carbohydrate metabolism and signaling in squashhexoses (davis et al., 1998). other species, such as...

Carbohydrate Metabolism and Signaling in SquashNectaries and Nectar Throughout Floral Maturation1[OPEN]

Erik M. Solhaug, Elizabeth Johnson, and Clay J. Carter2,3

Department of Plant & Microbial Biology, University of Minnesota, St. Paul, Minnesota

ORCID ID: 0000-0001-9726-1722 (C.J.C.).

Floral nectar is a sugary solution produced by plants to entice pollinator visitation. A general mechanism for nectar secretion hasbeen established from genetic studies in Arabidopsis (Arabidopsis thaliana); however, supporting metabolic and biochemicalevidence for this model is scarce in other plant species. We used squash (Cucurbita pepo) to test whether the genetic model ofnectar secretion in Arabidopsis is supported at the metabolic level in other species. As such, we analyzed the expression andactivity of key enzymes involved in carbohydrate metabolism in squash nectaries throughout floral maturation and theassociated starch and soluble sugars, as well as nectar volume and sugar under different growth conditions. Here we showthat the steps that are important for nectar secretion in Arabidopsis, including nectary starch degradation, Suc synthesis, and Sucexport, are supported by metabolic and biochemical data in C. pepo. Additionally, our findings suggest that sugars importedfrom the phloem during nectar secretion, without prior storage as starch, are important for generating C. pepo nectar. Finally, wepredict that trehalose and trehalose 6-P play important regulatory roles in nectary starch degradation and nectar secretion. Thesedata improve our understanding of how nectar is produced in an agronomically relevant species with the potential for use as amodel to help us gain insight into the biochemistry and metabolism of nectar secretion in flowering plants.

Floral nectar is a plant secretionmade predominantlyof sugars that serves as a reward for pollinators and isessential for efficient reproduction in many plants.Nectar is secreted from specialized glands called nec-taries and involves a number of steps that are tightlyregulated in order to increase pollination success whileconserving precious carbon resources (Roy et al., 2017).The composition of sugars in nectar varies greatlydepending on the species. Some species, such as Ara-bidopsis (Arabidopsis thaliana), produce nectar rich inhexoses (Davis et al., 1998). Other species, such as to-bacco (Nicotiana tabacum; Ren et al., 2007) and squash(Nepi et al., 2001; Solhaug et al., 2019), produce nectarrich in Suc. Previous research has shown that squash(Cucurbita pepo) cultivars with a higher Suc/hexose ra-tio in their nectar have increased pollinator visitscompared with cultivars with lower Suc relative tohexoses (Roldán-Serrano and Guerra-Sanz, 2005), sug-gesting that composition of nectar sugar can influence

pollinator visitation, which has also been shown tohave a direct impact on fruit yield (Motzke et al., 2015;Pereira et al., 2015; Zou et al., 2017).

Early stages of nectary maturation are typically as-sociated with build-up of starch in the nectary. Indeed,presecretory accumulation of starch in nectaries hasbeen reported for many flowering species (Nepi et al.,1996; Peng et al., 2004; Ren et al., 2007; Lin et al., 2014;Solhaug et al., 2019). As secretion proceeds, nectarystarch is degraded and nectar sugar is produced con-currently (Solhaug et al., 2019).

In Arabidopsis, Suc synthesis during secretion isimportant for efflux of sugar from nectar-secreting cells.Plants silenced for SUCROSE PHOSPHATE SYNTHASE1F and 2F (sps1f/2f) fail to secrete nectar and accumu-late more starch in their nectaries than wild type,suggesting synthesis of Suc from starch breakdownproducts is essential for nectar secretion in plants thataccumulate starch in nectariferous tissues (Lin et al.,2014).

Although the connection between starch degradationand nectar secretion has been well established in anumber of species (Peng et al., 2004; Ren et al., 2007; Linet al., 2014; Solhaug et al., 2019), the proportion ofnectar sugar coming from starch versus sugar deriveddirectly from the phloem is not well understood. Insome species, such as tobacco (Ren et al., 2007) andAnigozanthos flavidus (Wenzler et al., 2008), phloemsugar can be directly transported into nectar (bypassingstarch). It is likely that the source of nectar sugar (i.e.nectary starch or direct phloem sugar) varies betweenspecies and may partially depend on coevolution ofspecific plant-pollinator interactions (Heil, 2011).

1This work was supported by the U.S. National Science Founda-tion (grant 1339246 to C.J.C.).

2Author for contact: [email protected] author.The author responsible for distribution of materials integral to the

findings presented in this article in accordance with the policy de-scribed in the Instructions for Authors (www.plantphysiol.org) is:Clay J. Carter ([email protected]).

E.M.S. and C.J.C. designed the experiments and wrote the articletogether; E.M.S. performed the majority of the experiments and ana-lyzed the data, with assistance from E.J.

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During secretion in Arabidopsis nectaries, Suc isexported predominantly through SUGARS WILLEVENTUALLY BE EXPORTED TRANSPORTERS9(AtSWEET9), a nectary-specific plasma membrane-localized Suc uniporter (Lin et al., 2014). Similar toatsps1f/2f plants, atsweet9 mutants lack nectar and ac-cumulate large amounts of starch in nectar-secretingcells (Lin et al., 2014), suggesting that Suc synthesisand export are essential for maintenance of nectarystarch degradation and nectar secretion in Arabidopsis.Because AtSWEET9 is a bidirectional transporter (Linet al., 2014), the transport of Suc from nectar-secretingcells is contingent on the development and mainte-nance of a concentration gradient of Suc between theinside and outside of the cells. One way that this gra-dient can be generated is by cleavage of Suc in the ex-tracellular space by nectary-specific CELL WALLINVERTASE4 (AtCWINV4). atcwinv4 mutants do notmake any nectar, but still accumulate starch in thenectary (Ruhlmann et al., 2010), suggesting that extra-cellular hydrolysis of Suc by AtCWINV4 is essential fornectar secretion, but is not necessary for import ofphloem sugar into Arabidopsis nectaries. AlthoughCWINV4 is essential for nectar secretion in Arabi-dopsis, other species, particularly those that utilizevesicles via a granulocrine model of nectar secretion(such as Echinacea purpurea; Wist and Davis, 2008) orthose that produce a Suc-rich nectar (such as tobacco;Ren et al., 2007), may secrete nectar via a mechanismalternative to CWINV4.In summary, the most supported model of nectar

secretion in Arabidopsis involves build-up and degra-dation of nectary starch, resynthesis of Suc from starch-derived hexoses, export of Suc by AtSWEET9, andextracellular hydrolysis of Suc, which maintains a highintracellular-to-extracellular concentration gradient ofSuc while also increasing the solute potential, causingwater to flow out into the developing nectar droplet(Lin et al., 2014; Roy et al., 2017). Although this modelhas been highly useful in improving our understandingof the genes and proteins that are important for nectarsecretion in starch-accumulating nectaries, it has not yetbeen fully tested from a biochemical and metabolicperspective. The small size of Arabidopsis nectary tis-sue makes such studies nearly impossible. In order tobiochemically test the primary model of nectar secre-tion in Arabidopsis, we used C. pepo. C. pepo is an idealsystem for studying metabolism in nectaries before,during, and after nectar secretion because the plantsmake many flowers with a typical flower producingover 1000-fold more nectar sugar than a typical Arabi-dopsis flower (;25 mg sugar/flower for C. pepo [Nepiet al., 2001]; 1.5 mg sugar/flower in Arabidopsis [Daviset al., 1998]).Transcriptomic studies have suggested that the

mechanism of nectar secretion in C. pepo is similar tothat in Arabidopsis (Solhaug et al., 2019). However,additional experiments are warranted to test whetherthis geneticmodel is reflected by the enzymatic patternsand flux of metabolites through the nectary system. In

this study, we sought to examine how carbohydratesare trafficked, partitioned, and secreted into the nectarof C. pepo flowers. Here we show that the genetic modelof nectar secretion in Arabidopsis is supported byphysiological and biochemical experiments in C. pepo.We next show numerous pieces of evidence thatphloem-derived sugars play an important role in nectarsecretion in C. pepo. Finally, we examine the role oftrehalose metabolism in regulation of starch degrada-tion and production of nectar sugar. These data repre-sent an important step in improving our understandingof how floral nectar is produced from a metabolic andbiochemical perspective.

RESULTS

Most floral nectaries are nonphotosynthetic, beingfully dependent on phloem-derived sugars, primarilystored as starch, as the precursors to nectar (Roy et al.,2017). As such, we first examined starch and solublesugar accumulation from 3 d (272 h) to 1 d (224 h)before secretion (see Fig. 1, A and B for developmentalseries) in order to determine how nectaries store energythroughout development. Nectary starch increasedover 10-fold from272 to224 h (Fig. 1C). There was nochange in levels of nectary Suc, although nectary glu-cose decreased about 2-fold from 272 to 224 h(Fig. 1D). Over the same time course, we found a 3-foldincrease in nectary mass (Supplemental Fig. S1C).These data suggest that most of the carbohydrates en-tering the nectaries between272 and224 h, henceforthtermed the starch-filling stage, are broken down andeither used to synthesize starch or metabolized to pro-duce ATP for other processes essential for growth (e.g.,cell wall acidification and loosening via H1-ATPase;Majda and Robert, 2018).Because C. pepo uses raffinose-family oligosaccha-

rides (RFOs), such as stachyose, as its primary transportsugars (Zhang et al., 2010), we used a-GALase as ameasure of sink strength. a-GALase hydrolyses galac-tosyl units from stachyose (tetrasaccharide) and raffi-nose (trisaccharide) to form Suc (Carmi et al., 2003). Wesought to determine whether there was a temporalsimilarity between a-GALase activity and starch-fillingin nectaries at different time points. Total neutrala-GALase activity (cytosolic) in nectaries increasednearly 3-fold from 272 to 224 h (Fig. 1E). However,over the same time period, the acid a-GALase activity(vacuolar) was not significantly different. At each timepoint measured, the neutral a-GALase activity washigher than the acid activity (Fig. 1E). In order to de-terminewhethera-GALase activity induction is specificto the nectary, we examined a-GALase activity in nec-tary, receptacle, and peduncle tissues (diagrammed inSupplemental Fig. S2A). The neutral a-GALase activitywas over 2-fold higher in the nectary when comparedwith the peduncle or the receptacle at all three timepoints measured (Supplemental Fig. S2B). In contrast,acid a-GALase activity was not significantly different

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Sugar Metabolism in Squash Nectaries and Nectar

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throughout the starch-filling stage (SupplementalFig. S2C).

In order to further dissect the breakdown andutilization of soluble sugars during nectary devel-opment, we looked at invertase activity from 272 to224 h. Acid (vacuolar) invertase (vINV) activity in-creased nearly 4-fold from 272 to 224 h, whereasneutral (cytosolic) invertase (cINV) increased 3-foldover the same time (Fig. 1F). vINV activity wasconsistently higher than cINV activity at all timepoints measured (Fig. 1F). Taken together, these datashow that the starch-filling stage is accompanied byincreases in cytosolic a-GALase and cINV, as wellas vINV.

Previous studies have shown that degradation ofnectary starch occurs simultaneously with the produc-tion of nectar sugar in many species (Nepi et al., 1996;Peng et al., 2004; Ren et al., 2007; Lin et al., 2014;Solhaug et al., 2019). In order to further examine starchmetabolism in C. pepo nectaries, we first asked whetherthe starch was degraded during the dark period im-mediately preceding nectar secretion. Starch content innectaries remained relatively constant from215 h until23 h (dawn523 h; Fig. 2A). However, from23 to 0 h,the starch was degraded rapidly (P , 0.005; Fig. 2A).Because we saw an apparent dawn-induced degrada-tion of starch beginning at23 h, we wondered whetheramylase (a group of major starch degradative enzymes)activity showed the same pattern. Amylase activity wasnot significantly different from 215 to 29 h (Fig. 2B).However, from 29 to 26 h, there was a significant in-crease in amylase activity (P 5 0.039), which thenremained constant until 0 h (Fig. 2B). These data showthat amylase activity is strongly induced 3 h be-fore the onset of dawn, suggesting the regulation ofstarch degradative enzymes is at least partially light

independent, although it may be controlled viaentrained circadian rhythms.

Sink tissues, such as maize kernels or potato tubers,often contain a portion of their starch that is more re-sistant to degradation than transitory leaf starch

Figure 1. Carbohydrate partitioning dur-ing the starch-filling stage in C. peponectaries. A and B, Example imagesof three developmental stages for whole(A) and dissected (B) flowers. Scalebars 5 1 cm. Nectaries are denoted byan arrow in (B). C, Starch content innectaries from 272 to 224 h beforepeak secretion. D, Levels of Suc and Glcin nectaries at the same time points. E,Neutral (pH 7.4) and acid (pH 5.5)⍺-galactosidase (a-GALase) activity. F,Acid (pH 4.8) and neutral (pH 7) inver-tase activity (micromole Glc produced).For the activity assays, activity was nor-malized to total protein added (milli-gram protein). Bars that share a letter arenot significantly different from one an-other (a, b, c were tested together; x, y, zwere tested together; P , 0.05, Tukeypost hoc test). Error bars represent SE(n 5 4). 4-NP, 4-nitrophenol.

Figure 2. Starch and sugar metabolism as nectaries progress towardnectar secretion. Starch (A) and total amylase activity [B; micromolereducing (red.) sugar produced] in nectaries are shown. Error barsrepresent SE (n 5 8). Columns that share a letter are not significantlydifferent from one another (Tukey post hoc test, P , 0.05). The bottombar indicates light and dark periods.

1932 Plant Physiol. Vol. 180, 2019

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(McCleary and Monaghan, 2002). In order to confirmthat we were detecting all of the starch with our ana-lytical method, we used an additional KOH treatmentto measure starch that is resistant to hydrolysisby boiling with thermo-stable amylolytic enzymes(Megazyme, resistant starch protocol). There was nodetectable difference when analyzing starch using aresistant starch protocol at 224 h, (SupplementalFig. S3) indicating all of the starch is fully hydrolyzedduring the normal starch assay.To further examine the role of starch degradation in

nectar production, we analyzed starch, nectar volumeand sugar, nectary sugar, and total combined sugarover a 9-h period starting from 3 h before dawn (Fig. 3;26 h relative to peak nectar secretion). There was con-sistently no nectar present at dawn (23 h; Fig. 3A),when starch is still present in nectaries (Fig. 2A;Fig. 3D). Nectar volume increased more than 2-foldfrom 0 to 13 h, but after 13 h the nectar volume didnot significantly change (Fig. 3A). The period of great-est change in nectar volume was from 23 to 13 h(Fig. 3A), which coincided with near complete degra-dation of nectary starch (Fig. 3D), supporting previousresults showing that nectary starch degradation is in-timately connected to nectar secretion (Nepi et al., 1996;Peng et al., 2004; Ren et al., 2007; Lin et al., 2014;Solhaug et al., 2019). Our nectar volume data were alsosupported by previous data showing that most C. peponectar is produced in the early morning (Nepi et al.,2011).Previous studies in Arabidopsis suggested that ex-

tracellular hydrolysis of Suc by AtCWINV4 is requiredfor the generation of a concentration gradient to drive

Suc export (Ruhlmann et al., 2010). However, ourrecent study suggested there was low expression ofCpCWINV4 in C. pepo nectaries at 0 h (Solhaug et al.,2019). In order to examine the mechanism of nectarsecretion in squash, we sought to compare concentra-tions of sugar in the nectar and nectary throughout theprocess of nectar secretion. At 0 h (peak nectar secre-tion), the concentration of nectar Suc was 3-fold higherthan Glc (Fig. 3C). From 0 to 13 h, nectar Suc concen-tration ([Suc]) stayed constant, whereas nectar Glcconcentration ([Glc]) increased by 20% (Fig. 3C). From13 to 19 h, nectar [Suc] decreased by 36%, whereasnectar [Glc] decreased by only 18% (Fig. 3C). However,an additional experiment testing the nectar sugar froma 124 h flower showed that [Suc] and [Glc] both de-crease equally from 0 to 124 h (Supplemental Fig. S4),suggesting that resorption of Suc and Glc may occursimultaneously in C. pepo nectaries.Based on a mass-flow model of nectar secretion pre-

dicted by Suc export via SWEET9 (Lin et al., 2014), twoscenarios are possible for the export of nectar sugar.One possibility involves complete or near completehydrolysis of Suc by CWINV4 in the extracellular spaceto create a concentration gradient facilitating continuedexport of nectary Suc, generating a hexose-rich nectar.Alternatively, the nectary could maintain a similarlyhigh [Suc] as present in the nectar, allowing newlysynthesized (or phloem-derived) Suc to be transporteddown the concentration gradient into the nectar with-out complete hydrolysis, leading to a nectar rich in Suc.In order to compare the [Suc] between the nectary

and the nectar, we estimated nectary volume using atwo-half-sphere volume equation (Supplemental Fig. S5).

Figure 3. Carbohydrate partitioning during nectarsecretion in C. pepo. A, Nectar volume correctedfor nectary size (microliter per gram FW). B,Nectary Suc concentration (mole per liter). C,Nectar Suc andGlc concentrations (mole per liter)throughout the day. D and E, Total nectary starch(D) and total sugar (E, nectary1nectar) throughoutsecretion. Error bars represent SE [n 5 8 for alldata presented, except for (D) and (E), where n5 7for 23 and 13 h and (A), where n 5 9 for 0 h].Columns that share a letter are not significantlydifferent from one another (a, b, c were testedtogether; x, y, z were tested together; P , 0.05,Tukey post hoc test). The bottom bars indicateslight and dark periods.









At 0 h, during the highest rate of increase in nectarvolume, the [Suc] in the nectary was ;0.6 M (Fig. 3B),which was only 17% lower than the [Suc] in the nectarat 0h (;0.7 M), suggesting that similar [Suc] in thenectary and nectar may be important to allow SWEET9-facilitated Suc export into the nectar. The nectary [Suc]decreased 36% from 0 to19 h (Fig. 3B), almost perfectlymatching the decrease in nectar [Suc], which also de-creased 36% from 13 to 19 h (Fig. 3C). The strikingsimilarity between nectary (cytosolic) and nectar (apo-plastic) [Suc] further suggests that the creation andmaintenance of a high cytosolic [Suc] in C. pepo nec-taries is required for the initiation and continued flow ofSuc into the nectar.

The close temporal alignment of nectary starch deg-radation with the accumulation of nectary Suc andnectar secretion led us to question what proportion ofthe total system (nectary1nectar) sugar comes fromstarch breakdown versus sugars supplied directly fromthe phloem without prior storage. In order to answerthis question, we calculated total carbon balance com-paring the total starch, Suc, Glc, maltose, and malto-oligosaccharides (maltoOS) in the nectary and nectarthroughout the process of nectar secretion (Table 1). Wefound a maximum of;830 mmol of Glc equivalents pergram fresh weight (mmol Glc eq/g FW) existing asnectary starch (26 h; Fig. 3D). At this time, there was;185 mmol Glc eq/g FW in total soluble sugar (Fig. 3E;Table 1). Because we did not find substantial degrada-tion of starch before 26 h (Fig. 2A), we assumed thatany soluble sugar present at 26 h did not come fromnectary starch degradation. From26 to23 h, the starchdecreased by;270mmolGlc eq/g FW (Fig. 3E; Table 1),whereas the total sugar in the nectary increased bynearly 2-fold more (;502 mmol Glc eq/g FW; Fig. 3E),suggesting at least some of this sugar does not comefrom starch degradation (Table 1). The total system(nectary1nectar, summed together) sugar increased by;1190 mmol Glc eq/g FW from 26 to 0 h, whereas thestarch only decreased by ;758 mmol Glc eq/g FW(Table 1). Additionally, our estimate of the total systemsugar at 13 h was 1454 mmol Glc eq/g FW (Table 1),which is nearly 2-fold higher than the maximum starchwe estimated to be in the nectary at 26 h (Table 1).Taken together, these data suggest that a majority(;59%) of the total sugar in the nectary/nectar systemat 13 h (time of peak Suc content in nectary1nectarsystem; Supplemental Fig. S6) comes from starch.

Previous data has suggested that Suc is synthesizedfrom starch-derived hexoses via AtSPS1F/2F duringnectar production in Arabidopsis (Lin et al., 2014). Inorder to determine whether Suc synthesis is also acti-vated during secretion in C. pepo, we analyzed SPS ac-tivity at four stages of nectary maturation (224,215, 0,and 19 h; examples in Supplemental Fig. S1A). SPSactivity was nearly 3-fold higher at 0 h compared to224 h (P 5 0.003; Fig. 4A), suggesting that de novosynthesis of Suc from hexoses plays an important roleduring nectar secretion. SPS activity decreased from 0 hto19 h by nearly 20%; however, this difference was notstatistically significant (Fig. 4A).

Because we found evidence that the maintenance of ahigh cytosolic Suc concentration is important for gen-erating C. pepo nectar (Fig. 3), wewanted to see whetherSuc degradation by intracellular invertase (Suc hydro-lase) and Suc synthase (SuSy; important for Suc catab-olism and sink strength; Angeles-Núñez and Tiessen,2010; Ferreira and Sonnewald, 2012) were decreased inorder to maintain intracellular [Suc]. Activity of acidinvertase (vacuolar, vINV) increased by nearly 66%from 224 to 0 h (Fig. 4B), suggesting vacuolar hydrol-ysis of stored Suc increases during secretion. However,neutral invertase (cytosolic, cINV) activity did notchange throughout secretion (Fig. 4B). Additionally,SuSy activity increased gradually from 224 to 19 h,but these differences were not statistically significant(P . 0.18; Fig. 4C).

In order to further test whether sugar imported fromthe phloem is a substantial portion of the total carbonbalance in nectaries during nectar secretion, we exam-ined whether the activity of a-GALase, as a mea-sure of sink capacity, varied throughout the samestages. Neither neutral (cytosolic) nor acid (vacuolar)a-GALase activity were significantly different at any ofthe four stages measured (Fig. 4D), suggesting thatimport and breakdown of phloem-derived RFOs byC. pepo nectaries is unchanged during nectar secretion.

We next attempted to reduce the amount of sugarcoming from phloem-derived photoassimilates in orderto determine the extent to which phloem sugars(without prior storage as starch) contribute to C. peponectar sugar production. Toward this end, we exposedwhole plants to a 5-h extended night to induce starva-tion in source tissues (and nectaries) before analyzingnectar(y) carbohydrates. Nectary starch was 2.5-foldhigher in plants exposed to an extended night compared

Table 1. Total Glc eq in the nectary plus nectar system during the secretory processa

Stage Total Sucb Total Glcb Total Maltosec Total MaltoOSc Total Sugard Total Starch Total GLC Eqe

26 h (n 5 8) 117 6 11 13 6 2 34 6 4 21 6 1 185 6 16 829 6 46 1,014 6 5723 h (n 5 7) 455 6 48 71 6 16 128 6 17 34 6 4 688 6 78 558 6 59 1,246 6 760 h (n 5 8) 855 6 37 369 6 11 133 6 23 19 6 8 1,375 6 43 71 6 6 1,446 6 4513 h (n 5 7) 1,065 6 89 233 6 22 122 6 7 18 6 3 1,439 6 103 16 6 6 1,455 6 103

aContent of different sugars and starch in micromole of Glc eq per gram FW. bSum of nectary and nectar (micromole Glc eq per grams FW).Total Suc (in Glc eq) was calculated as two times the total Suc measured. cObtained from nectaries alone. Total Glc equivalents in maltose wascalculated as two times the total maltose measured. dSum of Suc, Glc, maltose, and maltoOS at individual time points. eSum of total sugarsand starch in the nectary and nectar system (in micromole Glc eq per grams FW).


Solhaug et al.





with control nectaries harvested the previous morning(P 5 0.0045; Fig. 5A); however, this difference repre-sented less than 2% of the total starch present in thenectary (assuming ;830 mmol Glc eq/g FW at peak of

nectary starch), suggesting most of the starch is stilldegraded in the absence of the light stimulus. Totalnectar volume was 49% higher in the control nectariescompared with the extended night (Fig. 5B), but therewas no significant difference in nectar [Suc] or [Glc](Fig. 5C). We next examined total Suc, Glc (in micro-mole per gram FW), and total sugar (Suc1Glc, in Glceq) in nectar, nectary, and total system (nectary1nectar,summed together) in order to determine whether re-duction of phloem-derived sugar led to differences incarbohydrate partitioning. The total nectar Suc was58% higher in the control versus the extended nightsamples (Fig. 5D), whereas the total system Suc was28% higher in the control compared with extendednight (Fig. 5D). However, the total nectary Suc was notsignificantly reduced upon treating the plants with anextended night (Fig. 5D). Similar to nectar Suc, the totalnectar Glc was 62% higher in the control versus theextended night samples (Fig. 5E), but there was nodifference in nectary or total system Glc (Fig. 5E). Ad-ditionally, we found 59% more total nectar sugar incontrol versus extended night samples (Fig. 5F). Al-though there was more nectar sugar in control samples,total system sugar did not vary between control versusextended night samples (Fig. 5F). Interestingly, therewas actually 15% more total nectary sugar in extendednight samples compared with control (P 5 0.032;Fig. 5F). In summary, exposing plants to an extendednight causes flowers to store more sugar in their nec-taries rather than secreting sugar into the nectar.We next sought to alter the amount of phloem sugars

entering the nectaries by removing flowers from plantsat215 h (the evening before secretion) and placing theirpeduncles in water. We compared carbon balance at11 h (total 16-h starvation) in flowers left on the plantfor the night (attached) to flowers removed and put intowater (detached). Removing the flowers from theplants led to no change in nectary starch (Fig. 6A) ornectar volume (Fig. 6B). However, the nectar [Suc] was39% higher in the attached flowers compared with thedetached flowers (P 5 0.0008; Fig. 6C), whereas thenectar [Glc] was actually higher in the detached flow-ers, although this difference was not statistically sig-nificant (Fig. 6C). The total nectar Suc was nearly 2-foldhigher in the attached samples compared with de-tached (Fig. 6D), whereas total system Suc was alsohigher in attached flowers, but to a lesser extent (41%;Fig. 6D). Similar to the response seen when extendingthe night, nectary Suc did not change as a result of re-moving the flowers from the plants (Fig. 6D). The totalnectar Glc and total system Glc (micromole per gramFW) were not significantly different between detachedand attached samples, but nectary Glc was nearly 67%higher in the attached samples compared with the de-tached samples (Fig. 6E). Total sugar in the nectar ornectary was not statistically different between attachedand detached flowers (Fig. 6F). However, there wasmore total system sugar in attached flowers comparedwith detached flowers (P 5 0.037; Fig. 6F). Taken to-gether, these data suggest that removing the flowers

Figure 4. Activity of sugar metabolism enzymes during nectary matu-ration. A, SPS activity is induced during secretion. B, Acid (pH 4.8)invertase increases during secretion (P , 0.1), whereas neutral (pH 7)invertase activity stays constant throughout secretion. C, SuSy activityincreases throughout secretion. D, Acid (pH 5.5) and neutral (pH 7.4)a-GALase assays show no change in activity throughout secretion. Errorbars represent SE (n5 4, except for SuSy activity at215 h, where n5 3).Columns that share a letter are not significantly different from one an-other ([Tukey post hoc test, P, 0.05 for (A), (C), and (D); P, 0.1 for (B)].4-NP, 4-nitrophenol.





from plants and treating them with water overnightcauses a highly similar response to exposing them to anextended night, in that the flowers retain more sugar intheir nectaries rather than secreting the sugar into thenectar.

Because Suc metabolism plays a central role in nectarsecretion, we were interested in whether sugar signalsplay regulatory roles in essential secretory processessuch as starch degradation and Suc synthesis. In par-ticular, we previously found up-regulation of tran-scripts for genes encoding enzymes involved intrehalose metabolism during secretion (Solhaug et al.,2019). In order to validate these data, we re-examinedexpression using reverse transcription-quantitativePCR (RT-qPCR) for Trehalase1 (CpTRE1) and Trehalose-phosphate phosphatase-J (CpTPP-J), which code forimportant enzymes in the catabolism of trehalose andTre-6P, respectively (Figueroa and Lunn, 2016). Ex-pression of CpTRE1 was nearly 5-fold higher duringsecretion (0 h) compared with 224 and 215 h before

secretion, and were nearly 6-fold higher at 0 h com-pared with 19 h (Fig. 7A). Additionally, CpTPP-Jtranscripts were over 5-fold induced during secretioncompared with224 and215 h, then decreased slightlyfrom 0 to 19 h (not significant; Fig. 7B). We next ex-amined whether the increase in CpTRE1 expression ledto a difference in enzymatic activity for trehalase. Tre-halase activity was nearly 2-fold higher at 0 h relative to215 h (P5 0.039; Fig. 7C), further suggesting inductionof trehalose-related catabolism may be an importantstep in C. pepo nectar secretion. Given the induction oftrehalose metabolic enzymes during secretion, wewondered whether trehalose and trehalose-6P (Tre6P)levels changed in the lead up to secretion. Both treha-lose (Fig. 7D) and Tre6P (Fig. 7E) remained constantfrom 26 to 23 h, then increased by nearly 3-fold from23 to 0 h (Fig. 7, D and E). Numerous studies havedemonstrated that Tre6P levels are directly propor-tional to Suc across different species, growth conditions,and tissues (Wingler et al., 2000; Debast et al., 2011;

Figure 5. Carbohydrate partitioningduring an artificially extended night.Dawn was delayed for 5 h before har-vesting nectar and nectary tissue andanalyzing sugar and starch. A and B,Nectary starch (A) and nectar volume (B)corrected for grams FW of nectary. C,Concentration (mole per liter) of Suc andGlc in the nectar. D to F, Total Suc (D),Glc (E), and total sugar (Suc1Glc) in Glcequivalents (F) of nectary and nectar,corrected for FW of nectary. *P # 0.05(2-sample t test). Error bars represent SE(n 5 8). Ext Night, Extended Night.

Figure 6. Carbohydrate partitioning af-ter removing flowers from plants. A andB, Nectary starch (A) and nectar volume(B) corrected for grams FWof nectary. C,Concentration (mole per liter) of Suc andGlc in the nectar. D to F, Total Suc (D),Glc (E), and total sugar (Suc1Glc) in Glcequivalents (F) content of nectary andnectar, corrected for FW of nectary.Detached, removed at 215 h andplaced in water; Attached, removedfrom the plant the following morning.*P # 0.05 (2-sample t test). Error barsrepresent SE (n 5 8 for Detached, n 5 7for Attached).


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Martins et al., 2013; Nuccio et al., 2015; Figueroa et al.,2016; Figueroa and Lunn, 2016), so we also included areanalysis of nectary Suc accumulation (same data as inFig. 3B, except expressed in micromole per gram FW)during the same time points. The nectary Suc increasednearly 4-fold from 26 to 23 h (Fig. 7F), then increasedfurther from 23 to 0 h, but to a much lesser extent (notsignificant; Fig. 7F). These data show that Tre, Tre6P,and trehalose catabolic enzymes accumulate alongwithSuc in nectaries during secretion (0 h), although Sucaccumulation precedes that of Tre and Tre6P (23 h inFig. 7).In order to further examine the role of trehalose in

regulation of nectar production, we treated detachedflowers (215 h) overnight with either 10 mM trehalose(Tre) or 10 mM sorbitol (Sorb). Tre treatment led to2-fold higher nectary starch at 11 h when comparedwith the sorbitol treatment; however, the differencewas not statistically significant (P5 0.122; Fig. 8A). Tre-treated flowers also produced nearly 2-fold lower nec-tar volume comparedwith the sorbitol control (Fig. 8B).There was no significant difference in nectar [Suc] or[Glc] of trehalose-treated flowers compared with sor-bitol (Fig. 8E). The total nectar Suc was over 2-foldlower in the trehalose-treated flowers compared withsorbitol controls, whereas the nectary Suc and totalsystem Suc was not significantly different between thetwo treatments (Fig. 8F). The total nectary Glc wasnearly 2-fold higher in trehalose-treated samples com-pared with sorbitol; however, neither nectar Glc nor

total system Glc were significantly different (Fig. 8G).Finally, the total sugar in the nectar was more than2-fold lower in the trehalose-treated samples com-pared with control, whereas the total nectary sugarwas 33% higher in the trehalose-treated samplescompared with control (Fig. 8H). The total systemsugar was remarkably similar between the sorbitoland trehalose-treated samples (Fig. 8H). In orderto determine whether the inhibited starch degrada-tion and nectar secretion was due to alterations ineither trehalose or Tre6P, we analyzed Tre6P andtrehalose content in trehalose- and sorbitol-treatedsamples. There was nearly 3-fold more trehalose inthe trehalose-treated samples compared with sorbitolcontrol (P 5 0.041; Fig. 8C), but there was onlyslightly more Tre6P (Fig. 8D). Taken together, thesedata show that trehalose treatment inhibits nectarystarch degradation and production of nectar sugar inC. pepo nectaries, possibly via a Tre6P-independentmechanism.

DISCUSSION

This work provides substantial insight into howcarbohydrates are trafficked and partitioned by nec-taries during nectarymaturation and nectar secretion inC. pepo. We have presented metabolic evidence sup-porting the eccrinemodel of nectar secretion discoveredin Arabidopsis, suggesting the mechanism of nectar

Figure 7. Trehalose metabolism in nectaries. Aand B, Gene expression of Trehalase1 (CpTRE1;A) and Trehalose-phosphate phosphatase J(CpTPP-J; B) measured in nectaries by RT-qPCRthroughout maturation; normalized to CpRING(XM _008439865.1) and presented as relativeexpression (0 h 5 1; n 5 3). C, Trehalase activitythroughout nectary maturation (n 5 4). D to F,Accumulation of trehalose (D), Tre6P (E), andSuc (F) throughout nectary maturation (n 5 7 fortrehalose; n 5 3 for Tre6P; n 5 8 for Suc). Barsthat share a letter are not statistically significantlydifferent from one another (Tukey post hoc test,P , 0.05). Error bars represent SE.





secretion is conserved between these two species. Itappears that phloem-derived sugars, without priorstorage as starch, are important for the generation ofnectar, but that nectar can still be produced in theirabsence. Finally, trehalose metabolism is likely impor-tant for regulation of starch degradation and nectarsecretion in C. pepo.

An Updated Model of Secretion for a Suc-Rich Nectar

In Arabidopsis, previous genetic experiments haveidentified a number of steps that are important fornectar secretion, including starch accumulation anddegradation, Suc synthesis by AtSPS1F/2F, Suc exportby AtSWEET9, and extracellular hydrolysis of Suc byAtCWINV4 (Ruhlmann et al., 2010; Lin et al., 2014). In

this study, we tested various steps of this mechanism tosee if they are conserved in C. pepo.

Starch Accumulation and Degradation

In C. pepo nectaries from 272 until 224 h (beforesecretion), the nectaries accumulate starch whereas thelevels of soluble sugars do not change (Fig. 1). Duringthis same time period, the nectaries grow drastically,although they continue to grow throughout secretion toa lesser extent (Supplemental Fig. S1). These findingssuggest that most of the carbohydrate coming into thenectary is converted into starch or metabolized to fuelother energetically demanding biosynthetic processesessential for continued growth of nectaries. During the

Figure 8. Exogenous trehalose inhibits nectar pro-duction. To examine a potential role for trehalose inregulating nectar production, the peduncles of ex-cised flowers were placed in solutions containing ei-ther 10 mM sorbitol (Sorb; osmotic control) or 10 mM

trehalose (Tre). A to D, Nectary starch (A), nectarvolume (B), nectary trehalose (C), and nectary Tre6P(D), all corrected for grams FWof nectary, are shown.E, Concentration (mole per liter) of Suc and Glc in thenectar. F to H, Total Suc (F), Glc (G), and total sugar(Suc1Glc) in Glc equivalents (H) of nectary andnectar, corrected for FW of nectary. *P # 0.05(2-tailed t test). Error bars represent SE (n 5 3 forTre6P; n 5 6 for trehalose; n 5 6 for Sorb samples instarch data; n 5 7 for all other samples).


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starch-filling stage (272 to 224 h), nectaries also in-crease in both acid (vacuolar) invertase and neutral(cytosolic) a-galactosidase activity (Fig. 1), indicating amechanism by which sink strength of the nectary in-creases throughout the starch-filling stage.The close temporal relationship between degradation

of nectary starch and production of nectar sugar hasbeen demonstrated in a number of species (e.g. Ren et al.,2007; Lin et al., 2014), but very little is known about themechanism behind this process. Previous data hasshown that the expression of b-amylase1 (CpBAM1) isinduced during secretion inC. pepo (Solhaug et al., 2019).InArabidopsis,AtBAM1plays an important role in light-induced starch degradation in leaf guard cells (Horreret al., 2016). Although AtBAM1 is regulated by light inArabidopsis leaves, and CpBAM1 likely plays a crucialrole in nectary starch degradation, we have shown evi-dence that nectary starch degradation in C. pepo is reg-ulated independently of light. First, total amylaseactivity (a and b) is induced 3 h before dawn and thebeginning of nectar secretion (Fig. 2B). Second, tran-scripts coding for other enzymes crucial for starch deg-radation, such as a-glucan phosphorylase 2 (CpPHS2) anddisproportionating enzyme 2 (CpDPE2; Streb and Zeeman,2012), are expressed at high levels in the evening beforenectar secretion (Supplemental Fig. S7). Finally, a largeportion of the starch is still degraded even if the night isextended, although it is likely degraded at a slower rate(Fig. 5A). Taken together, these data indicate that nectarystarch degradation probably occurs in response to de-velopmental and/or circadian signals, but ismostly lightindependent.

Suc Synthesis

Previous research has shown that Suc synthesisby AtSPS1F/2F is important for nectar secretion inArabidopsis (Lin et al., 2014). In C. pepo, CpSPS3FmRNA is high at 224 h and remains high throughoutsecretion, before decreasing at 19 h (Solhaug et al.,2019). Although CpSPS3F transcript does not changemuch from 224 to 0 h, we show here that the SPS ac-tivity is significantly higher at 0 h compared to 224 h(Fig. 4A), suggesting SPS may be regulated post-transcriptionally and/or post-translationally. SPS hasbeen shown to be regulated by reversible phosphoryl-ation in leaves (Winter and Huber, 2000), altering af-finity for substrates without impacting maximumcatalytic activity (McMichael et al., 1995).Whereas Suc synthesis activity via SPS is induced

during secretion, the activities of Suc degradation en-zymes do not substantially change (cINV and SuSy;Fig. 4, B and C). Because SPS typically exists in the cy-tosol (Champigny and Foyer, 1992), it is possible that anincrease in SPS activity without a concomitant increasein cINV and SuSy could favor Suc synthesis (rather thandegradation), leading to high cytosolic [Suc] and pas-sive transport of newly synthesized Suc out of the cellvia SWEET9. While cytosolic Suc degradation (relative

to synthesis) is likely lower during secretion, activity ofvacuolar invertase was induced slightly (Fig. 4B). vINVmay be important for liberating hexoses from storedvacuolar Suc during secretion, as transcripts encodingtwo predicted vacuolar hexose transporters (Poschetet al., 2011), ERD6-like 6 (CpEL6) and ERD6-like 16(CpEL16), are highly expressed in nectaries during thebuild-up to secretion (Supplemental Fig. S7).

Suc export via SWEET9

SWEET9 is a uniporter (Lin et al., 2014; Eom et al.,2015); therefore, transport of Suc into nectar is contin-gent on the generation of a concentration gradientbetween the nectar-secreting cells and the nectar (anal-ogous to the apoplast). In species that produce a hexose-rich nectar, like Arabidopsis, this intracellular:apoplasticSuc gradient is likely generated by extracellular hy-drolysis of Suc by CWINV4 (Ruhlmann et al., 2010);however, species that produce a Suc-rich nectar cannotrely on complete hydrolysis of Suc to generate a Sucgradient. Our estimate of intracellular Suc concentra-tion ([Suc]; Fig. 3B) at the time of maximum nectar se-cretion (0 h; Fig. 3A)was remarkably similar to the [Suc]in the nectar (Fig. 3, B and C). If nectary and nectar [Suc]are highly similar, any additional Suc produced fromSuc synthesis in the nectary (or degradation of phloem-derived RFOs) during secretion would be transporteddown the Suc gradient and into the nectar. Taken to-gether, these data indicate that maintaining an in-tracellular [Suc] that is similar to the nectar may beimportant for sustaining Suc export in species thatproduce a Suc-rich nectar.Even though the [Suc] in the nectar and nectary are

similar, we did see a slightly lower [Suc] in the nectarycompared with the nectar (Fig. 3), which seeminglydetracts from our hypothesis of mass-flow transport viaSWEET9. However, it is possible that our estimate of[Suc] may be different than what is actually present inthe nectar-secreting cells. In determining nectar [Suc],we calculated cytosolic volume based on estimatesfrom scanning electron microscopy imaging data(Nepi et al., 1996) that reported the relative volumes ofvacuole, mitochondria, and plastids in C. pepo nectaryparenchyma cells (Supplemental Table S1). Our calcu-lations are thus partially dependent on the accuracy ofthose estimates. Additionally, different parts of nec-taries likely play different roles in nectar secretion. Nepiand others found that epidermal cells are predomi-nantly composed of vacuoles (which may be importantfor sugar storage), whereas parenchyma cells storedmost of the nectary starch (Nepi et al., 1996). Data fromNepi and others suggest that epidermal cells in C. peponectaries may be important for storing and eventuallyproducing the sugar destined for secretion in the nectar.Our estimate of nectary sugar concentration is based onwhole nectaries (epidermis and parenchyma). Becauseparenchyma cells near the epidermis (or possibly epi-dermal cells themselves) are likely the ones secreting








the nectar, it is possible that the [Suc] near or at theepidermis is actually much higher than what we mea-sured merely because we measured all parenchymacells (some of which may have a lower [Suc]), therebydiluting the [Suc] of the cells that are actively secretingnectar.

In Arabidopsis, the expression and activity of AtC-WINV4 is induced during secretion and is required forthe eccrine model of nectar secretion via SWEET9(Ruhlmann et al., 2010). Previous studies have shownthat CpCWINV4 is also highly expressed in C. peponectaries, although it shows a slightly differentexpression pattern than in Arabidopsis. CpCWINV4expression is decreased during secretion in C. peponectaries (Solhaug et al., 2019), suggesting that down-regulation of extracellular hydrolysis may be importantfor generating a Suc-rich nectar. However, previousresults have shown that C. pepo nectar likely containsendogenous invertase activity (Nepi et al., 2012), sug-gesting Suc hydrolysis may partially drive export ofnectar sugar. Our data suggest that from 0 h to13 h, thelevels of Glc in the nectar increase whereas the levels ofSuc stay the same (Fig. 3C). This increase in Glc is notaccompanied by a change in Suc (Fig. 3C), suggestingapoplastic (or nectar) Suc hydrolysis may be occurring.If extracellular hydrolysis by CpCWINV4 (or nectar-derived invertases) is important for nectar productionin C. pepo, it is surely to a lesser extent than in speciesproducing a hexose-rich nectar.

Recovery of Nectar Sugar by Resorption

Previous research has suggested that nectar sugarcan be resorbed from C. pepo flowers (Nepi et al., 2001).Recovery and re-utilization of nectar sugar may par-tially ameliorate the energetic cost of producing nectar.We show here that the [Suc] in both the nectar andnectary falls drastically throughout the day (Fig. 3C),and [Glc] may also decrease to a similar extent based onour measurements of nectar sugar in flowers at 124 hafter secretion (Supplemental Fig. S4A). We see sus-tained activity of SuSy and INV in the nectary post-secretion (19h; Fig. 4), suggesting these enzymes mayplay a role in breaking down nectar-derived Suc aftersecretion. Hexoses produced from Suc breakdown canthen be used for biosynthesis of new RFOs, or to pro-duce energy for senescence-related processes or growthof seeds in the case of female flowers. The exact mech-anism of nectar resorption and the role it plays inoverall plant health is currently unknown and requiresfurther testing.

The Role of Nonstarch Sugar in Nectar Production

Here we present numerous pieces of evidence tosuggest that direct transport of phloem sugar, withoutprior storage as starch, plays an important role inthe production of nectar in C. pepo. First, the total

sugar in the nectar and nectary system is substantiallymore than exists in nectary starch at the peak ofstarch accumulation. Second, nectary sink strength ismaintained throughout nectar secretion, showing thatimport of phloem sugars into nectaries likely remainsconstant during secretion. Finally, reducing the amountof phloem sugar coming to the nectary results in lowernectar sugar, further suggesting phloem sugars play animportant role in nectar production.

Total Sugar in Nectary and Nectar Exceed Total Starch

The similar timing of nectar sugar production andbreakdown of nectary starch (Fig. 3D) suggests thatnectar sugar may come from hexoses derived fromstarch. Conversely, Nicotiana nectaries have shown theability to uptake Suc from media during secretion,suggesting a bypass of nectary starch and direct trans-port of phloem sugar into the nectar is possible (Renet al., 2007). We have presented evidence that the totalsugar in the nectar and nectary system is substantiallymore than exists only in starch (Table 1).

Although these data suggest that at least some of thenectar and nectary sugar comes from sources other thanstarch, it is also possible that there are other forms ofcarbon that come from the starch that we did not detect.For instance, it is possible that other long-chain mal-toOS (Critchley et al., 2001) derived from starch arepresent, but were not measured in this study. Addi-tionally, it is possible that we may be underestimatingthe level of sugar in the nectary. For example, we didnotmeasure Fru, which likely represents a large portionof the total soluble sugar present in the tissue andnectar. However, assuming that at least some Fru ispresent, this would only increase the estimate of totalsugar (which is already higher than starch; Fig. 3E),lending more evidence to support the hypothesis thatphloem-derived sugar makes up a substantial portionof total system sugar.

Sink Strength Is Maintained in Nectaries duringNectar Secretion

We have presented evidence that activity of enzymesimportant for sink strength, including INV, a-GALase,and SuSy, do not decrease during secretion (Fig. 4),suggesting that sink status and sugar import is main-tained. In Arabidopsis, the import and degradation ofSuc from phloem during secretion is difficult to recon-cile with the mounting evidence that nectaries activelysynthesize Suc during secretion (Lin et al., 2014;Solhaug et al., 2019). However, the breakdown pro-ducts of starch (hexose-phosphates) can be incorpo-rated into Suc directly, preventing futile degradationand synthesis of Suc in the nectary. From an energeticsperspective, the current eccrine model in Arabidopsismay require a reduced role of direct phloem sugars


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(and an increased role of degraded nectary starch) inproducing nectar sugar.Although direct phloem sugars may be less impor-

tant for generating nectar sugar in species that trans-port Suc in their phloem, the story may be different forspecies that transport RFOs in their phloem, such asC. pepo (Beebe and Turgeon, 1992). Stachyose, thepredominant phloem sugar in C. pepo (Zhang et al.,2010), contains two galactosyl moieties attached toSuc (Holthaus and Schmitz, 1991). Stachyose degra-dation by a-GALase produces one Suc and two Gal,which can be converted into UDP-Glc (UDP-glc) by anumber of enzymes, of which UDP-Glc/UDP-Gal-4-epimerase (UGE) is an important step. CpUGE5 ex-pression is high during the build-up to secretion in C.pepo nectaries (Supplemental Fig. S7). Because the deg-radation of RFOs liberates hexoses without degradingSuc, import and degradation of phloem sugar can coexistwith nectary Suc synthesis without futile synthesis/degradation cycles. Even though our data suggest thatimport of phloem-derived sugars during secretion isimportant for nectar production in C. pepo, it is possiblethat other species (particularly those that transport Sucvia the phloem) might rely more on nectary starch togenerate nectar sugar due to the conflict betweenbreaking down incoming phloem Suc and synthesizingSuc from starch-derived hexoses.

Reducing Phloem Sugar Leads to a Reduction inNectar Sugar

We have shown that two methods of reducingphloem sugar lead to decreases in nectar sugar. Ex-posing the plants to prolonged darkness led to de-creased Suc and total sugar in the nectar (Fig. 5, D andF), likely due to reduced photosynthate from leaves andthus lower transport of phloem sugars to the nectary.Interestingly, extended night samples also had morenectary sugar than the control (Fig. 5F), suggesting thatsugars were being retained more by the nectaries inresponse to darkness. Removing the flowers from theplants led to somewhat similar responses, namely areduction in total system sugar (Fig. 6F) and a reduced[Suc] in the nectar (Fig. 6C). The removed (detached)flowers likely had a lower amount of phloem sugaravailable to them during secretion compared with theattached flowers, which likely caused the production ofnectar to be inhibited. Both of these results show thatreduction or removal of phloem-derived sugar nega-tively impacts nectar secretion, lending further supportto our hypothesis that import of phloem-derived sugarduring secretion is important for nectar production.One interesting finding from the flower removal

study is that the total sugar in the system is reduced by;200 mmol Glc eq in detached flowers versus attached(Fig. 6F), which is remarkably similar to the differencein starch content (also ;200 mmol Glc eq; Fig. 2A) be-tween215 h (when the flowers were excised) and26 h,the time of maximum nectary starch accumulation

based on our data. It is, therefore, not out of the realm ofpossibility that the detached nectaries were deprived offull starch accumulation, which may have led to thereduction in total system sugar and [Suc] in the nectar.However, the difference of 200 mmol Glc eq also existsbetween control and extended night samples (Fig. 5F).In this case, the flowers had been attached to the plantfor the whole night andwere likely able to fully fill theirnectaries with starch. Therefore, even though it is pos-sible that the flower removal experiment was also af-fecting the ability of nectaries to accumulate starch, thefact that we saw the same response in extended nightexperiment supports the important role of phloem Sucin maintaining high intracellular [Suc] and Suc exportby SWEET9.Although these data clearly suggest that sugar pro-

duced from starch is supplemented by sugar derivedfrom phloem in C. pepo nectaries during secretion, thefact remains that most of the sugar (;59%) in the nectarcomes from starch. The role of starch in nectar secretionmay be to facilitate rapid production of high concen-trations of intracellular sugar, which likely would notbe possible if C. pepo nectar was produced via onlyexport of phloem-derived sugar. Future experiments,possibly incorporating mutation of key genes such asBAM1 (Solhaug et al., 2019), may allow us to betterunderstand the extent towhich nectary starch facilitatessecretion of nectar sugar in C. pepo.

A Role for Trehalose and Sugar Signaling duringNectar Secretion

Trehalose-6 phosphate (Tre6P) is a well-studiedsignal of Suc status in plants (Figueroa and Lunn,2016). Here we analyzed the accumulation and ca-tabolism of trehalose and Tre6P throughout secretionin order to determine their role(s) in C. pepo nectarsecretion. Expression and activity of Tre6P- andtrehalose-degradation enzymes is induced duringsecretion, whereas trehalose and Tre6P levels are in-creased at the same time (Fig. 7). Additionally, trehalosetreatment leads to an inhibition of starch degradationand secretion of nectar sugar (Fig. 8), suggesting tre-halose may play an inhibitory role in nectar sugarproduction and secretion.Previous research has implicated Tre6P in regula-

tion of starch degradation and maintenance of Suchomeostasis in both source and sink tissues (Wingleret al., 2000; Debast et al., 2011; Martins et al., 2013;Nuccio et al., 2015; Figueroa et al., 2016; Figueroa andLunn, 2016). The rosette leaves of soil-grown Arabi-dopsis accumulate trehalose and Tre6P up to ;20nmol/gFW and ;0.5 nmol/gFW, respectively (Carilloet al., 2013), with potato tubers accumulating Tre6P tosimilar levels (nearly 1 nmol/gFW; Debast et al., 2011);however, the endosperm of developing wheat grainshave among the highest reported values for Tre6P (up to119 nmol/g FW; Martínez-Barajas et al., 2011). Ourmeasurements for trehalose and Tre6P inC. pepo nectaries






were considerably higher than these previous studies(;1 mmol/gFW for Tre6P,;0.2mmol/gFWforTre; Fig. 7).However, nectaries also accumulate substantially more Suc(270 mmol/gFW; Fig. 7) than tubers (17 mmol/gFW; Debastet al., 2011), Arabidopsis rosette leaves (;1-10 mmol/gFW;Carillo et al., 2013), and developing wheat endosperm(;74 mmol/gFW; Martínez-Barajas et al., 2011), whichmay account for the relatively high levels of trehaloseand Tre6P in squash nectaries. Alternatively, thethreshold at which trehalose and Tre6P impact meta-bolic processes may be higher in nectaries comparedwith other plant tissues, which is currently unknownbecause the role of trehalose and Tre6P metabolismin nectaries had not yet been investigated beforethis study.

Our findings from carbohydrate analyses of C. peponectaries support the predicted role of Tre6P in regu-lation of starch degradation and Suc homeostasis fromprevious studies (Debast et al., 2011; Nuccio et al.,2015). Starch degradation occurs predominantly from23 to 0 h (Figs. 2 and 3), and accumulation of nectarySuc occurs mostly from 26 to 23 h (Figs. 4 and 7). Al-though Tre6P increases drastically from 23 h to 0 h(Fig. 7E), this increase occurs after both starch degra-dation (Figs. 2 and 3) and Suc accumulation (Fig. 3;Supplemental Fig. S6) are mostly complete; any po-tential inhibitory effects from increased Tre6P likelywould not impact these processes. At 0 h, C. pepo nec-taries are likely transitioning from a “Suc-accumulatingstate” (a combination of Suc synthesis and breakdownof phloem-derived RFOs) to a “Suc-export state” inwhich Suc is exported into the nectar. One way thatTre6P (or possibly Tre) can reduce intracellular Suc levelsis by positively regulating Suc export (Lunn et al., 2014),although there is currently no evidence to support thishypothesis. It would be interesting to examine the role ofTre or Tre6P on regulation of the Suc exporter SWEET9 infuture studies in order to test the hypothesis that accu-mulation of Tre6P at 0 h leads to induction of Suc export inorder to reduce the levels of intracellular Suc in nectaries.

Although previous studies have shown that Tre6P,not trehalose, is the true signal for Suc-mediatedchanges in morphology and carbohydrate partitioning(Schluepmann et al., 2003), it is possible that Tre6P-independent signaling may also occur. In our study,exogenous trehalose treatment leads to inhibition ofnectary starch degradation (Fig. 8A) and lower nectarsugar (Fig. 8E), and only slightly higher Tre6P in nec-taries (Fig. 8D). However, there was significantlymore nectary trehalose in the trehalose-treated samplescompared with the sorbitol control (Fig. 8C). Becausewe did not see a significant difference in Tre6P inthe trehalose-treated sample compared with sorbitol(Fig. 8D), it is unlikely that increased Tre6P is causingthe reduced nectar in the trehalose-treated samples. Itshould also be noted that trehalose plays a highlyconserved role in the regulation of cellular responses toosmotic stress, ranging from bacteria to plants andanimals (Iturriaga et al., 2009; Chen et al., 2017). Al-though a molecular mechanism for the involvement of

trehalose within the context of our study is unclear,secretory nectaries are exposed to a tremendousamount of osmotic stress due to a high intracellularconcentration of Suc (Fig. 3B). It is thus possible thatexogenously applied trehalose could cause nectaries tohave an altered response in nectary starch degradationand nectar secretion via amechanism related to osmoticpotential. Finally, AtTRE1 (encoding Arabidopsis tre-halase) has been shown to be induced during carbonstarvation conditions (Garapati et al., 2015; Sun et al.,2019), which leads to activation of catabolic processessuch as starch breakdown (Garapati et al., 2015).However,AtTRE1 expression is unaffected by feeding ofSuc (Schluepmann et al., 2004), suggesting that AtTRE1may be specifically important for initiating low-carbonmetabolic responses (such as starch degradation).

While the simultaneous accumulationof bothTre/Tre6Pand their catabolic enzymes may seem counterintuitive,there is a well-known positive relationship between [Suc]and [Tre6P/Tre] (Yadav et al., 2014; Figueroa et al., 2016).Thus, the extremely high levels of Suc in nectaries maynecessitate the expression of Tre/Tre6P catabolic enzymesin order to prevent overaccumulation of Tre6P andTre. It ispossible that catabolism of Tre/Tre6P in C. pepo nectariesmay partially lessen the inhibitory effects that Tre6P andTrehaveonstarchdegradation (Martins et al., 2013;Garapatiet al., 2015) and osmotic stress tolerance (Van Houtte et al.,2013), respectively, during starch degradation and sugaraccumulation in C. pepo nectar-producing cells (Lin et al.,2014; Solhaug et al., 2019). Taken together, these data sug-gest a conserved role for trehalose and Tre6P in inhibitingbreakdown of starch and keeping Suc levels constant, link-ing carbohydrate partitioning and energy storage to theamount of Suc available. The exact mechanism of this re-sponse in C. pepo nectaries is currently being researched.

CONCLUSION

These results represent an important examination ofhownectar is produced froma biochemical andmetabolicperspective in squash. We have presented evidence thatnectary starch accumulation during the starch-fillingstage coincides with an increase in sink strength andsink activity. Nectary starch degradation is an importantstep in the secretion of nectar, and the regulation of thisprocess may be mostly light independent in squash. Wehave shown that Suc synthesis is active and inducedduring secretion; however, direct phloem sugar alsoplays an important role in determining the content andcomposition of nectar sugar in C. pepo. Finally, wehave shown that trehalose metabolism plays an im-portant role in C. pepo nectar production and carbonpartitioning. Our predicted model of nectar secretionin C. pepo (summarized in Supplemental Fig. S8) opensup numerous avenues for future research that willcontinue to improve our understanding of how nectaris produced, which may lead to important agronomicadvancements to improve yields in crops that rely onpollinator visitation.


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MATERIALS AND METHODS

Plant Material and Growth Conditions

Staminate (male) flowers fromCrookneck Yellow Squash (Cucurbita pepo vartorticollia) plants were used for all studies. Plants were grown either undergreenhouse conditions at.200mmol/m2s and 28°C to 30°C or on a growth rackat ;250 mmol/m2s and 24°C.

Description of Experimental Design and Replication

Unless otherwise stated, biological replicates were defined as flowers takenfromagroupof fourplants.All fourplantswere at the samedevelopmental stageand grownunder the same conditions. For eachfigure, the data represent resultsfrom one experiment with indicated biological replicates.

Excised Flower Treatment Experiments

For all excised flower treatment experiments, flowers were excised fromplants at 215 h and placed in 1 to 2 mL of solution overnight. After the initialcut, the peduncle was fully immersed in deionized (DI) water and a second cutwas made on the peduncle of the flower. This was done in order to avoidembolism on xylem vessels from the initial removal from the plant. Nectar andnectaries were harvested the following morning at;4 h after dawn (11 h, total;16-h treatment). For figures using excised flowers (Figs. 6 and 8), the flowerswere taken from a total of six plants.

Starch Assays

Total starchwasquantitativelydeterminedusingakit fromMegazyme (TotalStarch Assay Kit, K-TSTA). Samples were ground in 0.25 mL of 80% (v/v)ethanol, then an additional 0.25 mL of 80% (v/v) ethanol was added. Sampleswere incubated at 80°C for 5 min, and then an additional 0.5 mL of 80% (v/v)ethanol was added. Samples were centrifuged for 10 min at 1,800g, and su-pernatants were removed. The samples were resuspended in 1 mL 80% (v/v)ethanol followed by repeating centrifugation at 1,800g for 10 min and removalof supernatant. From each wash, 0.5 mL was saved for soluble sugar analyses(1mL total). After removal of soluble sugars, 300 mL of a-amylase (100 U/mL in100 mM sodium acetate buffer, pH 5.0) was added to each sample, followed byincubation at 100°C for 6 min with vortexing at 2-min intervals. Then 10 mL ofamyloglucosidase (AMG; 3,300 U/mL)was added to each sample, and sampleswere incubated at 50°C for 30min. Samples were made up to 1 mL final volumeand then centrifuged at 1,800g. Supernatants were then assayed for D-Glc usingthe Glc oxidase/peroxidase (GOPOD) method. To get the concentration ofD-Glc in the range of detection for theGOPODassay,248 through23 h sampleswere diluted 10-fold. Supernatant (33 mLl) was added to 1 mL of GOPOD so-lution (made according to kit protocol) in duplicate, and tubes were incubatedat 50°C for 20 min. Absorbance was read at 510 nm (ΔA510) using a visiblespectrophotometer. Starch content (percentage, w/w) was calculated using theequation for solid samples provided in the Megazyme protocol supplied withthe kit. These values were then converted to micromole of Glc eq per grams FWusing the ΔA510 for 100 mg of D-Glc standard and the molar mass of D-Glc.

Resistant starch was analyzed by protocol “c” in the total starch assayprotocol procedure for the Megazyme Total Starch Assay Kit (K-TSTA). Briefly,soluble sugars were removed as with the normal starch assay. A magnetic stirbar and 0.2 mL of 2 M KOH were added to each tube, and samples were incu-bated for 20min in an ice/water bath over a stirrer. Then 0.8mL of 1.2 M sodiumacetate (pH 3.8), 0.3 mL of a-amylase, and 0.01 mL of AMGwere added to eachsample, followed by incubation for 30 min at 50°C. Samples were made up to afinal volume of 10 mL, before Glc produced from the resistant starch was an-alyzed by using the same method as in the normal starch assay (GOPODmethod, described above).

Soluble Sugar Assays

Nectar and nectary sugars were quantified using an AmpRed/Glc oxidase/horseradish peroxidase method as described previously by Bethke and Busse(2008). Suc was measured by pretreatment of samples with invertase. Briefly,two 25mL aliquots of sugarwere prepared. To one (Atotal), 25mL of invertase [38U/mL in 80 mM sodium acetate (pH 4.8)] was added; to the other sugar aliquot(Aglc), 80 mM sodium acetate (pH 4.8) was added. All samples were then

incubated at 22° to 23°C for 15 min. Then 25 mL of sodium phosphate buffer(150mM, pH 7.4) was added to each sample. Finally, 25mL of Glc assaymixwasadded to each sample consisting of 400 mMAmp-red (stock solution dissolvedin dimethyl sulfoxide), 0.1 U of horseradish peroxidase and 0.1 U of Glc oxidase,and 34 mM sodium phosphate buffer (pH 7.4). Samples were incubated at 22° to23°C for 25 min before reading A570. Asuc was determined by (Atotal 2 Aglc).Absorbance readingswere comparedwith a standard curve of Glc (25–500mM),and Suc standardswere also included to ensure that Suc hydrolysis by invertasewas proceeding adequately.

Maltose was assayed by the same method as with Suc, but 25 mL maltase[20 U/mL in 25 mM NaPO4 (pH 7)] was included instead of invertase. Nectarysugar was assayed from ethanol extracts prepared as described above for thestarch assay. The concentration of nectary sugar in the ethanol extract wasmultiplied by the volume of the extract (2mL) to obtain themicromoles of sugarpresent in each sample, before normalization to the amount of FW added. Thetotal amount of sugar in the nectar was determined by multiplying the con-centration by the nectar volume (microliter) to get amount of sugar present,then dividing by FW to get the total amount of sugar in micromole per gramsFW.All samples were tested before being assayed to determine the dilution thatwould yield absorbances that were in the linear range of the sugar assay (be-tween 25 and 500 mM).

Trehalose Assay

Trehalose was assayed using a Trehalose assay kit (Megazyme, K-TREH).Briefly, 20 mL of sample was added to 20 mL of alkaline borohydride solution(10 mg/mL NaBH4 in 50 mM NaOH). Samples were then incubated at 40°C for30min to remove reducing sugars from samples. To the samples, 50mL of aceticacid (200 mM) was added and then 20 mL of buffer 1 (supplied with kit). Thissolution was used in the detection of trehalose. Due to the low concentrations oftrehalose in our samples, the method of the trehalose assay was modifiedslightly. Sugar sample (50 mL; from NaBH4 treatment) was added to 50 ml ofreaction mix containing 20 mL solution 1, 10 mL solution 2 (NADP1/ATP), 2 mLsuspension 3 (HK/G-6-PDH), 2–4mL suspension 4 (trehalase), and 14–16 mLDIwater. Trehalase (2 mL) was added when assaying trehalose-treated samples(Fig. 8C), and 4 mL of trehalase was added when assaying overnight samples(Fig. 7D) because 26 and 23 h had a much lower level of soluble sugars. Bothmethods produced similar results for secretory (0 h) flowers (;0.15 mmol/gFW). A negative (Glc) control was included with 2 mL of water added insteadof trehalase. The A340 was measured after 20 min, and the amount of trehaloseproduced was determined using a standard curve of trehalose (5–250 mM).

Assay for Malto-Oligosaccharides

Malto-oligosaccharides were assayed in ethanol extracts using enzymessupplied with the Megazyme starch assay kit described above. a-Amylase[100 U/mL in 100 mM sodium acetate (pH 5.0)] and AMG (3,300 U/mL) werecombined 1:1 and diluted 10-fold. Sugar (25 mL; diluted to ,500 mM Glc) wasadded to 25 mL of a-amylase/AMG enzyme mix; then samples were incubatedat 50°C for 30 min. Next, 25 mL of 150 mM NaPO4 (pH 7.4) was added, and then25 mL of AmpRed/GlcOx/HRP was added (components described above inthe “Soluble Sugar Assays” section) and Glc produced from degradation ofmaltoOS was assayed as described above (AmpRed/GlcOx/HRP method;A570).

Determination of Nectary Sugar Concentration

Nectary sugar concentration was determined by calculating nectary volumeanddividing the totalmicromoles in each sample by the calculated volume. Volumewas estimatedbyfirst determining radius of inner cup (ri) and the radius of the entirenectary (ro) and using these values to determine the volume of the nectary (VN)based on the equation provided in Supplemental Fig. S5. We also measured themass of the same nectaries and used the volume to calculate the density of thenectary. Using this density value,we estimated nectary volume based on FWvaluesthat we had measured for normalization of nectary sugar data.

We performed an additional correction in order to estimate the relativevolume of the cytosol, based on percentage of cell component estimates fromimaging studies done by Nepi et al. (1996). Briefly, we set a cell radius of 10 mm,and then calculated the volume based on estimates of 30% vacuole (r 5 3 mm),25%mitochondria (r5 2.5mm), and 25%plastid (r5 2.5mm) and subtracted the






volume of each from the total cell volume to get volume of cytosol(Supplemental Table S1).

Enzyme Activity Assays

All absorbances were measured on a BioTek PowerWave HT MicroplateSpectrophotometer. All enzymatic activity assays were normalized per gramsFW. For each enzyme activity assay,mass of tissue varied from 50 to 300mg. Fornectaries that were drastically (.2-fold) different in weight, heavier proteinsamples were prediluted accordingly before adding to the assay.

Amylase Assay

Total amylase activity was assayed as described previously by Laby et al.(2001), with a few alterations. Samples were ground in 150 mL Tris-HCl, pH 8.0,on ice and then centrifuged at ;17,000g for 20 min at 4°C. Supernatant (10 mL)was added to a new tube containing 150mL of 0.1 M sodium acetate (pH 4.6) and75 mL of amylopectin (20 mg/mL in 0.2 M KOH). Each reaction was then madeup to 300 mL with DI water then incubated at 37°C for 60 min. The reactionswere stopped by incubation at 100°C for 3 min. Ten microliters of each reactionwas added to 750 mL of p-hydroxybenzoic acid hydrazide solution, and DIwater was added to a final volume of 1 mL. Samples were incubated at 100°Cfor 5 min, and then A410 was read for each sample blanked against a reagentblank with Tris-HCl grinding buffer added in place of crude protein. A stan-dard curve of maltose (20–2500 mM) was used to express amylase activityas mmol of reducing sugar produced (representingmaltose and Glc) per minuteper gram FW. Amylopectin stocks were prepared by boiling amylopectin/KOH solution for 10min until the solution became clear, and then aliquots werestored at 220°C before use. p-hydroxybenzoic acid hydrazide solution wasprepared by mixing one part 5% (w/v) p-hydroxybenzoic acid hydrazide in0.5 M HCl with four parts 0.5 M NaOH.

a-Galactosidase Assay

a-Galactosidase activity was assayed as described previously in Miao et al.(2007), with some changes. Samples were ground in extraction buffer con-taining 50 mM HEPES-NaOH, 2 mM MgCl2, 1 mM EDTA, and 5 mM dithio-threitol (DTT). Samples were then centrifuged at 17,000g for 30 min at 4°C.Protein extracts were diluted 5-fold in extraction buffer for the neutral activityassay, and 2-fold for the acid activity assay. For the neutral assay, we usedHEPES-NaOH (pH 7.4), whereas for the acid assay, we usedHEPES-NaOH (pH5.5). Reaction mixtures contained 240 mL of 125 mM HEPES NaOH (pH 7.4 orpH 5.5), 30 mL of 20 mM paranitrophenyl galactopyranoside, and 30 mL of di-luted protein extract (300 mL total). Samples were then incubated at 37°C for20 min, and then reactions were stopped by addition of 1 mL of 5% (w/v)Na2CO3, which also caused color to develop. Absorbance was measured at410 nm and compared against a standard curve of 4-nitrophenol. Standardswere prepared by adding 1 mL of 5% (w/v) Na2CO3 to 300 mL of 4-nitrophenolat concentrations ranging from 25 to 5000 mM.

Invertase Assay

Invertase activity was assayed as described previously in Ruhlmann et al.(2010), with a few alterations. Protein was extracted by grinding nectary tissue(100–200 mg) in 250 mL of buffer containing 50 mM HEPES-NaOH (pH 8.0),5 mM MgCl2, 2 mM EDTA, 1 mM MnCl2, 1 mM CaCl2, 1 mM DTT, and 0.1 mM

phenyl-methyl-sulfonyl fluoride. Extracts were centrifuged at 17,000g at 4°C,and the supernatant was used in the assay. Soluble sugars were removed fromprotein extracts using a SpinOUT GT-600 (3 mL) desalting column (G-Biosci-ences). Briefly, columns were centrifuged for 1 min at 1000g to compact theresin, and then buffer was removed by centrifuging again at the same speed for2 min. The column was washed with five columns of protein extraction buffer,centrifuging for 2 min at 1000g. Extracts were applied to the columns, and thenthe columns were centrifuged at 1,000g for 6 min to collect the desalted extracts.Ten microliters of desalted extract was added to 90 mL of either acid [80 mMsodium acetate (pH 4.8), 5 mM Suc] or neutral [75 mM NaPO4 buffer (pH 7.4),5 mM Suc] assay solution. Samples were incubated at 30°C for 30 min, and thenGlc produced was quantified as described above (AmpRed/GlcOx/HRPmethod; A570). Acid assays were diluted 1:5, and neutral assays were mea-sured undiluted for the Glc assays.

SPS Assay

SPS activitywasmeasured as described previously in Stitt et al. (1988),with afew modifications. Nectaries were ground in 250 mL extraction buffer con-taining 50 mM HEPES-NaOH (pH 7.4), 4 mM MgCl2, 1 mM EDTA, 10% (v/v)glycerol, and 0.1% (v/v) Triton X-100. Extracts were then centrifuged at 17,000gat 4°C and then desalted as described in the “Invertase Assay” section toremove soluble sugars. Assay mixture contained 10 mL desalted protein, 50 mM

HEPES-NaOH (pH 7.4), 4 mMMgCl2, 1 mM EDTA, 10mMGlc-6-P, 2 mM Fru-6-P(Fru6P), and 2 mM UDP-Glc in a final volume of 100 mL. The reactions wereincubated at 22° to 23°C for 15 min before boiling the samples for 3 min to stopthe reaction. UDP formed from Suc-6-P synthesis was then quantified by anadditional reaction. This reaction contained (final volume 100 mL) 50 mM

HEPES (pH 7.4), 5 mM MgCl2, 0.3 mM NADH, 0.8 mM phosphoenolpyruvate,10 mL of initial SPS reaction, and 2 mL of pyruvate kinase/lactate dehydro-genase enzyme mix (900-1400 U/mL; Sigma). Reactions were incubated at 22°to 23°C for 30 min, and ΔA340 (end point) was measured and calculated from astandard curve of NADH (50–500 mM).

SuSy Assay

SuSy activity was assayed as described previously by Sun et al. (1992), withsome minor differences as outlined below. Nectary protein was extracted bygrinding in 300 mL extraction buffer containing 200 mM HEPES-KOH (pH 7.0),3mMMg acetate, 0.5mM EDTA, 0.5mM phenylmethyl sulphonyl fluoride, 5mM

DTT, and 20 mM 2-mercaptoethanol. Extracts were centrifuged for 15 min at17,000g and at 4°C, before being desalted as described above in the “InvertaseAssay” section. SuSywas assayed in the synthesis direction in a 100 mL reactioncontaining 100 mM Tris HCl (pH 8.5), 25 mM Mg acetate, 75 mM KCl, 0.2 mM

UDP-Glc, 4 mM phosphoenolpyruvate, 15 mM Fru, 0.3 mL of pyruvate kinase/lactate dehydrogenase mix (900–1400 U/mL, Sigma), and 10 mL of protein.Disappearance of NADH was monitored at ΔA340 per min for 5 min, and therate was calculated from a standard curve of NADH (50–500 mM).

Trehalase Activity Assay

Trehalase activity was assayed as described previously by Garapati et al.(2015), with some differences. Nectary tissue was ground in 400 mL of proteinextraction buffer containing 0.1 M MES-KOH (pH 6), 1 mM EDTA, 1 mM phenylmethyl sulphonyl fluoride, 1% (w/v) polyvinylpolypyrrolidone, and 1 mM

DTT. Protein extracts were desalted as described in the “Invertase Assay”section to remove Glc in the extract. Desalted protein extract (10 mL) was addedto 100 mL of reaction buffer containing 62.5 mM MES-KOH (pH 7), 125 mMCaCl2, and 100 mM trehalose. The reaction was brought up to 250 mL with DIwater, and then samples were incubated at 30°C for 30 min. Reactions werestopped by boiling for 3 min before assaying Glc produced from trehalase re-action using AmpRed/GlcOx/HRP method described above in the “SolubleSugar Assays” section. Reaction solution (25 mL) was added to 25 mL of 150 mM

NaPO4 (pH 7.4), 25 ml of DI water, and 25 ml of Glc assay solution. Glc wasquantified spectrophotometrically as described above in the “Soluble SugarAssays” section (AmpRed/GlcOx/HRP method; A570) and used to determinetrehalase activity.

Analysis of Gene Expression

Nectary RNA was extracted by Trizol method (Catalog #15596026), andcomplementary DNA was prepared using the Promega GoScript ReverseTranscription System (Catalog #A5000), with 1 mg of RNA used for comple-mentary DNA preparation. Expression of key genes in trehalose metabolismwas analyzed via RT-qPCR using Agilent Brilliant III Ultra-fast SYBR GreenQPCR Master Mix (Catalog #600882). Expression values are expressed as foldchange relative to the 0 h time point and are based on the ΔΔCt values obtainedfrom the normalized Ct values for each gene. Gene expression was normalizedto a gene encoding a RING (Really Interesting New Gene)/U-Box ligase su-perfamily protein (CpRING; C. melo hit5 gij65907532 8jrefjXM _008439865.1j).This gene was chosen as the internal reference based on its stable expressionlevel in all nectary samples in our RNA-seq dataset (Solhaug et al., 2019). Primersequences for each gene are provided below:CpRING, F5GGGGAAGCCCAAAGCAAAGCCATGA, R 5 GCCTTCGAGGAGGGGCTTGGC; CpTRE1, F 5ACCCTTGTGAGAGCCATTATC, R 5 GGAGGCTTCACTAGTCAGAAAG;


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CpTPP-J, F5 TCCCACTCTTTCTCTCTCTACTT, R5 CCTTCCGAGGTTATGCACTT.

Measurement of Tre6P by LiquidChromatography-Tandem Mass Spectrometry

Samples were extracted in 80% (v/v) ethanol as described in the “StarchAssays” section for soluble sugar analyses. Samples were diluted in 80% (v/v)acetonitrile at a 1:10 dilution. Samples (10mL) for Selective ReactionMonitoring(SRM) analysis were subjected to separation using a Shimadzu guard columnfollowed an analytical SeQuant ZIC-pHILIC column (15034.6mm; MilliporeSigma) at 30°C connected to the Applied Biosystem 5500 iontrap fitted with aturbo V electrospray source run in negative mode with declustering potentialand collision energies in Supplemental Table S2. The samples were subjected toa linear gradient of 80% B to 25% B over an 18 min gradient [A: 10 mM am-monium acetate (pH 7), 20% acetonitrile; B: 100% acetonitrile] at a column flowrate of 400 mL/min. The column was cleared with 25% B for 2 min and thenequilibrated to 80% B for 10min. Transitionsmonitored as in Supplemental TableS2 were established using the instrument’s compound optimization mode withdirect injection for each compound. The data were analyzed using MultiQuant(ABI Sciex Framingham) providing the peak area. A standard curve was con-structed using 10 mL. Samples were run in triplicate and concentrations deter-mined from the standard curve. Sample chromatograms of standards and asecretory stage (0 h) nectary are shown in Supplemental Figure S9.

Statistical Analysis

All statistical analyseswere performedusingR (RCore Team, 2014). For eachexperiment, a 1-way ANOVA was conducted in addition to a Tukey post hoccomparison of means. Statistical difference was determined at the P# 0.05 levelunless otherwise stated.

Accession Numbers

The accession numbers for genes analyzed in this study were derived fromthe recently published C. pepo genome (Montero-Pau et al., 2018), including:Trehalase 1 (CpTRE1), XM_023678639.1; Trehalose phosphate phosphatase-J(CpTPP-J), XM_023657393.1; Maltose excess protein 1 (CpMEX1), XM_023678269.1;a-glucan phosphorylase (CpPHS2), XM_023656828.1; ERD6-like transporter 6(CpEL6),XM_023690129.1;Disproportionatingenzyme2 (CpDPE2),XM_023687667.1;ERD6-like transporter 16 (CpEL16), XM_023670748.1; Hexokinase 1 (CpHXK1),XM_023690103.1; UDP-Glc/UDP-Gal-4-epimerase 5 (CpUGE5), XM_023697437.1.

Additional accession numbers for genes mentioned but not analyzed in thispaper are SWEET9 (CpSWEET9), XM_023681010.1; b-amylase 1 (CpBAM1),XM_023671176.1; Cell wall invertase 4 (CpCWINV4), XM_023689815.1; Sucphosphate synthase 3F (CpSPS3F), XM_023658084.1.

SUPPLEMENTAL DATA

The following supplemental materials are available.

Supplemental Figure S1. Representative floral stages and nectary masses.

Supplemental Figure S2. a-Galactosidase activity in floral tissues.

Supplemental Figure S3. Comparison of soluble and resistant starch inC. pepo nectaries.

Supplemental Figure S4. Nectar sugar composition at 0 and 124 h.

Supplemental Figure S5. Method for calculation of nectary volume anddensity.

Supplemental Figure S6. Accumulation of total Suc (nectary 1 nectar)during secretion in C. pepo.

Supplemental Figure S7. Gene expression for select genes involved incarbohydrate metabolism during nectar secretion in C. pepo.

Supplemental Figure S8. Predicted model of enzymes and pathways in-volved in carbohydrate metabolism in C. pepo nectaries.

Supplemental Figure S9. Sample chromatogram of Tre6P and Suc6P.

Supplemental Table S1. Estimation of relative cytosolic volume for C. peponectary cells.

Supplemental Table S2. Transitions for Tre6P in LC MS/MS analysis.

ACKNOWLEDGMENTS

The authors thank Neil Olszewski, Adrian Hegeman, and John Ward of theUniversity of Minnesota for helpful comments on this work and BruceWitthuhn and the University of Minnesota Center for Mass Spectrometry andProteomics for assistance with Tre6P analyses.

Received April 17, 2019; accepted June 5, 2019; published June 18, 2019.

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carbohydrate metabolism and signaling in squashhexoses (davis et al., 1998). other species, such as...

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