Introduction {#s1}
============

In plants, sugars are the key carbon source for energy production and macromolecule biosynthesis, and they are important for carbohydrate accumulation in crop organs consumed as food by humans ([@CIT0007]). Thus, sugar allocation between source leaves (net exporters) and sink organs (net importers), such as young leaves, flowers, and storage organs, is of primary importance for organ development and plant growth, and as a sustainable food supply ([@CIT0001]; [@CIT0055]). For long-distance allocation, sugars are primarily synthesized by photosynthetic carbon fixation in source leaves and are loaded, mostly as sucrose, into phloem-specific companion cell (CC)--sieve element (SE) complexes. This loading occurs either symplasmically via plasmodesmata or via the apoplasmic route, with fine-tuned regulation of sugar partitioning ([@CIT0078]; [@CIT0053]; [@CIT0086]). Regarding the apoplasmic route, plasma membrane-localized sugar transporters facilitate sucrose efflux and import between adjunct CCs and parenchyma cells of minor veins, promoting mass flow in the phloem for continuous sugar allocation ([@CIT0059]; [@CIT0013]; [@CIT0034]).

During sucrose loading, SWEET ([s]{.ul}ugar [w]{.ul}ill [e]{.ul}ventually be [e]{.ul}xported [t]{.ul}ransporters) proteins, especially clade III SWEET members localized in the plasma membrane of phloem parenchyma cells, are probably the main efflux mechanism to facilitate sucrose export into the apoplast in both dicot ([@CIT0016]; [@CIT0020]) and monocot plants ([@CIT0005]). An absence of SWEET exporters significantly suppresses sucrose loading and plant growth, with accumulation of sugars and starch in source leaves ([@CIT0016]; [@CIT0040]; [@CIT0005]). The final loading step is mediated by SUT1/SUC2-type ([su]{.ul}crose [t]{.ul}ransporter; H^+^/[suc]{.ul}rose symporter) in the plasma membrane of phloem cells, that actively imports apoplasmic sucrose into CC--SE complexes against a concentration gradient ([@CIT0059]; [@CIT0034]). Plant species with decreased SUT1 uptake ability were stunted, due to lack of sugar loading from source leaves ([@CIT0025]; [@CIT0069]; [@CIT0067]).

Similarly, sucrose unloading from phloem complexes to sink organs can be symplasmic and apoplasmic depending on organ types and developmental stages ([@CIT0050]; [@CIT0002]; [@CIT0034]). The initial phloem unloading includes sucrose fluxes across SE, CC, and adjacent phloem parenchyma cells. Along the post SE--CC unloading pathway, a fraction of the sucrose could be unloaded from the CCs to the apoplast, probably by simple or facilitated diffusion ([@CIT0050]; [@CIT0034]). The apoplasmic sucrose can be either directly imported into nearby parenchyma cells by a sucrose transporter, or hydrolyzed by a cell wall invertase into hexoses that are subsequently imported by monosaccharide transporters ([@CIT0002]; [@CIT0048]; [@CIT0034]). Although many sugar transporter families have been characterized in Arabidopsis and other crops, transport mechanisms involved in initial sugar unloading are largely unknown. Some SUT-type sucrose transporters were implicated in sucrose export from CCs ([@CIT0006]). Monosaccharide transporters (MSTs) ([@CIT0063]), such as STP ([s]{.ul}ugar [t]{.ul}ransport [p]{.ul}rotein) transporters ([@CIT0064], [@CIT0063]), may be involved in hexose import; however, experimental evidence for their physiological roles is lacking. SWEET transporters not only mediate sucrose efflux for phloem loading, but are also major sugar unloaders mediating sugar efflux, as well as import, from maternal connecting tissues to reproductive sinks ([@CIT0041]; [@CIT0015]; [@CIT0085]). SWEETs have been characterized as low-affinity hexose and sucrose bidirectional transporters (exporters or importers), depending on physiological demands. *At*SWEET9 specifically localizes in the plasma membrane of inflorescence parenchyma cells to facilitate sucrose efflux for nectar formation ([@CIT0041]). Clade III SWEETs expressed in maternal cells of developing seeds catalyzed sucrose efflux and import, supporting embryo development in Arabidopsis ([@CIT0015]) and rice ([@CIT0085]). Moreover, the hexose export activity of *Zm*SWEET4c in corn and *Os*SWEET4, its ortholog in rice, is of primary importance for seed filling and appears to be linked to seed size ([@CIT0068]). These studies highlight the possibility that SWEETs could have a fundamental role in sugar unloading in sink organs as early as the seedling development stage ([@CIT0020]).

During vegetative stages, young leaves and roots are major sink organs ([@CIT0079]), with up to 60% of newly synthesized sucrose exported to sink leaves. Unlike simple rosette leaves in Arabidopsis, tomato (*Solanum lycopersicum*) has sequentially developed compound leaves, with multiple leaflets that require an extended morphogenetic phase ([@CIT0065]), suggesting that a controlled, fine-gated sugar unloading mechanism is needed through the phloem to developing leaves. Tomato is an important vegetable crop worldwide (United Nations Food and Agriculture Organization; <http://www.fao.org/>), with many efforts focused on sugar allocation to improve fruit size, sweetness, and quality ([@CIT0057]; [@CIT0023]). However, vigorous vegetative growth is the primary determinant of tomato biomass and fruit yield ([@CIT0077]). Thus, adequate leaf production could increase flower numbers and fruit weight up to 60% ([@CIT0032]; [@CIT0077]). However, transport mechanisms regulating initial sugar unloading to sink leaves are largely unknown in tomato. Early studies using either radiotracers, a membrane-impermeable fluorescent carboxyfluorescein, or green fluorescent protein (GFP) fusion proteins have demonstrated that initial phloem unloading in sink leaves is largely via a symplasmic pathway ([@CIT0060]; [@CIT0074]; [@CIT0054]). However, these data do not exclude an apoplasmic component. For instance, developing maize leaves have been found to switch dynamically from symplasmic to apoplasmic phloem unloading ([@CIT0003]). These results imply that putative, as yet unidentified, membrane carriers which reside in unloading cells may participate in the apoplasmic sugar unloading into tomato sink leaves.

To date, only two sugar transporters, *Le*SUT1 and *Le*SUT2 sucrose transporters, have been implicated in source to sink sugar distribution during vegetative growth of tomato plants ([@CIT0004]; [@CIT0081]; [@CIT0048]). *Le*SUT1 proteins are localized in the SEs and appear to have a major role in sucrose loading in source leaves ([@CIT0004]; [@CIT0081]), as impaired *LeSUT1* expression increased accumulation of soluble sugars and starch in mature leaves ([@CIT0029]). Although SE-localized *Le*SUT2 has been implicated in sucrose unloading to sink leaves, *LeSUT2* antisense mutants reduced fruit fertility, with no apparent changes in sugar concentrations in leaves or vegetative tissues ([@CIT0029]). Therefore, *Le*SUT2 is probably not the major sucrose unloader required for sugar unloading in young leaves. Based on the insensitivity of sucrose unloading to a metabolic inhibitor ([@CIT0060]), and a steep sucrose concentration gradient between SE--CC complexes and the apoplast in most sink organs performing apoplasmic unloading ([@CIT0050]), a passive sucrose facilitator, such as SWEET sucrose carriers, has been suggested to mediate the efflux ([@CIT0048]; [@CIT0034]). Yet, the genetic evidence is still missing.

On the other hand, a hexose facilitator may also be required to act in series to re-load apoplasmically hydrolyzed sucrose into the downstream phloem parenchyma cells ([@CIT0050]). For instance, radiotracer analysis indicated that glucose can be retrieved from the apoplasm of sink leaves by phloem parenchyma cells ([@CIT0060]). Thus, a glucose facilitator (e.g. SWEETs) on the plasma membrane of parenchyma cells could be a possible candidate to perform this function driven by the relatively low glucose concentrations in the cytoplasm, compared with the apoplasm ([@CIT0066]; [@CIT0056]).

We deduced that the sink leaf-expressed *Sl*SWEET transporter could be a promising sugar unloader for vegetative growth. Although the complete SWEET gene family has been identified in tomato 'Heinz 1706' and its expression pattern characterized by RNA sequencing (RNA-seq) ([@CIT0021]; [@CIT0062]), the role of SWEET proteins in phloem unloading during the vegetative stage in tomato is unknown. The objective in this study was to characterize transport properties of the most highly expressed *Sl*SWEET1a (Solyc04g064610) in young developing leaves of tomato plants. Dominant expression of *Sl*SWEET1a--β-glucuronidase (GUS) fusion proteins in unloading veins and high correlation between *SlSWEET1a* transcripts and carbon importing activity in sink leaves implicated that it had a specific role in sugar unloading. Furthermore, reduced hexose accumulation in young leaves consistently occurred in *SlSWEET1a*-silenced tomato plants. Thus, a putative role for *Sl*SWEET1a in apoplasmic sugar unloading to sink leaves is discussed.

Materials and methods {#s2}
=====================

Plant materials and growth conditions {#s3}
-------------------------------------

The Micro-tom cultivar of tomato (*Solanum lycopersicum*) was used, except where indicated. Tomato seeds were surface-sterilized for 8 min using diluted bleach solution (30% CLOROX bleach and 0.1% Triton X-100, v/v) and then washed three times with sterilized water. To propagate tomato plants, sterilized seeds were directly germinated in water at room temperature for 2 d and then transferred to a hydroponic system or pots (diameter, 11.5 cm) containing a mixture of soil and peat moss (1:1). All plants were grown in a controlled chamber (25 °C and 16/8 h light/dark regime with \~100 μmol m^−2^ s^−1^ illumination) before harvest or experiments. To collect various organs (except fruits), tomato plants were grown hydroponically for 3 weeks (for vegetative organs) to 5 weeks (for reproductive organs), then roots, stems, mature leaves (the terminal leaflet \~4 cm), young leaves (the terminal leaflet \<2 cm), flower buds (developing green buds), and flowers (first day of anthesis) were collected ([Supplementary Fig. S1](#sup1){ref-type="supplementary-material"} at *JXB* online). For leaves, all leaflets from one leaf were pooled together as one sample. Various stages of fruits were collected from soil-grown plants after 14, 21, 35, or 42 d of flowering. To analyze expression in different regions of leaves, the terminal leaflets of young leaves that were 80% of the final size (\~3--3.5 cm) were harvested from 5-week-old plants. Then, segments of 1 cm in length from the leaf base to tip were separated for mRNA analysis. For mesophyll protoplast isolation, *Arabidopsis thaliana* ecotype Col-0 plants were grown in peat moss, vermiculite, and perlite (6:1:1) under short-day conditions (22 °C, 10/14 h light/dark regime, \~80 μmol m^−2^ s^−1^ illumination) for 4--5 weeks before isolation.

Analysis of mRNA transcripts {#s4}
----------------------------

Total RNA was isolated from tomato organs with TRIsure reagent (Bioline, <http://www.bioline.com/>) with an RNA purification column (GeneMark, <http://www.genemarkbio.com/>), whereas RNA samples from fruits were prepared using the cetyltrimethylammonium bromide (CTAB) method ([@CIT0049]). Total RNA transcripts were reverse-transcribed, and gene-specific primers for all 31 *SlSWEET* genes (primer sequences are listed in [Supplementary Table S1](#sup1){ref-type="supplementary-material"}) were used for real-time quantitative PCR (qRT-PCR) as described ([@CIT0011]). Expression of the reference gene *SlActin7* (Solyc11g005330) was used to determine relative expression levels according to the following equation: 1000×2^−^(^Ct^~*SlSWEET*~^−Ct^~*SlActin7*~) (Ct=threshold cycle; [@CIT0011]).

Localization of GUS fusion proteins {#s5}
-----------------------------------

To generate *Sl*SWEET1a--GUS fusion proteins for observation, the entire DNA fragment of *SlSWEET1a* (1467 bp), including all introns and deletion of the stop codon, was amplified from genomic DNA prepared from Micro-tom leaves with specific primers (forward, 5\'-CCGCGGATGGGTGTTGTTCATACTCTG-3\'; and reverse, 5\'-CTGCAGTCCAGCGCCAACTT GCTCAAGCCTTGACTT-3\'). The resulting fragments were first cloned into a pGM-T vector using AT cloning technology and then cloned into pUTKan (including the GUS fragment) binary vector by *Sac*II and *Pst*I restriction sites (pUTKan-gSlSWEET1a). Subsequently, the native promoter of *SlSWEET1a* (2000 bp) was amplified with specific primers (forward, 5\'-TTAAACCTGATCGTCACATAAA-3\'; and reverse, 5\'-CAGAGTATGAACAACACCCATGTC-3\') and cloned into pGM-T, and then cloned into a pUTKan-gSlSWEET1a binary vector, using *Sac*I and *Kpn*I restriction sites. Then, Micro-tom plants were transformed with the resulting pUTKan-PSlSWET1a::gSlSWEET1a--GUS construct in the Academia Sinica Transgenic Plant Core Lab (<http://transplant.sinica.edu.tw/en/aboutus/intro/index3.htm>). In total, five T~1~ transgenic tomato plants were obtained. Due to limited numbers of T~1~ transgenic tomato plant leaves, three young leaves were collected from three independent T~1~ transgenic lines and three mature leaves were collected from other independent lines. Flowers were collected from T~2~ lines. All samples were examined by histochemical staining, 16 h for young leaves and mature leaves, and 9 h for flowers, as described ([@CIT0028]). Stained organs were imaged with a LEICA Z16 APO microscope.

Translocation of carboxyfluorescein {#s6}
-----------------------------------

To explore the phloem unloading fluxes in tomato sink leaves, the abaxial surface of the terminal leaflet of mature source leaves from 2-week-old tomato plants was gently abraded with fine sandpaper. A drop of 200 µl of CFDA \[5(6)-carboxyfluorescein diacetate, Sigma-Aldrich, USA\] solution (300 µg ml^−1^) was applied to each leaf, which was then covered with plastic wrap to prevent evaporation. After 3 h, fluorescence in the terminal leaflets (\<2 cm) of attached sink leaves on intact plants was observed and imaged using a Leica fluorescence microscope equipped with a SOLA light engine (Lumencor, USA) and a Nikon EOS700d digital camera. Images were taken under a GFP filter set and exported by the EOS utility software.

Subcellular localization of GFP fusion proteins {#s7}
-----------------------------------------------

To generate *Sl*SWEET1a--GFP fusion proteins, the cDNA fragment of *SlSWEET1a* (741 bp) without the stop codon was amplified from the pDRf1-GW-SlSWEET1a plasmid (see 'Expression of clade I *Sl*SWEET1a in yeast') with specific primers (forward, 5\'-CACCATGGGTGTTGTTCATACT-3\'; and reverse, 5\'-GGCGCCAACTTGCTCAAGCCTTGACTTGC-3\') and cloned into a pENTR-D-TOPO vector using TOPO cloning technology (Invitrogen, USA). Subsequently, the cDNA was transferred into pUBN-GFP and pUBC-GFP via an LR reaction using Gateway technology ([@CIT0026]). Arabidopsis and tobacco protoplasts were then transformed with the resulting pUBN-SlSWEET1a and pUBC-SlSWEET1a constructs to express N- and C-terminal translational GFP fusion proteins, respectively.

Arabidopsis mesophyll protoplasts were isolated as described ([@CIT0083]). A portion of the protoplast preparation (200 µl at a cell density of 2×10^5^ ml^−1^) was mixed with 10--20 μg of plasmids and 220 μl of polyethylene glycol (PEG) solution prior to incubation for 15 min, at 25 °C in the dark. Protoplasts were then washed and re-suspended in 1 ml of W5 solution (154 mM NaCl, 125 mM CaCl~2~, 5 mM KCl, and 2 mM MES, pH 5.7) and incubated for 24--48 h in the dark before observation. To determine the position of the plasma membrane, protoplasts were either co-transformed with the membrane marker pRT101-AtPIP2A--red fluorescent protein (RFP) in a 1:1 ratio ([@CIT0046]) or stained with 0.5 μl of 10 μM FM4-64 dye just before observation ([@CIT0011]). Fluorescence was observed on a Carl Zeiss LSM780 confocal microscope (Instrument Development Center, National Cheng Kung University) as described ([@CIT0011]).

Expression of clade I *Sl*SWEET in yeast {#s8}
----------------------------------------

To generate expression constructs for yeast, cDNA sequences of clade I *SlSWEET* were amplified using Phusion polymerase (New England Biolabs, USA) with gene-specific primers (primer sequences are given in [Supplementary Table S2](#sup1){ref-type="supplementary-material"}) from leaf cDNA derived from tomato cultivar 'TS19'. The cDNA sequences were confirmed by sequencing and had 100% identity with those in the Sol Genomics Network (<https://solgenomics.net/>). Then, cDNA was first cloned into the pDONR221-f1 vector and subsequently transferred to the pDRf1-GW vector using Gateway technology. The yeast strains YSL2-1 ([@CIT0011]) and EBY4000 were then transformed with the resulting pDRf1-GW-SlSWEET plasmids using the lithium acetate (LiAC) method ([@CIT0024]). Transformants were selected on synthetic deficient medium without uracil (SDM-U medium, containing 1.7 g of yeast nitrogen base without amino acids, 5 g of ammonium sulfate, 2% maltose, 2% agar, and 0.01% histidine, leucine, and tryptophan in 1 liter). The pDRf1 empty vector with a Gateway cassette deleted and a plasmid pDRf1-Hxt5 that harbored a yeast hexose transporter were also transformed as a negative and positive control, respectively, of transport activity ([@CIT0011]).

Yeast growth assay {#s9}
------------------

Yeast cells from single colonies were grown at 30 °C overnight and then refreshed with new SDM-U medium until they reached early mid-log phase growth (OD~600~ of 0.8--1.0). Cells were collected and diluted with SD-U medium (SDM-U medium without 2% maltose) to an OD of 0.2. Then, serial dilutions (1, 10^--1^, 100^--1^, and 1000^--1^) of all desired cells were prepared and 5 μl from each dilution was spotted on SDM-U medium or SD-U medium supplemented with 2% glucose, galactose, mannose, fructose, or sucrose. To maintain the media at pH 5 or 7, Tris-MES buffers (6.054 g of Tris and 9.76 g of MES in 1 liter) were included. Plates were incubated at 30 °C for 3--6 d and then scanned.

Radiotracer uptake assay in yeast {#s10}
---------------------------------

One portion of overnight yeast culture was diluted with 50 ml of fresh SDM-U medium to an OD~600~ of 0.2. After incubation for 4 h at 30 °C, yeast cells (\~0.5 OD~600~) were collected, washed, and re-suspended in 50 mM sodium phosphate buffer (112 mg of Na~2~HPO~4~ and 6.8 g of NaH~2~PO~4~ in 1 liter, pH 5) to an OD~600~ of 5 before assay. For a time-dependent uptake assay, uptake buffer containing 2 mM glucose and 1.5 μCi of \[^14^C\]glucose ml^−1^ in 50 mM sodium phosphate buffer (pH 5) was prepared. For a concentration-dependent uptake assay, uptake buffers with 11iter glucose concentrations (0.2, 1, 2, 5, 10, 20, 40, 100, 200, and 500 mM glucose in 50 mM sodium phosphate buffer, pH 5) with same molar ratio of μCi of \[^14^C\]glucose were prepared. For competition assays, uptake buffers (2 mM glucose with 2 μCi of \[^14^C\]glucose ml^−1^) without other sugars (control) or supplemented with 10-fold concentrations of various cold sugars (20 mM glucose, galactose, fructose, mannose, sucrose, and maltose, respectively) were prepared. For analysis under various pH values, uptake buffer medium with various pH values were prepared using 50 mM sodium phosphate containing 2 mM glucose with 2 μCi of \[^14^C\]glucose ml^−1^. Before the uptake assay, cells were washed once with corresponding cold pH buffers. For treatment with the protonophore NH~4~Cl, yeast cells were pre-treated with 10 mM NH~4~Cl for 10 min at 30 °C, then uptake buffer of pH 5 (2 mM glucose with 2 μCi of \[^14^C\]glucose ml^−1^) was added. To start the uptake assay, 110 μl of corresponding uptake buffers were always added into an equal volume of cells that was incubated at 30 °C. At the indicated time points in the time-dependent assay or at 10 min in all other assays, cells were collected on filter paper (MCE Membrane Filter, 0.45 μm) and washed three times with ice-cold 50 mM sodium phosphate buffer by vacuum filtration. Filter paper (with cells) was transferred into scintillation vials containing 4 ml of Rotiszint^®^ eco plus (Carl Roth, Germany) and incubated at room temperature for 16--18 h. Radioactivity of lysed cells was quantified in a Tri-Carb 4810TR scintillation counter (Perkin Elmer, USA) and a kinetic curve determined using single rectangular regression (Sigmaplot Version 13).

Induction of VIGS (virus-induced gene silencing) in tomato leaves {#s11}
-----------------------------------------------------------------

VIGS was carried out as described ([@CIT0044]). In general, the 300 bp fragment of *SlSWEET1a* for gene silencing was selected via the VIGS tool on the Sol Genomics Network (<https://solgenomics.net/>) and amplified from the pDONR221-f1-SlSWEET1a plasmid using Phusion polymerase with specific primers (forward, 5\'-CACCATGGGTGTTGTTCATACT-3\'; reverse, 5\'-CTTCTTCTCTTTAGTTGG-3\'). The resulting fragment was first cloned into pENTR-D-TOPO (Invitrogen, USA) via TOPO cloning technology, then transferred into pTRV2 via an LR reaction ([@CIT0044]). Then, *Agrobacterium tumefaciens* GV3101 was transformed with the resulting pTRV2-SlSWEET1a construct via the freeze--thaw method ([@CIT0030]). Positive transformants were selected on LB medium (5 g of yeast extract, 10 g of tryptone, 10 g of NaCl in 1 liter) supplemented with kanamycin (50 mg l^−1^) and gentamycin (50 mg l^−1^). A portion of overnight culture (2.5 ml) derived from an *Agrobacterium* transformant was added to 25 ml of LB fresh medium containing antibiotic, 10 mM MES, and 200 μM acetosyringone (Sigma-Aldrich, USA) and grown overnight at 30 °C. Cells were collected by centrifugation at 4000 *g* for 5 min and re-suspended to an OD~600~ of 1.5 with infiltration medium (10 mM MgCl~2~, 10 mM MES, and 200 μM acetosyringone). An aliquot (1 ml of culture) was injected into cotyledons of 2-week-old tomato plants with a 1 ml syringe. To estimate silencing efficiency, *Agrobacterium* cells carrying the silencing construct, pTRV2-SlPDS-VIGS that would result in reduced expression of the phytoene desaturase gene so as to inhibit chlorophyll biosynthesis and cause etiolation ([@CIT0043]), was also injected into other plants.

Analysis of sugar contents {#s12}
--------------------------

Young leaves (the terminal leaflet \<2 cm, \<50% of the final size) and mature leaves (the terminal leaflet \~4 cm), including all three leaflets, from VIGS-silenced tomato plants were harvested 2 weeks after injection and stored at −70 °C before extraction. Frozen leaves (0.2--0.5 g) were ground into fine powder and 480 μl of extraction buffer medium \[containing 460 μl of methanol (HPLC grade) and 20 μl of 0.2 mg ml^−1^ ribitol\] was added. Mixtures were incubated at 70 °C for 10 min and centrifuged at 14 000 *g* for 10 min at room temperature. The supernatant was transferred into a glass vial, with 250 μl of cold chloroform (HPLC grade) and 500 μl of cold MQ water added in the given order. Mixtures were vortexed for 15 s and then centrifuged at 4000 *g* for 15 min at 20 °C. An aliquot of 200 µl of supernatant was transferred into a new 1.5 ml Eppendorf tube and freeze-dried in an LyoLab 5 LT freeze dryer (LSI, USA). Dry samples were derivatized with 20 μl of methoxyamination solution (0.02 g of methoxyamine hydrochloride in 1 ml of pyridine) at 30 °C for 90 min and then the mixture was added to 100 μl of 99:1 BSTFA+TMCS solution (Macherey-Nagel, Germany) and incubated at 70 °C for another 120 min. All extracts were then transferred into 250 μl pulled conical glass insert vials (Agilent, Germany) and analyzed with an Agilent 7890A/5975C GC-MS System with a 60 m×0.250 mm column (Agilent, Germany), as described ([@CIT0058]; [@CIT0075]). Sugars were quantified relative to the internal ribitol standard, using the following retention times: glucose, 23.286 min and 23.521 min; fructose, 23.001 min and 23.093 min; galactose, 23.235 min and 23.487 min; mannose, 23.168 min and 23.378 min; and sucrose, 30.869 min.

Results {#s13}
=======

*Sl*SWEET1a was highly expressed in aerial sink organs {#s14}
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The genome of tomato cultivar 'Heinz 1706' was sequenced in 2012 ([@CIT0072]) and 31 tomato *SlSWEET* genes have been identified ([@CIT0021]; [@CIT0062]). However, these genes were not well annotated to the SWEET family, which makes it difficult to compare orthologous genes. Therefore, the phylogenic tree of all 31 *SlSWEET* genes was generated and classified into four clades ([Supplementary Fig. S2](#sup1){ref-type="supplementary-material"}). To identify the key *SlSWEET* member responsible for sugar unloading into sink tomato leaves, qRT-PCR was performed for all 31 *SlSWEET* genes using cDNA isolated from whole new developing leaves (the terminal leaflet was smaller than 2 cm) of 4-week-old hydroponic-grown tomato plants. Based on relative expression levels in young tomato leaves, transcript levels of *SlSWEET1a* (Solyc04g064610) were markedly higher compared with other genes ([Fig. 1A](#F1){ref-type="fig"}). Interestingly, *SlSWEET1a* was also highly expressed in reproductive sink organs, such as flower buds and flowers ([Fig. 1B](#F1){ref-type="fig"}), but only weakly expressed during the maturation stage of tomato fruits ([Fig. 1C](#F1){ref-type="fig"}). Nevertheless, expression of *SlSWEET1a* was increased \>10-fold during the maturation stage (35--42 d after flowering) of fruit development.

![Expression of *SlSWEET* genes in tomato organs. (A) Stable mRNA transcripts of 31 *SlSWEET* genes in young leaves. (B) Developmental expression of *SlSWEET1a* in young leaves (YL), mature leaves (ML), roots (R), stems (S), flowers (F), and flower buds (FB) from 3- (vegetative organs) to 5-week- (reproductive organs) old hydroponically grown tomato plants. (C) Gene expression of *SlSWEET1a* in several stages of tomato fruits from soil-grown plants. Total RNA was isolated from various organs, and cDNA was used for qRT-PCR with gene-specific primers. Relative expression level by normalizing to an internal control, *SlActin7*, is shown. Results are means ±SE from three independent biological repeats. DAF=days after flowering.](erz154f0001){#F1}

*Sl*SWEET1a proteins are highly abundant in unloading tissues {#s15}
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To explore tissue-specific localization of *Sl*SWEET1a proteins, we generated transgenic tomato plants expressing a C-terminal translational GUS gene fusion with the complete genomic *SlSWEET1a* DNA sequence, containing all introns driven by the native *SlSWEET1a* promoter (*Sl*SWEET1a--GUS). After shoot regeneration and transfer to potting mix for 1.5 months, mature and new leaves from T~1~ tomato plants were stained to detect GUS activity. Predominant blue staining was observed in the first- to third-order veins in both young ([Fig. 2A, B](#F2){ref-type="fig"}) and mature leaves ([Fig. 2C, D](#F2){ref-type="fig"}). Dominant expression of the *Sl*SWEET1a--GUS fusion in leaf veins was also observed in the other four independent T~1~ transgenic tomato plants ([Supplementary Fig. S3A](#sup1){ref-type="supplementary-material"}). The same localization to leaf veins was also observed in the three T~2~ transgenic plants, although with less staining (data not shown). Low expression of *Sl*SWEET1a--GUS in T~2~ plants may be the result of an unknown silencing effect, occurring in the tomato Micro-tom cultivar ([@CIT0037]). In flowers derived from T~2~ plants, distinct staining was obvious in stamens ([Fig. 2E, F](#F2){ref-type="fig"}). To determine whether the expression of *Sl*SWEET1a--GUS fusion coincides with sugar unloading, we exposed a nearby mature terminal leaflet to CFDA that, on accumulation in the phloem, is de-esterfied into the membrane-impermeant and fluorescent carboxyfluorescein (CF), to trace carbohydrate unloading in developing leaves ([@CIT0054]). After 3 h of CF loading in the nearby mature leaves, fluorescence was clearly observed in the first- (midrib), second-, and third-order veins in a young leaflet (\<2 cm) ([Fig. 2G, H](#F2){ref-type="fig"}), but not in mature leaves ([Supplementary Fig. S3B](#sup1){ref-type="supplementary-material"}). The overall fluorescence intensities exhibited a high to low gradient from leaflet base to tip, indicating that the highest sugar unloading activity was located in the base of the young leaflet. Interestingly, relatively higher mRNA levels of *SlSWEET1a* were measured in the leaflet base compared with the middle and tip that had reached 80% of their full size ([Fig. 2I](#F2){ref-type="fig"}) and probably undergone sink--source transition ([@CIT0054]).

![Tissue-specific expression of *Sl*SWEET1a proteins. (A--F) Histochemical staining of GUS activity in transgenic tomato plants expressing *Sl*SWEET1a--GUS fusion proteins driven by the *SlSWEET1a* native promoter. Independent transgenic plants were grown in soil for \>4 weeks, and selected organs were harvested for staining for 16 h in (A, B) young leaves and (C, D) mature leaves, and 9 h for (E, F) flowers. Leaves and flowers were harvested from T~1~ and T~2~ plants, respectively. Representative terminal leaflets are shown in (A--D). Images in (B), (D), and (F) are magnified images of (A), (C), (E), respectively. (G and H) Fluorescence images of (H) CF dye unloaded into the terminal leaflet of a young leaf (G). The same pattern had been observed in three independent plants. (I)Transcript levels (by qRT-PCR) of *SlSWEET1a* in terminal leaflets of tomato young leaves. Relative expression levels by normalizing to an internal control, *SlActin7*, is shown. B, M, and T indicated samples harvested from the leaflet base, middle, and tip, as shown in (G). Results are means ±SE from four independent biological repeats. Significant differences from expression in the leaflet base (B) were determined by Student's *t*-test and indicated by one (*P*\<0.05) or two (*P*\<0.01) asterisks. Scale bars are 1 mm in (B), (D), and (F); 2 mm in (A), (C), (E), (G), and (H).](erz154f0002){#F2}

Plasma membrane-specific localization of *Sl*SWEET1a proteins {#s16}
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To observe the subcellular localization of *Sl*SWEET1a proteins at a single-cell resolution, the C-terminal *Sl*SWEET1a--GFP was transiently expressed in isolated Arabidopsis mesophyll protoplasts. Green fluorescence derived from *Sl*SWEET1a--GFP enclosed cytosolic chloroplasts indicated by red autofluorescence, and largely overlapped with red fluorescence derived from the plasma membrane dye, FM4-64 ([Fig. 3A--C](#F3){ref-type="fig"}; [@CIT0080]). Furthermore, when Arabidopsis protoplasts were co-transformed with *Sl*SWEET1a--GFP and a plasma membrane marker, AtPIP2--RFP, based on path-analysis of fluorescence intensities, there was co-localization of green and red fluorescence ([Fig. 3D--H](#F3){ref-type="fig"}). Similarly, plasma membrane-specific localization of *Sl*SWEET1--GFP proteins was present in isolated tobacco BY2 cells ([Supplementary Fig. S4A](#sup1){ref-type="supplementary-material"}). However, N-terminal fusion of GFP to *Sl*SWEET1a resulted in mislocalization and aggregation of the fusion proteins in the cytosol ([Supplementary Fig. S4B](#sup1){ref-type="supplementary-material"}), suggesting the localization signal was at the N-terminus of *Sl*SWEET1a.

![*Sl*SWEET1a proteins co-localized with plasma membrane proteins. (A--C) The *Sl*SWEET1a--GFP fusion proteins were transiently expressed in Arabidopsis mesophyll protoplasts driven by the 35S promoter. Fluorescence derived from (A) *Sl*SWEET1a--GFP fusion proteins, (B) FM4-64 staining, and (C) merged signals from the same focal plane are shown. (D--F) Fluorescence images of mesophyll protoplast expressing both *Sl*SWEET1a--GFP fusion proteins and the Arabidopsis membrane marker, AtPIP2A--RFP fusion proteins, are shown. (G and H) Intensity analysis of the indicated line path in the protoplast shown in (F). Scale bar=10 µm.](erz154f0003){#F3}

*Sl*SWEET1a activity complements glucose uptake deficiency {#s17}
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Based on genome structure analysis, the four clade I genes, *SlSWEET1a*, *1b*, *1c*, and *1d* were clustered in a tandem repeat, implying that these clade I homologs could have been derived from gene duplication ([@CIT0062]). Furthermore, due to high identity (31--81%) between amino acid sequences of clade I *SlSWEET* genes ([Supplementary Fig. S5](#sup1){ref-type="supplementary-material"}), perhaps all clade I members have similar transport properties. Therefore, cDNAs of clade I *SlSWEET* genes, 10 genes in total, were isolated and expressed in the baker's yeast (*Saccharomyces cerevisiae*) mutant YSL2-1, which lacks all endogenous hexose transporters but expresses a cytosolic invertase and can grow only on medium containing maltose, but not monosaccharides or sucrose ([@CIT0028]). Expressing the yeast hexose transporter HXT5 allows YSL2-1 to grow on media containing glucose ([Fig. 4A](#F4){ref-type="fig"}). Under acidic growth conditions (pH 5), expression of only *Sl*SWEET1a, 1b, and 3 complemented growth deficiency of YSL2-1 on glucose- and galactose-containing media. However, there was no growth on media with fructose, mannose, or sucrose ([Fig. 4A](#F4){ref-type="fig"}; [Supplementary Fig. S6](#sup1){ref-type="supplementary-material"}). There were similar trends on medium buffered at neutral pH (pH 7), suggesting that the transport activity of clade I *Sl*SWEETs was not dependent on a pH gradient.

![Transport activity of *Sl*SWEET1a in yeast. (A) Complementary growth assay in the YSL2-1 yeast mutant line. Yeast transformants expressing clade I *SlSWEET* genes, yeast hexose transporter (Hxt5), or the empty vector (Vector) were cultured on solid media supplemented with 2% maltose, 2% glucose, or 1% maltose together with 0.2% 2-deoxyglucose (2-DG) under pH 5 and 7 conditions. Images were taken after incubation at 30 °C for 4--6 d. (B) Time-dependent \[^14^C\]glucose uptake activity was measured in YSL2-1 yeast cells expressing *SlSWEET1a* or the empty vector. Result are means ±SE (*n*=3). Significant differences from cells expressing the empty vector were determined. (C) Kinetics of \[^14^C\]glucose uptake activity of *Sl*SWEET1a are shown. Results are means ±SE (*n*=4--5). (D) Competition binding ability of other sugars was measured by incubating cells with both \[^14^C\]glucose and 10-fold concentrations of the indicated sugars (Glu, glucose; Gal, galactose; Fru, fructose; Man, mannose; Suc, sucrose; Mal, maltose). Relative uptake rate was calculated compared with cells incubated with only 1 mM hot glucose (control). The background uptake of non-transformed cells (empty) was also shown. Results are means ±SE (*n*=5). Significant differences from the control were determined. (E) Effect of various pH conditions and treatment of NH~4~Cl on *Sl*SWEET1a uptake activity in EBY4000 cells. Yeast cells expressing the empty vector, *At*SWEET1, or *Sl*SWEET1a were analyzed. Results are means ±SE (*n*=3). Significant differences from the pH 5 condition were determined. All statistics were performed by Student's *t*-test and indicated by one (*P*\<0.05) or two (*P*\<0.01) asterisks.](erz154f0004){#F4}

YSL2-1 cells transformed with an empty vector grew on 2-deoxyglucose (2-DG) medium. In contrast, cells expressing HXT5 had no detectable growth due to uptake of 2-DG that is a poorly metabolized glucose analog and causes cell death in low amounts ([Fig. 4A](#F4){ref-type="fig"}; [@CIT0011]). Similarly, only yeast cells expressing *Sl*SWEET1a, 1b, 2b, and 3 had a significant 2-DG-dependent growth inhibition, especially on medium buffered at neutral pH (pH 7). Furthermore low uptake activity of *Sl*SWEET1c and 1e was only observed at neutral pH. These results indicated that *Sl*SWEET1a, as well as some homologs in clade I, probably functions as a glucose-specific passive transporter.

SWEET1 functions as a glucoses-specific low-affinity carrier {#s18}
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To confirm the glucose-specific transport activity of *Sl*SWEET1a, indicated by the growth assay in YSL2-1 yeast cells ([Fig. 4A](#F4){ref-type="fig"}), a time-dependent glucose uptake assay was done using \[^14^C\]glucose in YSL2-1 yeast cells expressing *Sl*SWEET1a. Radioactivity had accumulated in transformed yeast cells after 5 min, with a linear increase until 30 min, whereas there was limited glucose uptake in cells expressing the empty vector ([Fig. 4B](#F4){ref-type="fig"}). The kinetics of the glucose transport activity of *Sl*SWEET1a were determined by exposure to glucose concentrations of 0.1--400 mM for 10 min. Significant uptake of \[^14^C\]glucose occurred with exposure to 0.1 mM, with saturation of glucose uptake at a concentration close to 200 mM ([Fig. 4C](#F4){ref-type="fig"}). Therefore, the estimated *K*~m~ and *V*~max~ of *Sl*SWEET1a activities for glucose were 119.3 mM and 28.9 nmol 10^7^ cells^−1^ h^−1^, respectively, indicating a low-affinity transport mode.

To address further the substrate specificity of *Sl*SWEET1a, a 10-fold excess of unlabeled sugars was used to compete with \[^14^C\]glucose. Only a 10-fold excess of unlabeled glucose, but no other hexoses or sucrose, competed the binding and significantly reduced the amount of \[^14^C\]glucose imported ([Fig 4D](#F4){ref-type="fig"}). The lower uptake rate to \[^14^C\]glucose under 10-fold higher maltose was attributed to background uptake activity in the YSL2-1 yeast strain ([@CIT0014]). The specific competition ability of unlabeled glucose was also measured by treatment with a 30-fold excess of sugar substrates ([Supplementary Fig. S7](#sup1){ref-type="supplementary-material"}). To verify whether *Sl*SWEET1a proteins functioned as other known SWEET transporters, the EBY4000 yeast strain was used. Similar to *At*SWEET1 ([@CIT0014]), glucose uptake activity of *Sl*SWEET1a was not repressed by alkaline pH ([Fig. 4E](#F4){ref-type="fig"}) and only slightly decreased by NH~4~Cl (10 mM) ([Fig. 4E](#F4){ref-type="fig"}), perhaps due to indirect toxicity.

*Sl*SWEET1a contributes to sugar distribution to sink leaves {#s19}
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To explore the physiological role of *Sl*SWEET1a in leaves, VIGS was used to transiently silence expression of *SlSWEET1a* in leaves. The efficiency of VIGS was confirmed by expressing VIGS-*Sl*PDS in parallel plants with leaf etiolation due to silenced expression of the phytoene desaturase gene ([Supplementary Fig. S8](#sup1){ref-type="supplementary-material"}). Samples collected from non-silenced and control plants (injected with infiltration buffers only, Control/NS) were compared with silenced plants in which *SlSWEET1a* expression in mature leaves was reduced by 50% ([Fig. 5A](#F5){ref-type="fig"}). In silenced young leaves (VIGS-YL), glucose contents were significantly reduced (\>50%) compared with control leaves ([Fig. 5B](#F5){ref-type="fig"}). The same reduction was also observed for fructose, but not for sucrose. In contrast, glucose and fructose contents in silenced mature leaves were increased \>2-fold compared with controls ([Fig. 5C](#F5){ref-type="fig"}).

![Effect of *Sl*SWEET1a deficiency on sugar allocation ability in tomato leaves. (A) Gene expression of *SlSWEET1a* in matures leaves of tomato plants injected with buffer (Control) or *Agrobacterium* cells containing the VIGS--*Sl*SWEET1a construct (non-silencing and silencing). Total RNA were isolated and cDNA was used for qRT-PCR with gene-specific primers. Relative expression by normalizing to an internal control, *SlActin7*. Numbers indicate fold changes of *SlSWEET1a* expression compared with control leaves. Results are means ±SE (*n*=5--11). (B and C) Sugar contents in young (B) and mature (C) leaves of tomato plants subjected to the VIGS-*Sl*SWEET1a construct. Two samples from control plants and non-silenced plants in (A) were used as the control/NS for comparison. Results are means ±SE (*n*=3--4). Significant differences from the control (A) or control/NS leaves (B and C) were determined by Student's *t*-test and indicated by one (*P*\<0.05) or two (*P*\<0.01) asterisks.](erz154f0005){#F5}

Discussion {#s20}
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For decades, possible membrane carriers involved in apoplasmic sugar unloading in developing leaves have been the subject of debate. In our study, distinct high expression of *SlSWEET1a* was detected (qRT-PCR) in young leaves of vegetative tomato plants ([Fig. 1](#F1){ref-type="fig"}), suggesting that the *Sl*SWEET1a proteins may mainly function in young leaves compared with source (mature) leaves. In addition, expression of a whole-gene *Sl*SWEET1a--GUS translational fusion further demonstrated that *Sl*SWEET1a proteins were co-localized in leaf vein tissues ([Fig. 2](#F2){ref-type="fig"}; [Supplementary Fig. S3](#sup1){ref-type="supplementary-material"}). The GUS staining was most evident in the first- to the third-order veins, representing major sites for unloading of photoassimilates from the transport phloem in both tomato sink leaves ([Fig. 2G](#F2){ref-type="fig"}, H) and developing tobacco leaves ([@CIT0074]; [@CIT0054]). Moreover, the carbohydrate unloading activity in a sink leaflet co-occurred with relatively high levels of *SlSWEET1a* transcripts in the leaflet base, compared with the tip ([Fig. 2G--I](#F2){ref-type="fig"}), which is the first region to switch to sugar export during sink/source transition ([@CIT0074]; [@CIT0054]). Based on these results, we suspect that *Sl*SWEET1a probably functions in mediating sugar flows along the phloem unloading pathway in sink leaves. The phloem unit is composed of SE--CC complexes and phloem parenchyma, with central control of sugar loading, transport, and unloading in plants ([@CIT0047]; [@CIT0039]). Before post-phloem unloading to neighboring sink cells, sucrose needs to be exported out of SE--CC complexes and into phloem parenchyma ([@CIT0050]). Sucrose movement between SEs and CCs occurs mostly symplasmically ([@CIT0074]; [@CIT0036]; [@CIT0050]; [@CIT0039]). In contrast, the sucrose efflux between CCs and phloem parenchyma can be mediated in parallel via sym- and apoplasmic routes (with the latter requiring sugar-transporting proteins, such as *Sl*SWEET1a).

To confer apoplasmic sugar unloading to sink leaves, the plasma membrane would be the primary site to control sugar fluxes. Co-localization of *Sl*SWEET1a--GFP fusion protein with a plasma membrane protein, namely *At*PIP2--RFP ([@CIT0046]), at single-cell resolution was evidence that *Sl*SWEET1a was located at the plasma membrane ([Fig. 3](#F3){ref-type="fig"}; [Supplementary Fig. S4](#sup1){ref-type="supplementary-material"}). The specific subcellular localization of *Sl*SWEET1a in single cells was consistent with transient expression of *Sl*SWEET1a--GFP in leaves of tobacco and Arabidopsis ([@CIT0062]). All major sugar loaders, SUCs/SUTs and STPs, are located in the plasma membrane ([@CIT0029]; [@CIT0002]; [@CIT0017]; [@CIT0003]; [@CIT0051]). In contrast, SWEETs expressed in sink organs were present in plasma or vacuolar membranes ([@CIT0014]; [@CIT0010]; [@CIT0012]; [@CIT0011]). Similar to key SWEET exporters to supply sugars in reproductive organs, *Sl*SWEET1a locates at the correct cellular site, the plasma membrane, to be involved in sugar unloading. Taken together, dominant accumulation of *Sl*SWEET1a proteins in the plasma membrane of vein tissues in young leaves supported its putative role to mediate sugar fluxes across the apoplasm between CCs and phloem parenchyma during sugar unloading to sink leaves.

Although strong *SlSWEET1a* RNA accumulation also occurred in flowers during the reproductive stage, most *Sl*SWEET1a protein accumulated in stamens ([Fig. 2E](#F2){ref-type="fig"}, F), suggesting that *Sl*SWEET1a is probably not involved in phloem sugar unloading during the reproductive stage. Furthermore, *AtSWEET8* (also named *AtRPG1*) has been demonstrated to be strongly expressed in Arabidopsis pollen, possibly to mediate import of apoplasmic hexoses required for microspore development ([@CIT0027]). Therefore, whether *Sl*SWEET1a participates in post-phloem unloading required for microsporogenesis in flowers requires further investigation. Interestingly, induced expression of *SlSWEET1a* during fruit maturation implies that *Sl*SWEET1a is involved in sugar homeostasis during fruit maturation. This conclusion correlates with the previous observation that variation of *SlSWEET1a* expression correlates with hexose composition of ripening tomato fruits ([@CIT0062]). Taken together, these results suggest that *Sl*SWEET1a probably has multiple physiological functions, depending on tissue localization and developmental stages.

Transport analyses on yeast cells using complementary growth assays or a radiotracer confirmed that *Sl*SWEET1a was a glucose-specific transporter, consistent with its closest Arabidopsis homolog, *At*SWEET1 ([Fig. 4](#F4){ref-type="fig"}; [Supplementary Fig. S6, S7](#sup1){ref-type="supplementary-material"}; [@CIT0014]) and previous speculations ([@CIT0062]). In contrast, many hexose transporters, such as TMT/TST (tonoplast monosaccharide transporter/tonoplast sugar transporter) or some STPs, have a broad substrate range ([@CIT0066]). For example, Arabidopsis *At*TST1 transports glucose, fructose, and sucrose ([@CIT0082]; [@CIT0061]). Sugar beet *Bv*TST1 also transports both glucose and sucrose ([@CIT0035]), whereas *Cs*HT1 (hexose transporter from *Cucumis sativus*) transports glucose, galactose, mannose, and xylose ([@CIT0017]). Functional redundancy between MSTs would be consistent with MST proteins not having a confirmed physiological role, despite their dominant expression in heterotrophic sink tissues ([@CIT0064]; [@CIT0066]; [@CIT0084]). Thus, high specificity of *Sl*SWEET1a to glucose reflects its putative importance in hexose transport during sugar unloading in young leaves.

In contrast to most plasma membrane-localized MSTs exhibiting a low *K*~m~ (2--10 mM) ([@CIT0008]; [@CIT0009]; [@CIT0038]; [@CIT0052]), *Sl*SWEET1a import activity exhibited a relatively high *K*~m~ (119 mM) for glucose ([Fig. 4C](#F4){ref-type="fig"}). Although this low-affinity feature may be due to a high turnover rate of *Sl*SWEET1a proteins ([@CIT0013]), it also highlights the possibility that *Sl*SWEET1a actively operates under high physiological concentrations of apoplasmic glucose. Due to significantly higher expression and activities of cell wall invertases observed in young leaves compared with mature leaves ([@CIT0063]; [@CIT0033]), sucrose unloaded to the leaf apoplasm could be hydrolyzed and glucose taken up via the hexose carrier. These observations further support the need for a low-affinity glucose importer, such as *Sl*SWEET1a, operating along the initial sugar unloading path from CCs to parenchyma cells in developing leaves.

Moreover, *Sl*SWEET1a was a proton-independent bidirectional uniporter ([Fig. 4A](#F4){ref-type="fig"}, E), in agreement with all other *At*SWEET members ([@CIT0016]; [@CIT0028]). In that case, a passive glucose carrier, such as *Sl*SWEET1a, would possess a more efficient transport system in response to instant sugar fluxes to assist continuous sugar unloading without the delay for energy biosynthesis. This was nicely in line with passive sugar fluxes observed in young leaves, where the sucrose unloading is insensitive to anoxia and PCMBS, a metabolite inhibitor ([@CIT0060]; [@CIT0074]; [@CIT0022]). Furthermore, proton ATPase is not expressed in the protophloem of growing sinks, such as root tips and expanding region of leaves ([@CIT0070]; [@CIT0073]), indicating a lack of energized transport. Taken together, all transport features of *Sl*SWEET1a supported its putative role in participating in initial sugar unloading from SE--CC complexes to phloem parenchyma in sink leaves.

Our physiological evidence supported the assertion that *Sl*SWEET1a import activity regulated hexose accumulation in young leaves. During the vegetative growth stage of tomato plants, suppressed expression of *Sl*SWEET1a by VIGS significantly reduced accumulation of glucose and fructose in young leaves, with concurrent increased concentrations of these sugars in source leaves ([Fig. 5](#F5){ref-type="fig"}). Since sucrose is the dominant type of phloem-transported sugar ([@CIT0071]; [@CIT0042]), it follows that apoplasmic glucose in sink leaves is mainly derived from hydrolysis of sucrose unloaded into the sink leaf apoplasm. Based on its low affinity, vein localization, and glucose-specific uniport feature, our results were best explained by the model that some *Sl*SWEET1a on the plasma membrane of phloem parenchyma significantly contributes to hexose import in sink leaves. The high glucose concentrations derived from sucrose hydrolysis during phloem unloading drive the import activity of *Sl*SWEET1a to enable glucose uptake. Once in the cytosol, glucose can be quickly distributed to sink cells via symplasmic connections ([@CIT0060]), or it can be compartmentalized into corresponding vacuoles ([@CIT0051]). The finding that knock out of *Sl*SWEET1a resulted in hexose concentrations being reduced in sink leaves is consistent with *Sl*SWEET1a playing a key role in the phloem unloading of hexoses. Similar conclusions have been reached for knock down of the hexose symporter that drives hexose uptake from the apoplasm into storage parenchyma cells of developing tomato fruit ([@CIT0045]) and of *Zm*SWEET4, located on the membrane of basal endosperm transfer cells, mediating hexose uptake into the endosperm of developing maize seeds ([@CIT0068]).

These results also concur nicely with the report that reduced sink strength by removing active sinks, such as developing leaves and fruits, stimulated sucrose accumulation in mature leaves of a citrus tree ([@CIT0031]), and with the observation that removal of young axillary leaves increased the size of mature leaves and the number of nodes in tomato plants ([@CIT0019]). Furthermore, as observed in wheat grains, once sugar import into sink efflux was diminished by detaching the sink, sucrose efflux out of the SE--CC complex declined immediately ([@CIT0022]). As a consequence, the resulting high sucrose contents in the phloem will suppress expression of the phloem loader, SUC/SUT2 symporter ([@CIT0018]; [@CIT0076]), and thus reduce whole-plant sugar loading and lead to increased carbohydrates in sucrose mesophyll cells ([@CIT0076]). We inferred that the import activity of *Sl*SWEET1a may be an indicator for sink strength and is thus a novel element in whole-plant carbon partitioning from source leaves to sink leaves.

Summary {#s21}
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Herein, we uncovered important functions of the plasma membrane-located uniporter *Sl*SWEET1a in glucose uptake during apoplasmic phloem unloading to young leaves ([Supplementary Fig. S9](#sup1){ref-type="supplementary-material"}). We propose that in vegetative tomato plants, once sucrose is allocated to sink leaves via phloem pressure flow, it will passively exit out of the SE--CC complexes (symplasmically or apoplasmically). In the apoplasmic route, some sucrose may be exported by an unknown sucrose exporter and be directly taken up through an unidentified sucrose carrier, possibly a SUT member. Yet, a significant portion of sucrose is hydrolyzed to glucose and fructose by extracellular invertase in the apoplasm around phloem parenchyma, partially contributing to formation of a sucrose gradient from SE--CC complexes to facilitate efflux ([@CIT0048]). Then, *Sl*SWEET1a, partially located in the plasma membrane of unloading cells, such as phloem parenchyma cells, promotes glucose import to increase apoplasmic solute potential so as to maintain lower turgor pressure toward the SE--CC symplast for continuous sucrose unloading. The fructose-specific carrier remains to be identified. Further studies will be needed to dissect how *Sl*SWEET1a import activity is coordinated with sink strengths during transition of leaf development and transition from the vegetative stage to the reproductive stage. We also speculated that efficient unloading to sink leaves was important for whole-plant growth. Thus, engineering-efficient unloading of hexoses in phloem parenchyma, for example by increasing *Sl*SWEET1a import activity, coupling symport with higher source import capacity ([@CIT0001]), may be key for future crop enhancement.

Supplementary data {#s22}
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Supplementary data are available at *JXB* online.

Table S1. Primers used for qRT-PCR.

Table S2. Primers used to generate yeast-expressing constructs for clade I *Sl*SWEET genes.

Fig. S1. Images of the tomato organs analyzed.

Fig. S2. Phylogenetic tree of 31 tomato *SlSWEET* genes.

Fig. S3. Tissue-specific expression of *Sl*SWEET1a proteins in five T~1~ plants.

Fig. S4. Subcellular localization of *Sl*SWEET1a in tobacco and Arabidopsis protoplasts.

Fig. S5. Amino acid identity between clade I *Sl*SWEET genes.

Fig. S6. Transport activities of clade I *Sl*SWEETs for various sugars in yeast.

Fig. S7. Competition binding ability of sugars to *Sl*SWEET1a transport.

Fig. S8. Images of tomato plants subject to VIGS.

Fig. S9. Proposed model of *Sl*SWEET1a function.

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Click here for additional data file.

We thank Dr Yi-Min Chen at the National Cheng Kung University for his assistance in the GC-MS analysis. We acknowledge the confocal core facility at the National Cheng Kung University for fluorescence observations, and the transgenic plant core lab in Academia Sinica for generating transgenic tomato plants. This work was financially supported by grants from the Ministry of Science and Technology, Taiwan (MOST 105-2628-B-006-001-MY3) and from the DAAD/NSC PPP program (106-2911-I-006-506; 107-2911-I-006-506) to WJG.
