabidopsis thaliana database to the panel of fasta files and run database searching.
10. Load the txt file of Phospho (STY) sites to the Perseus software as the search is done.
    NOTE: The detailed tutorials of Perseus software can be found at https://www.maxquant.org/perseus/.
11. Filter out the reverse phosphorylation sites. Filter out the unlocalized phosphorylation sites using localization probability 75% as the cut-off.
12. Use the intensities of phosphorylation sites for statistical analysis.

To demonstrate the performance of this workflow, we exploited IMAC stage tip coupled with Hp-RP stage tip fractionation to measure the phosphoproteomic changes in wild-type and snrk2-dec mutant seedlings with or without mannitol treatment for 30 minutes. Each sample was performed in biological triplicates, and the experimental workflow is represented in Figure 1. The digested peptides (400 µg) of each sample were labeled with one TMT-6plex channel, pooled and desalted. The phosphopeptides were further enriched using an IMAC stage tip, and the purified phosphopeptides were subsequently fractionated into eight fractions by a Hp-RP stage tip. Each fraction was analyzed by a 90 min LC gradient analysis. The raw files were searched using a search engine against Arabidopsis thaliana database.

A total of 11,077 unique phosphopeptides were identified corresponding to 3,630 phosphoproteins with 6,852 localized phosphorylation sites (Class I, localization probability > 0.75), indicating the wide coverage of Arabidopsis phosphoproteome. A total of 8,107 and 7,248 phosphopeptides were identified from wild-type and snrk2-dec mutant sample, respectively. This illustrates the efficiency of the workflow in providing in-depth coverage for delineating the global view of signal transduction in Arabidopsis. We compared the number of identified phosphopeptides across 8 fractions. A few phosphopeptides were identified in the first two fractions, the fraction 1 and 2. However, majority of phosphopeptides were evenly distributed in the rest 6 fractions (Figure 2A), suggesting this approach provides the capability to separate complex phosphopeptides from the plant phosphoproteome. To further demonstrate the separation efficiency of this workflow, we evaluated the overlap of phosphopeptides between two adjacent fractions (eq. F1-to-F2). Less than 5% phosphopeptides overlap in the adjacent fractions, indicating robust fractionation efficiency of the Hp-RP stage tip (Figure 2B).

Using the data obtained by the workflow, we compared the phosphoproteomic profiles of wild-type and snrk2-dec mutant upon mannitol treatment. A total of 433 and 380 phosphorylation sites were increased after mannitol treatment in wild-type and snrk2-dec mutant sample, respectively (Figure 3). Among that, 312 phosphosites showed induction (FDR < 0.01) in wild-type, but not in snrk2-dec mutant plants. The Gene Ontology (GO) analysis revealed that the function of phosphorylation and activation of protein kinase, regulation of phosphate metabolic process, signal transduction, are significantly enriched in the SnRK2-dependent phosphoproteins. Interestingly, GO term related to root development was also enriched in SnRK2-dependent group, consistent with the phenotype of root growth retardation under osmotic stress⁶. We also identified 116 phosphosites up-regulated by mannitol treatment in both wild-type and snrk2-dec mutant. These phosphoproteins were independent of SnRK2, or candidates that mediate osmotic stress-triggered signaling prior to SnRK2s activation. We observed that several B4 subgroup RAFs such as RAF18 (AT1G16270), RAF24 (AT2G35050), and RAF42 (AT3G46920), were significantly up-regulated by osmotic stress. Further study revealed that RAF kinases are quickly activated by osmotic stress and required for phosphorylation and activation of SnRK2s²⁰.

Figure 1: Workflow of stage tip-based phosphoproteomic method. Protein was extracted and digested from wild-type and snrk2-dec mutant seedlings treated with or without mannitol. The digested peptides of each replicate were labeled with a unique TMT6-plex channel. Phosphopeptides were pooled and then enriched using an IMAC stage tip. The purified phosphopeptides were separated by a Hp-RP stage tip. Each fraction was analyzed by mass spectrometer. Please click here to view a larger version of this figure.

Figure 2: Phosphoproteomic profiling of wild-type and snrk2 decuple mutant seedlings by IMAC stage tip and Hp-RP fractionation. (A) The number of identified phosphopeptides per fraction. (B) The separation efficiency of the stage tip-based Hp-RP chromatography. The overlap between the adjacent fractions is represented by the percentage of the same peptide identified in the adjacent fractions. Please click here to view a larger version of this figure.

Figure 3: Quantification of the phosphoproteomic changes in response to osmotic stress. Volcano plots show the log2 fold change of phosphorylation sites in (A) wild-type and (B) snrk2-dec mutant seedlings in response to mannitol treatment. The black circle represents the kinase phosphorylation sites induced by mannitol. The red circle indicates the phosphorylation sites of RAFs up-regulated in response to mannitol. Please click here to view a larger version of this figure.

Table 1: Buffers for Hp-RP stage tip fractionation.

The dynamic range and complexity of plant proteome and phosphoproteome are still a limiting factor to depth of phosphoproteomics analyses. Despite the capability of single run LC-MS/MS analysis to identify 10,000 phosphorylation sites^21,22, the coverage of the whole plant phosphoproteome is still limited. Therefore, a phosphoproteomic workflow that provides high sensitivity and superior separation efficiency is required in profiling the global view of plant signaling networks in response to environmental stress. Commercial high-pressure liquid chromatography (HPLC) column-based chromatography is a common method for reducing the peptide complexity prior to MS analysis^23,24. However, tedious sample collection steps and large elution volume are the major challenges of HPLC-based methods in terms of sensitivity and throughput. stage tip is an alternative approach to execute peptide pre-fractionation or enrichment for high-sensitivity and high-throughput works. This new phosphoproteomic workflow exploiting isobaric labeling coupled with phosphopeptide enrichment and fractionation provides better coverage and depth of plant phosphoproteomics over other phosphoproteomic workflows^25,26.

The pH value of samples is the crucial factor that determines the enrichment specificity of phosphopeptides. The pH value may be affected by the previous step of sample preparation, so it is essential to check the pH value before sample loading. If the pH is shifted, adding acetic acid or sodium hydroxide to samples to adjust the pH to 3.0. We execute this protocol for simultaneous elution and loading of phosphopeptides onto C18 stage tip. Therefore, it is important to equilibrate the IMAC stage tip prior to phosphopeptide elution to ensure the concentration of ACN is below 5%.

The number of fractions is dependent on the complexity and sample sizes. Typically, 5 to 8 fractions are enough to provide broad coverage of phosphopeptide identification in plants. The concentration of ACN in fractionation buffers can be adjusted according to the hydrophobicity of samples. Most of the phosphopeptides are eluted from C18 beads using the range of 5% to 25% ACN concentration. Phosphopeptides typically are more hydrophilic compared to non-phosphorylated peptides due to the additional negative charges of the phosphate group. However, tagging TMT reagent on N-terminus of peptide leads to an increase of hydrophobicity of labeled peptides²⁷, that may explain why fewer phosphopeptides were identified in the first two fractions in our case (Figure 2A). Thus, a test using a TMT-zero reagent and an aliquot of samples can be used to evaluate the number of fractions and the ACN concentration in each fractionation buffers. The optimized buffers may result in better separation efficiency of TMT-6plex labeled phosphopeptides.

The limitations of stage tip-based fractionation are the number of fractions and the capacity of tips. It is difficult to expand the number of fractions in stage tip separation because discontinuous gradient buffers are used for stage tip fractionation. On the other hand, a continuous gradient can be applied to HPLC columns to achieve a high fraction number. The capacity of tips is another factor that limits the application of stage tips. For large-scale phosphoproteomics analysis (> 10 mg proteins) it is better to use HPLC-based enrichment with longer column length packed with more C18 beads and fractionation, which is beyond the analytical scale of stage tips. Taken together, stage tip-based phosphopeptide enrichment and fractionation is a useful method for small to medium scale of phosphoproteomics analyses. This approach can be integrated with isobaric labeling to achieve multiplexed quantification. The results also demonstrated that it could be used for in-depth plant phosphoproteomics profiling and quantification. This workflow is applicable to other species and different tissue types for the delineation of specialized signaling networks.