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Plant Science 255 (2017) 40–50

Contents lists available at ScienceDirect

Plant Science

j ourna l ho me pa g e: www.elsev ier .com/ locate /p lantsc i

ucrose phloem unloading follows an apoplastic pathway with highucrose synthase in Actinidia fruit

heng Chen1, Yulin Yuan1, Chen Zhang, Huixia Li, Fengwang Ma, Mingjun Li ∗

tate Key Laboratory of Crop Stress Biology in Arid Areas/College of Horticulture, Northwest A&F University, Yangling 712100, Shaanxi, China

r t i c l e i n f o

rticle history:eceived 1 June 2016eceived in revised form4 November 2016ccepted 25 November 2016vailable online 25 November 2016

eywords:iwifruit

a b s t r a c t

Phloem unloading plays a pivotal role in photoassimilate partitioning and the accumulation of sugars insink organs, e.g. fruit. Here, we investigated the pathway of sucrose unloading in kiwifruit (Actinidia delici-asa cv. Qinmei) using a combination of electron microscopy, transport of the phloem-mobile symplastictracer carboxyfluorescein and enzyme activity and gene expression assays. Our structural investiga-tion revealed that the sieve element-companion cell complex of bundles feeding the fruit flesh wassymplastically isolated from its surrounding parenchyma cells throughout fruit development, whereasnumerous plasmodesmata were present between the phloem parenchyma cells. Consistent with this,carboxyfluorescein unloading showed that the dye remained confined in the phloem strands during fruit

nloadingucroseucrose synthaseell wall invertase

development. The activities and expression of cell wall acid invertase in fruit flesh were lower than thoseof other enzymes that catalyze sucrose dissociation. However, sucrose synthase showed higher enzymeactivities and mRNA expression in fruit flesh compared with other detected enzymes. These results implythat, in kiwifruit flesh, phloem unloading of sucrose is predominantly an apoplastic pathway during fruitdevelopment, and that sucrose synthase is a key enzyme for sucrose post-unloading pathways.

© 2016 Elsevier Ireland Ltd. All rights reserved.

. Introduction

In the majority of plants, sucrose (Suc) is produced in, andranslocated from, photosynthetically active leaves (source, load-ng) to support non-photosynthetic tissues (sinks, unloading), suchs developing seeds, fruits, and tubers. Phloem unloading includesransfer across the sieve element-companion cell (SE/CC) com-lex boundary (SE unloading) and subsequent transport through

diverse range of sink parenchyma cells (PCs) (post-phloem trans-ort) [1,2]. It is now well accepted that phloem unloading playsn important role in the partitioning of photoassimilates, therebyetermining to a large extent crop output and quality [2]. Moreover,arbohydrate accumulation in the fruit relies upon the phloem net-ork to transport sugars from the source to sink organs. Phloemnloading is generally considered to play a key role in the parti-ioning of photoassimilate [2–4]. For providing neoteric methods

o improve fruit quality and increase food security, it is critical tonderstand the pathway of Suc transfer from the phloem to theruit. Suc in the phloem moving into the fruit is regarded as the

∗ Corresponding author at: College of Horticulture, Northwest A&F University,angling 712100, Shaanxi, China.

E-mail address: limingjun@nwsuaf.edu.cn (M. Li).1 These authors contributed equally to this work.

ttp://dx.doi.org/10.1016/j.plantsci.2016.11.011168-9452/© 2016 Elsevier Ireland Ltd. All rights reserved.

transfer of Suc from SE-CC complexes to sites of utilization/storagein the recipient sink cells [1,2]. There are two routes of Suc unload-ing: the symplastic pathway and the apoplastic pathway. Sucroseunloading can utilize the symplastic pathway, the apoplastic path-way, or both. If Suc unloads via the symplastic pathway, it movesfrom the SE-CC complex directly to the surrounding PCs via plas-modesmata, driven by a concentration gradient. However, if Sucunloads via the apoplastic pathway, it is transported directly fromthe SE-CC complexes into the apoplastic space between cells inde-pendently of plasmodesmata. In the apoplastic space, Suc can betransported by sucrose transporters (SUCs or SUTs) into the PCs,meanwhile Suc also can be converted into fructose (Fru) and glu-cose (Glc) by cell wall invertases (CWINVs), and then be assimilatedby hexose transporters (HTs) into the PCs. Different plants can uti-lize distinct mechanisms to transport Suc from the phloem sievetubes to PCs in fruit [6]. For example, apple and pear fruit usean apoplastic phloem unloading pathway over the course of fruitdevelopment [7,8]. In contrast, for grape berries and tomatoes, Sucunloading shifts from a symplastic to an apoplastic pathway [9,10].

Irrespective of the pathway, Suc in the sink PCs is transferredinto vacuoles for storage or enzymatically degraded into Fru and

Glc/UDP-Glc to support the growth of the sinks, which affectsthe ability of Suc post-unloading. In the PCs, Suc imported intothe cytoplasm must rapidly be cleaved to decrease the chemical

dx.doi.org/10.1016/j.plantsci.2016.11.011http://www.sciencedirect.com/science/journal/01689452http://www.elsevier.com/locate/plantscihttp://crossmark.crossref.org/dialog/?doi=10.1016/j.plantsci.2016.11.011&domain=pdfmailto:limingjun@nwsuaf.edu.cndx.doi.org/10.1016/j.plantsci.2016.11.011

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radients in the sink cells, thereby promoting unloading. Hence,ucrose-cleaving enzymes are important factors for determiningink strength of fruit. Two different types of enzymes are respon-ible for such cleavage: sucrose synthase (EC 2.4.1.13) (SUSY) andnvertases (EC 3.2.1.26). SUSY catalyzes the reversible conversionf Suc to Fru and UDP-Glc. The activity of neutral invertase (NINV) isuch lower than that of SUSY in apple [11] and kiwifruit (Actinidia

eliciosa var. Hayward) [12], suggesting that SUSY is the key enzymeatalyzing the progress of Suc degradation in fruit. It has beeneported that the activity of SUSY is much more important thanhat of other enzymes (e.g. invertase) in many biological process,.g. sugar unloading [13], cellulose and callose synthesis [14,15]nd sink strength [16]. Therefore, it is important to understand thectivity and expression pattern of SUSY during fruit development.

Kiwifruit is an emerging commodity in international marketsue to its unique flavor and nutritional value (e.g., high contentf vitamin C, amino acids and minerals). As the health benefitsf kiwifruit become better appreciated by consumers, demands on the rise for cultivars with good size, texture, flavor andutritional value. All of these characteristics are related to thebility of carbohydrate unloading in fruit. Although Suc, myo-nositol and planteose are the major soluble carbohydrates in theeaves of kiwi [17], Suc is the main accumulated photosynthateroduced by the crowns in kiwifruit [18]. During kiwifruit devel-pment, there are three stages characterized by the dominatingetabolism: (1) cell division, (2) starch accumulation and (3) fruitaturation [19]. These stages have been studied in terms of non-

tructural carbohydrate content [20,21]. Carbohydrate metabolismuring development, ripening and the post-harvest period has beenescribed in detail, as have the dynamics of the related enzymectivities [12,22,23], but the unloading pathway of imported carbonnd sink strength have received limited attention.

In this work, we conducted a study of the unloading of carbo-ydrate that is imported from the source leaves into kiwifruit. Ourbjective was to determine the unloading pathway during kiwifruitevelopment and to identify the activities, and expression patternf the enzymes responsible for metabolizing the imported Suc.

. Material and methods

.1. Plant materials

Nine-year-old Actinidia deliciosa var. ‘Qinmei’ kiwi vines weresed in this study. The vines were grown under natural condi-ions in an experimental orchard at the Horticultural Experimentaltation of Northwest A&F University, Yangling, Shaanxi, China. At4 (early developmental stage), 75 (middle developmental stage)nd 135 (late developmental stage) days after blooming (DAB), fivendependent fruit samples were obtained from the south side ofhe tree canopy between 4:00 PM and 6:00 PM under full sun expo-ure; each replicate contained eight fruit harvested from four trees.he samples were weighed immediately, cut into small pieces, androzen in liquid nitrogen. To compare the expression patterns ofelevant genes in the source and sink tissues, mature leaves andhoot tips were obtained at 44 DAB from the vines. All of the frozenamples noted above were stored at −80 ◦C until use.

.2. Tissue preparation for ultrastructural observation

The method of tissue preparation for ultrastructural observa-ion was adapted from Zhang et al. [7] with some minor changes.

ruit were harvested from the south side of the trees. The fruitas cut transversely and the vascular bundle zone was cut into

mall columns (approximately 1 × 2 × 3 mm3), which were fixedmmediately with 4% (v/v) glutaraldehyde in 100 mM precooled

e 255 (2017) 40–50 41

phosphate buffer (pH 7.0). The samples were subjected to a vac-uum for 0.5 h and then incubated at 4 ◦C for at least 6 h. Afteran extensive rinse with precooled phosphate buffer (pH 7.0), thesamples were postfixed in 1% (w/v) OsO4 for 2–4 h at room tem-perature. Following another extensive rinse with the precooledphosphate buffer (pH 7.0), the samples were dehydrated using agraded ethanol series (30–100%) at room temperature. The detailsof the process were as follows: 30% ethanol for 10 min, 50% ethanolfor 10 min, 70% ethanol for 10 min, 80% ethanol for 10 min, 90%ethanol for 10 min and 100% ethanol for 20 min. Next, the sampleswere infiltrated with a graded LR White resin at room temperature.The details of the process was as follows: 3:1 ethanol and LR Whiteresin for 3 h, 1:1 ethanol and LR White resin for 6 h, 1:3 ethanoland LR White resin for 12 h, and 100% LR White resin for 24 h.Then, the tubes containing the samples were placed in capsulesand incubated at 55 ◦C for 2–3 days. The capsules were placed ina dessicator until use. Ultrathin sections (approximately 60–80 nmin thickness) were obtained using an Ultracut microtome (RMC,USA) and mounted on 100-mesh copper grids for ultrastructuralobservation. At least five observations were made for each ultrathinsection. Plasmodesmata were counted at all cell interfaces, includ-ing the interfaces between SE and CC, SE and PP, CC and PP, andPP and PP in each selected field. The results of the plasmodesmalcounting were given as the number of plasmodesmata per micronof specific cell–cell interface length on transversal section, whichis referred to as plasmodesmal density (no. of plasmodesmata �m−1).

2.3. Carboxyfluorescein diacetate labeling

The method of membrane-permeable 5(6)carboxyfluoresceindiacetate (CFDA) loading into the fruit through the pedicel wasadapted from Zhang et al. [7] with some modifications. First, theCFDA was dissolved in DMSO (50 mg mL−1) as the master stockand stored at −20 ◦C in a light-resistant container until use. Fora working solution, it was diluted to 1 mg mL−1 with 0.1 M phos-phate buffer (pH 7.0). Approximately 100 �l of CFDA solution wasinjected into the cotton lines around the outer phloem region of thepedicel, which was gently scratched with a saw without woundingthe xylem, and then silver paper was used to surround the cotton.After allowing the plant to transport the dye for 72 h, the fruit wasremoved and immediately taken into the lab at 4 ◦C for sectioningand microscopy. Free hand sections, including transverse and lon-gitudinal sections, were obtained; the microscopic examinationswere conducted using a fluorescence microscope (Olympus Corpo-ration, Tokyo, Japan) under a blue light (488 nm). Eight fruit wereexamined on each experimental date.

2.4. Measurements of soluble sugars

Soluble sugars were obtained and derivatized as describedby Wang et al. [24]. Briefly, the samples (0.1 g) were extractedin 1.4 ml of 75% methanol, with ribitol added as the internalstandard. After the non-polar metabolites were fractionated intochloroform, 5 �l of the polar phase was transferred into 2.0 mlEppendorf vials to measure the metabolites (Suc, Fru, Glc, and myo-inositol) in each sample. These were dried under vacuum withoutheating and then derivatized sequentially with methoxaminehydrochloride and N-methyl-N-trimethylsilyl-trifluoroacetamide[25]. The metabolites were analyzed using a Shimadzu GC/MS-2010SE (Shimadzu Corporation, Tokyo, Japan). These metaboliteswere identified by comparing their fragmentation patterns with

those from a mass spectral library generated on our GC/MS sys-tem, and from an annotated quadrupole GC/MS spectral librarydownloaded from the Golm Metabolome Database (http://csbdb.mpimp-golm. mpg.de/csbdb/gmd/msri/gmd msri.html). Quantifi-

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ations were based on standard curves generated for eachetabolite and internal standard.

.5. Detection of sugar in the phloem exudate

Phloem exudate was collected from the petiole of mature sourceeaves or pedicel of 40 DAB fruit using an adaptation of the methodeveloped by Liu et al. [26]. Following excision of the organ, a por-ion of the petiole or pedicel (about 5 mm) was removed underater, the sample was rinsed and the cut end was transferred

o 1.5 ml Eppendorf tubes with 500 �l 15 mM EDTA (pH 7.0). Theubes were then placed in a closed chamber for 5 h under low lightonditions and high humidity to reduce transpiration. For controlamples run in parallel, the petiole or pedicel was incubated in

mM CaCl2 (pH 7.0) to induce callose gelation and reduce exu-ation [27]. At the end of the incubation, the tubes with facilitatedxudation were immediately frozen in liquid nitrogen and storedn a freezer at −20 ◦C. To detect sugar by the GC/MS method, 200 �lxudate was used, as described above.

.6. Enzyme analysis

The method to extract SUSY, CWINV, NINV, and vAINV wasimilar to that used by Moscatello et al. [12]. One gram of frozenesh powder was homogenized in 4 ml extraction buffer contain-

ng 200 mM Hepes-KOH (pH 8.0), 5 mM MgCl2, 2 mM EDTA, 2 mMGTA, 2.5 mM DTT, 2 mM Benzamidine, 2 mM �-aminocaproic acid,.1 mM AEBSF, 1% BSA (w/v), 2% glycerol (v/v), 0.05% triton-X 100v/v) and 2% PVP (w/v). The extract was clarified by centrifugationt 16,000 × g for 20 min at 4 ◦C, immediately desalted in a Sephadex25 PD-10 column (GE Healthcare, UK) and then equilibrated with

he extraction buffer at a concentration of 200 mM Hepes-KOH (pH.4) but without triton-X 100, BSA or PVP. For the CWINV, the pre-ipitate was resuspended with extraction buffer containing 1 MaCl. The protein on the cell wall was solubilized via incubationt 4 ◦C overnight. Then, the extract was centrifuged and desalted asbove.

Activity of SUSY in the cleavage direction was determined asescribed by Dancer et al. [28] with some modifications. Thenzyme extract (20 �l) was incubated at 27 ◦C for 20 min in 100 �lfinal volume) of assay medium containing 20 mM Hepes-KOH (pH.0), 100 mM Suc and 4 mM UDP. The reaction was stopped by heat-

ng in boiling water for 3 min. The blanks contained the same assayixture, but the extract was boiled for 5 min before being mixed.

he amount of Fru produced from Suc was determined by GC/MS.For CWINV and vAINV, extracts were incubated for 40 min at

7 ◦C in a 200 �l assay mixture containing 100 mM phosphate-itrate buffer (pH 4.8), 100 mM Suc, and 50 �l of the desaltedxtract. The assays were stopped by boiling for 3 min. The con-itions for NINV activity assay were the same as for CWINV andAINV, except that the assay mixture contained acetate-K3PO4 (pH.5) as the buffer. The blanks contained the same assay mixture, buthe extract was boiled for 5 min before being mixed. The amountf Fru produced from Suc was determined by GC/MS. Five �l of theeaction mixture was derivatized directly as described above andhen analyzed using GC/MS.

.7. Identification of candidate genes

We identified candidate genes (Table S1) by performinglastp analysis against a kiwifruit open reading frame translationataset (Kiwifruit Genome Database; http://bioinfo.bti.cornell.edu/

gi-bin/kiwi/blast.cgi) [29], using an Expect (E)-value thresholdf 1.00 × 10−4. Gene sequences were retrieved from the Kiwifruitenome Database. The corresponding sequences of candidateenes were then used in a BLAST search against a Kiwifruit EST

e 255 (2017) 40–50

database (National Center for Biotechnology Information; http://www.ncbi.nlm.nih.gov/) to confirm that each predicted gene wasexpressed in the kiwifruit transcriptome (score > 300 bp; iden-tity > 98%).

2.8. Transcript analysis

Total RNA was isolated from 0.5 g of leaves, shoot tips andfruit tissues as described by Chang et al. [30]. Genomic DNA wasremoved and CDNA was synthesized using PrimeScriptTMII ReverseTransciptase (Takara). Gene-specific primers were designed usingPrimer5 software. Primer specificity was determined by RT-PCRand Melt Curve analysis. PCR products were quantified on aLightCycler

®96 real-time PCR detection system (Roche) using

the LightCycler UltraSYBR Mixture (CWBIO). Elongation factor 1�(EF1˛) was used for normalization for target gene transcripts usingthe 2−�Ct method [31].

3. Results

3.1. Sugar accumulation in developing kiwi fruit

We determined the concentration of the major sugar during dif-ferent growth stages of kiwi fruit (Fig. 1, A, B). The growth rate ofcv. ‘Qinmei’ kiwifruit exhibited a double sigmoidal curve with twogrowth peaks. The growth rate was significantly faster between 15and 45 DAB than during the remaining time period; after 45 DABthere was a decreased growth rate until 75 DAB. Fig. 1B shows thechanges in carbohydrate concentration during fruit development;the Suc concentration increased rapidly from 15 to 30 DAB, fol-lowed by a drop at 75 DAB. Subsequently, Suc increased until thefruit ripening period. Fru concentration decreased from 15 to 30DAB and then slowly increased until a period of rapid increase nearmaturity after 105 DAB. We found that Glc also exhibited a rapidincrease toward harvest after 105 DAB. However, inositol concen-tration was always sustained at a very low level over the entireperiod of fruit development.

We determined the concentrations of Suc, Fru and Glc at 44-, 75-and 135-DAB in four parts of the fruit: the epicarp, outer pericarp,peripheral pericarp and central placenta [32] (Fig. 2). The concen-trations of Suc and Fru were the highest in the central placenta; at135 DAB, Suc was also at a higher level in the epicarp comparedwith those in other tissues. The central placenta had similar con-centrations of Fru at 75 and 135-DAB. At 44 DAB, Glc exhibited amuch lower concentration in the epicarp compared to other parts;at 75 DAB, Glc concentration in the peripheral pericarp exhibited adecrease. However, at 135 DAB, the Glc concentration in both theepicarp and peripheral pericarp increased to the level of the centralplacenta.

3.2. Carbohydrate with long-distance transport in the phloem

To identify the carbohydrate transported from the leaves to fruit(e.g., fruit), we analyzed the soluble carbohydrate in leaves andthe leaf petioles. Suc was the largest peak compared with Fru, Glcand inositol (Fig. S1, Table 1). When we collected exudates in thepresence of CaCl2 instead of EDTA, Suc was strongly reduced dueto the reduction in phloem exudation caused by callose gelation.However, concentrations of Fru, Glc and inositol did not exhibitdifferences between the EDTA and CaCl2 exudation buffers.

3.3. The SE-CC complex is symplastically isolated from the

surrounding PCs during fruit development

Kiwi fruit is a typical true fruit originating from a compound pis-til ovary [32]. The flesh portion derives from the ovary wall, which

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C. Chen et al. / Plant Science 255 (2017) 40–50 43

Fig. 1. Growth rate of kiwifruit. B, Sucrose,fructose,glucose and inositol concentrations during kiwifruit development.Values are means of three replicates ± SD.

Fig. 2. Sucrose, fructose and glucose concentrations in different tissues of kiwifruit, at 44 DAB, 75 DAB and 135 DAB. Values are means of three replicates ± SD.

Table 1Sucrose, fructose, glucose and myo-inositol concentrations of exudation from fruit stalk and leaf stalk. The fruit stalk and leaf stalk were treated with EDTA or CaCl2.

Carbohydrate (mg d−1 stalk−1) Sucrose Fructose Glucose Myo-Inositol

Fruit stalk-EDTA 0.729 ± 0.12 0.094 ± 0.05 0.238 ± 0.08 0.060 ± 0.01Fruit stalk-CaCl2 0.075 ± 0.02 0.175 ± 0.06 0.352 ± 0.11 0.174 ± 0.06Leaf stalk-EDTA 0.721 ± 0.15 0.359 ± 0.14 0.601 ± 0.13 0.037 ± 0.01Leaf stalk-CaCl2 0.029 ± 0.01 0.037 ± 0.01 0.377 ± 0.05 0.030 ± 0.01

Fig. 3. Longitudinal mid-section of kiwifruit (left) according to Guo et al. [32], showing the network of the vascular bundles, the loading site of CFDA into the axis of thepedicel (indicated by red arrows). The sampling site for structural observations (indicated by red circle with cross) was amplified at right, the violet- stained semithin sectionsof placental vascular bundles and ovary wall vascular bundles. CB: central placenta vascular bundle, LB: lateral branch bundle, PB: placental vascular bundle, OB: ovary wallvascular bundle, OT: ovule trace, SB: septum vascular bundle. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version ofthis article.)

44 C. Chen et al. / Plant Science 255 (2017) 40–50

Fig. 4. Connectivity of parenchyma cells and the conducting phloem in the peripheral and central bundles of kiwifruit. A, C, E, Numerous plasmodesmata present betweenPCs at the early (44 DAB), middle (75 DAB) and late (135 DAB) developmental stage respectively. B, A transverse section of the phloem in the central bundle at the earlyd the mb are sieC

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evelopmental stage. D, A transverse section of the phloem in the central bundle atundle at the late stage. Solid arrows indicate plasmodesmata, while broken arrows C, companion cell.

ontains the epicarp, outer pericarp, peripheral pericarp and centrallacenta. The developing flesh of kiwi fruit is fed with photoassim-

lates through the carpellary vascular bundles, which are dividednto vascular bundles distributed in distinct regions (Fig. 3). Sincehe placental vascular bundles and ovary wall vascular bundles play

critical role in providing nutrients [32], we examined both placen-al vascular bundles and ovary wall vascular bundles at the early (44

AB), middle (75 DAB) and late (135 DAB) stages of the fruit devel-pment. The violet-stained semi-thin sections of placental vascularundles and ovary wall vascular bundles are shown in Fig. 3. Weound that numerous plasmodesmata connect storage PCs (Fig. 4A,

iddle developmental stage. E, A transverse section of the phloem in the peripheralve pore and sieve-pore plasmodesma units. PC, parenchyma cell; SE, sieve element;

C and E; Table 2). Additionally, the plasmodesmata were enrichedat the interface between SE and CC during fruit development (Fig. 4,D). However, nearly no plasmodesmata were observed at the inter-face between SE and PCs, or between CC and PCs, throughout thedevelopment of the fruit (Fig. 4, Table 2), which resulted in sym-plastic isolation in phloem unloading domains.

3.4. Carboxyfluorescein is confined to the phloem strands

Carboxyfluorescein (CF) is used as a fluorescent marker ofphloem transport and symplastic phloem unloading [33]. CF

C. Chen et al. / Plant Science 255 (2017) 40–50 45

Fig. 5. Fluorescence microscope images of CF unloading during development of kiwifruit. Berries were allowed to transport CF for 72 h, and then were collected and sectionedby free hand for observations under a fluorescence microscope. A to C, Transverse and longitudinal sections of a kiwifruit at the early developmental stage(44 DAB + 72 h).D to F, Transverse and longitudinal sections of a kiwifruit at the middle developmental stDAB + 72 h).

Table 2Plasmodesmal densities between different cells at the early (44 DAB), middle (75DAB) and late (135 DAB) developmental stage of kiwifruit respectively.

Developmental stages SE/CC SE/PP CC/PC PC/PC

early 2.69 ± 0.27 – – 1.72 ± 0.12Middle 2.55 ± 0.36 – – 1.57 ± 0.28Late 2.29 ± 0.19 – – 1.27 ± 0.16

Unit of plasmodesmal densities, number of plasmodesmata �m−1. Each value is themean ± SD of 20 replicates at least. − means no plasmodesmata were observed. CC,companion cell; FP, flesh parenchyma cell; PP, phloem parenchyma cell; SE, sieveelement.

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fruit than other tissues. Additionally, SUT4 was expressed stronglyin mature leaves and 135-DAB fruit. Interestingly, we found thatSUT5 was expressed more in mature leaves and shoot tips than in

ehaves similarly to assimilate unloading as determined by autora-iography [4]. When loaded into cells, the membrane-permeable,onfluorescent CFDA is degraded into membrane-impermeableuorescent dye (CF), which is symplastically unloaded to the sinkells with the flow of assimilates. Fluorescence microscope imagesf CF movement in the fruit sampled 48 h after CFDA was suppliedo fruit pedicel are shown in Fig. 5. The fluorescent signal could beetected less than 4 h after the CFDA treatment, suggesting thathe CF had been unloaded into the fruit. A series of experimentsonducted during the growing season showed that CF was con-ned to the phloem strands along the phloem pathway in vascularundles without apparent diffusion to the surrounding tissues, atarly (44 DAB), middle (75 DAB) and late (135 DAB) stages of fruit

evelopment (Fig. 5).

age(75 DAB + 72 h). G to I, Longitudinal sections of a kiwifruit at the late stage(135

3.5. Activities of key enzymes in source and sink tissues

We determined the activity of Suc post-unloading-relatedenzymes SUSY, CWINV, vANIV and NINV in mature leaves, shootapex and fruit at three developmental stages (Fig. 6). The activityof SUSY was the highest in 44-DAB fruit; there was no clear dif-ference among other samples. On the other hand, the activity ofCWINV in fruit significantly increased during fruit development,and exceeded the activity of the other enzymes in the leaves andshoot apices. Both the vAINV and NINV activities exhibited a simi-lar pattern (i.e., decreased during fruit development and had highexpression in the shoot apex). Additionally, the activity of SUSY wasmuch higher than that of CWINV, vANIV and NINV in the leaves,shoot apex and fruit.

3.6. Expression of genes in source and sink tissues

We surveyed the gene transcription patterns in leaves, shoot tipsand whole fruit tissues at different developmental stages (Fig. 7).The expression level of SUT1 and SUT6 in mature leaves and 135-DAB fruit was nearly the same, and both expression levels wereincreased during fruit development. The expression level of SUT2 inthe leaves was much higher than in other tissues. On the other hand,the expression level of SUT3 was much higher in 75- and 135-DAB

fruit. The expression level of CWINV1 was seven times higher in

46 C. Chen et al. / Plant Science 255 (2017) 40–50

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ig. 6. Activities of key enzymes involved in sugar metabolism during kiwifruit denvertase; NINV: Neutral invertase; ML: Mature leaves; SA: Stem apex. Values are m

eaves than in shoot tips and ∼200 times higher than in 75-DABruit. At the same time, the expression level of INH1 was also highn leaves. However, INH1 exhibited a smaller difference between

ature leaves and other tissues compared with CWINV1. NIV wasxpressed in all of the tissues, with the highest expression level in35-DAB fruit. The expression levels of SUSY1 in 44- and 135-DABruit were much higher than in 75-DAB fruit, leaves and shoot tips.

eanwhile, the expression level of SUSYA was highest in 135-DABruit. SUSY6 expression was detected in all tissues and the highestxpression level was observed in leaves. SUSY6 in fruit decreasedrom 44 DAB to 75 DAB and increased slightly near maturity.

.7. Identification of genes expressed differently in different partsf fruit

To classify the expression pattern of key genes involved in thenloading pathway and post-phloem transport, we divided theiwifruit into six parts: The epicarp (EP), outerpericarp (OP), ovaryall vascular bundles (OB), peripheral placenta (PP), placental vas-

ular bundles (PB), and central placenta (CP) (Fig. 8). Compared withhe PP and PB, the areas containing the OP and OB showed higherxpression levels of SUT1, SUT3, INH and SUSY1. By contrast, CWINV,T7 and HT14 exhibited high expression in the area containing theP and PB. However, the core had the highest expression levelsf SUT1, SUT3, SUT6 and SUSY6. SUT4 and SUT5 exhibited a slightxpression difference. The expression levels of HT14 in the OP andP were higher than in the OB and PB. For NIV, the expression levelas higher in the PB and CP than in other parts. (Fig. 8).

. Discussion

.1. Suc is a major transported carbohydrate in kiwifruit

Suc represents the predominant phloem translocated sugar in

he majority of higher plant species studied thus far, although sugarlcohols (mannitol or sorbitol) are also important phloem trans-orted carbohydrate in species like celery [35], olives [36], peachesnd apples [37]. In apples and other Rosaceae trees, both Sorbitol

ment. SUSY: Sucrose synthase; CWINV: Cell wall invertase; vAINV: Vacuolar acid of three replicates ±SD.

and Suc are translocated to and utilized in fruit; sorbitol accountsfor roughly 60–70% of the photosynthates produced in leaves andtransported in the phloem [38]. Although in kiwifruit leaves Suc,myo-inositol and planteose are the major soluble carbohydrates[17], it had been suggested that myo-inositol can not be transportedfrom Actinidia leaves to fruit through the phloem system [17,18]. InA. deliciosa leaves, Suc is the predominant soluble carbohydrate. Inphloem exudates from petioles and pedicels of kiwi, Suc was a pri-mary sugar. Although Fru, Glc and inositol were detected in phloemexudates, their concentrations in the EDTA buffer was not markedlydifferent than in stopped phloem exudation efflux buffer caused bycallose gelation induced by CaCl2 [27]. Those soluble carbohydratesin the CaCl2 buffer may be from other tissue cell types of the sam-ple, but not from phloem cells, as reported by Liu et al. [26]. Theseresults indicate that Suc is the major carbohydrate transported fromleaves to fruit in kiwi.

4.2. Phloem unloading follows an apoplastic pathway in kiwifruit

Our cytological studies reveal that no plasmodesmata exist atthe interface between the SE-CC complex and its surroundingPCs during the entire course of kiwifruit development, whereasthe phloem is symplastically interconnected with the surroundingphloem parenchyma cells, as shown in developing fruit of apple [9]and pear [8] (Fig. 4). The SE-CC complex in the phloem of periph-eral and central bundles feeding the fruit flesh with assimilates issymplastically isolated during fruit development. The scarcity ofplasmodesmata indicates symplastic disjunction; apoplastic trans-port is then required [39]. This was shown by an in vivo functionalinvestigation of CF movement, which can only be transportedthrough the plasmodesmata and not across membranes [9]. Thefluorescein is pH dependent, and it is normally transferred into thevacuole after 3–6 h presence in the cytosol [4], which would lead toquenching of fluorescein because of acidic pH values in the vacuole.

In the present study, to detect whether there was fluorescence sig-nal in the kiwi fruit parenchyma cells, fluorescence was checked at12 h, 1 d, 2 d, and 4 d after carboxyfluorescein feeding. At all timepoints, the fluorescence signal was confined strictly to the phloem

C. Chen et al. / Plant Science 255 (2017) 40–50 47

Fig. 7. Sugar concentration and relative mRNA expression for genes encoding sugar transporters(including SUT, HT, TMT) and enzymes involved in sugar metabolism duringk formew using a d from

sa(dtndfp

iwifruit development(including CWINV, NIV, SUSY). Quantitative RT-PCR was perith those of EF1�, and the relative expression levels of each gene were obtained

re means of three technical replicates of the reverse transcribed RNA sample poole

trands of peripheral and central bundles in fruit fresh withoutpparent diffusion to the surrounding tissues and parenchyma cellsFig. 5), confirming the absence of symplastic phloem unloadinguring development of kiwifruit. Both observations are consis-ent with extensive apoplastic Suc phloem unloading in kiwifruit,

amely, from the peripheral and central bundles, throughout fruitevelopment (Fig. 4, 5). Soluble sugars increase rapidly during lateruit development in kiwifruit. If phloem unloading were sym-lastic and driven mainly by bulk flow and passive diffusion via

d with gene-specific primers. For each sample, transcript levels were normalizedthe 2�Ct method while expression in mature leaves was designated as ‘10′ . Values

5 biological replicates ± SD.

plasmodesmata, the concentration gradient from the phloem to theflesh cells would result in back-diffusion to the SEs [2]. The apoplas-tic pathway of phloem unloading may be an efficient mechanismto hinder potential back flow out of the kiwifruit.

4.3. Post-phloem transport of Suc in kiwifruit

Suc unloaded into apoplastic space can be transported into fruitPCs via two routes. Suc can be converted into Fru and Glc by

48 C. Chen et al. / Plant Science 255 (2017) 40–50

Fig. 8. Relative mRNA expression for genes encoding sugar transporters (including SUT, HT) and enzymes involved in sugar metabolism (including CWINV, INH, NIV, SUSY)in different parts of kiwifruit. Quantitative RT-PCR was performed with gene-specific primers. For each sample, transcript levels were normalized with those of EF1�, andt ile exv l placs

CpbbwCtalieal

he relative expression levels of each gene were obtained using the 2-�Ctmethod whascular bundles; PP, peripheralplacenta; PB, placental vascular bundles; CP, centraample pooled from 5 biological replicates ± SD.

WINV located in the apoplast, with the resulting hexose trans-orted into PCs by HTs. Alternatively, Suc in apoplastic space cane directly transported into PCs by SUT located on the plasma mem-rane [7,40]. In the placenta phloem parenchyma of tomato fruit,here Suc is unloaded apoplastically, the transcript of fruit-specific

WINV, SlCWINV1 (Lin5), is abundant [41]. Furthermore, CWINV isypically considered to be a sink-specific enzyme in plants withpoplastic Suc unloading, and its activity is often low in sourceeaves [42,43]. However, Li et al. [11] found that all three MdCWINVsn apple with apoplastic Suc unloading had lower expression lev-

ls in fruit than in the leaves. In kiwifruit, CWINV enzyme activitynd gene expression in growing fruits is very low compared witheaves. It has been suggested that CWINV may not necessarily

pression in PE was designated as ‘10′ . EP, epicarp; OP, outerpericarp; OB, ovary wallenta. Values are means of three technical replicates of the reverse transcribed RNA

relate to apoplastic unloading of Suc in fruit [12,23]. Both HT7and HT14, the two key hexose transporters involved in apoplas-tic sugar unloading, also showed low expression during kiwifruitdevelopment. Meanwhile, we detected pronounced SUT expres-sion, especially SUT3, in kiwifruit (Fig. 7), as previously reportedby Nardozza et al. [23]. In apple, MdSUT1, a homolog of kiwifruitSUT3, was suggested to play a key role in Suc unloading with highaffinity to Suc [40]. These results indicate that Suc unloaded intoapoplastic space in kiwifruit is mainly transported into the PCs bySUT and then hydrolyzed by enzymes in the cytosol.

Suc transported into PCs by SUT can be hydrolyzed by NINV orSUSY, both of which affect unloading capacity and sink strength inthe post-unloading stage of Suc [6]. It was reported that, in Ara-

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idopsis, activities of NINV regulate sugar metabolic homeostasis,nd would allow coordination of sucrose catabolism with carbonemand in nonphotosynthetic cells [44]. However, the enzymectivity of SUSY in kiwifruit was higher than that of NINV (Fig. 5),s previously reported by Moscatello et al. in kiwifruit [12] and byi et al. [11] in apple. In the present study, we also detected threeUSY genes in fruit, and SUSY1 showed much higher expressionevels in fruit than in leaves, consistent with the enzyme activityattern. The activity of SUSY is correlated with sink strength oftorage organs, e.g. citrus fruit [5] and kiwifruit [12]. Higher SUSYctivity in kiwifruit may be due to the following reasons. (1) NINVatalyzes break the irreversible conversion of Suc into Glc and Fru,nd SUSY produces fructose and UDP-glucose, enabling reversibleuc synthesis [6]. Higher SUSY activity would better maintain Sucomostasis in fruit cells. (2) During development of kiwi fruit,xcess carbon is mainly stored as starch, which can be synthe-ized from UDP-Glc. SUSY has been suggested to be involved in theirect conversion of Suc into UDP-Glc linked to starch biosynthesis

n autotrophic organs [45]. (3) SUSY is an integral component ofhe cellulose synthesis machinery [14,46]. The higher SUSY activ-ty may explain why kiwifruit contains abundant cellulose and itserivatives [34].

In conclusion, in kiwifruit, when Suc is transported into fruitrom leaves, it can be unloaded via the apoplastic pathway. Most ofpoplasmic Suc is transported into the cytoplasm directly by SUTsn the outerpericarp. Cytoplasmic Suc is mainly hydrolyzed intoru and UDP-Glc by SUSY. However, compared with the outer peri-arp, the peripheral placenta of kiwifruit had higher CWINV1 andTs expression, whereas SUT1 and SUSY1 expression levels were

ower (Fig. 8). These results imply that, compared with the outer-ericarp, more apoplasmic Suc in the peripheral placenta can beydrolyzed into Glc and Fru via CWINV before being taken up intohe cytoplasm, with hexose transported into the cytoplasm by HTs6]. According to the model, the transporters of hexose and Suc

ay regulate phloem unloading in kiwifruit. Sucrose transporterSUT1) and two hexose transporters (HT7 and HT14) have shownotential function. Additionally, SUSY1 plays an important role inaintaining Suc concentration in the cytoplasm of kiwifruit [12].

urthermore a reverse genetic approach will facilitate the deter-ination of the importance of these genes, which will allow the

oluble sugars unloading mechanism to be fully characterized ando regulate sink strength as well as fruit sweetness.

cknowledgements

This work was supported in part by the Program for the Nationalatural Science Foundation of China (No. 31372038) and the Projectf Shaanxi Province Youth Science and Technology New Star (No.14KJXX-43). The authors would like to thank Prof. Steven Vanocker (Michigan State University) for help in revising our Englishomposition, and Mr. Xuanchang Fu and Zhengwei Ma for main-aining the plants.

ppendix A. Supplementary data

Supplementary data associated with this article can be found, inhe online version, at http://dx.doi.org/10.1016/j.plantsci.2016.11.11.

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