1 Short title: Role of NTRC in developing tomato fruits...2019/09/16 · NTRC acts as central hub...

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Short title: Role of NTRC in developing tomato fruits 1

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Corresponding author: 3

Liang-Yu Hou 4

Address: Ludwig-Maximilians-University Munich, Department Biology I, 82152 5

Planegg-Martinsried, Germany 6

Tel/Fax: +49 (089) 2180 74710 / +49 (089) 2180 74599 7

E-mail: [email protected] 8

Plant Physiology Preview. Published on September 16, 2019, as DOI:10.1104/pp.19.00911

Copyright 2019 by the American Society of Plant Biologists

www.plantphysiol.orgon October 18, 2020 - Published by Downloaded from Copyright © 2019 American Society of Plant Biologists. All rights reserved.

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Article title: NTRC plays a crucial role in starch metabolism, redox balance and tomato fruit 9

growth 10

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Liang-Yu Hou*, Matthias Ehrlich, Ina Thormählen, Martin Lehmann, Ina Krahnert, Toshihiro 12

Obata, Francisco J. Cejudo, Alisdair R. Fernie, Peter Geigenberger 13

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Ludwig-Maximilians-University Munich, Department Biology I, 82152 Planegg-Martinsried, 15

Germany (L.-Y. H., M.E., I.T., M.L., P.G.); Max-Planck-Institute of Molecular Plant 16

Physiology, 14476 Potsdam-Golm, Germany (I.K., T.O., A.R.F.); Instituto de Bioquímica 17

Vegetal y Fotosíntesis, Universidad de Sevilla and Consejo Superior de Investigaciones 18

Científicas, 41092 Seville, Spain (F.J.C.) 19

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One sentence summary: 21

An established regulator of photosynthetic processes in leaves influences tomato fruit quality 22

by regulating starch accumulation, redox balance, and fruit growth. 23



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Footnotes: 24

25

P.G. conceived the project; L.-Y.H., M.E., I.T., M.L., I.K., T.O., F.J.C. A.R.F., P.G. designed 26

the research; L.-Y.H. performed most of the experiments; M.E. performed the cloning; M.L. 27

performed GC-MS measurements; I.K. performed tomato transformation; L.-Y.H., I.T., M.L., 28

T.O., P.G. analyzed the data; I.T., A.R.F., P.G. supervised the experiments; L.-Y.H., P.G. 29

wrote the article. 30

31

This work was supported by the Deutscher Akademischer Austauschdienst (grant to L.-Y.H. 32

and P.G.) and the Deutsche Forschungsgemeinschaft (TRR 175, grants to P.G, A.R.F and 33

M.L.). 34

35

The author responsible for distribution of materials integral to the findings presented in this 36

article in accordance with the policy described in the Instructions for Authors 37

(www.plantphysiol.org) is: Peter Geigenberger ([email protected]). 38

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* Address correspondence to [email protected] 40



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Abstract 41

NADPH-thioredoxin reductase C (NTRC) forms a separate thiol-reduction cascade in 42

plastids, combining both NADPH-thioredoxin reductase and thioredoxin activities on a single 43

polypeptide. While NTRC is an important regulator of photosynthetic processes in leaves, its 44

function in heterotrophic tissues remains unclear. Here, we focus on the role of NTRC in 45

developing tomato (Solanum lycopersicum) fruits representing heterotrophic storage organs 46

important for agriculture and human diet. We used a fruit-specific promoter to decrease 47

NTRC expression by RNA interference in developing tomato fruits by 60-80% compared to 48

the wild type. This led to a decrease in fruit growth, resulting in smaller and lighter fully ripe 49

fruits containing less dry matter and more water. In immature fruits, NTRC down-regulation 50

decreased transient starch accumulation, which led to a subsequent decrease in soluble sugars 51

in ripe fruits. The inhibition of starch synthesis associated with a decrease in the 52

redox-activation state of ADP-glucose pyrophosphorylase and soluble starch synthase, which 53

catalyze the first committed and final polymerizing steps of starch biosynthesis, respectively. 54

This was accompanied by a decrease in the level of ADP-glucose. NTRC down-regulation 55

also led to a strong increase in the reductive states of NAD(H) and NADP(H) redox systems. 56

Metabolite profiling of NTRC-RNAi lines revealed increased organic and amino acid levels, 57

but reduced sugar levels, implying that NTRC regulates the osmotic balance of developing 58

fruits. These results indicate that NTRC acts as central hub in regulating carbon metabolism 59

and redox balance in heterotrophic tomato fruits, affecting fruit development as well as final 60

fruit size and quality. 61

62

Introduction 63

Reduction-oxidation (Redox) regulation appears to be a fundamental integrator of 64

metabolic pathways in different subcellular compartments (Geigenberger and Fernie, 2014). 65

In plant chloroplasts there are two different thiol redox systems, the ferredoxin 66

(Fdx)-thioredoxin (Trx) system, which depends on the reduction of Fdx by photosynthetic 67

electron transport in response to light, and the NADPH-dependent Trx reductase C (NTRC) 68

system, which relies on NADPH and thus may be linked to Fdx-NADPH reductase (FNR) in 69

the light or sugar metabolism in the dark (Buchanan and Balmer, 2005; Zaffagnini et al., 70

2018). NTRC is an unusual protein since it harbors both NADPH-Trx reductase (NTR) and 71

Trx domains on the same polypeptide (Serrato et al., 2004). This feature allows NTRC to use 72

NADPH as a source of electrons to regulate different chloroplast target proteins via 73

thiol-disulfide modulation (Spínola et al., 2008; Geigenberger et al., 2017). 74

In Arabidopsis (Arabidopsis thaliana) T-DNA insertion mutants, NTRC is involved in 75



5

various chloroplast functions. There is in-planta evidence that NTRC is important to reduce 76

2-Cys peroxiredoxins (2-Cys Prxs) for the removal of hydrogen peroxide (Pérez-Ruiz et al., 77

2006; Pérez-Ruiz and Cejudo, 2009; Kirchsteiger et al., 2009) and to facilitate starch 78

synthesis via redox regulation of ADP-glucose pyrophosphorylase (AGPase) (Michalska et 79

al., 2009; Lepistö et al., 2013). Furthermore, NTRC has overlapping functions with the 80

Fdx-dependent Trx systems to optimize photosynthetic processes, such as non-photochemical 81

quenching of light energy (Thormählen et al., 2015; Naranjo et al., 2016; Yoshida and 82

Hisabori, 2016; Thormählen et al., 2017), Calvin-Benson cycle activity (Thormählen et al., 83

2015; Nikkanen et al., 2016; Yoshida and Hisabori, 2016), chlorophyll biosynthesis (Richter 84

et al., 2013; Pérez-Ruiz et al., 2014; Da et al., 2017) and ATP synthesis (Carrillo et al., 2016; 85

Nikkanen et al., 2016). In this context, NTRC indirectly maintains the reducing capacity of 86

the pool of Fdx-TRXs by restricting their re-oxidation via an oxidation loop involving 87

hydrogen peroxide and oxidized 2-Cys Prxs (Pérez-Ruiz et al., 2017). NTRC is, therefore, of 88

crucial importance to balance oxidation and reduction pathways in the chloroplast to optimize 89

photosynthesis, overall plant growth and development in response to light (Thormählen et al., 90

2015; Carrillo et al., 2016; Yoshida and Hisabori, 2016; Thormählen et al., 2017). 91

In addition to its role in chloroplasts, NTRC is also present in non-photosynthetic 92

plastids together with Fdx-dependent Trxs (Kirchsteiger et al., 2012). However, as yet little is 93

known on the role of NTRC in heterotrophic plant tissues that lack photosynthetic functions. 94

There is evidence that NTRC affects starch synthesis in Arabidopsis roots (Michalska et al., 95

2009), but this does not contribute to the growth phenotype of the Arabidopsis ntrc mutant 96

(Kirchsteiger et al., 2012). Complementation of the ntrc mutant by NTRC overexpression 97

under the control of a leaf-specific promoter led to wild-type (WT) phenotypes, but the 98

mutant phenotype remained unaltered when a root-specific promoter was used (Kirchsteiger 99

et al., 2012). These data thus indicate a role of NTRC in photosynthetic leaves, rather than in 100

non-photosynthetic roots. 101

Tomato (Solanum lycopersicum) is an important horticultural crop and major component 102

of the human diet serving as the primary model system to study fruit physiology and 103

development (Kanayama, 2017). Throughout the fruit developing process, a myriad of 104

changes in color, firmness, taste and flavor occur that are associated with developmentally 105

regulated metabolic shifts (e.g. changes in the levels of carbohydrates, acids and volatile 106

compounds) (Carrari et al., 2006). Notably, the abundance of carbohydrates is a key 107

determinant for fruit yield and quality (Kanayama, 2017). During development, immature 108

tomato fruits act as strong sink tissues, into which massive amounts of carbon are imported as 109

sucrose from leaf tissues (Osorio and Fernie, 2014). Most of the sucrose is mobilized to 110



6

glucose and fructose and subsequently converted to starch as an intermediate carbon store to 111

promote the sink activity of the immature fruits (Osorio and Fernie, 2014). In later ripening 112

stages, starch is degraded to soluble sugars, which are key determinants of fruit taste and 113

quality (Davies and Hobson, 1981; Gierson and Kader, 1986). 114

Alterations of the transient starch pool of immature fruits have been proposed to be 115

important for the accumulation of soluble sugar in ripening fruits, which determines the final 116

fruit size and quality (Schaffer et al., 2000; Petreikov et al., 2009). In this context, AGPase 117

serves as a rate-limiting enzyme for starch synthesis in developing tomato fruits (Petriekov et 118

al. 2009). AGPase is a heterotetrameric enzyme located in the plastid comprised of two large 119

and two small subunits, which are subject to allosteric activation by glycerate-3-phosphate 120

(3-PGA) and inhibition by inorganic phosphate (Pi) (Preiss, 1973). AGPase is also subject to 121

redox regulation involving reversible reduction of a regulatory disulfide linking the small 122

subunits, which affects the allosteric properties of the enzyme (Fu et al., 1998; Tiessen et al., 123

2002). Manipulation of the cellular redox poise during tomato fruit development has 124

previously been demonstrated to lead to changes in the NADPH/NADP+ ratio, which were 125

accompanied by changes in the redox-activation states of NADP-malate dehydrogenase 126

(MDH) and AGPase of the developing fruits (Centeno et al., 2011). The changes in AGPase 127

redox state were additionally associated with alterations in starch synthesis in immature fruits 128

and subsequent alterations in soluble sugar content in mature fruits (Centeno et al., 2011). 129

Although these studies provide evidence that redox-regulation via thiol-disulfide modulation 130

is involved in tomato fruit development, the underlying fruit-specific thiol-redox systems 131

remain unresolved. 132

In this study, we down-regulated NTRC gene expression specifically in fruit tissues by 133

generating a RNA interference construct under the control of the fruit-specific patatin B33 134

promoter. The NTRC-RNAi lines were characterized by a 60-80% decrease in NTRC 135

transcripts and protein levels in developing fruits. In immature fruits, NTRC down-regulation 136

decreased transient starch accumulation by decreasing the redox-activation state of AGPase 137

and the activity of soluble starch synthase, which subsequently led to a decreased 138

accumulation of soluble sugars during ripening and to decreased fruit yield and quality in 139

fully ripe fruits. This was accompanied by an increased reduction state of the NAD(H) and 140

NADP(H) redox couples. These results provide evidence for a previously unknown function 141

of NTRC as a central hub in regulating carbon metabolism and redox balance in developing 142

fruits. 143



7

Results 144

Generation of transgenic tomato plants with decreased expression of NTRC under 145

control of a fruit-specific promoter 146

To silence NTRC gene expression specifically in fruit tissues, we generated a 147

NTRC-RNAi construct under the control of the patatin B33 promoter, which has previously 148

been identified to confer fruit specific expression in tomato plants (Rocha-Sosa et al., 1989; 149

Frommer et al., 1994; Obiadalla-Ali et al., 2004), with kanamycin serving as a marker for 150

selection (Fig. 1A). The resulting construct was transformed via an Agrobacterium 151

tumefaciens-mediated process into tomato plants. To identify functional NTRC-RNAi lines, 152

samples from 35 and 65 day-after-flowering (DAF) fruits were harvested for genotyping. As 153

shown in Figure 1B, the NTRC-RNAi lines 2, 26 and 33 all harbored the neomycin 154

phosphotransferase II (nptII) gene, indicating the presence of the NTRC-RNAi cassette 155

within their genome. The transcript level of NTRC gene (LOC101254347) decreased by 156

60-80% in comparison to WT in the three RNAi lines, both in 35 (Fig. 1C) and 65 DAF fruit 157

samples (Supplemental Fig. S1A). Furthermore, the NTRC protein level of RNAi-2, 26 and 158

33 decreased by 50-80% compared to WT, both in 35 (Fig. 1, D and E) and 65 DAF fruit 159

samples (Supplemental Fig. S1, B and C). The expression of the second NTRC gene 160

(LOC101266017) (Nájera et al., 2017) was also analyzed, but its expression was too low to 161

be detectable in fruit tissues. Thus, we concluded the NTRC-RNAi lines 2, 26 and 33 were 162

appropriate to study the function of NTRC in tomato fruit. 163

164

Fruit-specific silencing of NTRC has no substantial effects on the process of fruit 165

development 166

The different stages of tomato fruit development are characterized by changes in 167

pigmentation and ripening. To analyze whether decreased expression of NTRC affects the 168

process of tomato fruit development, we analyzed chlorophyll and light-harvesting complex 169

II (Lhcb2) protein levels as well as the expression of genes involved in ethylene synthesis and 170

ripening in 35 DAF and 65 DAF fruits. In 35 DAF fruits, chlorophyll content and protein 171

level of Lhcb2 were not significantly changed in NTRC-RNAi lines compared to WT, while 172

both parameters were not detectable in 65 DAF fruits in all genotypes (Supplemental Fig. S2, 173

A and B). There were no visible changes in fruit color at 35 DAF (Fig. 2A) and 65 DAF 174

(Supplemental Fig. S3A) when NTRC-RNAi lines were compared to WT. To further analyze 175

whether NTRC down-regulation affected the ripening processes, we assayed the expression 176

of genes involved in ethylene production (S-adenosylhomocysteine hydrolase, SAHH2) 177

(Yang et al., 2017) and ripening (NON-RIPENING, NOR) (Ma et al., 2019) in both 35 DAF 178



8

and 65 DAF fruits. Both genes were up-regulated when fruits became mature, but their 179

expression levels were not significantly changed between WT and NTRC-RNAi lines 180

(Supplemental Fig. S2, C and D). Taken together, NTRC down-regulation did not affect the 181

process of fruit development. While 35 DAF fruits represent the mature green stage and 65 182

DAF fruits represent the fully ripened stage, there were no stage-dependent differences 183

between NTRC-RNAi lines and WT. 184

185

Fruit-specific silencing of NTRC leads to smaller and lighter fruits with decreased dry 186

weight-to-fresh weight ratios 187

To investigate whether silencing of NTRC affects overall fruit growth, fully mature 188

fruits (65 DAF) were harvested for the measurement of final fruit size and weight. As shown 189

in Figure 2A, although both the WT and RNAi lines were fully ripe, the final fruit size of 190

NTRC-RNAi lines was clearly smaller than WT. Furthermore, the NTRC-RNAi lines 191

exhibited a 20% reduction in fruit height and 15% reduction in fruit width compared to WT 192

(Fig. 2B). The final tomato fruit fresh weight in NTRC-RNAi lines was 40% lower than WT 193

(Fig. 2C), and the NTRC-RNAi tomato fruits also contained less dry matter and more water 194

than WT (Fig. 2D). The effects of NTRC silencing on fruit growth and dry-matter content 195

were also observed at 35 DAF, indicating that they were not caused by developmental effects 196

(Supplemental Fig. S3). Taken together, fruit-specific silencing of NTRC led to the 197

generation of smaller and lighter fruits consisting of less dry matter in tomato plants. 198

199

Fruit-specific silencing of NTRC leads to lower accumulation of starch and soluble 200

sugars during fruit development 201

Tomato fruit development is characterized by a transient accumulation of starch in 202

immature fruits, which is subsequently mobilized to contribute to the accumulation of soluble 203

sugars, specifically glucose, in mature fruits (Schaffer et al., 2000). To investigate whether 204

the inhibition of fruit growth in NTRC-RNAi lines is due to a decrease in the transient starch 205

pool, we analyzed the levels of starch and soluble sugars during fruit development. In 206

immature fruits (35 DAF), NTRC down-regulation led to a strong decrease in starch 207

accumulation down to 25-50% of WT level (Fig. 3A). In mature fruits (65 DAF), where 208

starch has been almost completely mobilized in all genotypes, the NTRC-RNAi lines still 209

exhibited a trend of decreased residual starch levels compared to WT (Fig. 3B). In contrast to 210

this, NTRC silencing did not affect the protein content of the fruits at both developmental 211

stages (Supplemental Fig. S4). 212

To investigate whether the inhibition of starch accumulation was accompanied by an 213



9

increase in soluble sugars, we analyzed the levels of glucose, fructose, sucrose and maltose in 214

NTRC-RNAi lines compared to WT. At both developmental stages, glucose level, the major 215

soluble sugar in tomato fruits, was strongly decreased in NTRC-RNAi lines compared to WT, 216

while fructose, sucrose and maltose levels showed only slight decreases (Fig. 3, C and D; Fig. 217

4A; Supplemental Table S1). Thus, interfering with NTRC expression in tomato fruits greatly 218

compromised transient starch accumulation and final soluble sugar accumulation during their 219

development, while there were no effects on the protein content of the fruits. 220

221

Interfering with NTRC expression in tomato fruits has minor effects on the levels of 222

phosphorylated intermediates and adenine nucleotides 223

To investigate whether the decrease in starch and sugar accumulation is related to 224

changes in sugar and energy metabolism, we further assayed the level of phosphorylated 225

metabolites and adenine nucleotides in developing tomato fruits. In NTRC-RNAi lines, the 226

level of glucose-6-phosphate (G6P), fructose-6-phosphate (F6P), and glucose-1-phosphate 227

(G1P) showed no consistent changes at 35 and 65 DAF when compared to WT 228

(Supplemental Table S1 and Table S2). Also, no substantial changes were detected in the 229

level of the glycolytic intermediate 3-PGA between WT and NTRC-RNAi lines at both 230

developmental stages. Triose phosphates were also assayed, but the level was under detection 231

limit, which is characteristic for the glycolytic pathway operating in heterotrophic tissues 232

(Hatzfeld and Stitt, 1990). Furthermore, the levels of ATP and ADP and the ATP/ADP ratio 233

in NTRC-RNAi lines were not significantly changed during fruit development when 234

compared to WT (Supplemental Table S2). The data also show the levels of the substrates 235

(G1P, ATP) and the allosteric regulators (3-PGA, Pi) of AGPase were not substantially 236

changed in the transgenic fruits. Taken together, these results demonstrate that interfering 237

with NTRC expression does not perturb glycolytic metabolism or adenylate energy state in 238

developing fruits. 239

240

Interfering with NTRC expression in tomato fruits leads to global changes in their 241

metabolite profile 242

To obtain in-depth insight into more global metabolite changes in NTRC-RNAi tomato 243

lines, we next performed metabolite profiling via GC-TOF-MS. We firstly confirmed the 244

NTRC-RNAi lines showed generally lower sugar levels compared to WT (Fig. 4A, left panel), 245

which was the case for glucose (55-80% of WT level), fructose (65-85% of WT level), and 246

sucrose (40-55% of WT level) as well as other sugars. Moreover, upon NTRC silencing, the 247

levels of several sugars decreased more strongly at 65 DAF than 35 DAF (Fig. 4A, right 248



10

panel), including gentiobiose (3.6-fold), maltose (2-fold) and ribose (2.3-fold). With 249

exception of myo-inositol, there was no great change in sugar alcohols in NTRC-RNAi lines 250

(Fig. 4A). 251

By contrast, levels of several organic acids increased in NTRC-RNAi lines compared to 252

WT plants, including 2-imindazolidone-4-carboxylic acid, trans-aconitic acid, trans-caffeic 253

acid and quinic acid, which are involved in the synthesis of phenolic compounds, while 254

others, such as 1-pyrroline-2-carboxylate and gulonic acid, decreased (Fig. 4B). The decrease 255

in gulonic acid level might be attributable to lower glucose content, since this acid is formed 256

by oxidation of glucose. The intermediates of the tricarboxylic acid (TCA) cycle also showed 257

an increasing pattern compared to WT. Levels of citric acid and 2-oxo-glutaric acid were 2 to 258

4 times increased in NTRC-RNAi lines at both stages of fruit development, while malic acid, 259

fumaric acid and succinic acid only showed slight changes compared to WT (Fig. 4C). 260

Interestingly, the TCA cycle precursor, pyruvic acid, increased only in immature fruits of 261

NTRC-RNAi lines compared to WT (Fig. 4C, left panel). 262

In addition to organic acids, levels of amino acids also showed an increasing pattern 263

while NTRC expression was reduced. Asparagine, 3-cyano-alanine and homoserine were 1.5- 264

to 2-fold higher in NTRC-RNAi lines in immature fruits (Fig. 4C, left panel), and this change 265

became more severe while fruits ripened (Fig. 4C, right panel). Other amino acids, such as 266

tryptophan, histidine, glutamine and methionine, showed mild increases in NTRC-RNAi lines, 267

compared to WT. Metabolites, such as urea, amines, nucleosides and their derivate, were also 268

analyzed, but the results revealed only minor differences between the NTRC-RNAi lines and 269

WT at both stages (Fig. 4D). Overall, these results indicate interfering with NTRC expression 270

affects the balance between sugars, organic acids and amino acids during fruit development. 271

272

Interfering with NTRC expression in tomato fruits leads to an increase in the cellular 273

redox state of pyridine nucleotides 274

NAD(H) and NADP(H) redox couples are involved in various metabolic processes in 275

different subcellular compartments (Berger et al., 2004; Geigenberger and Fernie, 2014) and 276

also serve as cofactors for NTRC to reduce and regulate downstream plastidial proteins 277

(Pérez-Ruiz and Cejudo, 2009). To investigate whether NTRC down-regulation affects the 278

levels of these pyridine nucleotides, we next analyzed the levels of NAD(P)H and NAD(P)+ 279

in developing tomato fruits. In immature fruits (35 DAF), perturbing NTRC expression led to 280

a strong increase in the levels of the reduced forms, NADPH and NADH, while the levels of 281

the oxidized forms, NADP+ and NAD

+, decreased (Fig. 5A). This resulted in a 2- to 5-fold 282

increase in NADPH/NADP+ ratios and 1.5- to 2-fold increase in NADH/NAD

+ ratios in fruits 283



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of NTRC-RNAi lines in comparison to those of WT (Fig. 5C). 284

In fully ripe fruits (65 DAF), NTRC down-regulation also led to an increase in NADPH 285

and NADH, but the levels of the oxidized forms NADP+ and NAD

+ were not substantially 286

altered (Fig. 5B), which led to a minor increase in the ratios of NADPH/NADP+ and 287

NADH/NAD+ in NTRC-RNAi lines (Fig. 5D). Together, these results indicate reducing 288

NTRC expression caused an increased reduction state of the NAD(H) and NADP(H) redox 289

couples in developing tomato fruits with the effect being mitigated when fruits matured. 290

291

Interfering with NTRC expression in tomato fruits leads to a decrease in the 292

redox-activation state of AGPase and decreased levels of ADP-Glc 293

To investigate possible mechanisms involved in the inhibition of fruit starch 294

accumulation in response to decreased NTRC expression, we measured activities of enzymes 295

involved in starch metabolism. We first analyzed AGPase, which catalyzes the first 296

committed step of starch biosynthesis, since its activity is regulated via thiol-disulfide 297

modulation involving Trx and NTRC systems in the plastid (Geigenberger and Fernie, 2014; 298

Geigenberger et al., 2017). Since AGPase is activated by reduction of an intermolecular 299

disulfide bond between the two small subunits (AGPase small-subunit 1, APS1) of the 300

heterotetrameric holoenzyme (Ballicora et al., 2000; Tiessen et al., 2002; Hendriks et al., 301

2003), we assayed the changes in APS1 redox state by analyzing its monomerisation pattern 302

when NTRC expression was reduced. In developing fruits (35 DAF), perturbing NTRC 303

expression slightly decreased the monomerization ratios of APS1 (Fig. 6A) by approximately 304

20-30%, indicating an increased oxidation of the protein. Samples from ripe fruits (65 DAF) 305

were also tested, but there were no consistent differences between WT and RNAi lines in the 306

monomerization of APS1 (Supplemental Fig. S5A). 307

AGPase is also subject to allosteric regulation, being activated by 3-PGA and inhibited 308

by Pi (Kleczkowski, 1999; Hwang et al., 2005). Redox-regulation of AGPase alters the 309

kinetic properties of the holoenzyme, with oxidation of APS1 leading to an enzyme less 310

sensitive for allosteric activation (Tiessen et al., 2002; Hendriks et al., 2003). To investigate 311

whether perturbing NTRC expression in tomato fruits leads to a change in the kinetic 312

properties of AGPase, we assayed AGPase activity in the absence or presence of its allosteric 313

activator, 3-PGA, in 35 DAF fruits. As shown in Figure 6B, the NTRC-RNAi lines showed 314

comparable AGPase activity to WT when 3-PGA was omitted from the enzyme assay, but 315

there was a 30% decrease in AGPase activity compared to WT in the presence of 10 mM 316

3-PGA. To investigate 3-PGA activation kinetics in more detail, we performed AGPase 317

enzyme assays in the presence of varied concentrations of 3-PGA. Figure 6C shows that, in 318



12

WT, increasing concentrations of 3-PGA from 0.025-10 mM led to a progressive increase in 319

AGPase activity from 0.2-6.0 nmol min-1

mg protein-1

. AGPase from NTRC-RNAi lines was 320

less sensitive to 3-PGA activation leading to 30-50% lower activities when incubated in the 321

presence of 0.25-10 mM 3-PGA. This concentration range corresponds to the in vivo 322

concentration of 3-PGA in the plastid, estimated to be around 0.3-0.5 mM using non-aqueous 323

fraction of growing potato (Solanum tuberosum) tubers (Farré et al., 2001; Tiessen et al., 324

2002) and developing barley (Hordeum vulgare) seeds (Tiessen et al., 2012), and 0.7-1 mM 325

using the 3-PGA levels of Supplemental Table S1 (70-100 nmol g FW-1

), given that the 326

vacuole contains no 3-PGA and comprises 90% of the cell volume. The overall activities of 327

AGPase in fully ripe fruits (65 DAF), which are characterized by an overall suppression of 328

starch synthesis (Fig. 3B), were 10-times lower than in immature fruits (35 DAF) either in the 329

presence or absence of 3-PGA in the enzyme assay, and there was no significant difference 330

between WT and NTRC-RNAi lines (Supplemental Fig. S5B). 331

ADP-glucose (ADP-Glc) is the direct product of the reaction catalyzed by AGPase and 332

represents the ultimate precursor of the final polymerizing reactions of starch synthesis. In 35 333

DAF fruits, NTRC down-regulation decreased ADP-Glc concentration to 50-75% of WT 334

level (Fig. 6D). This provides independent confirmation that NTRC silencing leads to an 335

inhibition of AGPase activity in vivo. 336

337

NTRC down-regulation leads to deceased activity of soluble starch synthase 338

In addition to AGPase, we also assayed other enzymes involved in starch metabolism in 339

35 DAF fruits. Soluble and granule-bound starch synthases catalyze the final polymerizing 340

reactions of starch biosynthesis. As shown in Table 1, perturbing NTRC expression decreased 341

the activity of soluble starch synthase to 25-40% of WT level, while the activity of 342

granule-bound starch synthase only slightly decreased. Soluble starch synthase is subject to 343

post-translational redox regulation and a potential target of NTRC in vitro (Skryhan et al., 344

2015; Skryhan et al., 2018). In contrast, NTRC down-regulation had no significant effect on 345

the activities of starch phosphorylases and starch hydrolases, which are involved in starch 346

degradation (Table 1). Together these results imply down-regulation of NTRC expression 347

leads to decreased starch accumulation via a joint inhibition of both AGPase and soluble 348

starch synthase. 349

350

NTRC down-regulation has minor effects on the redox state of 2-Cys Prxs in tomato 351

fruits 352

2-Cys Prxs are well-known targets of NTRC in Arabidopsis leaves where they are 353



13

involved in hydrogen peroxide removal during photosynthesis (Cejudo et al., 2012). 354

Activation of 2-Cys Prxs involves reduction of an intermolecular disulfide bond linking the 355

two subunits of the homodimer (Pérez-Ruiz et al., 2006; Pérez-Ruiz et al., 2017). To 356

investigate the role of NTRC in reduction of 2-Cys Prxs in tomato fruits, we assayed the 357

monomer/dimer ratio of 2-Cys Prxs in tomato fruit tissues of NTRC-RNAi lines compared to 358

WT using non-reducing SDS gels. NTRC down-regulation slightly decreased the 359

monomerization ratio of 2-Cys Prxs in both 35 and 65 DAF tomato fruits (Supplemental Fig. 360

S6), even though the decrease was not consistent in all transgenic lines. The role of NTRC in 361

regulating the redox state of 2-Cys Prxs seems to be less important in tomato fruit than in 362

Arabidopsis leaves, where NTRC deficiency leads to a strong degree of 2-Cys-Prx oxidation 363

(Cejudo et al., 2012). However, other Trxs might have compensated for the deficiency of 364

NTRC with respect to 2-Cys-Prx reduction in tomato fruits. 365

366

Discussion 367

NTRC is a NADPH-dependent thiol redox enzyme, which has been found in 368

chloroplasts as well as in non-photosynthetic plastids (Kirchsteiger et al., 2012). In 369

photosynthetic tissues, the role of NTRC in carbon metabolism and redox balance of 370

chloroplasts in response to fluctuating light environments has been well characterized 371

(Geigenberger et al., 2017; Zaffagnini et al., 2018). Previous studies led to the perspective 372

that NTRC might function differently in heterotrophic tissues; however, our understanding as 373

to how remains fragmentary (Kirchsteiger et al., 2012). In this work, we used genetic and 374

biochemical approaches to directly test the role of NTRC in developing tomato fruits, which 375

represent heterotrophic tissues dependent on sucrose import with no substantial 376

photosynthetic carbon assimilation (Lytovchenko et al., 2011). Our results show NTRC does 377

not affect developmental and ripening programs of the fruit but promotes transient starch 378

accumulation by activating AGPase and soluble starch synthase to increase overall sink 379

activity and hence final fruit size, dry matter and sugar content. In contrast to this, NTRC 380

leads to a decreasing pattern of organic and amino acids and a decrease in the NAD(P)(H) 381

reduction state. This indicates NTRC serves as an important hub to orchestrate carbon 382

metabolism and redox balance in heterotrophic fruit tissues to optimize fruit growth and 383

quality in response to sucrose supply. 384

385

NTRC is important to optimize fruit size and quality 386

Tomato is an important horticultural crop and serves as major component of our diet 387

(Carrari and Fernie, 2006). There has been considerable interest to optimize tomato fruit yield 388



14

and quality by using genetic approaches to alter sink metabolism and ripening (Giovannoni, 389

2004; Carrari and Fernie, 2006), but there is a lack of studies investigating the role of redox 390

regulation in this context. In the present paper, NTRC expression was specifically decreased 391

by 50-75% in developing tomato fruits using RNA interference under the control of a 392

fruit-specific promoter (Fig. 1). Perturbing NTRC expression did not affect developmental 393

and ripening programs of the fruit (Supplemental Fig. S2) but led to a decrease in final fruit 394

size and weight with fully mature fruits containing less dry matter and more water than WT 395

(Fig. 2). The phenotype was accompanied by a decrease in the levels of soluble sugars (Fig. 396

3D; Fig. 4A; Supplemental Table S1) and an opposing increase in many organic acids and 397

amino acids as well as phenolic compounds (Fig. 4, B, C and D; Supplemental Table S1). 398

Domesticated tomatoes tend to accumulate high amounts of glucose and fructose. Thus, the 399

total abundance of glucose, fructose and other minor sugars determines the sweetness of 400

tomato fruits. The acid-to-sugar ratio determines the sourness of fruits, and it is also an 401

important quality parameter (Davies and Hobson, 1981; Gierson and Kader, 1986). Thus, it is 402

very likely the changes we observed in response to NTRC down-regulation would eventually 403

influence the taste and flavor of tomato fruits. 404

NTRC affects growth and development of Arabidopsis plants (Pérez-Ruiz et al., 2006; 405

Lepisto et al., 2009; Toivola et al., 2013; Thormählen et al., 2015), but this is mainly ascribed 406

to its role in photosynthesis and leaf metabolism, rather than in non-photosynthetic tissues 407

(Kirchsteiger et al., 2012). The present study provides direct evidence for a previously 408

unrecognized role of NTRC to regulate the development of heterotrophic storage organs to 409

optimize yield quantity and quality parameters. This shows NTRC has important functions 410

beyond photosynthesis that affect growth and development. Although immature fruits are 411

green and contain chlorophyll, it is unlikely the changes in fruit growth reported above are 412

due to a role of NTRC in regulating photosynthetic processes within the fruit (Supplemental 413

Fig. S2). Indeed, previous studies have shown knock-out of chlorophyll synthesis in pale 414

tomato fruits does not have significant effects on fruit physiology, growth and development 415

(Lytovchenko et al., 2011). 416

There are conflicting studies concerning the effect of NTRC overexpression on the 417

growth of Arabidopsis plants. While Toivola et al. (2013) provide evidence that mild 418

overexpression of NTRC results in bigger plants, there is evidence from Ojeda et al. (2018) 419

that strong NTRC overexpression may have opposite effects. It is tempting to speculate 420

whether increased expression of NTRC may lead to larger tomato fruits. However, given the 421

results in Arabidopsis, this may depend on the level of NTRC overexpression. More studies 422

will be necessary to clarify this question. 423



15

424

NTRC regulates transient starch accumulation in developing fruits 425

Tomato fruit development is characterized by a transient accumulation of starch in 426

immature fruits which serves as a carbon reserve for soluble sugar accumulation during 427

ripening and finally affects fruit size (Schaffer et al., 2000). In the present study, interfering 428

with NTRC expression resulted in a 50-75% decrease in the level of the transient starch pool 429

in immature fruits (Fig. 3A), which was accompanied by a decrease in soluble sugars during 430

maturation and decreased fruit size (Fig. 3, C and D; Fig. 4A). Our results show the decrease 431

in transient starch accumulation associates with a decrease in the redox-activation of AGPase 432

(Fig. 6), which is the rate-limiting enzyme in the pathway of starch synthesis in developing 433

tomato fruits (Schaffer et al., 2000) and other plant tissues (Geigenberger et al., 2004; 434

Vigeolas et al., 2004; Neuhaus and Stitt, 2006; Faix et al., 2012) and a known target of 435

NTRC (Geigenberger et al., 2017). NTRC down-regulation led to a decrease in the reduction 436

state of the small subunit of AGPase (Fig. 6A), which was accompanied by a 30-50% 437

decrease in the sensitivity of the AGPase holoenzyme for its allosteric activator 3-PGA (Fig. 438

6, B and C). While the subcellular concentration of 3-PGA in the plastid has been estimated 439

to yield 0.3-0.5 mM (Farré et al., 2001; Tiessen et al., 2002; Tiessen et al., 2012), the results 440

of Figure 6B indicate NTRC down-regulation will lead to a 30-50% lower AGPase activity 441

under in vivo conditions in developing tomato fruits. In confirmation to this, NTRC silencing 442

led to a 40% decrease in the in vivo level of ADP-Glc (Fig. 6D), which is the immediate 443

product of the reaction catalyzed by AGPase, providing independent in vivo evidence that 444

NTRC down-regulation restricts AGPase activity in developing fruits. 445

This is consistent with previous studies documenting the primary role of AGPase in 446

regulating starch accumulation, soluble sugar content and growth of tomato fruits. First, 447

increased AGPase activity in near-isogenic tomato lines led to elevated starch accumulation 448

in immature fruits, followed by increased soluble sugar content and larger size of fully ripe 449

fruits (Petreikov et al., 2009). Secondly, increased AGPase redox-activation by manipulation 450

of NAD(P)(H) redox states led to elevated rates of starch synthesis in immature fruits and a 451

subsequent increase in the content of soluble sugars in fully ripe fruits (Centeno et al., 2011). 452

Interestingly, NTRC down-regulation also led to a strong decrease in the activity of 453

soluble starch synthase catalyzing the subsequent step in the pathway of starch synthesis, 454

while enzyme activities involved in starch degradation were not significantly affected in 455

developing fruits (Table 1). Soluble starch synthase has been identified as a redox-regulated 456

enzyme and potential target of NTRC in vitro, with reduction via NTRC or Trx f leading to 457

an increase in its activity in vitro (Skryhan et al., 2015; Skryhan et al., 2018). Our results 458



16

provide evidence the relationship between NTRC and soluble starch synthase, which has 459

been shown in vitro, also extends to the in vivo situation of developing fruits. Intriguingly, 460

our data show a combined effect of NTRC on both AGPase and soluble starch synthase, 461

catalyzing the first committed and final polymerizing steps of starch biosynthesis, 462

respectively. This indicates a role of NTRC in stimulating starch accumulation and ultimately 463

growth and quality of tomato fruits by combined redox-activation of both enzymes. 464

NTRC is also involved in transient starch accumulation in leaves of Arabidopsis plants 465

during day-night cycles (Lepistö et al., 2013; Thormählen et al., 2015). NTRC deficiency in 466

Arabidopsis T-DNA insertion mutants leads to a 30-40% restriction in redox-activation of 467

AGPase and starch synthesis upon illumination, both parameters being further attenuated in 468

double mutants with combined deficiencies in NTRC and Trx f1 (Thormählen et al., 2015). In 469

leaves, NTRC is dependent on NADPH produced by the photosynthetic light reactions. 470

Furthermore, it is involved in the regulation of the Calvin-Benson cycle, and therefore in the 471

provision of the substrates and the allosteric activator of AGPase, which will indirectly 472

contribute to the regulation of starch synthesis (Thormählen et al., 2015). This phenomenon 473

contrasts to the situation in heterotrophic tissues where AGPase redox-state and starch 474

synthesis are both regulated by the momentary supply of sugars via the phloem (Tiessen et al. 475

2002; Michalska et al. 2009). Sugar translocation from leaves to heterotrophic organs has 476

been shown to vary in response to development (Walker and Ho, 1977) and diurnal 477

alterations (Bénard et al., 2015). Changes in sugar availability may affect the activity of the 478

oxidative pentose phosphate pathway (OPPP) (Geigenberger and Fernie, 2014), leading to 479

alterations in the NADP(H) redox state and, thus, in NTRC activity. Little is known about 480

changes in NADP(H) levels in plant tissues. There is evidence for diurnal alterations in 481

NADP(H) levels and redox states in Arabidopsis leaves (Thormählen et al., 2015). Moreover, 482

glucose - but not sucrose - feeding leads to an increase in the NADPH/NADP+ ratio in 483

heterotrophic potato tuber tissue (Kolbe et al., 2005). This implies that in developing fruits, 484

NTRC modulates starch synthesis via redox regulation of AGPase and soluble starch 485

synthase in response to changes in sugar import, which lead to alterations of NADPH by 486

affecting the OPPP. This is important to adjust the sink activity of the fruit to the supply of 487

sucrose from the leaves to optimize fruit growth. 488

489

NTRC regulates the redox-balance of NAD(P)H in developing fruits 490

In heterotrophic plastids lacking photochemical reactions, redox homeostasis depends 491

mainly on NADPH, so NTRC, which depends on NADPH, may play an essential role in 492

maintaining the plastidial redox poise. Our results show interfering with NTRC expression 493



17

strongly affects the cellular redox states of NAD(H) and NADP(H) in developing tomato 494

fruits (Fig. 5), which provides evidence that NTRC is a major regulator of cellular redox 495

homeostasis in this heterotrophic tissue. While, at the moment, the underlying mechanisms 496

are unresolved, NTRC uses NADPH as a substrate and, thus, may influence the 497

NADPH/NADP+ ratio directly or via regulation of plastidial pathways consuming or 498

producing NADPH, such as the OPPP. However, direct effects of NTRC on enzymes of the 499

OPPP have not been studied yet. A further possibility is that NTRC affects the redox shuttles 500

between plastids and mitochondria leading to changes in interorganellar redox regulation. 501

Previous studies in tomato fruits provide evidence that plastidial and mitochondrial 502

NAD(P)H systems are interconnected via the malate valve (Centeno et al., 2011). An 503

increase in the NADPH redox state in response to NTRC down-regulation in the plastid may 504

therefore be transmitted to mitochondria leading to an increase in NADH redox state and vice 505

versa. Since adenylate energy states were not significantly changed in NTRC-RNAi lines 506

(Supplemental Table S2), it is quite unlikely that the changes in redox states are due to 507

changes in respiration rates. 508

NTRC deficiency also leads to an increase in NADP(H) and NAD(H) reduction states in 509

Arabidopsis plants (Thormählen et al., 2015; Thormählen et al., 2017). However, this is most 510

probably due to NTRC regulating the balance between photosynthetic light reactions and the 511

Calvin-Benson cycle as well as NADPH export via NADP-dependent MDH of the malate 512

valve. Taken together, these results identify NTRC as a central hub controlling the cellular 513

redox poise in non-photosynthetic as well as photosynthetic tissues. 514

Based on the work in Arabidopsis plants, NTRC has been found to play a major role in 515

detoxification of hydrogen peroxide by providing reductants to 2-Cys Prxs, with NTRC 516

deficiency leading to a strong decreased in the 2-Cys Prxs reduction state in 517

photosynthesizing leaves (Pérez-Ruiz et al., 2006; Pérez-Ruiz et al., 2017). Compared to this, 518

our results show NTRC down-regulation leads to a relative minor decrease in 2-Cys Prxs 519

reduction in developing and ripe tomato fruits (Supplemental Fig. S6). However, it must be 520

noted that photosynthetic activity leads to increased rates of hydrogen peroxide production, 521

which requires increased antioxidant activity via the NTRC-2-Cys Prxs system (Pérez-Ruiz et 522

al., 2006; Pérez-Ruiz et al., 2017). These results suggest the antioxidant function of NTRC is 523

less important in non-photosynthetic than in photosynthetic tissues. 524

525

NTRC affects the carbon-nitrogen balance in developing fruits 526

While our results show NTRC down-regulation leads to a decrease in soluble sugar 527

levels, there is an opposing increase in the levels of several amino acids in developing fruits 528



18

(Fig. 4; Supplemental Table S1). Since total protein level remains unchanged (Supplemental 529

Fig. S4), it is more likely NTRC down-regulation leads to a stimulation of amino acid 530

synthesis, rather than protein degradation. Because amino acid biosynthesis requires reducing 531

equivalents as substrates, production of amino acids may have been promoted by the 532

increased NAD(P)H/NAD(P)+ ratios in developing fruits of the NTRC-RNAi lines (Fig. 5). 533

Increased levels of amino acids in the face of decreased sugar levels are also observed in 534

leaves of Arabidopsis mutants lacking NTRC (Lepisto et al., 2009; Thormählen et al., 2015), 535

indicating there are similar effects of NTRC on amino acid synthesis in photosynthetic and 536

non-photosynthetic tissues. Inverse relationships between sugars and amino acids have also 537

been found in other plant tissues, such as barley seeds (Faix et al., 2012) or potato tubers 538

(Trethewey et al., 1998; Fernie et al., 2003; Roessner-Tunali et al., 2003) across different 539

genetic and physiological manipulations. Overall, these results suggest NTRC is involved in 540

the regulation of the sugar-amino acid balance to prevent osmotic imbalances in the 541

developing fruit. 542

543

Conclusions 544

NTRC is a NADPH-dependent thiol redox system located in photosynthetic as well as in 545

non-photosynthetic plastids. While previous approaches were mainly performed in 546

Arabidopsis and photosynthetic leaves to date, there has been a lack of knowledge on the role 547

of NTRC in heterotrophic tissues and crop plants. By establishing NTRC-RNA interference 548

specifically in tomato fruit tissues, our results uncover the importance of NTRC in 549

developing tomato fruits. In immature fruits, NTRC promotes transient starch accumulation 550

by activating AGPase and soluble starch synthase to increase the carbohydrate stock for 551

soluble sugar accumulation during ripening to optimize final fruit yield and quality. NTRC 552

also plays a pivotal role in regulating the redox poise of NAD(P)(H) during fruit development, 553

which represents the primary redox system in non-photosynthetic tissues. In summary, NTRC 554

acts as a central hub in regulating carbon metabolism and redox balance in heterotrophic 555

tomato fruits, similar to its established role in photosynthetic tissues. Our results indicate the 556

activity of NTRC is indispensable to optimize fruit growth as well as tomato yield quantity 557

and quality. 558



19

Materials and Methods 559

Plant cultivation and harvest 560

Tomato (Solanum lycopersicum cv Moneymaker) were grown as detailed in the 561

literature (Carrari et al., 2003; Nunes-Nesi et al., 2005). Fruits were harvested at the mature 562

green (35 DAF) and ripe stage (65 DAF) with the intact pericarp being immediately frozen in 563

liquid nitrogen for further analysis. 564

565

NTRC-RNAi construct and plant transformation 566

The NTRC gene fragment was amplified from potato (Solanum tuberosum) cDNA with 567

attB1-RNAi-NTRC (5’-ggggacaagtttgtacaaaaaagcaggctcagatgtcaggaattttgatca-3’) and 568

attB2-RNAi-NTRC (5’-ggggaccactttgtacaagaaagctgggtcgcttgtcagatatctctccactgata-3’), and 569

cloned into modified pBINAR-RNAi vector (pBIN19 backbone containing a patatin B33 570

promoter and a GATEWAY cassette) using a commercial GATEWAY cloning kit (Invitrogen). 571

Tomato plants were transformed as described before (Tauberger et al., 2000) and the 572

transformants were selected by kanamycin. The patatin B33 promoter acts as a fruit-specific 573

promoter when expressed in tomato plants (Rocha-Sosa et al., 1989; Frommer et al., 1994; 574

Obiadalla-Ali et al., 2004). 575

576

Molecular characterization of NTRC-RNAi lines 577

Genomic DNA extraction followed the CTAB based protocol (Doyle and Doyle, 1987). 578

The kanamycin resistance gene (nptII) was amplified with primer pairs, Kana-F 579

(5’-aactgttcgccaggctcaag-3’) and Kana-R (5’-tcagaagaactcgtcaagaagg-3’), and the internal 580

control (CAC, SGN-U314153) was amplified with CAC-F (5’-cctccgttgtgatgtaactgg-3’) and 581

CAC-R (5’-attggtggaaagtaacatcatcg-3’). The RNAzol (Sigma-Aldrich) was used for RNA 582

extraction, and 500 μg of total RNA was used for preparing cDNA with Maxima Reverse 583

Transcriptase (Thermo). The SYBG reagent (Biorad) was used for real-time PCR, and the 584

signal was recorded via iQ5 Multicolor Real-Time PCR Detection System (Biorad). The 585

primers, NTRC-F (5’-aggtgtatttgcagctggagatg-3’), NTRC-R (5’-ttcttcagtagggggctggtggaa-3’), 586

NTRC-2F (5’-ggaccaactaagggcatcagcaa-3’) and NTRC-2R (5’-cgagcacagactcttctcctg-3’), 587

were used to amplify NTRC genes (LOC101254347; LOC101266017), respectively. For 588

detecting the expression of ethylene production and ripening genes, the primer pairs, 589

SAHH2-F (5’-gacgctcacagtggaacaacga-3’), SAHH2-R (5’-tcaactttccagcaaaatatcccc-3’), 590

NOR-F (5’-agagaacgatgcatggaggtttgt-3’) and NOR-R (5’-actggctcaggaaattggcaatgg-3’), were 591

used for amplification of SAHH2 and NOR genes, respectively (Yang et al., 2017; Ma et al., 592

2019). The CAC gene described above served as an internal control (Expósito-Rodríguez et 593



20

al., 2008). Total protein was extracted from 35 mg of pulverized sample with 150 μl of 594

double strength Laemmli buffer (Laemmli, 1970) containing 10 mM DTT, and 10 μl of 595

protein extract was used for SDS-PAGE and immunoblotting. The membrane was blocked 596

with 5% (w/v) non-fat milk and incubated with the first antibody of NTRC (1:1000 dilution), 597

Lhcb2 (Agrisera, 1:5000) or RbCL (Agrisera, 1:10000 dilution) at 4oC overnight, respectively. 598

The secondary antibody conjugating horseradish peroxidase (HRP) (Agrisera, 1:10000 599

dilution) was applied onto the membrane, and the membrane was incubated at 25oC for one 600

hour. The membrane reacted with Prime ECL solution (GE Healthcare), and the signal was 601

visualized with X-ray film (Fuji). 602

603

Spectrophotometric metabolite measurements 604

(a) Starch and soluble sugar quantification 605

The extraction and detection of starch and soluble sugar followed the method described 606

previously (Hendriks et al., 2003). In brief, 20 mg of pulverized sample was extracted with 607

250 μl of 80% ethanol twice followed by another 250 μl of 50% ethanol at 90oC for 30 min. 608

The supernatant was used for the detection of soluble sugar, and the pellet was dried out for 609

the detection of starch. For measuring starch level, the pellet was re-suspended with 400 μl of 610

0.1 M sodium hydroxide and incubated at 95oC for one hour. The solution was then 611

neutralized to pH 7 with HCl-acetate buffer (0.5 M HCl prepared in 0.1 M acetate-NaOH 612

buffer, pH 4.9), and 40 μl of aliquot was incubated with 110 μl of starch degradation buffer 613

(2.8 U ml-1

amyloglucosidase, 4 U ml-1

α-amylase, 50 mM acetate-NaOH buffer) at 37oC 614

overnight. Fifty microliters of digested sample was firstly mixed with 160 μl of glucose 615

determination buffer (3.4 U ml-1

G6PDH, 3 mM ATP, 1.3 mM NADP, 3 mM MgCl2, 100 mM 616

HEPES-KOH, pH 7.0). When the reaction saturated, and 0.45 U hexokinase was applied to 617

detect the formation of NADPH, which was proportional to starch amount, at 340 nm via a 618

spectrophotometer (FilterMax F5, Molecular Device). 619

For measuring soluble sugar level, 30 μl of supernatant was firstly mixed with 200 μl of 620

detection buffer (0.7 U ml-1

G6PDH, 1.7 mM ATP, 0.8 mM NADP, 5 mM MgCl2, 50 mM 621

HEPES-KOH, pH 7). When the reaction was stable, 0.5 U hexokinase, 1.2 U 622

phosphoglucoisomerase and 133 U invertase were applied to detect glucose, fructose and 623

sucrose, respectively, via the formation of NADPH at 340 nm. 624

For the detection of maltose, the method described previously was followed with several 625

modifications (Shirokane et al., 2000). Briefly, the above ethanol extract was dried out by 626

heating at 95oC and substituted with equal volume of double distilled water. Then, 30 μl of 627

extract was mixed with 200 μl of detection buffer described above with addition of 0.5 U 628



21

hexokinase. When the reaction was stable, 0.2 U maltose phosphorylase was applied to detect 629

maltose via NADPH formation in the florescence mode of spectrometer (Ex: 340 nm, Em: 630

465 nm). Varied concentration of maltose (Sigma-Aldrich) served as the standard for 631

calculation. 632

(b) Hexose phosphates and 3-phosphoglycerate (3-PGA) 633

The assay of hexose-phosphates and 3-PGA was performed as described before (Häusler 634

et al., 2000). Briefly, 50 mg of pulverized sample was suspended in 500 μl of ice-cold 0.5 M 635

HClO4 and incubated at 4oC for 10 min. Then, leaf residue was removed by centrifuging for 636

10 min at 20,000 g. The pH of the supernatant was adjusted to the range from 5.0 to 6.0 using 637

2.5 M K2CO3 solution, and the supernatant was then centrifuged for 10 min at 20,000 g to 638

remove formed precipitant. Around 5 mg of active charcoal was applied into solution, which 639

was incubated on ice for 20 min. Charcoal was further removed via centrifuging for 10 min at 640

20,000 g in 4oC. For measuring hexose phosphates, 50 μl of supernatant was mixed with 50 641

μl of water and reacted with 100 μl of detection buffer (200 mM Tris-HCl, pH 8.1, 1.6 mM 642

NADP, 10 mM MgCl2, 2 μM glucose-1,6-bisphosphate). When the reaction was stable, 0.1 U 643

glucose-6-phosphate dehydrogenase, 0.35 U phosphoglucoisomerase and 0.04 U 644

phosphoglucomutase were applied to detect G6P, F6P and G1P, respectively, via NADPH 645

formation in the fluorescence mode of the spectrometer (Ex: 340 nm, Em: 465 nm). For 646

detecting 3-PGA, 50 μl of supernatant was mixed with 50 μl of water and reacted with 100 μl 647

of detection buffer (200 mM HEPES-NaOH, pH 7.0, 2 mM MgCl2, 2 mM ATP, 8 μM NADH, 648

0.8 U ml-1

Glyceraldehyde 3-phosphate dehydrogenase). When the reaction was stable, 0.45 649

U phosphoglycerate kinase was applied to detect 3-PGA via NADH consumption in the 650

florescence mode of spectrometer described above. Varied concentration of NADPH and 651

NADH (Carl Roth) served as the standard for calculation. 652

(c) Pyridine nucleotides 653

The assay of pyridine nucleotides (NAD+, NADP

+, NADH and NADPH) was performed 654

as described previously (Lintala et al., 2014). In short, 25 mg of pulverized sample was 655

suspended in 250 μl of 0.1 M HClO4 for detecting NAD+ and NADP

+. Another 25 mg of 656

sample was suspended in 250 μl of 0.1 M KOH for detecting NADH and NADPH. Samples 657

were further incubated on ice for 10 min and then centrifuged at 20,000 g for 10 min at 4oC 658

to remove the pellet. The supernatant was incubated at 95oC for 2 min, and the pH value was 659

adjusted to the range from 8.0 to 8.5 with 0.1 M KOH prepared in 0.2 M Tris-HCl (pH 8.4) 660

or HClO4 prepared in 0.2 M Tris-HCl (pH 8.4), respectively. For measuring NAD(H), 50 μl 661

of extract was mixed with 50 μl of water and reacted with 50 μl of detection buffer 662

containing 300 mM Tricine-KOH (pH 9.0), 12 mM EDTA, 1.5 M ethanol, 0.3 mM phenazine 663



22

ethosulfate (PES), 1.8 mM methylthiazolydiphenyl-tetrazolium bromide (MTT), and 18 U 664

ml-1

alcohol dehydrogenase. For measuring NADP(H), another 50 μl of extract was mixed 665

with 50 μl of water and reacted with 50 μl of detection buffer containing 300 mM 666

Tricine-KOH (pH 9), 12 mM EDTA, 9 mM Glc-6-P, 0.3 mM PES, 1.8 mM MTT, and 9 U 667

ml-1

Glc-6-P dehydrogenase. The absorbance at wavelength 570 nm was recorded in a 668

spectrometer (FilterMax F5, Molecular Device). Pyridine nucleotides (Carl Roth) solutions 669

prepared in water were used to generate the standard curve for calculation. 670

(d) ADP-glucose (ADP-Glc) 671

The extraction of ADP-Glc was performed as described previously with minor 672

modification (Lunn et al., 2006). Briefly, 50 mg of pulverized sample was suspended in 375 673

μl of ice-cold chloroform/methanol mixture (3:7) and vortexed vigorously at 4oC for 10 min 674

followed by the addition of 300 μl of double distilled water for phase partitioning. After 675

centrifugation at 18000 g at 4oC for 10 min, the supernatant was transferred to a new tube and 676

dehydrated in a vacuum at ambient temperature. The dried pellet was re-suspended in 70 μl 677

of water, and 10 μl of the suspension was used for detecting ADP-Glc. 678

ADP-Glc was analyzed by LC-MS using a Dionex Ultimate 3000 UHPLC (Thermo 679

Fisher Scientific, Waltham, Massachusetts, USA) and a timsTOF QTOF (Bruker Daltonik, 680

Fahrenheitstr. 4, 28359 Bremen, Germany) in combination. The assay supernatant was 681

injected into a C18 column (Ultra AQ C18, 150 x 2.1 mm, 3 µm, Restek, Bellefonte, 682

Pennsylvania, USA) with 300 µl min-1

flow at 22°C. The solvents used were (A) water and 683

(B) acetonitrile, both including 0.1% (v/v) formic acid. The gradient started at 1% B for 1 684

min, followed by a ramp to 25% B within 10 min. After a 4 min washing step at 95% B, the 685

gradient turned back to 1% B and kept constant for 3 min equilibration. After the column 686

compartment, the analytes passed a diode array detector (DAD) at 254 nm before entering the 687

mass spectrometer. An electrospray ionization (ESI) source in negative mode was used at 688

5000 V capillary voltage and nitrogen as dry gas, 8 L min-1

at 200°C. The timsTOF mass 689

spectrometer was run in MS mode from 50-1300 m/z mass ranges. Compounds were 690

identified using the specific mass (m/z), isotope pattern, and retention time after standard 691

injection. Data were acquired by otofControl 4.0. The evaluation was performed by 692

DataAnalysis 5.1. All software tools were provided by Bruker (Bruker Daltonik, 693

Fahrenheitstr. 4, 28359 Bremen, Germany). All solvents were supplied in LC-MS-grade by 694

Biosolve. The ADP-Glc standard was bought from Sigma-Aldrich. 695

(e) Total chlorophyll 696

The quantification of total chlorophyll was performed according to a previous approach 697

(Porra et al., 1989). In short, 35 mg of pulverized sample was extracted with 1 ml of ice-cold 698



23

80% acetone and vortexed at 4oC for 2 min followed by centrifugation to remove insoluble 699

residues. Then, 250 μl of supernatant was mixed with 750 μl of 80% acetone and applied into 700

a quartz cuvette. The optical density of extract was determined by measuring the absorbance 701

at wavelength 664 and 647 nm. 702

703

Measurement of adenylate nucleotide 704

The extraction of adenylate nucleotides followed a previously published method (Jelitto 705

et al., 1992). Briefly, 50 mg of pulverized sample was extracted with 600 μl of extraction 706

buffer (16% TCA, 5 mM EGTA), and rigorously vortexed at 4oC for one hour. The 707

supernatant was then washed twice with 4 ml of cold water-saturated diethyl ether to remove 708

TCA. The lower water phase was transferred to a new microtube, and the pH was adjusted to 709

between 6.0 and 7.0 with neutralization buffer (5 M KOH, 1 M triethanoamine). The sample 710

was placed under a hood for one hour to let the remaining diethylether evaporate. The 711

detection procedure was described before (Haink and Deussen, 2003) with several 712

modifications: 85 μl of extract reacted with 20 μl of citrate buffer (62 mM Citric acid, 76 mM 713

KH2PO4, pH 5.2) and 20 μl of 50% chloroacetaldehyde at 80oC for 15 min. Then, 20 μl of 714

sample was applied into a NUCLEODUR 100-5 C18 ec column (MACHEREY-NAGEL) and 715

eluted with the mixture of mobile buffer A (10 mM KH2PO4, 5.7 mM 716

tetrabutylammoniumhydrogensulfate, pH 5.4) and mobile phase B (90% acetonitrile) under a 717

flow rate of 0.8 ml min-1

. The gradient was as described below: 0 min (100% A / 0% B), 18 718

min (86% A / 14% B), 36 min (56% A / 44% B), 38.4 min (17.5% A / 82.5% B), 39.6 min 719

(100% A / 0% B), 42 min (100% A / 0% B). The signal was analyzed by a fluorescence 720

detector (FINNIGAN SURVEYOR PDA Plus Detector, Thermo), which was set to 280 nm 721

excitation wavelength and 410 nm emission wavelength. Standards were provided by 722

Sigma-Aldrich. 723

724

Metabolite profiling: GC-TOF-MS 725

Metabolic profiling was performed using a method previously described with 726

modification (Roessner et al., 2001; Lisec et al., 2006; Erban et al., 2007). In brief, for the 727

extraction, 50 mg of tissue (fresh weight) was ground in 360 µl of cold (-20°C) methanol 728

containing internal standards (10 μl of ribitol, 0.2 mg ml-1

in water and 10 μl of 13C-sorbitol, 729

0.2 mg ml-1

in water) for the relative quantification. First, the samples were incubated at 4°C 730

for 30 min, and then the extract was carefully mixed with 200 μl of chloroform and 400 µl of 731

water. To force the separation of the polar and non-polar phase, a 15 min centrifugation step 732

at 25000 g was performed. For derivatization, 50 μl of the upper (polar) phase was dried in 733



24

vacuum. The pellet was resuspended in 10 μl of methoxyaminhydrochloride (20 mg ml-1

in 734

pyridine) and agitated for 90 min at 40°C. After the addition of 20 µl of BSTFA 735

(N,O-Bis[trimethylsilyl]trifluoroacetamide) containing 2.5 μl of retention time standard 736

mixture of linear alkanes (n-decane, n-dodecane, n-pentadecane, n-nonadecane, n-docosane, 737

n-octacosane, n-dotriacontane), the whole mixture was incubated at 40°C for 45 min. Finally, 738

1 μl of sample was injected into a GC–TOF–MS system (Pegasus HT, Leco, St Joseph, USA). 739

Samples were prepared by an auto-sampler system (Combi PAL, CTC Analytics AG, 740

Zwingen, Switzerland). Helium acted as carrier gas at a constant flow rate of 1 ml min-1

. Gas 741

chromatography was performed on an Agilent GC (7890A, Agilent, Santa Clara, USA) using 742

a 30 m VF-5ms column with 10 m EZ-Guard column. The injection temperature of the 743

split/splitless injector was set to 250°C, as well as the transfer line and the ion source. The 744

initial oven temperature (70°C) was permanently increased to a final temperature of 350°C 745

by 9°C min-1

. To avoid solvent contaminations, the solvent delay time was set to 340 s. 746

Compounds were ionized and fractionated at 70 eV. Mass spectra were recorded at 20 scans 747

s-1

with an m/z 35-800 scanning range. Chromatograms and mass spectra were evaluated 748

using ChromaTOF 4.5 and TagFinder 4.1 software (Luedemann et al., 2008). All solvents 749

were supplied in LC-MS-grade by Biosolve (Leenderweg 78, 5555 Valkenswaard, NL). 750

Standards were provided by Sigma-Aldrich. 751

752

Redox shift assay of APS1 and 2-Cys Prxs 753

To preserve the redox state, the total protein was extracted using a TCA-based method 754

(Hendriks et al., 2003). In short, 35 mg of frozen fruit tissue was extracted with 1 ml of 755

ice-cold 16% TCA prepared in diethylether and incubated at -20oC overnight. After 756

centrifugation, the supernatant was discarded, and the pellet was washed with ice-cold 757

acetone three times. The pellet was then suspended with double strength Laemmli buffer 758

(Laemmli, 1970) without reducing reagent and incubated at 90oC for 5 min with 1400 rpm 759

shaking speed. Then, 10 μl of protein extract was applied into 10 or 12% SDS-acrylamide gel 760

to perform immunoblot analyses detecting APS1 or 2-Cys Prxs, respectively. The 761

immunoblotting procedure was performed as described before (Hendriks et al., 2003; 762

Pérez-Ruiz et al., 2006), and Image-J was used for quantification of the protein signals. 763

764

Enzyme activity assay of starch metabolism 765

(a) ADP-glucose pyrophosphorylase (AGPase) 766

The AGPase activity assay followed a previously described method (Tiessen et al., 2002) 767

with several modifications. Briefly, 25 mg of pulverized frozen fruit tissue was extracted with 768



25

150 μl of 50 mM HEPES-KOH buffer (pH 7.5), and incubated on ice for 10 min followed by 769

centrifugation to remove leaf residue. The crude protein extract was applied into a Zeba 770

desalting column (Thermo) as described in the manual. Then, 20 μl of protein extract was 771

incubated with 180 μl of AGPase reaction buffer (50 mM HEPES-KOH, pH 7.5, 7 mM 772

MgCl2, 1.5 mM ATP, 0.5 mM G1P, varied concentration of 3-PGA) at 25oC for 30 min with 773

900 rpm shaking speed, and the reaction was terminated by mixing rigorously with 100 μl of 774

chloroform. Two microliters of the supernatant was used for quantification of ADP-Glc 775

produced during the enzyme assay. ADP-Glc was detected as described in the section of 776

ADP-Glc assay with modifications that an electrospray ionization (ESI) source in negative 777

mode was used at 4000 V capillary voltage and nitrogen as dry gas, 8 L min-1

at 250°C. 778

(b) Soluble starch synthase and granule-bound starch synthase 779

The assay was performed as previously described with modification (Kulichikhin et al., 780

2016). Briefly, 25 mg of pulverized sample was suspended in 150 μl of ice-cold buffer (100 781

mM HEPES-NaOH, pH 7.5, 8 mM MgCl2, 2 mM EDTA, 12.5% glycerol, 5% PVP-10, 782

protease inhibitor cocktail), and incubated on ice for 10 min followed by centrifugation to 783

separate the supernatant and insoluble pellet. For detecting soluble starch synthase, 50 μl of 784

the supernatant was applied into a self-packed Sephadex G25 column equilibrated with 50 785

mM HEPES-NaOH (pH 7.5) for desalting and mixed with 50 μl of double strength reaction 786

buffer (100 mM HEPES-NaOH, pH 7.5, 2 mM ADP-Glc, 0.2% amylopectin) followed by 787

incubation at 30oC for 30 min. The reaction was terminated by the addition of one-third 788

volume chloroform with vigorous vortex, and the upper phase was used for detecting the 789

production of ADP during the reaction. For the granule-bound starch synthase, the insoluble 790

pellet was washed twice with 1 ml of 50 mM HEPES-NaOH (pH 7.5) and re-suspended in 791

100 μl of reaction buffer (50 mM HEPES-NaOH, pH 7.5, 1 mM ADP-Glc) followed by 792

incubation at 30oC for 30 min with mild shaking. The reaction was terminated by the addition 793

of chloroform, and 10 μl of the upper phase was used for detecting ADP. ADP was analyzed 794

as described in the section of AGPase assay. ADP standards were provided by Sigma-Aldrich. 795

(c) Starch phosphorylase 796

The activity of starch phosphorylase was assayed by detecting the number of G1P 797

produced in the reaction (Zeeman et al., 1998; Häusler et al., 2000). In brief, 25 mg of 798

pulverized sample was extracted with 150 μl of extraction buffer (40 mM HEPES-NaOH, pH 799

7.5, 1 mM EDTA) and incubated on ice for 10 min followed by centrifugation to remove fruit 800

residues. Then, 25 μl of protein extract was mixed with 75 μl of water and 100 μl of double 801

strength reaction buffer (200 mM Tris-HCl, pH 8.1, 1.6 mM NADP+, 10 mM MgCl2, 2 μM 802

glucose-1,6-bisphosphate, 20 mM sodium phosphate buffer, pH 6.9, 0.1% amylopectin, 0.11 803



26

U glucose-6-phosphate dehydrogenase, 0.07 U phosphoglucomutase). The NADPH 804

formation was detected in the fluorescence mode of the spectrophotometer (Ex: 340 nm, Em: 805

465 nm). 806

(d) Starch hydrolase 807

The activity of starch hydrolase was assayed by detecting the number of reducing groups 808

released from starch with DNS reagent (Miller, 1959). In short, 25 mg of pulverized sample 809

was extracted with 150 μl of extraction buffer (20 mM sodium phosphate buffer, pH 6.9, 6 810

mM NaCl) and incubated on ice for 10 min followed by centrifugation to remove fruit 811

residues. The supernatant was applied to a self-packed Sephadex G-25 column for desalting. 812

Then, 50 μl of protein extract was mixed with equal volume of 1% soluble starch followed by 813

incubation at 60oC for 30 min, and protein extract without the input of starch served as the 814

blank reaction. The reaction was terminated by the addition of 100 μl of DNS reagent (1% 815

3,5-dinitrosalicylic acid, 30% sodium potassium tartrate tetrahydrate), and the whole mixture 816

was incubated at 95oC for 5 min. The amount of reducing sugar released from starch was 817

determined in the absorbance mode of the spectrophotometer (wavelength: 540 nm). Maltose 818

served as the standard for quantification. 819

The protein concentration was measured by Bradford reagent (Carl Roth) and Pierce 660 820

nm protein reagent (Thermo) according to the manual. 821

822

Statistical analysis 823

All figures and statistical analyses were made via Graphpad Prism 8. ANOVA and 824

Dunnett’s test were used to evaluate the difference between WT and NTRC-RNAi lines 825

unless otherwise stated. 826

827

Accession numbers 828

Sequence data from this article can be found in the GenBank/EMBL data libraries under 829

accession numbers SlNTRC (LOC101254347; LOC101266017), SlSAHH2 (Solyc09g092380), 830

SlNOR (Solyc10g006880), SlCAC (SGN-U314153). 831

832

Supplemental Data 833

Supplemental Figure S1. The molecular characterization in 65 DAF tomato fruits of 834

NTRC-RNAi lines compared to wild type (WT). 835

836

Supplemental Figure S2. Characterization of developmental stage in tomato fruits. 837

838



27

Supplemental Figure S3. Phenotypic analysis of mature green tomato fruits (35 DAF) of 839

NTRC-RNAi lines (RNAi-2, 26 and 33) compared to wild type (WT). 840

841

Supplemental Figure S4. Total protein content of fruits of NTRC-RNAi lines compared to 842

wild type (WT). 843

844

Supplemental Figure S5. Redox and allosteric properties of AGPase in 65 DAF tomato 845

fruits of NTRC-RNAi lines (RNAi-2, 26 and 33) compared to wild type (WT). 846

847

Supplemental Figure S6. The redox state of and 2-Cys Prxs in 35 and 65 DAF tomato fruits 848

of wild type (WT) and NTRC-RNAi lines. 849

850

Supplemental Table S1. Fold changes in metabolite profiles in tomato fruits of wild type 851

(WT) and NTRC-RNAi lines. 852

853

Supplemental Table S2. Changes in the level of hexose phosphates, 3-PGA and adenine 854

nucleotides in wild-type (WT) and NTRC-RNAi tomato fruits. 855



28

Acknowledgements 856

We thank Anne Bierling (LMU Munich) for excellent technical assistance. We are also 857

grateful to all greenhouse staff in the Max-Planck-Institute of Molecular Plant Physiology for 858

taking care of tomato plants. 859

860

Tables 861

Table 1. Changes in the activity of starch metabolism enzymes in 35 DAF tomato fruits of 862

wild type (WT) and NTRC-RNAi lines. Results are the mean with standard error from six 863

biological replicates. Statistical analysis used ANOVA with Dunnett’s test (*, different from 864

WT, P < 0.05). 865

866



29

Figure legends 867

Figure 1. Molecular characterization of NTRC-RNAi lines (2, 26 and 33) compared to wild 868

type (WT) in 35 DAF tomato fruits. A, The RNA interference construct targeting the NTRC 869

gene. B, Genotyping by PCR analysis for detection of nptII gene in NTRC-RNAi lines. C, 870

Detection of NTRC transcript level using RT-qPCR. D, Detection of NTRC proteins by 871

immunoblot analysis. E, Quantification of NTRC protein abundance using Image-J software. 872

The level of clathrin adaptor complexes (CAC) serves as a reference gene for genotyping and 873

RT-qPCR. The protein level of ribulose 1,5-bisphosphate carboxylase/oxygenase large 874

subunit (RbcL) served as a loading control for immunoblotting. Results are the mean with 875

standard error from six biological replicates. Statistical analysis used ANOVA with Dunnett’s 876

test (*, different from WT, P < 0.05). 877

878

Figure 2. Phenotypic analysis of fully-ripened tomato fruits (65 DAF) of NTRC-RNAi lines 879

(RNAi-2, 26 and 33) compared to wild type (WT). A, Visible phenotype of ripe fruits. B, 880

Fruit length and width. C, Fruit fresh weight. D, Ratio of fruit dry weight to fresh weight. 881

Results are the mean with standard error from six biological replicates (each replicate 882

contains data from 1 to 3 fruits). Statistical analysis used ANOVA with Dunnett’s test (*, 883

different from WT, P < 0.05). The red scale indicates 2 cm in length. 884

885

Figure 3. The accumulation of starch and soluble sugars in tomato fruits of wild type (WT) 886

and NTRC-RNAi lines (RNAi-2, 26 and 33). The levels of starch (A and B) and soluble 887

sugars (C and D) were measured in fruits harvested at 35 (A and C) and 65 DAF (B and D). 888

Results are the mean with standard error from six biological replicates. Statistical analysis 889

used ANOVA with Dunnett’s test (*, different from WT, P < 0.05). 890

891

Figure 4. Metabolite profile of NTRC-RNAi tomato fruits (RNAi-2, 26 and 33) compared to 892

wild type (WT). The metabolite profile was measured in fruits harvested at 35 (left panel) and 893

65 DAF (right panel) by using GC-TOF-MS. A. Sugars and sugar alcohols. B. Organic acids. 894

C. Amino acids and TCA cycle intermediates. D. Miscellaneous. Results are normalized to 895

WT with log2 transformation and visualized as a heatmap with hierarchical clustering done 896

by R software. Data are taken from Supplemental Table S1. 897

898

Figure 5. Changes in pyridine nucleotide level in tomato fruits of NTRC-RNAi lines in 899

comparison to wild type (WT). The levels of NADP+, NADPH, NAD

+ and NADH (A and B) 900

and the ratios of NADPH/NADP+ and NADH/NAD

+ (C and D) were measured in fruits 901



30

harvested at 35 (A and C) and 65 DAF (B and D). Results are the mean with standard error 902

from six biological replicates. Statistical analysis used ANOVA with Dunnett’s test (*, 903

different from WT, P < 0.05). 904

905

Figure 6. Redox and allosteric properties of AGPase and endogenous ADP-glucose content 906

in 35 DAF tomato fruits of NTRC-RNAi lines (RNAi-2, 26 and 33) compared to wild type 907

(WT). A, the redox state of intermolecular disulfide bond in AGPase small subunit 1 (APS1). 908

B, AGPase activity assayed with or without the activation of 10 mM 3-PGA. C, AGPase 909

activity kinetics in the presence of varied concentration of 3-PGA as indicated in the figure. 910

D, the content of endogenous ADP-glucose. The monomer/dimer ratios of the respective 911

proteins were quantified by Image-J software. Results are the mean with standard error 912

(n=5-6). Statistical analysis used ANOVA with Dunnett’s test (*, different from WT, P < 913

0.05). 914

915

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Figure 1. Molecular characterization of NTRC-RNAi lines (2, 26 and 33) compared to wild type (WT) in 35 DAF tomato fruits. A, The RNA interference construct targeting the NTRC gene. B, Genotyping by PCR analysis for detection of nptII gene in NTRC-RNAi lines. C, Detection of NTRC transcript level using RT-qPCR. D, Detection of NTRC proteins by immunoblot analysis. E, Quantification of NTRC protein abundance using Image-J software. The level of clathrin adaptor complexes (CAC) serves as a reference gene for genotyping and RT-qPCR. The protein level of ribulose 1,5-bisphosphate carboxylase/oxygenase large subunit (RbcL) served as a loading control for immunoblotting. Results are the mean with standard error from six biological replicates. Statistical analysis used ANOVA with Dunnett’s test (*, different from WT, P < 0.05).

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Figure 2. Phenotypic analysis of fully-ripened tomato fruits (65 DAF) of NTRC-RNAi lines (RNAi-2, 26 and 33) compared to wild type (WT). A, Visible phenotype of ripe fruits. B, Fruit length and width. C, Fruit fresh weight. D, Ratio of fruit dry weight to fresh weight. Results are the mean with standard error from six biological replicates (each replicate contains data from 1 to 3 fruits). Statistical analysis used ANOVA with Dunnett’s test (*, different from WT, P < 0.05). The red scale indicates 2 cm in length.

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Figure 3. The accumulation of starch and soluble sugars in tomato fruits of wild type (WT) and NTRC-RNAi lines (RNAi-2, 26 and 33). The levels of starch (A and B) and soluble sugars (C and D) were measured in fruits harvested at 35 (A and C) and 65 DAF (B and D). Results are the mean with standard error from six biological replicates. Statistical analysis used ANOVA with Dunnett’s test (*, different from WT, P < 0.05).

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Figure 4. Metabolite profile of NTRC-RNAi tomato fruits (RNAi-2, 26 and 33) compared to wild type (WT). The metabolite profile was measured in fruits harvested at 35 (left panel) and 65 DAF (right panel) by using GC-TOF-MS. A. Sugars and sugar alcohols. B. Organic acids. C. Amino acids and TCA cycle intermediates. D. Miscellaneous. Results are normalized to WT with log2 transformation and visualized as a heatmap with hierarchical clustering done by R software. Data are taken from Supplemental Table S1.

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Figure 5

Figure 5. Changes in pyridine nucleotide level in tomato fruits of NTRC-RNAi lines in comparison to wild type (WT). The levels of NADP+, NADPH, NAD+ and NADH (A and B) and the ratios of NADPH/NADP+ and NADH/NAD+ (C and D) were measured in fruits harvested at 35 (A and C) and 65 DAF (B and D). Results are the mean with standard error from six biological replicates. Statistical analysis used ANOVA with Dunnett’s test (*, different from WT, P < 0.05).

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Figure 6. Redox and allosteric properties of AGPase and endogenous ADP-glucose content in 35 DAF tomato fruits of NTRC-RNAi lines (RNAi-2, 26 and 33) compared to wild type (WT). A, the redox state of intermolecular disulfide bond in AGPase small subunit 1 (APS1). B, AGPase activity assayed with or without the activation of 10 mM 3-PGA. C, AGPase activity kinetics in the presence of varied concentration of 3-PGA as indicated in the figure. D, the content of endogenous ADP-glucose. The monomer/dimer ratios of the respective proteins were quantified by Image-J software. Results are the mean with standard error (n=5-6). Statistical analysis used ANOVA with Dunnett’s test (*, different from WT, P < 0.05).

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Soluble starch synthase

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Granule-bound starch synthase

(pmol min-1

mg FW-1

) 2.9±0.2 2.0±0.1* 2.0±0.1* 2.4±0.2

Starch phosphorylase

(nmol min-1

mg protein-1

) 49.6±10.9 76.5±18.3 30.7±4.2 40.0±11.3

Starch hydrolases

(μmol min-1

mg protein-1

) 2.4±0.4 2.3±0.7 1.1±0.2 1.0±0.4

Table 1. Changes in the activity of starch metabolism enzymes in 35 DAF tomato fruits of wild type (WT)

and NTRC-RNAi lines. Results are the mean with standard error from six biological replicates. Statistical

analysis used ANOVA with Dunnett’s test (*, different from WT, P < 0.05).

Table 1



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https://www.ncbi.nlm.nih.gov/pubmed?term=Ballicora MA%2C Frueauf JB%2C Fu Y%2C Schurmann P%2C Preiss J %282000%29 Activation of the potato tuber ADP%2Dglucose pyrophosphorylase by thioredoxin%2E J Biol Chem 275%3A 1315%26%23x&dopt=abstract

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https://scholar.google.com/scholar?as_q=Activation of the potato tuber ADP-glucose pyrophosphorylase by thioredoxin.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Ballicora&as_ylo=2000&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=B�nard C%2C Bernillon S%2C Biais B%2C Osorio S%2C Maucourt M%2C Ballias P%2C Deborde C%2C Colombi� S%2C Cabasson C%2C Jacob D%2C Vercambre G%2C Gautier H%2C Rolin D%2C G�nard M%2C Fernie AR%2C Gibon Y%2C Moing A %282015%29 Metabolomic profiling in tomato reveals diel compositional changes in fruit affected by source%2Dsink relationships%2E J Exp Bot 66%3A 3391%26%23x&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Bénard&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Metabolomic profiling in tomato reveals diel compositional changes in fruit affected by source-sink relationships.&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Metabolomic profiling in tomato reveals diel compositional changes in fruit affected by source-sink relationships.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Bénard&as_ylo=2015&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Berger F%2C Ram�rez%2DHern�ndez MH%2C Ziegler M %282004%29 The new life of a centenarian%3A Signalling functions of NAD%28P%29%2E Trends Biochem Sci 29%3A 111%26%23x&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Berger&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=The new life of a centenarian: Signalling functions of NAD(P).&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=The new life of a centenarian: Signalling functions of NAD(P).&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Berger&as_ylo=2004&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Buchanan BB%2C Balmer Y %282005%29 Redox regulation%3A A broadening horizon%2E Annu Rev Plant Biol 56%3A 187%26%23x&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Buchanan&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Redox regulation: A broadening horizon.&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Redox regulation: A broadening horizon.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Buchanan&as_ylo=2005&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Carrari F%2C Baxter C%2C Usadel B%2C Urbanczyk%2DWochniak E%2C Zanor M%2DI%2C Nunes%2DNesi A%2C Nikiforova V%2C Centero D%2C Ratzka A%2C Pauly M%2C Sweetlove LJ%2C Fernie AR %282006%29 Integrated analysis of metabolite and transcript levels reveals the metabolic shifts that underlie tomato fruit development and highlight regulatory aspects of metabolic network behavior%2E Plant Physiol 142%3A 1380%26%23x&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Faix B%2C Radchuk V%2C Nerlich A%2C H�mmer C%2C Radchuk R%2C Emery RJN%2C Keller H%2C G�tz KP%2C Weschke W%2C Geigenberger P%2C Weber H %282012%29 Barley grains%2C deficient in cytosolic small subunit of ADP%2Dglucose pyrophosphorylase%2C reveal coordinate adjustment of C%3AN metabolism mediated by an overlapping metabolic%2Dhormonal control%2E Plant J 69%3A 1077%26%23x&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Farr� EM%2C Tiessen A%2C Roessner U%2C Geigenberger P%2C Trethewey RN %282001%29 Analysis of the compartmentation of glycolytic intermediates%2C nucleotides%2C sugars%2C organic acids%2C amino acids%2C and sugar alcohols in potato tubers using a nonaqueous fractionation method%2E Plant Physiol 127%3A 685%26%23x&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Tiessen A%2C Nerlich A%2C Faix B%2C H�mmer C%2C Fox S%2C Trafford K%2C Weber H%2C Weschke W%2C Geigenberger P %282012%29 Subcellular analysis of starch metabolism in developing barley seeds using a non%2Daqueous fractionation method%2E J Exp Bot 63%3A 2071%26%23x&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Toivola J%2C Nikkanen L%2C Dahlstr�m KM%2C Salminen TA%2C Lepist� A%2C Vignols F%2C Rintam�ki E %282013%29 Overexpression of chloroplast NADPH%2Ddependent thioredoxin reductase in Arabidopsis enhances leaf growth and elucidates in vivo function of reductase and thioredoxin domains%2E Front Plant Sci 4%3A 1%26%23x&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Trethewey RN%2C Geigenberger P%2C Riedel K%2C Hajirezaei MR%2C Sonnewald U%2C Stitt M%2C Riesmeier JW%2C Willmitzer L %281998%29 Combined expression of glucokinase and invertase in potato tubers leads to a dramatic reduction in starch accumulation and a stimulation of glycolysis%2E Plant J 15%3A 109%26%23x&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Zeeman SC%2C Northrop F%2C Smith AM%2C Rees T %281998%29 A starch%2Daccumulating mutant of Arabidopsis thaliana deficient in a chloroplastic starch%2Dhydrolysing enzyme%2E Plant J 15%3A 357%26%23x&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Zeeman&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=A starch-accumulating mutant of Arabidopsis thaliana deficient in a chloroplastic starch-hydrolysing enzyme.&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=A starch-accumulating mutant of Arabidopsis thaliana deficient in a chloroplastic starch-hydrolysing enzyme.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Zeeman&as_ylo=1998&as_allsubj=all&hl=en&c2coff=1


1 Short title: Role of NTRC in developing tomato fruits...2019/09/16 · NTRC acts as central hub...

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