2.1 Plant material

Seeds of the common bean (Phaseolus vulgaris) cv V8025 were sown in hydroponic perlite and irrigated with a complete Hoagland solution as previously described (Bernal et al., 2005). Plants were cultivated in a greenhouse with natural light under 12-hour light and 12-hour dark periods. Daytime temperatures ranged from 24 to 26°C, while nighttime temperatures ranged from 16 to 18°C. The flowers were tagged at anthesis, and fruit age was defined by the number of days after anthesis (DAA). The pedicel phloem was removed using a sharp razor blade at 20 DAA (Li et al., 2019). Fruit pericarps were immediately frozen in liquid nitrogen and stored at −70°C until use.

2.2 Carbohydrate analysis

Soluble sugars (glucose, fructose, sucrose, and maltose) and starch were determined as previously reported (Bernal et al., 2005). The tissue was homogenized in 80% (v/v) ethanol and incubated for 30 min at 80°C. The extract was centrifuged at 12,000 g for 10 min, and the soluble sugars in the supernatant were detected using a Multiskan FC microplate photometer (Thermo Fisher Scientific). Quantification was performed in a sequential assay using NADH formation at 340 nm. In the first part of the reaction, 10–20 μL ethanolic extract was mixed with 150 μL of reaction mixture (50 mM HEPES [pH 7.4], 50 mM KCl, 1 mM ATP, 3 mM MgCl₂, 0.5 mM NAD⁺ and 1 U mL⁻¹ yeast hexokinase [EC 2.7.1.1]). An initial reading was taken, followed by the addition of 1.2 U mL⁻¹ glucose 6-P dehydrogenase (EC 1.1.1.49 from Leuconostoc mesenteroides) in 10 μL of 50 mM HEPES (pH 7.4). After 30 min, a second reading was obtained. The difference between the first and second readings was used to calculate glucose concentration. Next, 1.5 U mL⁻¹ phosphoglucose isomerase (EC 5.3.1.9 from yeast) in 10 μL of 50 mM HEPES (pH 7.4) was added and incubated for 30 min before taking the reading. The difference between the second and the third readings was used to calculate fructose concentration. Finally, 5 U mL⁻¹ invertase (EC 3.2.1.26) in 10 μL of 50 mM HEPES (pH 7.4) was added, and after 30 min, the final reading was registered. The sucrose concentration was calculated based on the difference in readings after the addition of phosphoglucose isomerase and invertase. For maltose quantification, ethanolic extracts were evaporated at 65°C and resuspended in water. Maltose was hydrolyzed in 90 mM HEPES (pH 7.5), 1 mM EDTA, and 2.5 U yeast maltase (EC 3.2.1.20) in a 50 μL final reaction volume for 30 min at 30°C. For maltase inactivation, the reaction mix was incubated for 5 min at 80°C, and glucose was quantified enzymatically. Starch was determined in the ethanol-insoluble fraction. Pellets were resuspended in 1 mL of distilled water and incubated at 80°C for 3 h. After cooling at room temperature, 250 U α-amylase (EC 3.2.1.1) and 270 U amyloglucosidase (EC 3.2.1.3, from Rhizopus) in 0.5 mL of 0.25 M Na acetate buffer (pH 4.5) were added to each sample. After gentle mixing, the samples were incubated for 12 h at 37°C. The next day, the samples were centrifuged at 12,000 g for 10 min, and glucose was measured in the supernatant using the enzymatic method described above.

2.3 ¹⁴CO₂ labelling

Fruit photosynthetic activity was evaluated by the incorporation of ¹⁴CO₂ following a previously reported procedure (Belmont et al., 2022). Five hours after the light period started, the fruits were placed into a sealed plastic bag, and ¹⁴CO₂ was liberated from 5 μCi (2.18 GBq mmol⁻¹) sodium [¹⁴C] bicarbonate by adding 200 μL of 3 M lactic acid. Fruits were exposed to ¹⁴CO₂ for 2 h before adding 800 μL of 3 M KOH to neutralize the acid and absorb any remaining ¹⁴CO₂. One hour later, the fruits were harvested. Seeds and pericarps were separated, frozen in liquid nitrogen and stored at −70°C. For analysis, samples were homogenized in 80% ethanol and incubated at 80°C for 10 min. An aliquot was used to estimate the total incorporation. Extracts were processed as described for carbohydrate analysis, and the amount of ¹⁴C in the fractions corresponding to ethanol-soluble sugars and starch was determined. The pellets after starch hydrolysis were resuspended in 500 μL of deionized water, and the incorporation of ¹⁴C in the insoluble material was also determined. In all cases, 100 μL was mixed with 3 mL of liquid scintillation Ultima-Gold XR, and the radioactivity was measured using a Beckman LS 6000Ic liquid scintillation counter and expressed as disintegrations per minute (dpm).

2.4 Enzymatic activities

Cytosolic and chloroplast FBPases (cFBPase and chlFBPase, respectively) were partially purified using a previously reported method (Lee and Hahn 2003) with minor modifications. Pods were extracted in an extraction buffer [50 mM MOPS (pH 7.5), 0.25 mM EDTA, 2.5 mM MgSO₄, 20% glycerol and complete protease inhibitor cocktail (Roche Diagnostics Gmbh)]. One gram of tissue was ground in a mortar with liquid nitrogen and homogenized with two volumes of extraction buffer (EB). The mixture was centrifuged at 12,500 g for 6 minutes. The supernatant (1.5 mL) was brought to 60% saturation with (NH₄)₂SO₄ and incubated for 30 minutes at 4°C. It was then centrifuged at 12,500 g for 10 minutes. The pellet was resuspended in EB (0.5 mL). This volume was mixed with DEAE-Sepharose resin (0.5 mL) previously washed and equilibrated with EB (1 mL × 3). The mixture was kept under rotary agitation for 15 minutes at room temperature and then centrifuged at 7,000 g for 3 minutes. The unbound fraction (cFBPase) was collected and kept at room temperature until the activity was measured. The resin was washed with EB (1 mL × 3) and eluted with 0.5 mL elution buffer [400 mM NaCl, 50 mM MOPS (pH 7.5), 0.25 mM EDTA, 2.5 mM MgSO₄, 20% glycerol and complete protease inhibitor cocktail (Roche Diagnostics Gmbh)]. The mixture was centrifuged at 7,000 g for 3 minutes. The bound fraction (chlFBPase) was collected.

Cytosolic and chloroplast FBPases were assayed at 30°C using a continuous assay coupled to NADH production. cFBPase was assayed in 200 μL final volume of reaction mix [100 mM Tris–HCl (pH 7.5), 0.2 mM EDTA, 5 mM MgCl₂, 0.3 mM NAD⁺, 1.2 U phosphoglucose isomerase, 0.6 U glucose-6-phosphate dehydrogenase]. The activity of chlFBPase was evaluated in a 200 μL final volume of reaction mix containing 100 mM Tris–HCl (pH 8.7), 0.5 mM EDTA, 12 mM MgCl₂, 0.3 mM NAD⁺, 1.2 U phosphoglucose isomerase and 0.6 U glucose-6-phosphate dehydrogenase. In both cases, the reaction was initiated by adding fructose-1,6-bisphosphate (1 mM for cytosolic and 10 mM for chloroplast FBPases). To evaluate the effect of reduction on cFBPase activity, partially purified fractions were incubated for 30 min at room temperature with 30 mM DTT (Dithiothreitol, threo-1,4-dimercapto-2,3-butanediol).

The activity of SPS was evaluated following a previously reported procedure (Huber et al., 1989) with minor modifications. Pod tissue was ground with liquid nitrogen and then homogenized with three volumes of extraction buffer [50 mM MOPS-NaOH (pH 7.5), 10 mM MgCl₂, 1 mM EDTA, 2.5 mM DTT, 0.1% Triton X-100, 25 mM NaF and complete protease inhibitor cocktail (Roche Diagnostics Gmbh)]. The extract was centrifuged at 12,000 g for 15 min at 4°C and the supernatant was precipitated with 15% PEG 10,000 for 1 h at 4°C. Protein was recuperated after centrifugation at 12,000 g for 15 min at 4°C and resuspended in a minimum volume of extraction buffer. SPS activity was assayed under limiting and V_max substrate concentrations. Under limiting conditions, PEG precipitated protein was incubated with 10 mM UDP-Glc, 3 mM Fru 6-P, 12 mM Glc 6-P, 10 mM Pi, 15 mM MgCl₂, 2.5 mM DTT and 50 mM MOPS-NaOH (pH 7.5) in a total volume of 70 μL. Under V_max condition, PEG precipitated protein was assayed in the presence of 10 mM Fru 6-P, 40 mM UDP-Glc, minus Pi using the same buffer and volumes as the limiting assays. Reactions were terminated after 30 min at 30°C by adding 70 μL of 30% KOH and unreacted Fru 6-P was destroyed by placing the tubes in boiling water for 10 min. After cooling, 1.0 mL of 0.14% anthrone in 13.8 N H₂SO₄ was added and the tubes were incubated at 40°C for 30 min before measuring the absorbance at 620 nm. The activation state of the enzyme was calculated as the ratio of the activities under limiting and V_max conditions X100.

2.5 PAGE and western blotting

Samples for electrophoresis were mixed with Laemmli denaturing buffer (Laemmli, 1970) and heated at 95°C for 5 min. Western blotting was performed according to Bernal et al. (2005). Rabbit antibodies against chloroplast and cytosolic FBPases (Agrisera) were used at 1:5,000 dilution, and goat anti-rabbit antibody conjugated with HRP (diluted 1:20,000) was used. The blots were developed with chemiluminescent HRP substrate (Millipore), and the antibody–antigen signal was detected using a ChemiDoc Image System from BioRad.

2.6 RNA extraction

RNA extraction was performed as previously reported (Bernal et al., 2022). Control samples were collected at 20, 21, 23, 25, 27 and 30 DAA. Fruits subjected to carbohydrate restriction were collected at 1, 3, 5, 7 and 10 days after phloem removal (corresponding to 21, 23, 25, 27 and 30 DAA). Plant material was immediately frozen in liquid nitrogen. Total RNA was isolated using TRIzol (Thermo Fisher Scientific), clean-up and concentration were performed using the RNA Clean and Concentrator TM-5 Kit (Zymo Research) according to the manufacturer's instructions. RNA concentration was measured using a Biodrop spectrophotometer, and the quality of the total RNA was assayed by agarose gel electrophoresis.

2.7 Transcript quantification by RT-qPCR

RT-qPCR was performed as previously reported (Bernal et al., 2022). Purified and DNase I-treated RNA was subjected to reverse transcription (RT) using oligo (dT) primers and the Ready Script Synthesis Mix (Sigma). A BLAST search in the Phytozome database (https://phytozome-next.jgi.doe.gov) was conducted using the At1g43670 sequence, which encodes cFBPase (Rojas-González et al., 2015), as well as the At3g54050 and At5g64380 sequences, which encode the redox-regulated and redox-independent chlFBPase isozymes, respectively (Serrato et al., 2009). Sequences encoding chloroplast FBPases (Phvul.008G112000 and Phvul.002G213400) and cytosolic FBPases (Phvul.004G095800, Phvul.004G128900 and Phvul.007G079500) were retrieved. Arabidopsis and Phaseolus sequences were then aligned using CLUSTALW (https://www.genome.jp/tools/clustalw). Specific primers for each selected target were designed using Primer3Plus (Untergasser et al., 2007) with qPCR settings activated (Table S1). qPCR was performed using Maxima SYBR Green/ROX qPCR Master Mix in a CFX96 Touch real-time PCR system (Bio Rad). Relative expression levels were calculated using the 2^-ΔΔCt method (Rao et al., 2013) with 20 DAA sample as a reference and ACTIN transcript as an internal housekeeping control. Targets were assayed with at least three biological replicates and three technical replicates each.

2.8 Statistical analysis

Data were analyzed using Prism software (GraphPad) and compared using one-way analysis of variance (ANOVA) with Tukey's multiple comparison post hoc test to determine significance.

3.1 Photosynthate restriction modifies seed development

Our recent work has shown that bean fruit pericarp utilizes surplus sucrose for starch synthesis and its later degradation helps to support the accumulation of reserve materials in seeds. This function makes the pericarp crucial for balancing seed growth with the supply of photosynthates. To identify metabolic adaptations in the fruit pericarp that could enhance seed development under reduced photosynthate availability, we removed the phloem at the pedicel level in fruits at 20 DAA (Figure 1A). At this developmental stage, seeds are about to begin the accumulation of reserve materials. The interruption of photosynthate flow resulted in a significant reduction in the number of seeds able to continue their growth (Figure 1B). Under normal conditions, seed growth progressed linearly from 20 to 30 DAA (Figure 1C). However, photosynthate restriction limited the growth of most seeds and five days later, it became evident that most seeds had stopped growing (Figure 1D). Despite this, 1–2 seeds per fruit continued to develop similarly to those under control conditions (Figure 1C, D). As expected, the disruption of phloem continuity caused an immediate reduction in sucrose levels in the fruit pericarp (Figure 2A), while a slight decrease in fructose and glucose levels became evident after five days (Figure 2B, C). The pericarp also accumulated a significant amount of starch that is degraded shortly after 20 DAA when seed growth accelerates. Interestingly, the reduction in photosynthate supply did not immediately affect the rate of starch degradation, which increased five days after the phloem was interrupted (Figure 3A). Maltose, a product of starch degradation that is converted in the cytosol into products transported to developing seeds, was consistently lower in the pericarp of fruits subjected to photosynthate restriction (Figure 3B), suggesting that this condition might accelerate maltose utilization.

3.2 Effect of photosynthate restriction on ¹⁴CO₂ assimilation and carbon partitioning

The photosynthetic activity of the fruit significantly contributes to the carbon supply for seed development (Bennett et al., 2011; Cho et al., 2023). To determine whether photosynthetic activity in the bean pericarp increased after phloem interruption, we assessed the incorporation of ¹⁴CO₂ by fruits between 20 and 27 DAA under both control and restricted conditions. In control condition, ¹⁴CO₂ assimilation gradually increased to reach a maximum at 25 DAA and then decreased; however, photosynthate restriction caused a significant reduction in assimilation at 25 DAA (Figure 4A). Rubisco activity was not significantly affected by phloem interruption (Figure 4B), suggesting that the observed reduction in ¹⁴CO₂ assimilation at 25 DAA in fruits with reduced photosynthate supply could be attributed to stomatal closure. Under control conditions, most of the ¹⁴C-products remain in the pericarp, with the proportion transferred to seeds gradually increasing to reach up to 15% at 27 DAA (Figure 4C). Analysis of the ¹⁴C-products retained in the pericarp revealed that at 20 DAA, up to 60% of the ¹⁴C-label was incorporated into starch, and this proportion gradually decreased to 30% at 27 DAA (Figure 4E). The incorporation into the soluble fraction followed an opposite pattern, increasing from 20% at 20 DAA to nearly 60% at 27 DAA. Approximately 10% of the ¹⁴C-incorporation was associated with insoluble pod material, and this proportion remained almost constant during the period analyzed (Figure 4E). On the other hand, photosynthate restriction led to a significant reduction in the proportion of the ¹⁴C-label that remained in the fruit pericarp, while the proportion transferred to seeds increased to 50% of the ¹⁴C-label incorporated at 27 DAA (Figure 4D). Further analysis of ¹⁴C-products remaining in the fruit pericarp indicated that this shift was accompanied by a substantial reduction in the proportion of the ¹⁴CO₂ incorporated into starch, while the incorporation into the soluble fraction increased to 80% (Figure 4F). This change was observed three days after photosynthate supply was reduced, and the ratio remained stable thereafter (Figure 4F), suggesting that metabolic adaptations might occur to facilitate the transfer of resources from the pericarp to developing seeds.

3.3 Photosynthate restriction modulates SPS and FBPase activities

In leaves, the synthesis of sucrose and starch is regulated by the activities of SPS and cFBPase. A reduction in SPS activity leads to decreased sucrose levels (Strand et al., 2000), while the absence of cFBPase results in increased starch accumulation and a decline in sucrose levels (Lee et al., 2008; Rojas-González et al., 2015). To investigate if a similar response occurs in bean fruit pericarp following phloem interruption, we evaluate both activities. The phosphorylation of SPS at Ser 158 inhibits its activity and changes its sensitivity to Glc6P (activator) and Pi (inhibitor); the effect of such modification can be evaluated by measuring the activity under V_max and substrate limited conditions (Toroser et al., 1999). Under control conditions, the activity of SPS measured under optimal conditions shows a transient decrease at 23 DAA and gradually increases afterward, while the reduction in photosynthate supply caused a significant reduction (Figure 5A); these differences disappeared when SPS activity was measured under substrate limited conditions (Figure 5B). The percentage of SPS activation increased in both conditions after 23 DAA (Figure 5C); the larger increase observed when photosynthate supply is reduced could compensate for the reduction observed when SPS activity was assayed under optimal conditions (Figure 5A). We also evaluated the activity of cytosolic and chloroplast FBPase isozymes (cFBPase and chlFBPase, respectively). Under control conditions, the activity of cFBPase shows a slight increase at 23 DAA, followed by a gradual decrease, while photosynthate restriction caused a consistent increase, reaching up to 20 times at 30 DAA (Figure 6A). The activity of chlFBPase results from the action of two isozymes, one regulated by redox and the other redox-insensitive (Serrato et al., 2009). When DTT was excluded from the assay, control conditions produced a small increase in chlFBPase activity at 25 DAA; the reduction in photosynthate supply led to a twofold increase in activity at 25 DAA, followed by a decrease (Figure 6B). When DTT was included in the assay, activity increased under both conditions; however, the response was more pronounced in control samples (Figure S1). Western-blot analysis revealed that under control conditions, the levels of both cFBPase and chlFBPase isoforms gradually decreased as fruit development progressed, whereas photosynthate restriction led to a significant increase in the levels of both isozymes (Figure 6C, D).

3.4 Photosynthate restriction alters cFBPase and chlFBPase gene expression

In Arabidospsis, cFBPase is encoded by the gene At1g43670 (Rojas-González et al., 2015), whereas the redox-regulated and redox-independent chlFBPase isozymes are encoded by At3g54050 and At5g64380, respectively (Serrato et al., 2009). A Blast search in the Phytozome v10.3 database (https://phytozome-nextjgj.doe.gov) indicated that in Phaseolus vulgaris cFBPase is encoded by three closely related genes (Phvul.004G095800, Phvul.004G128900 and Phvul.007G079500, Figure S2). The expression patterns of Phvul.004G095800 and Phvul.004G128900 were very similar in both conditions. Under control conditions, both genes exhibited expression peaks at 23 and 30 DAA. Photosynthate restriction increased the response, with the two expression peaks observed at 21 and 25 DAA (Figure 7A, B). The expression of Phvul.007G079500 differed; under control conditions, it gradually increased, reaching a 20-fold increase at 25 DAA before declining. Reduction in photosynthate supply increased expression, reaching a 20-fold increase from 21 to 23 DAA, then increasing to a 70-fold increment at 27 DAA (Figure 7C). The expression of the gene Phvul.008G112000, which encodes a redox-regulated chlFBPase, consistently increased during fruit development, to reach a 15-fold increase at 30 DAA. Under photosynthate restriction, expression increased five-fold at 23 DAA and remained almost constant afterward (Figure 8A). In contrast, under control conditions, the expression of Phvul.002 g213400, which encodes the redox-independent chlFBPase, increased nine-fold at 21 DAA, then gradually decreased, with a similar second peak at 30 DAA. Photosynthate restriction produced a large reduction in its expression, with a five-fold increase observed at 30 DAA (Figure 8B). It is important to point out that the substantial increases in gene expression were not fully reflected in the levels of cytosolic and chloroplast FBPase proteins detected by western-blotting (Figure 8C, D). This discrepancy may be partially explained by the regulation of cFBPase turnover through ubiquitination (Wang et al., 2022), suggesting that this post-translational modification could contribute to the differences observed between messengers and protein levels.