Physiologia Plantarum 2014

© 2014 Scandinavian Plant Physiology Society, ISSN 0031-9317

Sucrose transport and phloem unloading in peach fruit: potential role of two transporters localized in different cell types Laura Zanon, Rachele Falchi* , Simonetta Santi and Giannina Vizzotto Dipartimento di Scienze Agrarie e Ambientali, University of Udine, 33100 Udine, Italy

Correspondence *Corresponding author, e-mail: [email protected] Received 6 August 2014; revised 17 October 2014 doi:10.1111/ppl.12304

Several complex physiological processes, which include long-distance translocation in the phloem and unloading in sink tissues, govern the partitioning of sugars in economically important organs, such as peach fruit. In this study, we took advantage of a symplastic tracer, carboxyfluorescein (CF), providing evidence for an apoplastic sucrose transfer in the early (SI) and middle (SIII) phases of peach fruit development. Moreover, using a combination of in situ hybridization and laser microdissection-assisted expression analysis, three putative sucrose transporters encoding genes (PpSUT1, PpSUT2, PpSUT4) were transcriptionally analyzed to relate their expression with sucrose storage in this organ. Our study revealed that PpSUT2 and PpSUT4 are the genes predominantly expressed in fruit flesh, and the detailed analysis of their expression pattern in the different cell types enabled us to suggest a specialized role in sucrose distribution. Both PpSUTs transporters could be involved in the retrieval of sucrose lost from the symplastic continuum of the phloem and, when expressed in parenchyma cells, they could be active in the import of sucrose into sink tissues, via symport from the apoplast. An alternative hypothesis has been proposed and discussed for PpSUT4 because of its putative tonoplastic localization. Taken together, our results provide new insights into the molecular mechanisms underpinning sucrose unloading and accumulation in peach fruit.

Introduction Source leaves export sucrose that, by long distance transport, is released into non-photosynthetic tissues, such as roots, stems, flowers, fruits and seeds (Ludewig and Flügge 2013). The partitioning of sugars in economically important sink organs is governed by several complex physiological processes, including photosynthetic rate, phloem loading in the source leaf, long-distance translocation in the phloem, phloem unloading in sink organs, post-phloem transport and metabolism of

imported sugars in sink cells (Nie et al. 2010). Fleshy fruits are a class of terminal reproductive strong storage sinks, but the mechanism of phloem unloading in these organs has received little attention in comparison with vegetative tissues and developing seeds (Zhang et al. 2006). Peach fruit is an economically important sink in which sugar accumulation is a major determinant of yield and quality. Generally, the average values of soluble solids content obtained in peach commercial cultivars range between 9 and 15%, even if in some cases they reach

Abbreviations – CDS, coding DNA sequence; CCCP, carbonyl cyanide m-hlorophenylhydrazone; CF, carboxyfluorescein; CFDA, carboxyfluorescein diacetate; dafb, days after full bloom; DEPC, diethyl pryrocarbonate; DMDC, dimethyl dicarbonate; LMPC, laser microdissection pressure catapulting; PBS, phosphate-buffered saline; PEN, polyethylene–naphthalate; qRT-PCR, quantitative real-time polymerase chain reaction; SE-CC, sieve elements-companion cells; SSC, saline-sodium citrate; SUF, sucrose facilitator; SUT, sucrose transporter; UPGMA, unweighted pair group method with arithmetic mean.

Physiol. Plant. 2014

up to 20% or more (Layne and Bassi 2008, Byrne et al. 2012). Soluble sugars show different accumulation levels throughout fruit growth when individually analyzed. In detail, glucose and fructose are present in nearly equal amounts and mainly during the early stage of growth, sorbitol levels remain low throughout development, whereas sucrose accumulates and becomes the predominant sugar in mature fruit (Vizzotto et al. 1996, Falchi et al. 2013). Among the sugars synthesized in a plant, only a few move in the phloem over a long distance, regardless of the species and type of phloem loading considered. In all cases, sucrose is the main form of carbon found in the phloem. In addition to sucrose, polyols (mainly sorbitol and mannitol) and oligosaccharides of the raffinose family can also be found (Lemoine et al. 2013). In Prunus persica, like other Rosaceae species, sucrose and sorbitol are both present in phloem sap as form of translocated carbon. Sugars can exit the phloem through either a symplastic or an apoplastic pathway, but the unloading pathway depends on the particular sink involved and its developmental stage (Lemoine et al. 2013). Concerning the apoplastic pathway, the uptake of hexoses resulting from the hydrolysis of the apoplastic sucrose by cell wall invertases has been suggested to predominate in the sink tissues that undergo cell division and elongation, while sucrose transport predominates in the sink storage tissues (Weschke et al. 2000). In this context, sucrose transporters (SUTs) play pivotal roles in mediating sucrose transport within plants, and their activities have a major impact upon plant growth rates and crop yields. The classification of SUT proteins into different subfamilies reflects either an amino acids sequence homology, and similar substrate affinity and function throughout phloem pathway. The SUT genes found in higher plants exhibit distinct expression patterns, depending on tissue, development and plant species (Kühn and Grof 2010). In apoplastically loading plants, membrane-localized sucrose/H+ symporters mediate the loading of sucrose into the phloem in source tissues, but, interestingly, sucrose/H+ symporter transcripts and proteins have also been localized in sink tissues (Geiger 2011). According to a recent classification (Kühn and Grof 2010), SUTs of dicotyledonous species fall into three distinct groups: SUT1, SUT2 and SUT4 subfamilies. In fruit trees, the path of phloem unloading has been elucidated for a small number of species. These investigations often have taken advantage of the fluorescent marker of phloem transport 6(5)-carboxyfluorescein diacetate (CFDA), a membrane-permeable and nonfluorescent compound that, when degraded to 6(5)-carboxyfluorescein (CF) in live cells, becomes a membrane-impermeable fluorescent dye. This technique showed that symplastic and

apoplastic mechanism can be interchanged, depending on genotype and developmental stage. In apple, cherry and pear fruit, there is evidence for an apoplastic step in sucrose and sorbitol unloading, involving sugar transporters (Gao et al. 2003, Zhang et al. 2004, Zhang et al. 2014). A similar pattern has been observed in jujube fruit (Zizyphus jujuba), but in this case the predominantly apoplastic unloading mechanism is interrupted by a transient symplastic phase at the middle stage of growth (Nie et al. 2010). The shift of unloading routes in response to sink development has also been observed in grape fruit: a symplastic mechanism occurs at the early stages, while an apoplastic pathway operates at the late stage of development. In grape berry, the turning point has been observed at or just before the onset of ripening, when the fruit begins to store soluble sugars (Zhang et al. 2006). Moreover, in walnut fruit, distinct unloading patterns have been found in two types of tissues: symplastic in seed and apoplastic in fleshy pericarp (Wu et al. 2004). Although the possibility of apoplastic phloem loading through an active transport pathway has been hypothesized as the most important means of exporting carbohydrates from mature leaves in peach (Moing et al. 1997), unloading mechanisms are still a source of debate. However, carrier-mediated transport has been hypothesized for mesocarp of young peaches (Vizzotto et al. 1996). Other attempts to elucidate the mechanism of sugar accumulation in peach fruits have been carried out by the analysis of key enzymes involved in the main sugar conversion pathways (Moriguchi et al. 1990, Nonis et al. 2007, Morandi et al. 2008). Some authors have postulated a preferential utilization of sorbitol in vegetative sink tissues close to the source, such as young sink leaves and cambium (Moing et al. 1992), and a major role of sucrose in fruit growth (Lo Bianco et al. 1999). Recently, the distribution of sugars among different compartments (free space, cytoplasm and vacuole) of peach mesocarp at two different growth stages has been determined, providing evidence for a predominant cytoplasmic localization in young fruit and a vacuolar localization in mature fruit. Moreover, the same authors showed that the permeability through plasma membrane and tonoplast decreased with peach fruit maturation (Jiang et al. 2013). In this work, we provide the first description of the phloem unloading pathway in peach fruit, by means of microscopy coupled with the symplastic tracer dye CFDA. In addition, three genes encoding SUTs have been identified, and their transcriptional regulation has been examined in different fruit cell types. The results are discussed in order to present a model for the physiological roles that these carriers may play in carbohydrate partitioning in peach fruit. Physiol. Plant. 2014

Materials and methods Plant material Experiments were conducted on fruits harvested from peach plants of cultivar Redhaven [P. persica (L.) Batsch] cultivated in the Experimental Farm ‘A. Servadei’ of Udine University in northeastern Italy (46.01N, 13.13E), in 2010. Trees received standard horticultural care. Fruit growth was monitored weekly from about 30 days after full bloom (dafb) until harvest by measuring the fresh weight of a pool of fruits collected from 10 different plants. Three pools of mesocarp tissues for each date were frozen in liquid nitrogen and stored at −80∘ C for subsequent analysis. Flesh pieces of about 40 mm3 (or smaller) in size, representative of different growth stages, were produced for the following embedding procedures. Determination of mesocarp soluble sugar content Sugars were extracted from 1 g of pooled mesocarp or seed tissues with several washing steps with boiling ethanol-water solution, as previously described (Nonis et al. 2008). Determination of sugars was performed by enzymatic assay. In brief, sucrose, glucose and fructose contents were obtained by measuring the increase in NADPH absorbance at 340 nm, stoichiometric to the amount of glucose and fructose, after treatments with invertase, phosphoglucoisomerase, hexokinase and glucose-6-phosphate dehydrogenase (Vizzotto et al. 1996). Fixation, dehydration and embedding of tissues In order to maintain morphology and integrity of fruit tissues, mesocarp samples were fixed and embedded as previously described (Christensen et al. 1998), with some modifications. Briefly, the small flesh pieces, representative of different growth stages, were incubated in a fixative (45% ethanol, 5% formalin, 5% acetic acid) for 2 days at 4∘ C, with gentle shaking. Subsequently, tissues were dehydrated through a series of washing in ethanol solutions (50, 70, 96 and 100%). Each step was done twice for 1 h, and the last step overnight. The following day, the samples were first transferred in a solution containing 50% ethanol and 50% limonene (Roti®-Histol, Carl Roth Gmbh, Karlsruhe, Germany) and then two times in limonene for 2 h each. Half of this solution was replaced with liquid paraffin (Paraplast-plus, McCormick Scientific, St. Louis, MO, USA), and samples incubated at 60∘ C for 2 h. Afterwards, the limonene/paraffin solution was discarded, substituted with paraffin and the samples maintained at 60∘ C for 2 h. Finally, the samples were transferred in new paraffin and incubated overnight at Physiol. Plant. 2014

60∘ C. Inclusion blocks were prepared by means of a HistoStar Embedding Workstation (Thermo Scientific, Rockford, IL, USA). Methylene blue staining Tissue sections of 10 μm in thickness were cut by means of the rotary microtome Leitz 1512 (Leitz, Wetzlar, Germany) and placed on slides. Before staining with methylene blue, sections were dewaxed by two steps of 10 min each in limonene and rehydrated in ethanol solutions at decreasing concentration (100, 95, 70, 50 and 30%) and, finally, in deionized water. A drop of 0.1% aqueous methylene blue was placed on each section and left for 5 min. The dye was replaced with deionized water, and samples were dehydrated through washes in ethanol solutions at increasing concentration (30, 50, 70, 95 and 100%). Finally, a drop of mounting solution (CC/Mount™ from Sigma-Aldrich, Milan, Italy) was applied, and a cover slip was placed on each sample. CFDA labeling Analysis of 6(5)-CFDA transport was carried out on peach fruits collected at different growth stages, at about 50 and 85 dafb. A water solution containing 1 mg ml−1 5(6)-CFDA (Sigma-Aldrich, Milan, Italy) was prepared. A branch carrying some leaves and one fruit was fed with CFDA aqueous solution for at least 5 h. This membrane permeable and non-fluorescent precursor is degraded to CF by intracellular esterases as soon as it is loaded into cells, becoming fluorescent and membrane impermeable. Treated fruits were subsequently hand sectioned and examined directly by inverted fluorescence microscope (Axiovert35, Zeiss, Jena, Germany). Sequence analysis A whole genome tBLASTn search on peach GBrowse site (http://services.appliedgenomics.org/fgb2/iga/prunus_ public/gbrowse/prunus_public/), using SUT proteins from Arabidopsis thaliana as a query, allowed the identification of peach SUT genes. Proteins belonging to the same families from different plant species were selected to carry out a multiple alignment and used to create a sequence similarity UPGMA (Unweighted Pair Group Method with Arithmetic Mean)-tree by means of MEGA version 5 (Tamura et al. 2011). The putative amino acids sequences were analyzed by taking advantage of the TMHMM Server v. 2.0 web-based service (Krogh et al. 2001) (http://www.cbs.dtu.dk/services/TMHMM-2.0/) to calculate the probability of a region to form a transmembrane domain.

Laser microdissection pressure catapulting Sections for laser microdissection pressure catapulting (LMPC) were obtained from mesocarp samples fixed overnight in 3:1 (v/v) ethanol:acetic acid at 4∘ C and embedded (Santi and Schmidt 2008). Ten-μm-thick slices were sectioned on a rotary microtome (Leica, Bensheim, Germany). Sections were stretched at 42∘ C for a few seconds on a drop of DEPC (Diethyl pryrocarbonate) water that was delivered directly on the PEN (polyethylene-naphthalate)-covered glass slides (PALM Microlaser Technologies, Carl Zeiss MicroImaging GmbH, Jena, Germany), and then dried at 42∘ C for not more than 30 min. Sections were deparaffinized two times for 5 min each in xylene and then air-dried. Phloem and parenchyma cells were separately microdissected from the deparaffinized sections with a PALM Microbeam System (Carl Zeiss MicroImaging GmbH, Jena, Germany). Areas of at least 2×106 μm2 in total were cut and catapulted in 0.2 ml tubes with adhesive caps (PALM Carl Zeiss, Jena, Germany). RNA isolation and cDNA synthesis Total RNA was obtained from peach fruit mesocarp as previously described (Nonis et al. 2007). The final RNA pellet was resuspended in RNAse-free water and stored at −80∘ C. An aliquot of total RNA was treated with DNase (Promega Co., Madison, WI, USA) to remove contamination by genomic DNA, and it was finally purified and concentrated with an RNeasy MinElute cleanup kit (Qiagen Gmbh, Hilden, Germany), according to the manufacturer’s instructions. An aliquot of RNA was spectrophotometrically quantified (Nano Drop 1000, Thermo Scientific, Rockford, IL, USA), and electrophoretically separated on an agarose gel, to check the integrity. RNA from LMPC-captured cells was extracted using the Absolutely RNA Nanoprep Kit (Agilent Technologies, Santa Clara, CA, USA) with minor changes with respect to manufacturer’s instructions. DNase-treated RNA was eluted in 14 μl of RNase-free water heated to 60∘ C. T7 RNA polymerase-based RNA amplification was performed using the MessageAmp II aRNA Amplification Kit (Ambion, Life Technologies Co., Carlsbad, CA, USA) according to the manufacturer’s instructions, as already described (Santi et al. 2013). Nucleic acid quantity and integrity were evaluated using an RNA 6000 Pico Assay kit on an Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). RNA from both fruit flesh and LMPC-captured cells was retrotranscribed in a total volume of 20 μl using SuperScript® VILO™ cDNA Synthesis Kit (Invitrogen™, Life Technologies, Paisley, UK), following manufacturer’s

instructions, with the addition of 1 μl oligo dT (10 μM) to the reaction mix, and employed for expression analysis. Quantification of gene expression by quantitative real-time polymerase chain reaction PpSUT transcripts quantification was performed by quantitative real-time polymerase chain reaction (qRTPCR) with Opticon 2 (MJ Research, Bio-Rad Laboratoires Co., Hercules, CA, USA) detection system, using a Real Master Mix SYBR ROX (5 PRIME, Eppendorf, Hamburg, Germany) kit. Specific primers (Table 1) were designed, and their efficiency was tested by measuring the amplification of a 2× cDNA serial dilution. All quantifications were normalized to ubiquitin-conjugating enzyme (accession number BF717254), that we previously demonstrated to be unaffected by sugars and suitable for experiments carried out during fruit development (Nonis et al. 2007), amplified with the primers 5′ -CCCACCTGATTACCCTTTCA-3′ and 5′ -GGATCTGTC AGCAGTGAGCA-3′ . Detailed procedures have been already described (Falchi et al. 2010). In situ hybridization analysis Digoxigenin-labeled sense and antisense riboprobes were synthesized by in vitro transcription, using the DIG RNA Labeling Kit (Sp6/T7) (Roche Applied Sience, Penzberg, Germany). In detail, fragments of the three PpSUT genes were amplified from a pool of cDNA (from samples at different growth stages) with specific primers (Table 2) and Taq DNA polymerase High Fidelity (Invitrogen, Life Technologies, Paisley, UK). The products were gel purified using the Invisorb® Spin DNA Extraction Kit (Invitek, Berlin, Germany). The cleaned products were quantified by NanoDrop 1000 (Thermo Table 1. Primers used to quantify gene expression by qRT-PCR. Gene name

Forward primer

Reverse primer

PpSUT1

CTATGGCTCCCGCAATAAAG CAGTATTCCCTTCGCTCTGG

PpSUT2

AATACGGTGCAGGGACCAG AACCATCTGTGCCAACTTCC

PpSUT4

ATGTGACTCAGGGTCCTTGC ACCAACCGCCATAAACAGAG

Ubiquitinconjugating enzyme

CCCACCTGATTACCCTTTCA

GGATCTGTCAGCAGTGAGCA

Table 2. Primers used to amplify PpSUT regions selected as RNA probes. Gene name

Forward primer

Reverse primer

PpSUT1

GTGACGACCCGAAGAGAATG

GACATGAACCCCAAAACGAC

PpSUT2

TCGTGCTCTTCTGGCTGAT

TGCTGGTGGTAAATGCCTTA

PpSUT4

GCTGATCTCACTGCAAAGGA

CCGAGTTCAACATCAGACCA

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Scientific, Rockford, IL, USA), checked on 1% agarose gel and the fresh products were directly cloned into pCR®II dual promoter vector using TA cloning® kit Dual promoter from Invitrogen, according to manufacturer’s instructions. The plasmids were purified by kit Wizard® Plus Minipreps DNA Purification System (Promega Co., Madison, WI, USA) and sequenced to check their orientation in the vector, using M13 universal primers. Each vector was linearized (BamHI, KpnI, EcoRV from Promega Co., Madison, WI, USA) in order to transcribe both sense and antisense RNA probes in the presence of UTP conjugated with digoxigenin. Paraffin-embedded samples were cut into 8 μm thick slices, deparaffinized with limonene (Roti®-Histol, Carl Roth Gmbh, Karlsruhe, Germany) and hydrated through an ethanol series to DMDC (Dimethyl dicarbonate) water. Slices were treated with 2× SSC (Saline-Sodium Citrate, 0.3 M NaCl, 0.03 M trisodium citrate dehydrate, pH 7), washed with DMDC water two times for 2 min, incubated in TE buffer containing 0.1 μg ml−1 proteinase K for 30 min at 37∘ C, treated with 0.2 M HCl for 20 min and washed two times in DMDC water. Slides were transferred for 2 min in 1× PBS (Phosphate-buffered saline) buffer with 0.2% glycine to stop the action of proteinase K and were washed twice in 1× PBS for 2 min, then transferred in 1× PBS with 4% paraformaldehyde for 10 min, and washed in 1× PBS for 2 min. In order to reduce the background staining, a pre-treatment with 0.25% acetic anhydride and 0.1 M triethanolamine for 10 min with slow agitation was performed. After washing in 1× PBS for 5 min, sections were dehydrated. Afterwards, a step of pre-hybridization was carried out treating samples with hybridization buffer without probe for 1 h at 40∘ C in a humid chamber. Hybridization took place overnight in a humidified box, with 5 ng μl−1 of the specific digoxigenin-labeled RNA probe in hybridization solution, at the optimal temperature calculated from the Tm of each probe. The slides were washed twice in 0.2×SSC for 1 h at 40∘ C, treated with 1× PBS solution for 10 min and incubated with two blocking solutions following the manufacturer’s instructions. The hybrid mRNA/DIG-RNA probe was detected through an antibody anti-DIG alkaline phosphatase conjugate (Roche Applied Sience, Penzberg, Germany). The antibody was diluted 1:500 in block solution and several drops of this solution were placed onto tissue sections; samples were incubated for 2 h, then washed two times in block solution for 20 min each and maintained in block solution overnight at 4∘ C. Tissue sections were washed in buffer C (0.1 M NaCl, 0.1 M Tris-HCl pH 9.6, 0.05 M MgCl2 ) two times for 15 min. Subsequently, they were drained and treated with several drops of 1× NBT/BCIP solution (Roche), Physiol. Plant. 2014

1 mM Levamisole, in buffer. Samples were incubated until color development at room temperature in a dark chamber. Before detecting the signal, color solution was removed and samples were washed in buffer 3 (10 mM Tri-HCl pH 8, 10 mM EDTA) for 2 min and washed twice in DMDC water for 5 min. After drying, mounting solution (CC/Mount, Sigma-Aldrich, Milan, Italy) and a cover-slide were placed on the sections. Finally, sections were observed and photodocumented by optical microscopy (B-350 Optika).

Results Determination of peach fruit growth pattern and of sugar content during development The plotting of fruit fresh weight against time (dafb) shows a curve characteristic of drupaceous fruit growth (Bonghi et al. 2011, Falchi et al. 2013), with a double-sigmoidal trend (Fig. 1). In detail, four stages can be distinguished: two steps of rapid growth rate, from flowering to 66 dafb (SI), and from about 80–108 dafb (SIII), interrupted by a lag phase from 66 to 80 dafb (SII), and a final stage of growth slowdown from 108 dafb to harvest (SIV), taking place at 115 dafb. The developmental changes in sucrose, glucose and fructose were analyzed in fruit flesh. In the early period of peach fruit growth, the two hexoses were the most abundant soluble sugars in mesocarp tissue, showing a maximum around 57 dafb, and a constant decrease until harvest. On the other hand, sucrose was fairly absent in young peach fruits, but begun to be steadily accumulated starting from about 43 dafb until harvest, becoming the predominant soluble sugar in mature fruit (Fig.1). Structural studies of fruit mesocarp Sections of whole or part of fruit (depending on the developmental stage) were produced in order to follow their anatomical differentiation, with particular regard to vascular bundles arrangement. The staining of sections pointed out that the network of strands feeding the mesocarp tissue is uniformly distributed through the flesh, both in transverse and longitudinal direction, since the earliest stages of growth (Fig. 2A, B). Moreover, the examination of vascular bundles at higher magnification revealed their different structure according to the tissue localization. We found that, in the mesocarp, the majority of veins in the mesocarp present a cavity located between the two types of conductive cells (Fig. 2C, D). This structure was previously described by other authors (Masia et al. 1992, Nii et al. 2005, Liu et al. 2010) as a glandular cavity.

Fig. 1. Peach fruit (cv. Redhaven) sugar accumulation pattern (histograms, values are means ± standard deviations of three replicates) and growth expressed as fresh weight (line, values are means ± standard deviations of 10 fruits) plotted against dafb. As expected, fruit growth shows a double-sigmoid trend. In detail, four stages can be distinguished: rapid growth rate (SI and SIII), a lag phase (SII) and a final stage of growth slowdown (SIV). In the early phases of fruit growth, glucose and fructose are the most abundant soluble sugars in mesocarp. Nearly in correspondence of SIII beginning, sucrose initiates to be accumulated, becoming the predominant soluble sugar in mature fruits. Arrows indicate the time of sampling for CF experiments and in situ hybridization or LMPC analyses.

Fig. 2. Transverse (A) and longitudinal (B) sections of peach fruit during early period of development (about 5 mm diameter and 15 dafb) treated with methylene blue. The staining allows to distinguish, since the earliest stages, cells destined to form endocarp tissue and the network of vascular strands in mesocarp. A magnification of these structures is represented in transverse (C) and longitudinal (D) section. GC, glandular cavity; PH, phloem; X, xylem.

Distribution of phloem-imported CF Structural analysis and distribution of phloem-imported CF allowed us to study the unloading route from the phloem in fruit at two developmental stages (SI and SIII, 50 and 85 dafb, respectively). The treated fruits were sampled 5 h after CF loading, and longitudinal or transverse hand-cut sections were prepared from fleshy

pericarp. The fluorescence microscope images of CF movement in leaves, pedicel (data not shown) and fruits confirmed that the fluorescent marker was successfully transported throughout the whole network of peach vascular bundles. In young fruits, CF was restricted to phloem strands, as shown both in longitudinal and in transverse section Physiol. Plant. 2014

Fig. 3. CFDA distribution in peach fruit at SI (A, B) and SIII (C, E) stages. Observations were performed by fluorescent microscope. Transverse (A) and longitudinal (B) pictures of the whole fruit are feasible when its size is very small, whereas transverse (C) and longitudinal (E) images of single bundles are reported for middle stage of fruit development. (D, F) Transmitted light pictures allowed to recognize the localization of fluorescence circumscribed to vascular bundles.

(Fig. 3A, B), indicating a symplastic discontinuity between veins and parenchyma cells. The observation of the same sections in bright-field allowed us to substantiate the reliability of our results (data not shown). In fruits collected at 85 dafb, the mechanisms of transport appeared unaffected, being CF confined to the phloem strands along the phloem pathway both in major and minor bundles (Fig. 3C–F). Taken together, these findings pointed out that the SE–CC (sieve element-companion cell) complex could be symplastically isolated during the early and middle developmental stages. Identification and analysis of genes encoding SUTs The availability of peach genome sequence (http://www. rosaceae.org/species/prunus_persica/genome_v1.0) has allowed the identification of genes encoding SUT through BLAST searches using SUT proteins from A. thaliana. Three predicted genes were identified: ppa004033, ppa003041 and ppa004620. Physiol. Plant. 2014

Both ppa003041 and ppa004620 were located in pseudomolecule 1, whereas ppa004033 was positioned in pseudomolecule 8. Moreover, the comparison of the deduced amino acid sequence of the three genes with SUTs from other plant species showed a difference in length only for ppa004033, suggesting a more accurate control of its prediction. The comparison between transcript prediction and RNAseq profile, both available in the public genome browser (http://services.appliedgenomics.org/fgb2/iga/ prunus_public/gbrowse/prunus_public/), showed a discrepancy at the 5′ -terminus of the sequence, and the attempt to amplify the whole CDS (coding DNA sequence) from cDNA with different primers and reaction conditions was unsuccessful (data not shown). For these reasons, a manual correction of the predicted CDS was performed, and a putative start codon was identified in the first intron, 36 bp upstream of the second exon. In accordance with the Kozak consensus sequence (Kozak 1987), the new ATG identified presented a purine at positions −3 and +4 (Fig. S1 in Supporting Information). Comparison of the three SUTs identified in peach with SUTs from nine other species revealed that the peach predicted protein grouped into the three clades belonging to the dicots, as proposed by (Kühn and Grof 2010), and, as expected they show no similarity with any of the sucrose facilitators (SUFs) found in legumes. Accordingly, the genes from P. persica were renamed PpSUT1 (ppa004033), PpSUT2 (ppa003041) and PpSUT4 (ppa004620) (Fig. 4). Additional analyses were performed on PpSUTs nucleic and amino acidic sequences, in order to substantiate their assignment to the SUTs group. PpSUTs gene structures appeared consistent with those of other plant species, in relation to clade membership, as proposed by (Shiratake 2007). In detail, PpSUT1 presented a large first exon and three small exons. This gene structure is slightly different, but still more similar to other members of the SUC2/SUT1 subfamily than to the other two families. PpSUT4, similarly to all genes of the SUC4 subfamily, had large first and second exons and three smaller exons. PpSUT2 consisted of 14 exons, according to gene structure of the SUC3/SUT2 subfamily comprising more than 10 small exons (Fig. S2). The corresponding predicted proteins had 511, 609 and 499 amino acids for PpSUT1, PpSUT2 and PpSUT4, respectively. The putative amino acidic sequences were analyzed by TMHMM Server v. 2.0 (Fig. S3), pointing out that all three PpSUT proteins displayed 12 highly probable transmembrane regions, typical structure of the members of the major facilitator superfamily. Moreover, both N-terminal and C-terminal were localized in the

Fig. 4. UPGMA consensus tree of SUT proteins identified in peach and in other plant species. Prunus persica: PpSUT1 (ppa004033), PpSUT2 (ppa003041), PpSUT4 (ppa004620); Arabidopsis thaliana: AtSUC1 (At1g71880), AtSUC2 (At1g22710), AtSUC3/AtSUT2 (At2g02860), AtSUC4 (At1g09960), AtSUC5 (At1g71890), AtSUC6 (At5g43610), AtSUC7 (At1g66570), AtSUC8 (At2g14670), AtSUC9 (At5g06170); Oryza sativa: OsSUT1 (Os03g07480), OsSUT2 (Os12g44380), OsSUT3 (Os10g26740), OsSUT4 (Os02g58080), OsSUT5 (Os02g36700); Malus domestica: MdSUT1 (AAR17700.1); Lycopersicon esculentum: LeSUT1 (CAA57726), LeSUT2 (AAG12987), LeSUT4 (AAG09270); Solanum tuberosum: StSUT1 (CAA48915), StSUT2 (AY291289), StSUT4 (AAG25923); Vitis vinifera: VvSUT2 (AAL32020), VvSUC11 (AF021808), VvSUC12 (AF021809), VvSUC27 (AF0218010); Lotus japonicas (AJ538041); Zea mays: ZmSUT1 (NP_001104840), ZmSUT2 (NP_001146651); Hordeum vulgare: HvSUT2 (CAB75881). To support the classification reliability, SUF proteins have been added to the tree: Pisum sativum: PsSUF1 (ABB30163); PsSUF4 (A3DSX1); Phaseolus vulgaris: PvSUF1 (ABB30165). Rectangles are used to point out the five clades according to the classification of Kühn and Grof (2010). Peach proteins are indicated by asterisks.

cytosolic side. As expected, PpSUT2 presented extended domains in the central loop and N-terminus. Additionally, a dileucin-like motif (LRQLL), a specific sequence for vacuolar targeting (Aoki et al. 2003), was found in the N-terminal cytoplasmic domain of PpSUT4 sequence (position 23–27), allowing us to infer a tonoplastic localization for this protein. PpSUTs transcriptional analysis by qRT-PCR The expression of PpSUT genes on the whole fruit tissue was evaluated by qRT-PCR, on samples collected at

different developmental stages, from 36 dafb to harvest, at 115 dafb. Results for qRT-PCR analysis from three independent experiments (Fig. 5) indicated that the PpSUT4 expression levels were the highest compared with PpSUT1 and PpSUT2. In young fruits, PpSUT4 transcription was relatively low, but increased showing a two-peaks pattern from about 50 to 85 dafb. Finally, in mature fruits, the relative abundance of PpSUT4 mRNA exhibited a steady reduction up to the completion of fruit development. In contrast, PpSUT2 displayed low expression levels with minor increment at 50 and 78 dafb. As compared Physiol. Plant. 2014

Fig. 5. Relative abundance of PpSUTs transcripts in peach mesocarp of cultivar Redhaven detected by qRT-PCR. PpSUT1, PpSUT2 and PpSUT4 expression patterns are represented by closed circles, open circles and closed triangles, respectively. The bars represent standard deviation calculated from the mean of three replicates.

to the other two genes, PpSUT1 showed undetectable transcriptional levels. Taken together, data obtained encouraged more detailed analysis informative for drawing conclusions on the role of transporters in sucrose partitioning. For these reasons, in situ hybridization and laser microdissection addressed to provide cell type-specific expression details were pursued. The pattern of PpSUTs mRNA accumulation in the developing peach fruits In situ localization of the PpSUTs transcripts was performed with digoxigenin-labeled probes to obtain information about tissue-specific expression of the transporters. The analyses were carried out in peach fruits, at two stages of growth (50 and 85 dafb), representative of the earliest and the middle phases of rapid growth rate. In immature peach mesocarp (SI), all probes (Fig. 6), displayed a distinct staining, in particular in the phloem tissue. In detail, besides the weak diffused signal in all parenchyma cells (absent in negative control), present for all PpSUTs probes, the localization of PpSUT2 and PpSUT4 transcripts was highly circumscribed in specific cells, constituting phloem tissue, as shown by the dark and well defined spots. Conversely, mesocarp sections incubated with PpSUT1 probe led to weak and diffuse staining of phloem tissue. The hybridization carried out on samples collected during the latter phase of development (SIII) (Fig. 6) indicated a general decrease in gene expression levels, consistent with qRT-PCR results. The parenchyma cells appeared stained only in section treated with PpSUT4 probe, and the significant amount of chromogenic Physiol. Plant. 2014

Fig. 6. Localization of PpSUT1, PpSUT2 and PpSUT4 transcripts by in situ hybridization, in serial sections of mesocarp tissue collected during SI (upper panel, ×50 magnification) and SIII (lower panel, ×10 magnification) stages of fruit development. In each panel, hybridizations with sense probe control are presented below the respective antisense riboprobe results. Phloem cells are indicated by arrows.

product previously localized in phloem cells, in samples treated with PpSUT2 and PpSUT4 probes, appeared unnoticeable. Identification of genes expressed differentially in parenchyma or vascular tissues by LMPC LMPC was employed in order to validate and integrate the information developed by in situ hybridization with high spatial resolution analysis. Therefore, RNA was extracted, amplified and reverse-transcribed from parenchyma or phloem tissue of peach fruit mesocarp properly collected (Fig. 7). The expression pattern of PpSUT1 and PpSUT2 genes appeared consistent in both the developmental stages considered, with PpSUT1 transcripts almost absent in both the tissues and PpSUT2 preferentially expressed within phloem complex cells. On the contrary, PpSUT4 appeared at all times expressed in both cell types, with faintly higher levels in parenchyma or phloem cells in the early or late phases of development, respectively (Fig. 7).

Fig. 7. LMPC of peach fruit mesocarp cells. Representative cross section (12 μm) of fruit under ultraviolet (C) and bright field (A, B, D, E, F). Magnification of the targeted site before (A, B) and after cutting (D, E). Panels (A) and (D) represent the sampling of parenchyma cells (bars = 300 μm). Panels (B) and (E) show the selection of phloem tissue (bars = 150 μm). Captured areas (F). Bottom panels (G, H): relative gene expression levels of PpSUT1, PpSUT2 and PpSUT4 measured in phloem and parenchyma cells by combination of LMPC and qRT-PCR in SI (G) and SIII (H) developmental stages. The bars represent standard deviation calculated from the mean of three replicates.

Discussion The orchestrated production, transport and storage of sucrose are crucial for normal plant growth and development (Kühn and Grof 2010). In different organs, two distinct routes for photoassimilates release have been described: the symplastic and the apoplastic pathway. In the first case, carbohydrates movement from cell to cell is driven by passive diffusion, owing to the presence of plasmodesmatal connections. On the other hand, when symplastic continuity is absent, sugars are actively transported across biological membranes by sugar carriers. Phloem unloading in the fruit of some plant species has been found to switch from a symplastic to an apoplastic pathway at the onset of cell expansion or at the beginning of ripening (Wu et al. 2004, Zhang et al. 2004, Zhang et al. 2006, Nie et al. 2010, Zhang et al. 2014). This developmental shift isolates the pericarp cells (sink symplast) from the phloem sieve elements and companion cells (vascular symplast), allowing the fruit

to accumulate solutes without inhibiting phloem influx (Keller and Shrestha 2013). Nevertheless, limited information is still available for peach, the third most important fruit species after apple and pear in the world market (Byrne et al. 2012). Similarly to the other stone fruits, peach exhibits a double sigmoid growth kinetic, characterized by four stages of growth. The first (SI) and the third (SIII) phases show a rapid increase in fruit size determined mainly by active cell division, and cell enlargement, respectively. During the other two phases, fruit growth rate appears significantly reduced. In particular, the growth slowdown in the second phase is related to the endocarp lignification, embryo development and accumulation of reserves into the seed (Bonghi et al. 2011, Falchi et al. 2013). Each developmental stage is also characterized by a distinct and dynamic pattern of sugar accumulation. The contribution in term of fixed carbon, provided by immature fruits photosynthetic activity, is limited to 5–9% (Pavel and DeJong 1993), therefore sugars imported, deriving from phloem system Physiol. Plant. 2014

are essential for the growth of these organs. At this stage, when a rapid cell division takes place, the main sugars noticeable in peach are glucose and fructose, consistent with the hypothesis of a link between hexoses presence and mitotic activity (Weber et al. 1996). On the other hand, sucrose, probably involved in stimulating cell differentiation (Weber et al. 1996), accumulates during the final phases of fruit development. The symplastic tracer, CF, has previously been used to investigate unloading pathway in several sink organs. In this study, we have adopted this approach in peach fruit, providing evidence for an apoplastic sucrose transfer in the early (SI) and middle (SIII) phases of development. This information has been expanded with the study of transporters that regulate the movement of sucrose across membranes, as important control points in the pathway, being sucrose the main form of photoassimilate transported from source leaves to sink tissues in higher plants (Lalonde et al. 1999), and the predominant sugar accumulated in peach mature fruit (Moriguchi et al. 1990, Vizzotto et al. 1996, Nonis et al. 2007, Falchi et al. 2013). Three putative SUTs encoding genes (PpSUT1, PpSUT2, PpSUT4) were identified in the peach genome sequence and transcriptionally analyzed to relate their expression with sucrose partitioning. As expected, the determination of expression profiles by qRT-PCR revealed that PpSUT1 is barely expressed in fruit. This result is not surprising, being confirmed also in peach RNA-Seq data from the public database (http://services. appliedgenomics.org/fgb2/iga/prunus_public/gbrowse/ prunus_public/). On the other hand, SUTs of the SUT1/ SUC2 subfamily, classified as phloem loaders (Wippel and Sauer 2012), are usually localized at the plasma membrane of the SE–CC and facilitate the uptake of sucrose from the apoplast, thereby creating the osmotic potential which drives mass flow in the phloem system (Liesche et al. 2011). Therefore, we focused mainly on the characterization of PpSUT2 and PpSUT4 expressed in the sinks, in general poorly investigated. PpSUT4 displayed the highest expression level at all developmental stages. Other members of SUT4 subfamily have been found in fleshy fruit, such as MdSUT1, PbSUT1 and VvSUC11 suggesting an involvement of these proteins in sugar distribution (Davies et al. 1999, Peng et al. 2011, Zhang et al. 2013). Conversely, consistent with the behavior of other members belonging to its subfamily (Ayre 2011), a small amount of PpSUT2 transcripts has been detected in developing peach fruits. Besides, fruit flesh is an heterogeneous tissue, and only a complete spatial information of different mRNA abundance could help in elucidating their specific physiological tasks. To this purpose, appropriate experiments of in Physiol. Plant. 2014

situ hybridization, validated and improved by laser microdissection-assisted expression analysis, were set up. As expected, no significant signal was detected in tissue localization of PpSUT1 mRNA, determined by means of these techniques, both in SI and SIII stages, confirming the results obtained by qRT-PCR, and the hypothesis of its role in phloem loading. On the contrary, when antisense RNA probes for PpSUT2 and PpSUT4 were hybridized to peach fruit sections, strong signals were observed in the phloem cells and lighter staining was visible in parenchyma cells. On the other hand, laser microdissection-assisted expression analysis allowed to distinguish a remarkable presence of PpSUT2 transcripts in phloem bundles throughout fruit development, and an upregulation of PpSUT4 both in parenchyma and in bundles, with differences of steady-state mRNA levels of genes between the two cell types, according to the developmental stage. PpSUT2 is probably localized to the plasma membrane and exhibits a long N-terminal and large central hydrophilic loop, as other members of SUT2 clade (Gottwald et al. 2000). This structure most likely confers some unique function or activity to these proteins that are not shared by other SUTs, therefore a role in sucrose sensing and signaling has been attributed to SUT2-type transporters (Barker et al. 2000). However, the function of these SUTs is still subject of much debate (Eckardt 2003). Concerning peach fruit, we cannot exclude any of these hypotheses. PpSUT2, expressed at low and constant levels during development, might be transcribed in companion cells and function, in sieve elements, in the perception of carbohydrates availability. Otherwise, PpSUT2 might be involved in the retrieval of sucrose lost from the symplastic continuum into the phloem and, when expressed in parenchyma cells, it might be active in the import of sucrose into sink tissues, functioning as symporter from the apoplast (Fig. 8). Several members of SUT4 subfamily have been localized in the tonoplast, and the identification of a dileucin-like motif, a specific sequence for vacuolar targeting (Aoki et al. 2003), argued for a vacuolar localization of PpSUT4. All data obtained concerning PpSUT4 transcription were consistent: higher expression levels in SI, concomitant with rapid fruit growth, primarily due to the active cell division, and following downregulation at SIII stage, when a minor occurrence of transcription in phloem cells takes place. Because sucrose is rapidly stored into the vacuole, PpSUT4 probably has a role in the regulation of sucrose release to sustain cell metabolism. On the other hand, in the second phase of fruit development, the expression of PpSUT4 decreases, predominantly in parenchyma cells, allowing the beginning of sucrose accumulation into the

Fig. 8. Model proposed for the role of PpSUTs in phloem unloading and assimilates partitioning in peach fruit. PpSUT2, expressed at SI both in vascular bundles and in parenchyma, is most likely localized in plasma membrane, therefore PpSUT2 (green circle) could function in a mechanism of solute retrieval from apoplast and in distribution of sucrose, being parenchyma cells symplastically isolated from phloem. At this stage, two hypothesis have been proposed for PpSUT4, which could be involved in vacuolar mobilization of sucrose (dark blue square) or could have a localization and a role similar to PpSUT2 (light blue square). In SIII, an analogous model can be hypothesized except for the absence of PpSUT2 transcripts in parenchyma cells. The presence of PpSUT4 mRNA at this stage, when sucrose accumulates, is in agreement with the possibility that it is not localized in tonoplast, even if the low transcriptional levels encourage verifying every hypothesis. Sucrose effluxers (indicated by question marks) failed to be identified, but a role for the SWEET proteins (Chen et al. 2012), at this level, could be hypothesized.

vacuole. Functional characterization of cloned higher plant SUTs has currently failed to identify sucrose/H+ antiporters (Kühn and Grof 2010), consequently a role for PpSUT4 in sugar uptake and vacuolar compartmentation cannot be hypothesized. Moreover, the prediction of PpSUT4 localization is based on the simple sequence analysis and still uncertain. Therefore, the presence of gene transcripts in the vascular bundles, indicated by LMPC, suggests a role for PpSUT4 different from the vacuolar mobilization and its possible plasma membrane localization. Similar to PpSUT2, PpSUT4 could function in regulating extracellular sucrose by way of re-uptake or, if expressed outside of the phloem, it could function directly in sucrose uptake into sink cells, playing a role in determining sink strength, as suggested by (Weise et al.

2000). It is noteworthy that the description of sucrose distribution pathway is far from completely elucidated, being the identification of proteins responsible for the export of sucrose still lacking. We can speculate that, as hypothesized by other authors (Chen et al. 2012), SWEET proteins could be involved in the export from phloem parenchyma of sucrose, then taken up in all mesocarp cells by a secondary active proton-coupled SUT (Fig. 8). In this regard, we were able to identify 15 sequences related to SWEET proteins in peach genome, two of which display high levels of homology with Arabidopsis orthologs, known as sucrose carriers (Fig. S4). The transcript abundances of the two SWEETs, measured by RNA-seq, available in peach GBrowse, confirm their expression in fruits, supporting the hypothesis of an involvement in sucrose distribution, but additional investigations are clearly required before ascribing a role for these proteins in the process. Interestingly, none of the three SUTs identified presented a pattern of regulation consistent with sucrose accumulation, being all PpSUTs genes downregulated at the end of fruit development. Unfortunately, our inability to produce suitable section of ripe fruits had in all probability precluded a cell-specific expression analysis in peach flesh at the last stage of growth. Preliminary results indicate that, in this phase, CFDA is apparently released from the phloem strands into surrounding fruit tissues (data not shown). This is not uncommon, as the unloading route may vary according not only to sink type, but also to sink development, function and growth condition (Nie et al. 2010). Our evidence appear consistent with the low levels of PpSUTs transcripts and with previous studies on the energy dependence of sugar transport in peach fruit, indicating an unaltered uptake of sugars at the end of fruit development, even in presence of the proton motive force inhibitor CCCP (Carbonyl cyanide m-chlorophenylhydrazone) (Vizzotto et al. 1996). Taken together, our data suggest that two (PpSUT2 and PpSUT4) of the three SUTs identified in peach genome could have a pivotal role in the regulation of sucrose unloading and distribution in fruits. However, further investigations aiming to elucidate the function of these proteins at the end of peach development, their precise localization at subcellular level and to discover other transporters or facilitators acting in the process are required. The answer to these questions will enable a better understanding of sucrose accumulation mechanisms and, in the long run, will be supportive in designing strategies for manipulating this essential quality determinant. Acknowledgements – This work was supported by the Italian Ministry of Research and University, PRIN Project

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(20074AXSCA_002). We would like to thank Stefano Gustincich (SISSA, International School for Advanced Studies, Trieste, Italy) and Michele Bellucci (IBBR-CNR, Perugia, Italy) for the use of LMPC system.

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Supporting Information Additional Supporting Information may be found in the online version of this article: Fig. S1. Schematic representation of PpSUT1 transcript correction. In orange, the automatically predicted CDS in GBrowse, in yellow the new gene model lacking of the first exon and the new ATG starting codon. Fig. S2. PpSUT gene structures from Prunus persica. Physiol. Plant. 2014

Fig. S3. Prediction of transmembrane domain by TMHMM Server v.2.0 software for PpSUT1, PpSUT2 and PpSUT4. Fig. S4. UPGMA consensus tree of Arabidopsis and peach SWEET proteins. The tree topology was generated

Edited by C. Foyer

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by the neighbor-joining method of MEGA version 4 software (Tamura et al. 2011). The red box indicates the sucrose effluxers identified in Arabidopsis; in yellow the two peach SWEET proteins clustering in this group and expressed in fruit.