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. 2023 Oct 25;22(3):650–661. doi: 10.1111/pbi.14212

Differential heat‐response characteristics of two plastid isoforms of triose phosphate isomerase in tomato

Chong Chen 1,2, † , Meng Zhang 1, † , Xiaocui Ma 3, Qingwei Meng 1, Kunyang Zhuang 1,✉
PMCID: PMC10893939  PMID: 37878418

Summary

Heat stress causes dysfunction of the carbon‐assimilation metabolism. As a member of Calvin–Benson–Bassham (CBB) cycle, the chloroplast triose phosphate isomerases (TPI) catalyse the interconversion of glyceraldehyde 3‐phosphate (GAP) and dihydroxyacetone phosphate (DHAP). The tomato (Solanum lycopersicum) genome contains two individual SlTPI genes, Solyc10g054870 and Solyc01g111120, which encode the chloroplast‐located proteins SlTPI1 and SlTPI2, respectively. The tpi1 and tpi2 single mutants had no visible phenotypes, but the leaves of their double mutant lines tpi1tpi2 had obviously reduced TPI activity and displayed chlorotic variegation, dysplasic chloroplasts and lower carbon‐assimilation efficiency. In addition to altering carbon metabolism, proteomic data showed that the loss of both SlTPI1 and SlTPI2 severely affected photosystem proteins, reducing photosynthetic capacity. None of these phenotypes was evident in the tpi1 or tpi2 single mutants, suggesting that SlTPI1 and SlTPI2 are functionally redundant. However, the two proteins differed in their responses to heat stress; the protein encoded by the heat‐induced SlTPI2 showed a higher level of thermotolerance than that encoded by the heat‐suppressed SlTPI1. Notably, heat‐induced transcription factors, SlWRKY21 and SlHSFA2/7, which negatively regulated SlTPI1 expression and positively regulated SlTPI2 expression, respectively. Our findings thus reveal that SlTPI1 and SlTPI2 have different thermostabilities and expression patterns in response to heat stress, which have the potential to be applied in thermotolerance strategies in crops.

Keywords: Calvin–Benson–Bassham cycle, chloroplast triose phosphate isomerase, heat stress, photosynthesis, thermotolerance, tomato

Introduction

Chloroplasts serve as metabolic centres and play a key role in physiological processes for heat stress (HS) adaptation. Photosynthetic activity is inhibited by HS, which damages key components of metabolic processes, with concomitant reductions in biomass production and crop yield (Wang et al., 2018). As a key metabolic process in photosynthesis, the Calvin–Benson–Bassham (CBB) cycle is responsible for the fixation of CO2 into carbohydrates, as well as the assimilation, transport and utilization of photo‐assimilates. HS causes dysfunction in the carbon‐assimilation metabolism in the stroma of the chloroplasts (Sharkey, 2005).

As members of the carbon‐assimilation metabolic system, triose phosphate isomerases (TPIs) are fast, abundant and efficient enzymes involved in CBB cycle (Fridlyand, 1992; Fridlyand et al., 1999; Harris and Königer, 1997); however, the influence of HS on the TPIs requires further exploration. Plants contain both cytosolic and plastid TPIs, neither of which have been extensively characterized; because TPIs catalyse a reversible reaction [glyceraldehyde 3‐phosphate (GAP) to and from dihydroxyacetone phosphate (DHAP)], their products remain in equilibrium and are therefore presumed likely to be of regulatory importance (Chen and Thelen, 2010a). In Arabidopsis thaliana, the plastid TPI isoform (pdTPI) does not affect the expression level and enzymatic activity of the cytosolic TPI (Chen and Thelen, 2010a,b). Inhibiting pdTPI in Arabidopsis thaliana reduced TPI activity, resulting in a severely stunted and chlorotic seedling that accumulated DHAP, glycerol, glycerol‐3‐phosphate and methylglyoxal, but had reduced levels of GAP (Chen and Thelen, 2010b). The reduced number and size of chloroplasts and the shortage of carbon intermediates may account for the pale‐green colour of the pdtpi cotyledons (Chen and Thelen, 2010b); however, in pdTPI‐antisense rice (Oryza sativa), plant growth was not affected, whereas the overexpression of pdTPI only marginally improved photosynthesis to a level insufficient to improve biomass production in rice (Suzuki et al., 2022a,b). In pdTPI‐antisense rice lines, the pool sizes of both GAP and DHAP decreased to 60%–70% of those in the wild‐type (WT) plants. Decreases of a similar magnitude were observed for sedoheptulose 7‐phosphate (S7P) and fructose 6‐phosphate (F6P). Simultaneously, the pool sizes of the CBB cycle metabolites from pentose phosphate to 3‐phosphoglycerate (3‐PGA) increased by 125%–188% compared with the WT plants (Suzuki et al., 2022a). The reduced pdTPI limits GAP concentration, disturbing the carbohydrate metabolism in the chloroplast, but the effect of this on photosynthesis has not been directly analysed.

pdTPI has two splice variants (At2g21170.1 and At2g21170.2) in Arabidopsis (Chen and Thelen, 2010b). The At2g21170.2 splice variant contains a nine–amino acid deletion compared with At2g21170.1, the first three of which are highly conserved among all TPIs. Whereas the At2g21170.1 form was expressed in roots, leaves, stems, flowers and siliques, with the highest expression in leaves and flowers, At2g21170.2 expression was only weakly detected in the roots (Chen and Thelen, 2010b). The differential expression patterns for these two splice variants suggest organ‐specific roles for these gene products.

Heat stress generally impairs photosynthetic activity, changes the cell membrane lipids (which damages in membrane fluidity and permeability) and disrupts the balance between the production and scavenging of reactive oxygen species (ROS), that eventually restricting the growth and development and also reducing the yield of crops (Allakhverdiev et al., 2008; Grover et al., 2000; Hasanuzzaman et al., 2013; Niu and Xiang, 2018). Plants have developed complex and diverse systems to cope with HS; for example, HEAT SHOCK FACTOR A1 (HSFA1), a major heat response transcription factor, is indispensable in the activation of transcriptional networks under HS (Liu et al., 2011; Mishra et al., 2002; Ohama et al., 2017). In Arabidopsis thaliana, HSFA1s have been predicted to directly regulate the expression levels of genes encoding important heat‐response transcription factors (TFs), including DEHYDRATION‐RESPONSIVE ELEMENT BINDING PROTEIN 2A (DREB2A), HSFA2, HSFA7a and the HSFBs (Yoshida et al., 2011). HSFA2 is a direct target of the HSFA1s and is essential for the HS response in plants, with the hsfa2 knockout mutant displaying high sensitivity to HS due to its reduced expression of many HS‐inducible genes (Charng et al., 2007). As one of the largest families of TFs in land plants, WRKY proteins bind to a W‐box in the promoter of target genes and activate or repress the expression of downstream genes to regulate stress responses (Yamasaki et al., 2005). Some WRKY members are involved in the HS response (Jiang et al., 2017); for example, the overexpression of AtWRKY25, AtWRKY26 or AtWRKY39 enhanced HS resistance (Li et al., 2010, 2011). The specific roles of the WRKY and HSF members in the photosynthesis‐related response to HS require further investigation.

In tomato (Solanum lycopersicum), the functions of cytosolic TPI and pdTPI remain unclear. The tomato genome contains two genes, annotated as Solyc10g054870 and Solyc01g111120, that encode the chloroplast‐located proteins TPI1 and TPI2, respectively. We obtained their single and double mutants (tpi1, tpi2 and tpi1tpi2) to analyse their functions. The tpi1 and tpi2 single mutant lines had no visible phenotypes, whereas the leaves of the double mutant line tpi1tpi2, which had reduced TPI activity, showed chlorotic variegation. These findings suggest that tomato TPI1 and TPI2 are functionally redundant; however, TPI1 and TPI2 had different thermostability and expression patterns under HS. Our further analysis showed that TPI1 was suppressed by SlWRKY21, whereas TPI2 was activated by HSFA2 and HSFA7a under HS. These provide a potential strategy to protect the plant from the heat‐induced photodamage.

Results

Solyc01g111120 and Solyc10g054870 encode chloroplast‐located TPIs

The tomato genome contains four genes annotated as SlTPIs: Solyc01g111120, Solyc04g011510, Solyc06g005490 and Solyc10g054870. A protein sequence alignment showed that both Solyc10g054870 (SlTPI1) and Solyc01g111120 (SlTPI2) had a chloroplast‐signalling peptide (CTP) in their N‐terminal regions, which some of the residues are similar to the chloroplast‐located TPI (AtpdTPI) in Arabidopsis (Figure S1a). A phylogenetic analysis showed that SlTPI1 and SlTPI2 are more similar to AtpdTPI than AtcytTPI (Figure S1b). A transient expression assay in Nicotiana benthamiana further confirmed that both SlTPI1 and SlTPI2 were targeted to the chloroplasts (Figure S2). Considering the organ‐specific roles of the two splice variants of AtpdTPI, we detected the expression patterns of SlTPI1 and SlTPI2, revealing them both to be most highly expressed in the leaves and cotyledons, especially the SlTPI1 (Figure S3).

The tpi1tpi2 double mutant shows chlorotic variegation and stunted growth

To analyse the roles of SlTPI1 and SlTPI2, we generated tpi1 and tpi2 single mutant lines (tpi1‐1/2 and tpi2‐1/2) using a CRISPR/Cas9 system. These lines were identified using DNA sequencing, which revealed premature termination codons at the +212 site of the SlTPI1 coding sequence and the +47 site of the SlTPI2 coding sequence (the transcription initiation codon ATG was defined as +1; the blue letters indicated the extra added bases and * indicated the deleted bases) (Figure 1a). The chloroplast TPI activity was lower in the tpi1 and tpi2 mutants than in the WT (Figure 1b), but there were no visible differences between their phenotypes when grown at 25 °C (Figure 1c). In addition, the tpi1 and tpi2 mutants had less chlorophyll than the WT (Figure 1d). To further investigate the effect of the pdTPI deletion, we produced two double mutants by crossing tpi1‐1 to tpi2‐1 and tpi1‐2 to tpi2‐2, yielding tpi1*2‐1 and tpi1*2‐2, which were confirmed by DNA sequencing (Figure 1a). An obvious chlorotic variegation and stunted growth phenotype was observed in these double mutant lines (Figure 1c). Accordingly, the tpi1tpi2 double mutants had much lower chloroplast TPI activities and chlorophyll contents than the WT and single mutants (Figure 1b,d). These results indicate that the chloroplast TPI1 and TPI2 are functionally redundant, with the loss of both seriously affecting the function of chloroplasts.

Figure 1.

Figure 1

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Phenotypes of tpi1, tpi2 and tpi1tpi2 mutants. (a) Identification of the CRISPR/Cas9‐induced mutation in the tpi1, tpi2 and tpi1tpi2 double mutants by sequencing. The red letters indicated the termination codon at +212 site (the initiation codon ATG was defined as 0). The blue letters indicated the mutant base. (b) The chloroplast TPI activity of WT and tpi1, tpi2 and tpi1tpi2 double mutants. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05). (c) Phenotypes of WT and tpi1, tpi2 and tpi1tpi2 double mutants (5‐week‐old) under normal condition (25 °C). (d) Chlorophyll content of WT and tpi1, tpi2 and tpi1tpi2 double mutants was detected under normal conditions. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05).

Loss of the chloroplast TPIs seriously damages photosynthesis

TPIs are important members of the carbon‐assimilation metabolism system. Previous studies demonstrated that pdTPI inhibition affects the metabolic processes in the CBB in Arabidopsis and rice (Chen and Thelen, 2010a; Suzuki et al., 2022a). To explore this phenomenon in tomato, we first detected the CO2 assimilation rate in the WT, tpi1, tpi2 and tpi1tpi2 mutants. Based on the A‐Ci curves, the tpi1tpi2 double mutants had a much lower CO2 assimilation rate than the WT or the tpi1 and tpi2 single mutants (Figure 2a). In agreement with this, relative to the WT, the decreased levels of the starch, sucrose and glucose contents were significantly higher in the tpi1tpi2 double mutants than that in single mutants (Figure 2b–d). These results show that tomato chloroplast TPIs are also involved in carbon metabolism.

Figure 2.

Figure 2

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Knockout of SlTPI1 and SlTPI2 seriously affected photosynthesis. (a) A/Ci curves of the WT and tpi1, tpi2 and tpi1tpi2 double mutants under normal condition (25 °C). A, CO2 assimilation rate; Ci, intracellular CO2 concentration. Data are presented as means ± standard deviation of three biological replicates (**P < 0.01). (b–d) The starch, sucrose and glucose contents of the WT and tpi1, tpi2 and tpi1tpi2 double mutants under normal conditions. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05). (e) Chloroplast ultrastructure observation of the WT and tpi1, tpi2 and tpi1tpi2 double mutants. Bars, 1 μm. The red and yellow arrows pointed to the stacked grana and starch granules respectively. (f) BN‐PAGE analysis of chlorophyll‐protein complexes. Thylakoid membrane samples with equal concentration from the leaves of the WT and tpi1, tpi2 and tpi1tpi2 double mutants under normal condition were solubilized by treatment with 2% (w/v) DM (n‐Dodecyl‐beta‐D‐maltoside) and separated by BN‐PAGE. The assignments of macromolecular protein complexes of thylakoid membranes indicated at left. I, PSII supercomplexes; II, PSI + PSII dimer; III, PSII monomer; IV, CP43 free PSII; V, LHCII trimer; VI, LHCII monomer. (g, h) The Fv/Fm values and photosynthesis rate (Pn) of the WT and tpi1, tpi2 and tpi1tpi2 double mutants at 25 °C. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05).

To explore the effect of chloroplast TPI deletion on the photosynthetic machinery, we observed the chloroplast ultrastructure of the WT, tpi1‐1, tpi2‐1 and tpi1tpi2‐2 mutants using transmission electron microscopy. The tpi1tpi2 chloroplasts were smaller than those of the WT, tpi1‐1 and tpi2‐1 (Figure 2e). Whereas the WT, tpi1 and tpi2 chloroplasts had stacked grana and unstacked stroma thylakoid membranes, the stability of the stacked grana seemed poor in tpi1 and tpi2 (Figure 2e); by comparison, stacked grana were hardly observed in the tpi1tpi2 double mutants (Figure 2e). To further investigate the stability of the thylakoid protein complexes, we performed a blue native‐PAGE (BN‐PAGE) analysis. Notably, far fewer thylakoid protein complexes were observed in tpi1tpi2 than in the WT, tpi1‐1 and tpi2‐1, suggesting the double mutants may have a lower photosynthetic capacity (Figure 2f). The detection of the maximal photochemical efficiency of PSII in the dark (F v/F m) and the net photosynthetic rate (Pn) supports this speculation (Figure 2g,h).

To investigate the effect of the chloroplast TPI deletion on the levels of photosynthesis‐associated proteins, we performed a proteomics analysis of the leaves of the WT, tpi1‐1, tpi2‐1 and tpi1tpi2‐2. Our Venn analysis showed that tpi1tpi2‐2_vs_WT had 562 differentially expressed proteins (DEPs), whereas 598 DEPs were detected in the tpi1‐1_vs_WT comparison and 670 DEPs in tpi2‐1_vs_WT (Figure 3a). These DEPs were enriched in functions associated with the photosynthesis pathway, especially the photosystem pathway (Figure 3b), suggesting that the loss of chloroplast TPIs severely damaged the photosystem. Besides the chloroplast TPIs, a heatmap analysis of the DEPs showed that the levels of the Ribulose 1,5‐Bisphosphate Carboxylase/Oxygenase (RuBisCO) large subunit (RbcL), RuBisCO small subunits 1 and 2 (RbcS1 and RbsS2) and Glyceraldehyde‐3‐Phosphate Dehydrogenase 1 and 2 (GAPD1 and GAPD2) in the carbon‐fixation pathway were much lower in the tpi1tpi2 double mutant than the WT and single mutants (Figure 3c). In addition, the key chlorophyll biosynthesis proteins Protochlorophyllide Oxidoreductase (POR), Chlorophyllide a8‐Vinyl‐Reductase (DVR) and Chlorophyllide a Oxygenase (CAO) in tpi1tpi2 were also significantly decreased, which is the cause of its lower chlorophyll content (Figure 3c). The photosystem is the most affected pathway in the tpi1tpi2 double mutant. Many subunits of the photosynthetic complexes, including PSII (D1, D2, CP43, CP47, PsbF, PsbJ, PsbRl and PsbQ), PSI (PsaA, PsaC, PsaD, PsaE, PsaG and PsaH) and ATP synthase (AtpA, AtpB, AtpG, AtpH and AtpI), were less abundant in tpi1tpi2 than in the WT (Figure 3c). These results indicate that, at the protein level, the loss of chloroplast TPIs mainly affected the carbon‐fixation pathway through the subunits of RuBisCO, with a broader range of impacts on the photosystem pathway. In addition, the loss of SlTPI1 and SlTPI2 also caused an imbalance in the abundance of the amino acid‐ and lipid metabolism‐related proteins, which may be a consequence of the impaired photosynthesis (Figure S4).

Figure 3.

Figure 3

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Proteomics analysis of tpi1, tpi2 and tpi1*2 mutants and WT. (a) Venn diagram of the numbers of differentially expressed proteins in the tpi1‐1 versus WT, tpi2‐1 versus WT, tpi1tpi2‐2 versus WT, tpi2‐1 versus tpi1tpi2‐2 and tpi1‐1 versus tpi1tpi2‐2. (b) Gene ontology (Go) terms biological process analysis of differentially expressed proteins in tpi1‐1, tpi2‐1, tpi1tpi2‐2 and WT. (c) Expression analysis of the proteins involved in carbon fixation, photosystem and chlorophyll synthesis. Higher protein levels are shown in red (1–1.3) and lower protein levels are shown in blue (0.6–0.9).

Tomato TPI1 and TPI2 have different thermostabilities

Our previous proteomic data showed that the SlTPI1 protein decreased in WT leaves after a 12‐h heat treatment, but SlTPI2 seems more stable under HS (Figure S5). To explore the different roles of SlTPI1 and SlTPI2, we cloned their respective coding sequences and independently transformed them into pSuper1300‐GFP vectors for a transient expression assay in N. benthamiana leaves via an Agrobacterium‐mediated infiltration. After the heat treatment, we observed that the levels of SlTPI1‐GFP decreased more than those of SlTPI2‐GFP (Figure 4a). Consistent with this, the TPI activity in the leaves infiltrated with SlTPI1‐GFP decreased to a lower level than the leaves infiltrated by SlTPI2‐GFP under the heat treatment (Figure 4b). These results suggest that SlTPI2 may be more stable than SlTPI1 under HS.

Figure 4.

Figure 4

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SlTPI2 showed more thermostability than SlTPI1. (a) Western blot assay showed the SlTPI1‐GFP and SlTPI2‐GFP contents under heat treatment. LC indicated the loading control. Three biological replicates were performed in this assay. (b) The detection of TPI activity of N. benthamiana leaves expressing SlTPI1‐GFP or SlTPI2‐GFP. Experiments were conducted three times with similar results. Data are presented as means ± SD of three biological replicates (**P < 0.01). (c) RT‐qPCR analysis of SlTPI1 and SlTPI2 expression in SlTPI1 and SlTPI2 overexpression lines (TPI1‐OE and TPI2‐OE). The mRNA level was normalized to that of ACTIN. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05). (d) Phenotypes of SlTPI1 and SlTPI2 overexpression lines and WT plants (5‐week‐old) under normal conditions (25 °C) and heat stress (42 °C). (e–g) The REC values, chloroplast TPI activity and Fv/Fm of WT and SlTPI1 and SlTPI2 overexpression lines at 25 °C and 42 °C. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters indicate significant differences (P < 0.05) among tomato lines within 25 °C or 42 °C. (h) BN‐PAGE analysis of chlorophyll‐protein complexes. Thylakoid membrane samples with equal concentration from the leaves of the WT and SlTPI1 and SlTPI2 overexpression lines under normal conditions (25 °C) and heat stress (42 °C) were solubilized by treatment with 2% (w/v) DM (n‐Dodecyl‐beta‐D‐maltoside) and separated by BN‐PAGE. The assignments of macromolecular protein complexes of thylakoid membranes indicated at left. I, PSII supercomplexes; II, PSI + PSII dimer; III, PSII monomer; IV, CP43 free PSII; V, LHCII trimer; VI, LHCII monomer.

To further investigate the different thermostabilities of SlTPI1 and SlTPI2, we generated six independent tomato lines overexpressing SlTPI1 and five independent tomato lines overexpressing SlTPI2. These lines all showed higher SlTPI1 or SlTPI2 mRNA levels (Figure 4c). SlTPI1‐OE‐1, SlTPI1‐OE‐4, SlTPI2‐OE‐1 and SlTPI2‐OE‐2, all of which displayed about 40‐fold increases in TPI1 or TPI2 expression than the WT, were selected for further analysis. Under normal growth conditions (25 °C), we observed no obvious difference in the phenotypes of the TPI1‐OE lines and TPI2‐OE lines compared with the WT, suggesting that SlTPI overexpression was not sufficient to improve tomato biomass (Figure 4d). By contrast, the WT was much more severely wilted than the TPI1‐OE and TPI2‐OE lines after a heat treatment (42 °C for 24 h; Figure 4d). In agreement with this observation, the relative electrical conductivity (REC), an indicator of membrane damage, increased to a greater extent in the WT leaves than in the TPI1‐OE and TPI2‐OE leaves, suggesting that overexpressing SlTPI2 contributed to a higher thermotolerance than overexpressing SlTPI1 (Figure 4e, Table S2). At 25 °C, the SlTPI1‐OE lines showed similar chloroplast TPI activity to the SlTPI2‐OE lines, with both higher than the WT. After the heat treatment, the chloroplast TPI activity decreased, especially in the WT, but it was still higher in the TPI2‐OE lines than in the SlTP1‐OE lines (Figure 4f, Table S2). The F v /F m values, which its reduction reflects PSII photodamage, were significantly lower in the SlTPI1‐OE lines than in the SlTPI2‐OE lines, but higher than that of the WT, after HS (Figure 4g, Table S2). Accordingly, the thylakoid protein complexes in the TPI2‐OE lines were significantly more stable than those of the SlTPI1‐OE lines and the WT following the heat treatment (Figure 4h). We further analysed the thermotolerance of the various mutants, revealing that the tpi1tpi2 double mutants had the lowest thermotolerance, followed by tpi2, tpi1, and the greatest thermotolerance detected in the WT (Figure 5a). The TPI activity, REC, F v /F m and thylakoid protein complex stability were all consistent with these observations (Figure 5b–e, Table S3). Our results also indicate that the function of SlTPI2 under HS is more important than that of SlTPI1.

Figure 5.

Figure 5

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Analysis of tpi1, tpi2 and tpi1tpi2 mutants and WT under heat stress. (a) Phenotypes of WT and tpi1, tpi2 and tpi1tpi2 double mutants (5‐week‐old) under normal condition (25 °C) and heat treatment (42 °C). (b) The chloroplast TPI activity of WT and tpi1, tpi2 and tpi1tpi2 double mutants under heat stress (42 °C). The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05). (c) The REC values of WT and tpi1, tpi2 and tpi1tpi2 double mutants at 25 °C and 42 °C. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters indicate significant differences (P < 0.05) among tomato lines within 25 °C or 42 °C. (d) Fv/Fm of WT and tpi1, tpi2 and tpi1tpi2 double mutants at 42 °C. The data are presented as means ± SD of three biological replicates. Significant differences between means were compared using Fisher's least significant difference (LSD) test. Different letters above bars indicate significant differences (P < 0.05). (e) BN‐PAGE analysis of chlorophyll‐protein complexes. Thylakoid membrane samples with equal concentration from the leaves of the WT and tpi1, tpi2 and tpi1tpi2 double mutants under heat stress (42 °C) were solubilized by treatment with 2% (w/v) DM (n‐Dodecyl‐beta‐D‐maltoside) and separated by BN‐PAGE. The assignments of macromolecular protein complexes of thylakoid membranes indicated at left. I, PSII supercomplexes; II, PSI + PSII dimer; III, PSII monomer; IV, CP43 free PSII; V, LHCII trimer; VI, LHCII monomer. There was a repeating tpi1‐2 lane between WT lane and tpi1*2 lane.

SlWRKY21 and SlHSFA2/7 were roles as upstream regulatory factors of SlTPI1 and SlTPI2 under HS

To explore the response of SlTPI1 and SlTPI2 to HS, we first measured the transcript levels of SlTPI1 and SlTPI2 in WT plants under the heat treatment (42 °C) using RT‐qPCR. During the heat treatment, SlTPI1 expression was constantly suppressed over 24 h, whereas the SlTPI2 expression was significantly induced until 6 h and reduced to the initial levels after 12 h of HS (Figure 6a). We generated luciferase (LUC) reporter constructs driven by the SlTPI1 or SlTPI2 promoter, and transiently expressed them in heat‐stressed N. benthamiana leaves following an Agrobacterium‐mediated infiltration. During the heat treatment, we detected increased LUC activity driven by the SlTPI2 promoter fragments and a decreased LUC activity driven by the SlTPI1 promoter fragments (Figure 6b). These results suggest that SlTPI1 is suppressed by HS whereas SlTPI2 is induced.

Figure 6.

Figure 6

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Analysis of the heat response mechanism of SlTPI1 and SlTPI2. (a) RT‐qPCR analysis of SlTPI1 and SlTPI2 expression in WT during heat treatment. The mRNA level was normalized to that of ACTIN. Data are presented as means ± standard deviation of three biological replicates (**P < 0.01). (b) Agrobacterium cells containing vectors expressing SlTPI1‐pro::LUC and SlTPI2‐pro::LUC were injected into Nicotiana benthamiana leaves respectively. After injection, the plants were cultured for 2 days, incubated at 42 °C before detection. The relative expression of SlTPI1‐pro::LUC and SlTPI2‐pro::LUC was normalized to that of 35S::REN (internal control) (LUC/REN, mean ± SD, n = 3). Data are presented as means ± standard deviation of three biological replicates (**P < 0.01). (c) Above, diagram of the upstream regions of SlTPI1 (left) and SlTPI2 (right), and results of the yeast one‐hybrid assay. W‐box and HSF binding elements (HBE and HBE‐like) were shown in the region upstream of SlTPI1 and SlTPI2 (H1‐4) respectively. These fragments containing W‐box and HBE or HBE‐like were inserted into the pLacZi2μ expression vector. The lower panel shows the binding of the selected SlWRKY members (SlWRKY2, SlWRKY12, SlWRKY15, SlWRKY21, SlWRKY22, SlWRKY33, SlWRKY35, SlWRKY40, SlWRKY69 and SlWRKY70) and the selected SlHSF members (SlHSFA1, SlHSFA2, SlHSFB1, SlHSFB2A, SlHSFB2B and SlHSFA7) to the LacZ reporter gene driven by the indicated fragments in yeast. Blue staining indicates the activation of LacZ. (d) Electrophoretic mobility shift assay (EMSA). The W‐box element and H3 region were used as the probes (5′ biotin). The same probe without biotin was added as a competitive control (C‐probes). Mutant probe, which replaced the W‐box and HBE to AAAAAA, was used as negative control. (e, f) The pGreen0800 (a double LUC reporter system)‐mediated LUC assay is shown. Agrobacterium cells containing vectors expressing SlWRKY21‐FLAG, SlHSFA2‐FLAG and SlHSFA7‐FLAG and Agrobacterium cells containing vectors expressing SlTPI1pro(1 to −900 bp)::LUC or SlTPI2pro(1 to −1602 bp)::LUC were co‐injected into Nicotiana benthamiana leaves respectively. The relative expression levels of SlTPI1pro(1 to −900 bp)::LUC or SlTPI2pro(1 to −1602 bp)::LUC were normalized to that of 35S::REN (internal control) (LUC/REN, mean ± SD, n = 3). The relative expression levels of SlTPI1pro(1 to −750 bp)::LUC or SlTPI2pro(1 to −501 bp)::LUC were used as negative control, respectively. Data are presented as means ± standard deviation of three biological replicates (**P < 0.01).

To investigate the potential regulatory mechanisms of SlTPI1 and SlTPI2 under HS, we analysed the cis‐elements in their promoter regions. The promoter of SlTPI1 includes a W‐box element, which are known binding sites for WRKY family members, including AtWRKY1, AtWRKY4, AtWRKY11, AtWRKY33 or AtWRKY50, as described by Brand et al. (2013). The promoter of SlTPI2 includes HSF‐binding elements (HBEs) (Zhang et al., 2023). First, we analysed the response of the tomato WRKY members to HS, observing that SlWRKY2, SlWRKY12, SlWRKY15, SlWRKY21, SlWRKY22, SlWRKY33, SlWRKY35, SlWRKY40, SlWRKY69 and SlWRKY70 were significantly induced by the heat treatment (Figure S6). We selected the heat‐induced HSF members with high expression levels in the leaves, identified by Yang et al. (2016) and subjected them and selected WRKYs to a yeast one‐hybrid (Y1H) assay. SlWRKY21 (Solyc06g008610) could bind to the W‐box element (W region) in the promoter of SlTPI1, whereas SlHSFA2 (Solyc08g062960) and SlHSFA7 (Solyc02g090820) bound to the HBE (H2 region) in the promoter of SlTPI2 (Figure 6c). An electrophoretic mobility shift assay (EMSA) confirmed the above result (Figure 6d). To further explore the effect of SlWRKY21 and SlHSFA2/7 on SlTPI1 and SlTPI2 expression, respectively, we performed a transient expression assay, in which we introduced a LUC reporter construct driven by different SlTPI1 or SlTPI2 promoter regions into N. benthamiana leaves. When we co‐infiltrated the leaves with the SlWRKY21 or SlHSFA2/7 effector construct, we observed a significant suppression of SlTPI1pro(−1 to −900 bp):LUC expression and an activation of SlTPI2pro(−1 to −1602 bp):LUC expression, respectively, as determined by changes in the relative LUC activity (Figure 6e,f). These results indicate that SlHSFA2/7 positively and directly regulates SlTPI2 expression, whereas SlWRKY21 negatively and directly regulates SlTPI1 expression.

Discussion

As a member of the carbon‐assimilation metabolic system, TPIs catalyse the interconversion of GAP and DHAP (Chen and Thelen, 2010a). In tomato, we found that the knockout of the chloroplast TPIs led to an obvious decrease of the CO2 assimilation rate and the starch, sucrose and glucose contents, supporting that they also involved in carbon metabolism (Figure 2a–d). In pdTPI‐antisense rice lines, the pool sizes of both GAP and DHAP decreased to 60%–70% of those in WT plants, impacting the pool sizes of the CBB cycle metabolites, such as S7P and F6P (Suzuki et al., 2022a). Here, we in‐depth analysed the susceptible targets of chloroplast SlTPI deletion in the CBB cycle and photosynthetic machinery. Unlike the above extensive influence to the CBB cycle metabolites, in the protein level, we found that RbcL, RbcS1, RbcS2, GAPD1 and GAPD2 decreased more in the tpi1tpi2 double mutant lines than that in WT and the single mutants (Figure 3b,c). In addition, the loss of the tomato pdTPIs more broadly affected the photosystem pathway. The subunits of photosynthetic complexes, including PSII (D1, D2, CP43, CP47, PsbF, PsbJ, PsbR and PsbQ), PSI (PsaA, PsaC, PsaD, PsaE, PsaG and PsaH), ATP synthase (AtpA, AtpB, AtpG, AtpH and AtpI) and some key chlorophyll biosynthesis‐related proteins, including POR, DVR and CAO, were less abundant in the tpi1tpi2 double mutants than in the WT (Figure 3c). These could be the proximate causes of the dwarfed and chlorotic variegation phenotypes of the chloroplast tpi mutants, with the build‐up of the DHAP targets and methylglyoxal causing triose phosphate imbalance in the chloroplasts (Chen and Thelen, 2010b).

The chloroplast TPI has two different splice variants (At2g21170.1 and At2g21170.2) in Arabidopsis, with a T‐DNA insertion in this pdTPI resulting in a severely stunted and chlorotic seedlings (Chen and Thelen, 2010b). Similar phenotypes were shown in a mutant of YL2.1, which encodes a unique pdTPI in cucumber (Cucumis sativus) (Xiong et al., 2020). Unlike in Arabidopsis and cucumber, the suppression of pdTPI expression in rice did not cause a dwarf or chlorotic phenotype (Suzuki et al., 2022a), suggesting there may be other pdTPI transcript function in rice. We determined that the tomato genome contains two separate SlTPI genes, Solyc10g054870 and Solyc01g111120, which encode the chloroplast‐located SlTPI1 and SlTPI2 proteins, respectively. The tpi1 and tpi2 single mutants had reduced TPI activity, but no visible phenotypes, whereas their tpi1tpi2 double mutants showed dwarfed and chlorotic variegation phenotypes (Figure 1b,c). Chloroplast TPI is considered as an efficient enzyme (Fridlyand et al., 1999), thus the reduced TPI activity in single mutants is not enough to cause the metabolic disturbance in chloroplast. When we knockout one TPI, the other also can maintain the function of TPI in chloroplasts. This suggests that chloroplast SlTPI1 and SlTPI2 are functionally redundant in tomato under normal growth conditions. In addition, overexpressing SlTPI1 or SlTPI2 did not improve the growth or photosynthetic capacity of the tomato plants (Figure 4d,g). Similarly, although the overexpression of pdTPI marginally improved photosynthesis at elevated CO2 levels in rice, it was not sufficient to improve biomass production (Suzuki et al., 2022b). These findings suggest that the chloroplast TPIs are abundant and efficient in various plant species, thus their overexpression does little to magnify their functions.

In Arabidopsis thaliana, the two pdTPI splice variants were expressed in different organs. At2g21170.1 was expressed in roots, leaves, stems, flowers and siliques, with the highest expression in leaves and flowers, whereas At2g21170.2 was weakly detected only in the roots, suggesting organ‐specific roles for these gene products (Chen and Thelen, 2010b). In tomato, both SlTPI1 and SlTPI2 were expressed in the leaves and cotyledons (Figure S3). Although they are functionally redundant under normal growth conditions, we observed that they displayed functional differentiation in response to HS; SlTPI2 seems more heat‐stable than SlTPI1, displaying less of a reduction in its protein levels and activity in response to HS (Figure 4a,b). Overexpression SlTPI2 in tomato showed a higher thermotolerance than overexpression SlTPI1 (Figure 4d–h), which is a promising strategy to increase crop tolerance to HS. In addition, SlTPI1 expression was suppressed and SlTPI2 was induced by HS, suggesting that there were respective upstream regulatory factors in the HS response system (Figure 6a,b). HSFA2 and HSFA7 are important positive regulatory factors in the HS response system, both of which are activated by HSFA1 and suppressed by HSFB (Ohama et al., 2017). Our further analysis showed that both HSFA2 and HSFA7 were the direct upstream regulators of SlTPI2, and could bind to the HBE elements in the SlTPI2 promoter region and activate its expression (Figure 6c,d,f). We determined that SlWRKY21 was the direct suppressor of SlTPI1 through the screening of these heat‐induced SlWRKY members (Figures 6c–e and S6).

In summary, SlHSFA2/7 and SlWRKY21 bind to the promoters of SlTPI2 and SlTPI1, respectively, in response to HS, thereby activating SlTPI2 expression and suppressing SlTPI1 expression. The increased levels of thermotolerant SlTPI2 may play a positive role in photoprotection under HS (Figures 4g, 6 and 7). These provide a potential upgrade strategy to decrease the heat‐induced photodamage in plant.

Figure 7.

Figure 7

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Model of the response of SlTPI1 and SlTPI2 to heat stress. Under heat stress, both SlHSFA2/7 and SlWRKY21 were induced to bound to the promoter of SlTPI2 and SlTPI1 respectively, thus activating SlTPI2 expression and suppressing SlTPI1 expression, the increased thermotolerant SlTPI2 may play a positive role to maintain photosynthesis under heat stress.

Experimental procedures

Plant materials, growth conditions and experimental treatments

Tomato (Solanum lycopersicum, cv. Micro‐TOM) plants were used to generate all transgenic materials in this study. The tpi1 and tpi2 mutant lines were generated using a pHSE401‐based CRISPR/Cas9 system. Eight independent T0 mutant lines of SlTPI1 and six independent T0 mutant lines of SlTPI2. All of them were heterozygotes. From their T1 lines, we acquired two homozygous mutant plants of SlTPI1 and SlTPI2 respectively and then used them to produce the tpi1tpi2 double mutant lines by crossing. The T1 tpi1tpi2 double mutant lines and T2 tpi1 and tpi2 single mutants were used for further analysis. The resulting lines were confirmed by DNA sequencing. To generate the SlTPI1‐ and SlTPI2‐overexpression lines, the full‐length SlTPI1 and SlTPI2 coding sequences were individually cloned into the pSuper1300 plant expression vector with the cauliflower mosaic virus (CaMV) 35S promoter via the KpnI and SalI restriction sites. The tomato plants were transformed using the Agrobacterium tumefaciens (LBA4404)‐mediated leaf disk method. The genotypes of the Hygromycin‐resistant SlTPI1‐ and SlTPI2‐overexpression lines were confirmed by PCR using a vector‐specific forward primer and a reverse primer specific to SlTPI1 or SlTPI2 (35S‐F/SlTPI1‐R and 35S‐F/SlTPI2‐R).

Five‐week‐old tomato seedlings were exposed to heat treatment (42 °C) in an illuminated incubation chamber (E‐41L2; Percival) with a photon flux density (PFD) of approximately 350 μmol m−2 s−1 and a relative humidity of 60%–70% for 24 h. The same‐age tomato seedlings cultivating at 25 °C in another E‐41L2 chamber with the same photon flux density and relative humidity were used as control. The leaves were then harvested for analysis.

RT‐qPCR analysis

Total RNA was isolated from mature leaves using an RNAprep Pure kit (DP441) from Tiangen (Beijing, China) and reverse‐transcribed into first‐strand cDNA using a MonScript RTIII Super Mix with dsDNase (Two‐Step) Kit from Monad (Wuhan, China). The qPCR was conducted using a MonAmp SYBR Green qPCR Mix (Monad) using an Applied Biosystems quantitative PCR Q6 instrument (Thermo Fisher Scientific, Waltham, MA). The gene expression levels were normalized to ACTIN, and the values in the WT were set to 1. For each assay, at least three biological replicates were performed. All primers used in this study are listed in Table S1.

Measurement of physiological parameters

The net photosynthetic rate (Pn) was measured by a portable photosynthetic system (CIRAS‐3; PP Systems, Amesbury, MA) under ambient CO2 conditions (360 μL L−1), a PFD of 800 μmol m−2 s−1, and a relative humidity of 70%. Before measurement, these heat‐treated plants were kept at 25 °C in above E‐41L2 chamber to induce stomatal opening for approximately 30 min. A Handy Plant Efficiency Analyzer system (Hansatech Instruments, Pentney, UK) was used to detect the maximal PSII photochemical efficiency (F v /F m). Before measurement, the tomato leaves were adapted in dark for 20 min. CO2 assimilation was measured using a portable gas exchange system (LI‐6800, Li‐Cor, Lincoln, NE). This measurement was under 800 μmol m−2 s−1 PFD, and a relative humidity of 70%. The leaf temperature was maintained at 25 °C. Different concentrations of CO2 were supplied to detect a series of different intracellular CO2 concentrations and the corresponding photosynthetic rate (A), which were then used to make A/Ci curves. Ten fresh leaf discs (0.8 cm diameter) were harvested from different TPI‐transgenic tomato lines and the WT before and after heat treatment, and were used to measure the REC (Kong et al., 2014).

The chloroplast TPI activity, starch content, sucrose content and glucose content were determined using a TPI‐2‐G kit, DF‐2‐Y kit, ZHT‐2‐Y kit and PT‐2‐Y kit, respectively (all Comin Biotechnology, Suzhou, China). For the chloroplast TPI activity detection, the chloroplasts in tomato leaves were firstly extracted by differential centrifugation using reagents in TPI‐2‐G kit. Three biological replicates were performed for each assay.

LUC reporter assay

For the LUC reporter assay, the promoter regions of SlTPI1 and SlTPI2 were independently inserted into the pGreen0800 vector (a double LUC reporter vector) via the KpnI and SalI sites. The SlWRKY21, SlHSFA2, and SlHSFA7 coding sequences were independently ligated into the pZP211‐FLAG vector without the 35S promoter after an EcoRI/SalI digestion. These resulting constructs were introduced into the Agrobacterium strain GV3101 and transiently infiltrated in appropriate pairs into N. benthamiana leaves, using each promoter:LUC reporter construct and the empty vector as controls. The transient infiltration in N. benthamiana and its analysis were conducted as described by Zhuang et al. (2019).

Y1H assay

The Y1H was performed essentially as described by Zhuang et al. (2019). The promoter regions of SlTPI1 and SlTPI2 shown in Figure 6c were individually inserted into the KpnI and SalI sites of the pLacZi2u vector. The coding sequences of the heat‐induced SlWRKY members and the selected SlHSF members were individually cloned into the pGAD vector via an EcoRI and XhoI digestion. The appropriate plasmid pairs (according to Figure 6c) were co‐transformed into yeast strain EGY48. Cells were plated onto synthetic defined (SD) medium lacking Ura and Trp, and positive clones were grown in SD medium lacking Ura and Trp but containing 20% (w/v) galactose, 20% (w/v) raffinose, BU salts (0.27 M Na2HPO4 and 0.23 M NaH2PO4.2H2O) and 0.05 M X‐Gal.

EMSA

The EMSAs were performed as described by Zhuang et al. (2019). The W‐box and EBS regions were used as probes, which were synthesized by Sangon Biotech (Shanghai, China). The sequences encoding WRKY21, HSFA2 and HSFA7 were individually cloned into the pET‐30a (+) vector between the KpnI and SalI sites. The production and purification of recombinant WRKY21, HSFA2 and HSFA7 were carried out using a His‐tag Protein Purification Kit (Beyotime, Shanghai, China). The proteins were incubated with 0.5 μM of the corresponding biotin‐labelled ssDNA probes in a total volume of 10 μL for 20 min. Unlabelled probes were added at various concentrations for the competition experiments. A LightShift Chemiluminescent EMSA Kit (Thermo Fisher Scientific, Waltham, MA) was used for chemiluminescence detection.

Observation of the chloroplast ultrastructure

The mesophyll tissues of the tpi1‐1, tpi2‐1, tpi1*2‐2 and WT plants were collected, cut into small pieces 2–3 mm in length and width, fixed immediately in 3.5% (w/v) glutaraldehyde, and post‐fixed in 1% (w/v) aqueous osmium tetroxide. The tissues were dehydrated in an ethanol series and then infiltrated and embedded in epoxy resin. Ultra‐thin sections were obtained with an ultramicrotome and stained with uranyl acetate and lead citrate. Finally, the transmission electron microscopy observation of stained sections was performed with a JEM‐1200EX (Jeol, Tokyo, Japan), which was also used to photograph the samples.

BN‐PAGE analysis

Mature fresh tomato leaves (0.15 g) were collected from the tpi mutant lines and WT before and after the heat treatment and placed on ice. Chloroplasts were extracted from these samples with an extraction solution (0.33 M sorbitol, 30 mM tricine, 5 mM EGTA, 5 mM EDTA and 10 mM NaHCO3). The extracts were centrifuged at 20 000  g at 4 °C for 2 min; this step was performed four times to remove starch. The procedure was conducted under green light. The supernatant was discarded and mixed with 2% n‐Dodecyl‐beta‐D‐maltoside. The extracts were centrifuged at 20 000  g at 4 °C for 10 min, after which the supernatant was combined with a one‐tenth volume of 5% SERVA Blue G in 100 mM Bis Tris–HCl (pH 7.0), 0.5 M 6‐amino‐n‐caproic acid and 30% (w/v) glycerol and applied to 0.75‐mm‐thick 6%–12% acrylamide gradient gels in a Hoefer Mighty Small vertical electrophoresis unit connected to a cooling circulator (Hoefer, Holliston, MA). Other steps were conducted, following the protocol reported by Peng et al. (2006). The extracted proteins were quantified with a Bradford protein assay kit (Tiangen).

Proteomics analysis

Proteins were extracted from the leaves of tpi1‐1, tpi2‐1, tpi1*2‐2, and the WT for use in this analysis. Following protein extraction and digestion, SDS‐PAGE, labelling, high‐pH reversed‐phase fractionation, liquid chromatography (LC)–mass spectrometry (MS)/MS analysis and protein identification and quantitation were performed by APTBIO (Shanghai, China). The LC–MS/MS analysis was performed on a Q Exactive mass spectrometer (Thermo Fisher Scientific) that was coupled to an Easy nLC (Proxeon Biosystems, now Thermo Fisher Scientific) for 60/90 min. TMT (Tandem Mass Tag, Thermo Fisher Scientific, Waltham, MA) report ion peaks are generated in the low mass region of the mass spectrum, the intensity of which reflects the relative expression of the peptide in different samples. The raw MS data for each sample were searched using the MASCOT engine (Matrix Science, London, UK; version 2.2) embedded into Proteome Discoverer 1.4 software for identification and quantitation analysis. n = 3 in each transgenic plants and WT.

Protein isolation and immunoblot analysis

The total protein extraction and immunoblot analysis were performed as described by Kong et al. (2014). A Mini‐PROTEAN Tetra System (Bio‐Rad Laboratories, Hercules, CA) was used for the electrophoresis and a TransBlot SD transfer cell (Bio‐Rad Laboratories) was used for the protein transfer to polyvinylidene fluoride. Specific antibodies against GFP and a β‐ACTIN mouse monoclonal antibody from TransGen Biotech (Beijing, China) were used to detect the proteins.

Statistical analysis

The data represent the mean ± standard deviation (SD) of three biological replicates. The statistical significance of any differences in the heat‐induced assays between the control and each heat treatment point and the transcriptional activation assay were tested using a Student's t‐test performed in Excel (Microsoft, Redmond, WA). One‐way analysis of variance (ANOVA) with Fisher's least significant difference (LSD) test was used to examine the difference among tomato plants. The effects of HS and tomato plants on measured parameters were tested using two‐way ANOVA. Analysis of variance was conducted using the ‘aov’ function in the ‘stats’ package. The LSD test was applied to assess differences in measured parameters among tomato plants for control and HS using the ‘LSD.test’ in the ‘agricolae’ package. These statistical analyses were performed using R software (v.4.3.1) (http://cran.r‐project.org).

Author contributions

K. Zhuang and Q. Meng designed the research; K. Zhuang, C. Chen and M. Zhang performed research; K. Zhuang and X. Ma analysed data; K. Zhuang and Q. Meng wrote the paper.

Conflict of interest

The authors declare that there is no conflict of interest.

Supporting information

Figure S1 Amino acid sequence alignment and phylogenetic analysis of tomato TPIs and Arabidopsis TPIs.

Figure S2 Subcellular localization analysis of SlTPI1 and SlTPI2.

Figure S3 Expression analysis of SlTPI1 and SlTPI2 in different tomato organs.

Figure S4 Analysis of other differentially expressed proteins (DEPs) in WT and mutants of SlTPIs.

Figure S5 Analysis of SlTPI1 and SlTPI2 protein levels in WT under heat stress.

Figure S6 Analysis of the heat‐induced expression of selected SlWRKY members.

Table S1 Primers were used in this manuscript.

Table S2 Results of two‐way analysis of variance (ANOVA) for the effects of heat stress (HS) and tomato lines (WT, TPI1‐OE‐1, TPI1‐OE‐4, TPI2‐OE‐1 and TPI2‐OE‐2) on REC, Fv/Fm and TPI activity.

Table S3 Results of two‐way analysis of variance (ANOVA) for the effects of heat stress (HS) and tomato lines (WT, tpi1‐1, tpi1‐2, tpi2‐1, tpi2‐2, tpi1tpi2‐1 and tpi1tpi2‐2) on REC, Fv/Fm and TPI activity.

PBI-22-650-s001.docx (7.4MB, docx)

Acknowledgements

This work was supported by National Key Research and Development Program (2020YFA0907600), the National Natural Science Foundation of China (grant number 32100203) and the Natural Science Foundation of Shandong Province (grant no. ZR202103010453).

Data availability

The data supporting the findings of this study are available from the corresponding author upon request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1 Amino acid sequence alignment and phylogenetic analysis of tomato TPIs and Arabidopsis TPIs.

Figure S2 Subcellular localization analysis of SlTPI1 and SlTPI2.

Figure S3 Expression analysis of SlTPI1 and SlTPI2 in different tomato organs.

Figure S4 Analysis of other differentially expressed proteins (DEPs) in WT and mutants of SlTPIs.

Figure S5 Analysis of SlTPI1 and SlTPI2 protein levels in WT under heat stress.

Figure S6 Analysis of the heat‐induced expression of selected SlWRKY members.

Table S1 Primers were used in this manuscript.

Table S2 Results of two‐way analysis of variance (ANOVA) for the effects of heat stress (HS) and tomato lines (WT, TPI1‐OE‐1, TPI1‐OE‐4, TPI2‐OE‐1 and TPI2‐OE‐2) on REC, Fv/Fm and TPI activity.

Table S3 Results of two‐way analysis of variance (ANOVA) for the effects of heat stress (HS) and tomato lines (WT, tpi1‐1, tpi1‐2, tpi2‐1, tpi2‐2, tpi1tpi2‐1 and tpi1tpi2‐2) on REC, Fv/Fm and TPI activity.

PBI-22-650-s001.docx (7.4MB, docx)

Data Availability Statement

The data supporting the findings of this study are available from the corresponding author upon request.

Articles from Plant Biotechnology Journal are provided here courtesy of Society for Experimental Biology (SEB) and the Association of Applied Biologists (AAB) and John Wiley and Sons, Ltd

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