Skip to main content
NCBI home page
Plant Physiology logoLink to Plant Physiology
. 2017 Aug 31;175(2):734–745. doi: 10.1104/pp.17.00995

Etiolated Stem Branching Is a Result of Systemic Signaling Associated with Sucrose Level1

Bolaji Babajide Salam a,b,2, Siva Kumar Malka a,2, Xiaobiao Zhu c,2, Huiling Gong c,d, Carmit Ziv a, Paula Teper-Bamnolker a, Naomi Ori b, Jiming Jiang c, Dani Eshel a,3
PMCID: PMC5619910  PMID: 28860154

Hot or cold storage of potato tuber, a swollen stem, induces sucrose accumulation in the parenchyma that enhances branching during sprouting under dark conditions.

Abstract

The potato (Solanum tuberosum) tuber is a swollen stem. Sprouts growing from the tuber nodes represent loss of apical dominance and branching. Long cold storage induces loss of tuber apical dominance and results in secondary branching. Here, we show that a similar branching pattern can be induced by short heat treatment of the tubers. Detached sprouts were induced to branch by the heat treatment only when attached to a parenchyma cylinder. Grafting experiments showed that the scion branches only when grafted onto heat- or cold-treated tuber parenchyma, suggesting that the branching signal is transmitted systemically from the bud-base parenchyma to the grafted stem. Exogenous supply of sucrose (Suc), glucose, or fructose solution to detached sprouts induced branching in a dose-responsive manner, and an increase in Suc level was observed in tuber parenchyma upon branching induction, suggesting a role for elevated parenchyma sugars in the regulation of branching. However, sugar analysis of the apex and node after grafting showed no distinct differences in sugar levels between branching and nonbranching stems. Vacuolar invertase is a key enzyme in determining the level of Suc and its cleavage products, glucose and fructose, in potato parenchyma. Silencing of the vacuolar invertase-encoding gene led to increased tuber branching in combination with branching-inducing treatments. These results suggest that Suc in the parenchyma induces branching through signaling and not by excess mobilization from the parenchyma to the stem.

Plant shoot branching is determined by apical dominance (AD), a process in which the apical bud (shoot tip) inhibits the outgrowth of axillary buds farther down the stem to control the number of growing branches (Phillips, 1975). In response to this inhibition, plants have evolved rapid long-distance signaling mechanisms to release axillary buds and replenish the plant with new growing shoot tips (Mason et al., 2014). Since shoot branching is an important productivity-associated trait in agriculture, understanding the mechanism of AD is essential to breeding for and manipulating this yield-related characteristic.

AD and other forms of correlative bud inhibition have long been considered to be controlled by hormones, the plant’s nutritional status, or their interaction (Phillips, 1975). The classical hormone hypothesis proposes that the apical bud is the source of the correlative hormonal signal (auxin) that moves down the stem and restricts the development of axillary buds (Sachs and Thimann, 1964; Martin, 1987; Cline, 1994). Research centering on this hypothesis has shown that auxin moves throughout the plant and interacts with two other hormones, cytokinin, a promoter of bud outgrowth, and strigolactone, a branch-inhibiting hormone, forming a network of systemic signals that control lateral bud activation (Gomez-Roldan et al., 2008; Ferguson and Beveridge, 2009; Domagalska and Leyser, 2011; Dun et al., 2012). The nutrition hypothesis assumes that access to plant nutrients is the major factor regulating axillary bud growth (Phillips, 1975; Van den Ende, 2014; Buskila et al., 2016). Research centering on this hypothesis has shown that varying nitrogen supply can control the degree of AD (McIntyre, 1987, 1997), with nitrogen limitation delaying the activation of axillary buds (de Jong et al., 2014). This hypothesis has been narrowed down to the sugar nutrients, proposing that AD is maintained largely by the sugar demand of the shoot tip, which limits the amount of sugar available to the axillary buds (Mason et al., 2014; Rameau et al., 2015).

Sugars are the primary energy source produced by green plants via the process of photosynthesis. Small-molecule sugars also are known to act as signaling molecules in many physiological processes (Smeekens et al., 2010; Granot et al., 2013). From the site of their synthesis (source tissues), sugars are partitioned to sink tissues via the vascular system in a controlled manner. Buds are actively growing plant organs that act as strong sinks for sugars to meet their metabolic demands and support their growth. The growth capacity of the bud depends on its sink strength in terms of its ability to acquire and use sugars. Thus, buds have to compete for the sugars, which constitute the main source of carbon and energy (Maurel et al., 2004a, 2004b). The association between bud outgrowth and the mobilization of starch reserves in stem tissues has been well documented, especially in perennial plants (Decourteix et al., 2008; Rameau et al., 2015). High activity of sugar-metabolizing enzymes leads to increased sugar absorption in the bud (Marquat et al., 1999; Maurel et al., 2004a; Decourteix et al., 2008; Girault et al., 2010; Rabot et al., 2012). Plasma membrane H+-ATPase activity results in an electrochemical gradient that is required for H+/nutrient cotransport (Gévaudant et al., 2001; Alves et al., 2007; Pedersen et al., 2012). Recent studies have shown that buds compete for sugars during their development; in this process, apical buds acquire available sugars intensively over the axillary buds, which results in the sugar limitation related to AD (Morris et al., 2005; Mason et al., 2014).

The potato (Solanum tuberosum) tuber is a swollen underground stem formed by swelling of the subapical underground stolons (Harris, 1992). During tuber development, a growing number of axillary buds (termed eyes) are formed in a spiral arrangement on its surface, while the apical bud is located at the tip of the stolon (Goodwin, 1967). During sprouting, the tuber exhibits an AD phenomenon, similar to a normal stem (Eshel and Teper-Bamnolker, 2012; Teper-Bamnolker et al., 2012; Eshel, 2015). During tuber development, the storage parenchyma converts soluble assimilates (i.e. Suc and amino acids) into polymeric reserves (starch and storage proteins; Prat et al., 1990; Visser et al., 1994), which must be converted into transport-compatible solutes for sprouting initiation and growth (Sonnewald, 2001; Viola et al., 2007). Since the potato tuber exhibits classical stem behavior, apart from being a nutrient reservoir, it is an ideal model in which to study the role of sugars in AD and shoot branching under dark, etiolated conditions (Buskila et al., 2016). In this study, tubers and detached stems were induced to branch by temperature treatments. Branching was induced by exogenous sugar application as well. Using a grafting technique, we demonstrated that the branching signal is systemic in nature and correlated to parenchyma sugar level. Finally, we demonstrate that by silencing the gene encoding vacuolar invertase (VInv) in transgenic potato plants, we can induce overbranching of the tuber.

RESULTS

Heat Treatment Induces Stem Branching

The potato tuber, a swollen stem, stored at 14°C tends to sprout, when dormancy is released from the apical bud, in an AD form (Teper-Bamnolker et al., 2012). Sprouts growing from the tuber nodes (eyes) indicate loss of AD (Eshel, 2015). Seeking environmental factors that induce loss of AD, we heated nonsprouting tubers, after incubating them at 14°C for 4 to 5 weeks, when they are developmentally in shallow dormancy. Heating was performed at 33°C for 21 d, and then they were incubated at 14°C for sprouting. Sequential heating and 14°C incubation induced loss of AD and the growth of several stems simultaneously 2 weeks after heating, as compared with one apical stem in the nonheated tubers stored at 14°C for an additional 30 d (Fig. 1, A1 and B). These results suggested that tuber branching can be induced by short heating, similar to the branching induced by long cold storage. Heating of the sprouting tubers induced secondary branching (loss of AD type III; Teper-Bamnolker et al., 2012) on the growing stems as well, whereas the nonheated tubers produced only aerial roots (Fig. 1A2). Interestingly, tubers sprouted only after transferring them to 14°C and not during the heating process, which inhibited sprouting similar to cold storage (4°C).

Figure 1.

Figure 1.

Open in a new tab

Heat induces branching in a parenchyma-dependent manner. For heat treatments, tubers were stored at 14°C for 4 to 5 weeks after harvest, and tubers or detached sprouts were incubated at 33°C in the dark and 95% relative humidity for 21 d and then incubated at 14°C. A, Photographs of tubers (1 and 2) or detached buds (3 and 4) with or without parenchyma base, respectively. Top row, Nonheated; bottom row, heated. Asterisks indicate lateral bud growth from branching points of the tuber or stem. Bars = 1 cm. B, Number of buds induced to sprout over time after heating tubers. C, Heating-induced branching only in detached buds with parenchyma. Different letters represent significant differences between heated and nonheated stems, with or without parenchyma (P < 0.05). Data are means ± se of three repeats, each with 30 tubers/stems.

To examine whether the tuber parenchyma is essential for inducing stem branching, we heated detached sprouts with or without a cylinder of tuber parenchyma attached to their base. The sprouts, collected from tubers stored for 8 weeks at 14°C, were heated at 33°C for 7 d and then incubated at 14°C for further growth. Interestingly, detached sprouts were induced to branch only when the parenchyma cylinder was attached to their base (Fig. 1, A3 and C). Sprouts that were not attached to parenchyma produced only etiolated leaves on their stems (Fig. 1, A4 and C), suggesting that the tuber parenchyma is essential for bud branching following heat treatment. Nonheated stems tended to elongate much more in the presence of parenchyma at their base, but with no branching (Fig. 1A, 3 and 4).

Figure 3.

Figure 3.

Open in a new tab

Effects of in vitro sugar feeding on branching of detached tuber sprouts. Sprouts were detached from the tubers and supplemented with sugars (Suc, Glc, Fru, sorbitol, or mannitol) at different concentrations (0, 25, 50, 100, 300, and 600 mm) for 10 d at 14°C in 95% relative humidity under dark conditions. A, Number of branches after 10 d of treatment. Data represent averages of three experiments (seven replicates per treatment). B, Images showing sprouts with or without branches after 5 d of treatment with 300 mm solution. Bars = 100 µm. C, Content of simple sugars in stems fed with 300 mm Suc for 10 d. Sprouts were detached from tubers and supplemented with 300 mm Suc for 10 d at 14°C, in 95% relative humidity, under dark conditions. HPLC measurements of simple sugars were performed with samples extracted from the node and apex of the sprout. Data represent averages of three experiments, each performed with five replicates per treatment. Error bars represent se. FW, Fresh weight.

Figure 4.

Figure 4.

Open in a new tab

Contents of simple sugars in stems grafted on heated potato tubers. Sprouts from tubers stored at 14°C exhibiting AD were grafted onto heated tubers (33°C for 21 d) and vice versa, in two different combinations, namely, nonheated scion on heated stock (NH/H) and heated scion on nonheated stock (H/NH). Two other combinations, heated scion on heated stock (H/H) and nonheated scion on nonheated stock (NH/NH), served as controls. Grafted organs were then exposed to 14°C for 15 d. Data are means ± se of experiments done in 15 repeats. Different letters represent significant differences between treatments in each time point (P < 0.05). FW, Fresh weight.

Branching Signal Is Transmitted Systemically from Tubers

We performed grafting experiments to examine whether the signal for branching is systemic and can affect any bud attached to the treated parenchyma. Tubers or detached buds with a parenchyma base, which were heated at 33°C for 21 or 7 d, respectively, were used as stocks (the bottom part) for grafting. After treatment, they were incubated at 14°C for 1 to 2 weeks, until the bud was a few centimeters in length. Nonbranching buds detached from nontreated tubers were used as scions. Grafted organs were incubated at 14°C for 30 d. We investigated the translocation of solute after grafting using carboxyfluorescein diacetate (CFDA) at the base of the graft union. Intense fluorescent labeling in the vascular bundles of the scion 7 and 10 d after grafting confirmed phloem transport (Viola et al., 2007; Supplemental Fig. S1).

The scions were induced to branch only when they were grafted on previously heated tubers or parenchyma (Fig. 2A). The pattern and degree of branching of stems grafted on heated tubers or parenchyma were similar to those of the attached stems (Fig. 2B). These results showed that, upon heating, the signal for branching was produced in the parenchyma located under the stem and was transmitted to the scion to induce branching.

Figure 2.

Figure 2.

Open in a new tab

Systemic branching signals transmitted to the grafted stem. A, Before treatment, tubers were stored at 14°C for 4 to 5 weeks after harvest. Tubers (left) or detached buds with a parenchyma base (right) were heated at 33°C for 21 d and then used as stocks for grafting. Nonheated tubers were used as controls. Nonbranching buds detached from nontreated tubers were used as scions and grafted on the previously heated part. After grafting, organs were incubated at 14°C for 30 d. Arrows indicate the grafting point. Bars = 1 cm. B, Percentage of branching in the indicated treatments. Data are means ± se of three repeats, each with 30 tubers/stems. Different letters represent significant differences between heated and nonheated in grafted or nongrafted stems (P < 0.05).

Exogenous Sugars Induce Etiolated Stem Branching

We assumed that the products of starch degradation, specifically Suc and its hydrolytic products, would induce stem branching. We tested this hypothesis by replacing the tuber parenchyma with a solution containing Suc and its hydrolytic products, or sugar alcohols (mannitol and sorbitol), or water as alternative sugars and control, respectively. Tubers were incubated at 14°C until sprouting, and sprouts with two to three nodes were then detached manually, placed in Suc, Glc, Fru, sorbitol, or mannitol solution at six different concentrations each (0, 25, 50, 100, 300, and 600 mm), and incubated at 14°C for 10 d. Suc and its cleavage products Glc and Fru induced branching in a dose-dependent manner (Fig. 3, A and B). Branching was not induced by sorbitol, mannitol, or water (Fig. 3, A and B). These results suggested that branching induction might be due to Suc and its degradation products (Glc and Fru). To determine whether the effect is due to the transport of Suc or its hydrolytic hexose to the apical meristem and the branching node, stems were fed with 300 mm Suc for 10 d. Sugar levels were quantified in the apex and nodes at several time points during this experiment. During the 10 d of feeding, Suc, Glc, and Fru contents decreased gradually in the apex and Suc degradation products increased in the node (Fig. 3C). These results suggest that Suc from the potato parenchyma induced the branching of etiolated stems but likely did not transport into the stem as a whole molecule. To test this hypothesis, we fed radioactively labeled sugars to stems attached to the parenchyma cylinder. After 14 h of incubation, we were able to detect about 98% of all labeled sugars in the upper part of the parenchyma cylinder (stem base) but only 1.6% and 2.3% of the labeled Suc at the node and apex, respectively. The average amount of Glc and Fru was 0.2% and 0.4% at the apex and the node, respectively, suggesting a blockage at the stem vascular system (Table I).

Table I. Distribution of labeled sugar after feeding of detached stems with [U-14C]Suc, [U-14C]Glc, or [U-14C]Fru for 14 h under the dark condition.

Percentage of label indicates the label recovered in any particular tissue out of the total labeled sugar measured in the treated stem. Each value is the mean of three independent replicates ± se. Different letters indicate significant differences (P < 0.05) among treatments by Tukey-Kramer and ANOVA tests.

Tissue Suc Glc Fru
%
Apex 2.321 ± 0.001 b 0.233 ± 0.001 b 0.251 ± 0.001 b
Node 1.635 ± 0.001 b 0.371 ± 0.001 b 0.425 ± 0.001 b
Parenchyma 96.042 ± 0.005 a 99.394 ± 0.007 a 99.481 ± 0.001 a
Open in a new tab

Feeding of potato stems with palatinose or deoxyglucose, nonmetabolizable analogs of Suc and Glc, resulted in lower and no effect on stem branching, respectively (Supplemental Fig. S3). The palatinose effect was significantly higher than water application but lower than Suc, whereas deoxyglucose was phytotoxic and induced no branching (Supplemental Fig. S3). This experiment strengthened our conclusion that the Suc molecule may signal branching without the need to be metabolize.

Heat Treatment Induces Elevated Sugar Levels in Tuber Parenchyma

To test whether sugars are involved in the systemic signal transmitted from the stock to the scion, we examined whether sugars accumulate in heated tubers and whether they are transported from the parenchyma to the grafted scion. Heated (33°C for 21 d) or nonheated (14°C for 8 weeks) sprouts were detached from their original tuber and grafted onto heated or nonheated tubers in a total of four combinations. Following incubation at 14°C for 15 d, sugar levels were analyzed in the parenchyma and in two parts of the grafted scion: node and apex. Sugar quantification revealed significantly higher levels of Suc in the bud-base parenchyma of heated tuber stocks, which induced branching, compared with bud-base parenchyma of nonheated stock, which did not induce branching (Fig. 4). In contrast, Glc and Fru quantification in the bud-base parenchyma revealed no differences between heated and nonheated tubers (Fig. 4). Surprisingly, no correlation was found between sugar levels in the scion (node or apex) and the degree of branching after grafting in any of the treatments (Fig. 4). These results suggested that the main effect of heat treatment is on parenchyma Suc level; sugar transport to the scion in correlation with branching phenotype was not detected.

Long Cold Storage Induces Elevated Sugar Levels in the Parenchyma Correlated with Tuber Branching

Previous studies have shown that long cold storage of tubers also induces multiple stems and secondary branching of the growing stems (Teper-Bamnolker et al., 2012; Supplemental Fig. S2). Therefore, we utilized long cold storage to further understand the mechanism of stem branching. Sprouts from noncooled tubers (stored at 14°C for 8 weeks) or cooled tubers (90 d at 4°C) were detached from their original tuber and grafted onto noncooled or cooled tubers. Grafted organs were then incubated at 14°C for 20 d. Interestingly, branching was observed in the scion only when the stock was a cooled tuber, whereas the other graft combinations showed reduced or no branching (Fig. 5).

Figure 5.

Figure 5.

Open in a new tab

Long cold storage as an inducer of branching. Sprouts from tubers stored at 14°C (noncooled [NC]) and exhibiting AD were grafted onto long cold-stored tubers (cooled [C]) and vice versa, in two different combinations, namely, NC scion on C stock (NC/C) and C scion on NC stock (C/NC). Two other combinations, NC/NC and C/C, served as controls. Grafted organs were then exposed to 14°C for 20 d. A, Images showing the outcomes of the four different grafting combinations. B, Number of branches observed after grafting. Values for each combination and time interval represent averages of 10 repeats. Error bars represent se.

We observed significantly higher Suc and reducing sugar levels in the bud-base parenchyma of cooled stock, correlating with the branching phenotype, in comparison with noncooled tuber stock, which showed a nonbranching phenotype (Fig. 6). Surprisingly, we did not detect differences in sugar levels in the node or apex tissues in either branching or nonbranching combinations (Fig. 6), similar to the results with the heat-induced tubers. These results supported that there is no detectible transport of Suc, Glc, or Fru from the parenchyma to the grafted scions when a branching phenotype is exhibited. The Suc in the parenchyma thus likely induces branching via a downstream signal rather than being mobilized from the parenchyma to the stem.

Figure 6.

Figure 6.

Open in a new tab

Contents of simple sugars in stems grafted on potato tubers treated by long cold storage. Sprouts from tubers stored at 14°C (noncooled) exhibiting AD were grafted onto long cold-stored tubers (cooled) and vice versa, in two different combinations, namely, noncooled scion on cooled stock (NC/C) and cooled scion on noncooled stock (C/NC). Two other combinations, NC/NC and C/C, served as controls. Grafted organs were then exposed to 14°C for 20 d. HPLC measurements of simple sugars were performed with samples extracted from the parenchyma stock and node and apex of the scion. Data are means ± se of experiments done in 15 repeats. Different letters represent significant differences between treatments in each time point (P < 0.05). FW, Fresh weight.

Invertase Silencing Induces Tuber Branching

Since only the level of Suc was found to be affected by both branching inducers (i.e. hot and cold treatments), we analyzed the effect of altered Suc level on branching phenotype. Vacuolar invertase is a key enzyme in determining the level of Suc and its cleavage products, Glc and Fru, in potato parenchyma. Silencing of VInv results in effective prevention of Suc degradation (Bhaskar et al., 2010). To investigate the involvement of Suc in branching, we tested the effect of VInv silencing on branching phenotype using the treatments described above. We developed a set of five VInv-silenced lines of potato ‘Russet Burbank’ (RBK) using RNA interference (Zhu et al., 2014, 2016). The VInv gene in these five lines (RBK1, RBK22, RBK25, RBK27, and RBK46) showed different levels of silencing. Cold-stored tubers from these lines accumulate high levels of Suc and reduced levels of Glc and Fru (Zhu et al., 2014). Thus, these lines provide an ideal set of materials in which to study the relationship between Suc and tuber branching. By silencing VInv, we expected to reduce Glc and Fru elevation during hot and cold treatments and induce the accumulation of Suc in the tuber parenchyma. Then, if Suc is indeed involved in branching, we would expect the silenced lines to branch more than the wild type.

We conducted a branching study using the five RBK RNA interference lines as well as an RBK control line. Following branching-induction treatments (heat at 33°C for 3 weeks or storage at 4°C for 3 months), parenchyma sugars and tuber branching were found to be correlated to the level of VInv silencing among the RBK lines (Figs. 7 and 8). Our measurements, in accordance with previous studies, showed that VInv silencing correlated with higher Suc and lower hexose contents in the tuber parenchyma, especially after cold storage (Figs. 7 and 8; Zhu et al., 2014, 2016). Whereas the wild type and a weakly silenced line (RBK46; 21% silencing) produced one to two stems after induction, strong VInv-silenced lines (RBK22 and RBK1; 62% and 74% silencing, respectively) produced three to six stems (Figs. 7 and 8). These results further support that parenchyma Suc level during cold storage can serve as a predictor of stem number after replanting.

Figure 7.

Figure 7.

Open in a new tab

VInv silencing leads to increased parenchyma Suc content and stem sprouting following heat treatment. A, RBK lines were stored at 33°C for 21 d, and then the parenchyma of the apical bud base was used for sugar measurements. Each value is the mean of five independent replicates. Percentages in parentheses represent degrees of VInv silencing in tubers compared with the control RBK (0%). Error bars represent se. Different letters represent significant differences between transgenic lines in each sugar type (P < 0.05). Fw, Fresh weight. B, Tubers were transferred to 14°C for 13 weeks (until sprouting) under dark conditions (left column) or grown in the greenhouse (22°C; right column). Numbers on the pots represent the numbers of stems that sprouted from the planted tuber.

Figure 8.

Figure 8.

Open in a new tab

VInv silencing affects sugar levels and increases the number of stems following cold storage. A, RBK lines were stored at 4°C for 3 months, and then the parenchyma of the apical bud base was used for sugar measurements. Each value is the mean of five independent replicates. Percentages in parentheses represent degrees of VInv silencing in tubers compared with the control RBK (0%). Error bars represent se. Different letters represent significant differences between transgenic lines in each sugar (P < 0.05). Fw, Fresh weight. B, Tubers were transferred to 14°C for 7 weeks (until sprouting) under dark conditions (left column) or grown in the greenhouse (22°C; right column). Numbers on the pots represent the numbers of stems that sprouted from the planted tuber.

DISCUSSION

Parenchyma Is the Source of the Branching Signal

As already noted, the potato tuber is a swollen stem. The apical bud usually sprouts in an AD manner after physiological dormancy is released. Long cold storage usually leads to bud burst, loss of AD, and branching of multiple shoots (Teper-Bamnolker et al., 2012). In this study, heating of dormant tubers followed by their incubation at 14°C induced bud burst and loss of AD in the sprouting tuber, similar to the effect of long cold storage (Fig. 1). These results are in agreement with an earlier report on loss of AD in the tuber upon short-term exposure to a range of high temperatures (30°C–50°C) in potato (Juknevičienė et al., 2011). Taken together, these observations indicate a general role for high-temperature stress in lateral bud burst and loss of AD. Exposure of stems to extreme temperatures has been shown previously to induce early bud burst in tree species such as apple (Malus domestica; Wang and Faust, 1994), red-osier dogwood (Cornus sericea; Shirazi and Fuchigami, 1995), poplar (Populus nigra; Wisniewski et al., 1997), and nectarine (Prunus persica; Yue et al., 2013). Induction of primary and secondary branching upon prolonged exposure to high temperature (32°C) also has been observed in Arabidopsis (Arabidopsis thaliana; Antoun and Ouellet, 2013).

The branching signal is induced in the tuber parenchyma and transmitted systemically to the bud (Figs. 2 and 5). Grafting is often used as a tool to study systemic or long-range signaling processes in plants (Melnyk and Meyerowitz, 2015). For example, grafting of the branching mutants rms1, rms2, or rms3 (with increased bud outgrowth) in pea (Pisum sativum; Beveridge et al., 1994, 1997; Morris et al., 2001) and daad1 (decreased AD) in petunia (Petunia hybrida; Napoli, 1996) onto wild-type plants revealed the systemic nature of the branching signals in these species. Using Y-shaped grafts with wild-type and mutant shoots on a mutant rootstock revealed that, in pea, the branching signal seems to move only acropetally in shoots (Foo et al., 2001). This is consistent with the notion that sprouting of tuber buds is regulated by hormones within the tuber (Hemberg, 1985; Turnbull and Hanke, 1985; Suttle, 2004). Shoot branching is well accepted to be regulated primarily by a systemic network of plant hormones such as auxin, cytokinin, and strigolactones (Domagalska and Leyser, 2011).

Nutrients have been proposed as a possible signal for stem branching in plants. Simple mature leaf removal before decapitation in pea and selective defoliation in sorghum (Sorghum bicolor) demonstrated an absolute requirement for photoassimilates or endogenous sugars for loss of AD (Mason et al., 2014). Using an elegant set of experiments, Mason et al. (2014) demonstrated the systemic movement of Suc as a branching signal from the leaf to the lateral bud after decapitation. Up-regulation of Suc-starvation genes upon selective defoliation in the inhibited lateral buds of sorghum indicated the potential systemic movement of sugars between leaves and the lateral buds (Kebrom and Mullet, 2015). In potato tuber sprouting in the dark, removal of the parenchyma may leave the lateral buds with limited available nutrients, resulting in no bud outgrowth (according to the nutrient hypothesis; Phillips, 1975). Our grafting experiments confirmed that loss of AD may be regulated by a systemic branching signal originating from the tuber bud base. This hormone-like signal is transmitted to the lateral buds and activates their outgrowth.

Suc as a Potential Branching Signal

Similar to our findings, previous studies reported that cold storage of tubers induces Suc synthesis and elevates the levels of reducing sugars (Wiltshire and Cobb, 1996; Sowokinos et al., 2000). The elevation of Suc in the parenchyma bud base after both branching-induction treatments suggests a role for Suc as a branching signal. However, it was unclear whether sugars that accumulate in the parenchyma bud base following branching-induction treatments (long cold storage and heat) move upward to the sprout (scion). In our study, no clear disparities were noted in sugar levels between the stems that were induced to branch and those that were not (Figs. 4 and 6). These results suggest that the high level of Suc in the parenchyma is likely the branching inducer, but probably through signaling rather than excessive mobilization from the parenchyma to the stem. Therefore, even if the input flow is important, we do not necessarily expect an increase of the sugar levels in the sink. It should be noted that the inducing effect is probably not limited to Suc (Fig. 3) and that the stem-branching inducers could be, for example, hexoses or the Suc-to-hexoses ratio. Treatment with a nonmetabolizable analog of Suc showed a partial increase of branching as compared with Suc (Supplemental Fig. S3), suggesting the existence of a branching signal that is not necessarily related to Suc metabolism. 2-Deoxyglucose, a nonmetabolizable sugar analog for Glc, induces phytotoxicity in the apical meristem tissue, probably because it inhibits the glycolytic pathway of the cells (Shim et al., 1998). Future studies should consider screening the effect on branching of various nonmetabolizable analogs of Suc, such as lactulose and turanose, and of Glc, such as Man and 3-O-methylglucose.

Similar results were found in the herbaceous perennial Gentiana spp. plants during ecodormancy, where gentiobiose was found to be not seldom used as an energy source but is involved in signaling pathways (Takahashi et al., 2014). Feeding of detached stems with Suc showed an increase of its degradation products (Glc and Fru) specifically in the node, suggesting a rapid metabolism (Fig. 3C). However, feeding the bud-base parenchyma with labeled sugars showed that a very limited fraction of the Suc and hexoses is transported to the stem (Table I). Thus, Suc may effect etiolated stem branching by transport and be rapidly metabolized in the growing buds and/or may signal for stem branching through other secondary messengers. The requirement of sugars for bud outgrowth also was reported in Rosa hybrida, where a sugar supply was necessary to trigger bud outgrowth of in vitro-cultivated single nodes (Rabot et al., 2012; Barbier et al., 2015a). It was reported that Suc can modulate the dynamics of bud outgrowth in a dose-dependent manner, particularly the transition phase between bud release and sustained bud elongation (Barbier et al., 2015a). Moreover, different nonmetabolizable Suc analogs (palatinose, turanose, lactulose, and melibiose) also could trigger such outgrowth, suggesting that Suc can play a signaling role, as well as a nutritional role, in this process. This signaling effect also was reported with psicose, a nonmetabolizable Fru analog (Rabot et al., 2012). Similar to our findings, HXK1 overexpression in Arabidopsis lines showed enhanced branching despite a lack of elevated sugar levels (Kelly et al., 2012).

Within the tuber, a rapid shift from storage metabolism (starch synthesis) to reserve mobilization during postharvest storage suggests a transition from sink to source. Suc synthesis appears as a dominant anabolic pathway in the storage parenchyma of dormant and sprouting tubers (Viola et al., 2007). Cold-induced sweetening, resulting from the accumulation of reducing sugars in cold-stored potato tubers, has been studied mainly for its impact on potato processing (Sowokinos, 2001; Dale and Bradshaw, 2003). During cold-induced sweetening, Suc synthesis increases and some Suc is transported to the vacuole, where it is hydrolyzed to Glc and Fru (Isherwood, 1973; Isla et al., 1998; Sowokinos, 2001). This step is controlled predominantly by vacuolar invertase, an enzyme that is strongly associated with the accumulation of reducing sugars during cold storage (Matsuura-Endo et al., 2004). Silencing of VInv results in an effective control of cold-induced sweetening (Bhaskar et al., 2010; Zhu et al., 2016). Our experiments using VInv-silenced lines showed that tuber branching is correlated to the level of VInv silencing (Fig. 7). We believe that a higher level of Suc in the tuber parenchyma, resulting from VInv silencing, induces the sprouting of multiple stems from the tuber. Recent research has implicated a tight link between endogenous Suc and trehalose 6-phosphate (Tre6P) levels, and the latter was proposed to signal Suc availability and influence the relative amounts of Suc and starch (Lunn et al., 2014; Yadav et al., 2014). It is proposed that Suc increases Tre6P levels and that this increase will decrease the Suc levels by a negative feedback, which also can explain why we did not observe an increase of sugar levels in the scion. Moreover, transgenic lines overaccumulating Tre6P have been shown to overbranch (Yadav et al., 2014), suggesting that this nexus between Suc and Tre6P can take place in shoot branching (Barbier et al., 2015b). This result does not rule out that Suc can be the mobile signal.

In summary, in this study, we showed that etiolated stem branching is a result of systemic signaling associated with Suc level in the parenchyma. Further studies are required to determine the cascade of Suc signaling that leads to stem branching.

MATERIALS AND METHODS

Plant Material and Growth and Storage Conditions

Freshly harvested tubers of potato (Solanum tuberosum ‘Nicola’) were obtained from a potato-growing field in the northern Negev, Israel, and stored at 14°C for 3 weeks for curing. Tubers were then divided into three batches and transferred to 14°C or 33°C for 21 d (heat treatment) or 4°C for 3 months (long cold storage).

RBK potato tubers were harvested from our previously developed VInv-silenced lines (RBK1, RBK22, and RBK46) and control wild-type RBK (Zhu et al., 2014). For sprouting induction, tubers were transferred from 4°C to 14°C in the dark. In all experiments, sprouts with two to three nodes were selected unless stated otherwise. Sprouts with lateral bud growth, with or without tuber parenchyma (base), were excised using a cork borer (Ø 1 cm, 3 cm penetration) or detached manually, respectively. Sprouts were planted in vermiculite, irrigated with sterile double-distilled water, and grown at 14°C in the dark. Tubers and sprouts in all treatments were maintained at 95% relative humidity using an ultrasonic fogger (S.M.D.).

Grafting of Potato Sprouts

Sprouts developed from potato tubers under dark conditions were grafted onto other sprouts (as a scion) using the cleft-grafting method (Palauqui et al., 1997) with slight modifications. Briefly, for the scion, sprouts with two to three nodes were cut from tubers and trimmed on both sides, at the cut ends, to form a wedge shape using a blade. For the stock design, sprouts attached to tuber or tuber base were cut horizontally, approximately 2 to 3 cm above the base, and bisected vertically, using a sterile blade, to form an open cleft. The sprout (potential scion) was then gently inserted into the cleft of the sprout (potential stock) of another tuber, covered with Parafilm or held with grafting clips (Agron), and then incubated at 14°C and 95% relative humidity. After successful union of the stock and scion, grafting clips were removed. The root stock was pruned every few days.

Confirming Wound Healing of the Grafting Point

Solute translocation through the grafting point was confirmed by detaching the stock holding the scion from the tuber at different times and then exogenously supplying CFDA (Sigma) to the bud-base parenchyma to allow translocation from stock to scion. CFDA is membrane soluble, but endogenous esterases cleave off the acetate moieties to generate the fluorescent, membrane-impermeable carboxyfluorescein (CF) molecule (Viola et al., 2007). CF then acts as a marker of phloem transport and provides evidence for symplastic unloading (Roberts et al., 1997). To image phloem transport into grafted sprouts, CFDA (Sigma) was loaded into the stock, below the grafting point. The cut stock was immersed, to a depth of 5 mm, in 20 mg mL−1 aqueous CFDA solution for 16 h prior to examination of sprouts by confocal microscopy (SP8; Leica) equipped with an argon laser. Tissue was excited at 488 nm, and emission was collected at 517 to 552 nm for the CF signal.

Translocation of Labeled Sugars

To determine sugar translocation, tubers were cut and shaped, ensuring a minimum of 3 cm of parenchyma tissue between the cut surface and the sprout; the cut surface was incubated in 1 µCi of [U-14C]Glc, [U-14C]Fru, or [U-14C]Suc, to a depth of 1 cm, supplemented with 100 mm Glc, Fru, or Suc. Sugars were allowed to translocate for 14 h. A total of 200 mg of tissues was subsequently collected from the apex, node, or parenchyma. It is worth mentioning that the collected parenchyma tissue collected was not in contact with the labeled sugar solution. Radioactivity in Suc, Glc, and Fru was determined by liquid scintillation counting after crushing and diluting into an Ultima Gold liquid scintillation cocktail (PerkinElmer) using a Parkard Tri-Carb 2100TR counter analyzer (Packard BioScience).

Extraction and Quantification of Sugars

After storage for 21 d at 14°C or 33°C, or for 3 months at 4°C, tuber tissues were extracted for sugar quantification. For sugar analysis of the tuber parenchyma, 1 g of bud-base parenchyma was excised separately from heated, nonheated, or cold-stored tubers using a cork borer (Ø 1 cm, 3 cm penetration) and immediately frozen in liquid N2 and transferred to −80°C until use. To analyze sugars in the scion, 400 mg of apex or node tissue was cut using a sterile blade and immediately frozen in liquid N2 until use. Tissues were incubated three times in 80% ethanol at 80°C for 45 min each time. The solution was then dried using a speed vacuum (Centrivap concentrator; Labconco) and passed through a 0.2-µm membrane filter (Millex-GV filter unit; Merck Millipore). The filtrate was used for Suc, Glc, and Fru analyses by ultrafast liquid chromatography. An ultrafast liquid chromatography system (LC-10A UFLC series; Shimadzu) equipped with an SIL-HT automatic sample injector, pump system, refractive index detector (SPD-20A), and automatic fraction collector (FRC-10A) was employed. In addition, the ultrafast liquid chromatograph was equipped with a differential refractometer detector (Waters 410) and analytical ion-exchange column (6.5 × 300 nm; Sugar-Pak I; Waters). The mobile phase (ultrapurified deionized water; Bio Lab) was eluted through the system for 30 min at a flow rate of 0.5 mL min−1, and the column temperature was set to 80°C. The chromatographic peak corresponding to each sugar was identified by comparing the retention time with that of a standard. A calibration curve was prepared using standards to determine the relationship between the peak area and concentration.

Exogenous Supply of Sugar to Detached Sprouts

Sprouts were detached manually from tubers stored at 14°C and surface sterilized with 0.1% (v/v) sodium hypochlorite and 0.02% (v/v) Tween 20 for 5 min followed by washing with sterile water for 10 min. Sprouts were dried for 5 min and placed in 50-mL sterile tubes containing Suc, Glc, Fru, sorbitol, or mannitol solution at six concentrations (0, 25, 50, 100, 300, and 600 mm) and then incubated at 14°C in the dark for 10 d. After incubation, the sprouts were carefully rinsed with sterile water for 10 min, transferred to pots containing vermiculite, and incubated under the same conditions for 7 d. Branching was evaluated daily.

Statistical Analysis

Statistical analysis of the data was performed with JMP-in software (version 3 for Windows; SAS Institute).

Supplemental Data

The following supplemental materials are available.

Glossary

AD

apical dominance

CFDA

carboxyfluorescein diacetate

RBK

‘Russet Burbank’

Tre6P

trehalose 6-phosphate

CF

carboxyfluorescein

Footnotes

1

This research was supported by BARD (U.S.–Israel Binational Agricultural Research and Development fund) project IS-4864-15 F, by the USDA NIFA Specialty Crop Research Initiative and Hatch funds to J.J., and by Natural Science Foundation of China (NSFC) project 31360296 to H.G. The article is a contribution of the Agricultural Research Organization, The Volcani Center, Rishon LeZion, Israel, no. 792/17.

References

  1. Alves G, Decourteix M, Fleurat-Lessard P, Sakr S, Bonhomme M, Améglio T, Lacointe A, Julien JL, Petel G, Guilliot A (2007) Spatial activity and expression of plasma membrane H+-ATPase in stem xylem of walnut during dormancy and growth resumption. Tree Physiol 27: 1471–1480 [DOI] [PubMed] [Google Scholar]
  2. Antoun M, Ouellet F (2013) Growth temperature affects inflorescence architecture in Arabidopsis thaliana. Botany 91: 642–651 [Google Scholar]
  3. Barbier F, Péron T, Lecerf M, Perez-Garcia MD, Barrière Q, Rolčík J, Boutet-Mercey S, Citerne S, Lemoine R, Porcheron B, et al. (2015a) Sucrose is an early modulator of the key hormonal mechanisms controlling bud outgrowth in Rosa hybrida. J Exp Bot 66: 2569–2582 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Barbier FF, Lunn JE, Beveridge CA (2015b) Ready, steady, go! A sugar hit starts the race to shoot branching. Curr Opin Plant Biol 25: 39–45 [DOI] [PubMed] [Google Scholar]
  5. Beveridge CA, Ross JJ, Murfet IC (1994) Branching mutant rms-2 in Pisum sativum (grafting studies and endogenous indole-3-acetic acid levels). Plant Physiol 104: 953–959 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Beveridge CA, Symons GM, Murfet IC, Ross JJ, Rameau C (1997) The rms1 mutant of pea has elevated indole-3-acetic acid levels and reduced root-sap zeatin riboside content but increased branching controlled by graft-transmissible signal(s). Plant Physiol 115: 1251–1258 [Google Scholar]
  7. Bhaskar PB, Wu L, Busse JS, Whitty BR, Hamernik AJ, Jansky SH, Buell CR, Bethke PC, Jiang J (2010) Suppression of the vacuolar invertase gene prevents cold-induced sweetening in potato. Plant Physiol 154: 939–948 [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Buskila Y, Sela N, Teper-Bamnolker P, Tal I, Shani E, Weinstain R, Gaba V, Tam Y, Lers A, Eshel D (2016) Stronger sink demand for metabolites supports dominance of the apical bud in etiolated growth. J Exp Bot 67: 5495–5508 [DOI] [PubMed] [Google Scholar]
  9. Cline MG. (1994) The role of hormones in apical dominance: new approaches to an old problem in plant development. Physiol Plant 90: 230–237 [Google Scholar]
  10. Dale MFB, Bradshaw JE (2003) Progress in improving processing attributes in potato. Trends Plant Sci 8: 310–312 [DOI] [PubMed] [Google Scholar]
  11. Decourteix M, Alves G, Bonhomme M, Peuch M, Ben Baaziz K, Brunel N, Guilliot A, Rageau R, Améglio T, Pétel G, et al. (2008) Sucrose (JrSUT1) and hexose (JrHT1 and JrHT2) transporters in walnut xylem parenchyma cells: their potential role in early events of growth resumption. Tree Physiol 28: 215–224 [DOI] [PubMed] [Google Scholar]
  12. de Jong M, George G, Ongaro V, Williamson L, Willetts B, Ljung K, McCulloch H, Leyser O (2014) Auxin and strigolactone signaling are required for modulation of Arabidopsis shoot branching by nitrogen supply. Plant Physiol 166: 384–395 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Domagalska MA, Leyser O (2011) Signal integration in the control of shoot branching. Nat Rev Mol Cell Biol 12: 211–221 [DOI] [PubMed] [Google Scholar]
  14. Dun EA, de Saint Germain A, Rameau C, Beveridge CA (2012) Antagonistic action of strigolactone and cytokinin in bud outgrowth control. Plant Physiol 158: 487–498 [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Eshel D. (2015) Bridging dormancy release and apical dominance in potato tuber. In JV Anderson, ed, Advances in Plant Dormancy. Springer; International Publishing, Switzerland, pp 187–196 [Google Scholar]
  16. Eshel D, Teper-Bamnolker P (2012) Can loss of apical dominance in potato tuber serve as a marker of physiological age? Plant Signal Behav 7: 1158–1162 [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Ferguson BJ, Beveridge CA (2009) Roles for auxin, cytokinin, and strigolactone in regulating shoot branching. Plant Physiol 149: 1929–1944 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Foo E, Turnbull CG, Beveridge CA (2001) Long-distance signaling and the control of branching in the rms1 mutant of pea. Plant Physiol 126: 203–209 [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Gévaudant F, Pétel G, Guilliot A (2001) Differential expression of four members of the H+-ATPase gene family during dormancy of vegetative buds of peach trees. Planta 212: 619–626 [DOI] [PubMed] [Google Scholar]
  20. Girault T, Abidi F, Sigogne M, Pelleschi-Travier S, Boumaza R, Sakr S, Leduc N (2010) Sugars are under light control during bud burst in Rosa sp. Plant Cell Environ 33: 1339–1350 [DOI] [PubMed] [Google Scholar]
  21. Gomez-Roldan V, Fermas S, Brewer PB, Puech-Pagès V, Dun EA, Pillot JP, Letisse F, Matusova R, Danoun S, Portais JC, et al. (2008) Strigolactone inhibition of shoot branching. Nature 455: 189–194 [DOI] [PubMed] [Google Scholar]
  22. Goodwin P. (1967) The control and branch growth on potato tubers. I. Anatomy of buds in relation to dormancy and correlative inhibition. J Exp Bot 18: 78–86 [Google Scholar]
  23. Granot D, David-Schwartz R, Kelly G (2013) Hexose kinases and their role in sugar-sensing and plant development. Front Plant Sci 4: 44. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Harris PM. (1992) The Potato Crop: The Scientific Basis for Improvement. Chapman and Hall, London [Google Scholar]
  25. Hemberg T. (1985) Potato rest. In Li P, ed, Potato Physiology. Academic Press, Orlando, FL, pp 353–388 [Google Scholar]
  26. Isherwood FA. (1973) Starch-sugar interconversion in Solanum tuberosum. Phytochemistry 12: 2579–2591 [Google Scholar]
  27. Isla MI, Vattuone MA, Sampietro AR (1998) Hydrolysis of sucrose within isolated vacuoles from Solanum tuberosum L. tubers. Planta 205: 601–605 [DOI] [PubMed] [Google Scholar]
  28. Juknevičienė Z, Venskutonienė E, Pranaitienė R, Duchovskis P (2011) The influence of different temperatures and exposition time on potato tuber sprouting and development of plants. Žemdirbystė (Agriculture) 98: 131–138 [Google Scholar]
  29. Kebrom TH, Mullet JE (2015) Photosynthetic leaf area modulates tiller bud outgrowth in sorghum. Plant Cell Environ 38: 1471–1478 [DOI] [PubMed] [Google Scholar]
  30. Kelly G, David-Schwartz R, Sade N, Moshelion M, Levi A, Alchanatis V, Granot D (2012) The pitfalls of transgenic selection and new roles of AtHXK1: a high level of AtHXK1 expression uncouples hexokinase1-dependent sugar signaling from exogenous sugar. Plant Physiol 159: 47–51 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Lunn JE, Delorge I, Figueroa CM, Van Dijck P, Stitt M (2014) Trehalose metabolism in plants. Plant J 79: 544–567 [DOI] [PubMed] [Google Scholar]
  32. Marquat C, Vandamme M, Gendraud M, Pétel G (1999) Dormancy in vegetative buds of peach: relation between carbohydrate absorption potentials and carbohydrate concentration in the bud during dormancy and its release. Sci Hortic (Amsterdam) 79: 151–162 [Google Scholar]
  33. Martin G. (1987) Apical dominance. HortScience 22: 824–833 [Google Scholar]
  34. Mason MG, Ross JJ, Babst BA, Wienclaw BN, Beveridge CA (2014) Sugar demand, not auxin, is the initial regulator of apical dominance. Proc Natl Acad Sci USA 111: 6092–6097 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Matsuura-Endo C, Kobayashi A, Noda T, Takigawa S, Yamauchi H, Mori M (2004) Changes in sugar content and activity of vacuolar acid invertase during low-temperature storage of potato tubers from six Japanese cultivars. J Plant Res 117: 131–137 [DOI] [PubMed] [Google Scholar]
  36. Maurel K, Leite GB, Bonhomme M, Guilliot A, Rageau R, Pétel G, Sakr S (2004a) Trophic control of bud break in peach (Prunus persica) trees: a possible role of hexoses. Tree Physiol 24: 579–588 [DOI] [PubMed] [Google Scholar]
  37. Maurel K, Sakr S, Gerbe F, Guilliot A, Bonhomme M, Rageau R, Pétel G (2004b) Sorbitol uptake is regulated by glucose through the hexokinase pathway in vegetative peach-tree buds. J Exp Bot 55: 879–888 [DOI] [PubMed] [Google Scholar]
  38. McIntyre G. (1997) The role of nitrate in the osmotic and nutritional control of plant development. Funct Plant Biol 24: 103–118 [Google Scholar]
  39. McIntyre GI. (1987) The role of water in the regulation of plant development. Can J Bot 65: 1287–1298 [Google Scholar]
  40. Melnyk CW, Meyerowitz EM (2015) Plant grafting. Curr Biol 25: R183–R188 [DOI] [PubMed] [Google Scholar]
  41. Morris SE, Cox MC, Ross JJ, Krisantini S, Beveridge CA (2005) Auxin dynamics after decapitation are not correlated with the initial growth of axillary buds. Plant Physiol 138: 1665–1672 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Morris SE, Turnbull CG, Murfet IC, Beveridge CA (2001) Mutational analysis of branching in pea: evidence that Rms1 and Rms5 regulate the same novel signal. Plant Physiol 126: 1205–1213 [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Napoli C. (1996) Highly branched phenotype of the petunia dad1-1 mutant is reversed by grafting. Plant Physiol 111: 27–37 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Palauqui JC, Elmayan T, Pollien JM, Vaucheret H (1997) Systemic acquired silencing: transgene-specific post-transcriptional silencing is transmitted by grafting from silenced stocks to non-silenced scions. EMBO J 16: 4738–4745 [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Pedersen CN, Axelsen KB, Harper JF, Palmgren MG (2012) Evolution of plant p-type ATPases. Front Plant Sci 3: 31. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Phillips IDJ. (1975) Apical dominance. Annu Rev Plant Physiol 26: 341–367 [Google Scholar]
  47. Prat S, Frommer WB, Höfgen R, Keil M, Kossmann J, Köster-Töpfer M, Liu XJ, Müller B, Peña-Cortés H, Rocha-Sosa M, et al. (1990) Gene expression during tuber development in potato plants. FEBS Lett 268: 334–338 [DOI] [PubMed] [Google Scholar]
  48. Rabot A, Henry C, Ben Baaziz K, Mortreau E, Azri W, Lothier J, Hamama L, Boummaza R, Leduc N, Pelleschi-Travier S, et al. (2012) Insight into the role of sugars in bud burst under light in the rose. Plant Cell Physiol 53: 1068–1082 [DOI] [PubMed] [Google Scholar]
  49. Rameau C, Bertheloot J, Leduc N, Andrieu B, Foucher F, Sakr S (2015) Multiple pathways regulate shoot branching. Front Plant Sci 5: 741. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Roberts AG, Cruz SS, Roberts IM, Prior DA, Turgeon R, Oparka KJ (1997) Phloem unloading in sink leaves of Nicotiana benthamiana: comparison of a fluorescent solute with a fluorescent virus. Plant Cell 9: 1381–1396 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Sachs T, Thimann KV (1964) Release of lateral buds from apical dominance. Nature 201: 939–940 [Google Scholar]
  52. Shim H, Chun YS, Lewis BC, Dang CV (1998) A unique glucose-dependent apoptotic pathway induced by c-Myc. Proc Natl Acad Sci USA 95: 1511–1516 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Shirazi AM, Fuchigami LH (1995) Effects of “near-lethal” stress on bud dormancy and stem cold hardiness in red-osier dogwood. Tree Physiol 15: 275–279 [DOI] [PubMed] [Google Scholar]
  54. Smeekens S, Ma J, Hanson J, Rolland F (2010) Sugar signals and molecular networks controlling plant growth. Curr Opin Plant Biol 13: 274–279 [DOI] [PubMed] [Google Scholar]
  55. Sonnewald U. (2001) Control of potato tuber sprouting. Trends Plant Sci 6: 333–335 [DOI] [PubMed] [Google Scholar]
  56. Sowokinos J, Shock C, Stieber T, Eldredge E (2000) Compositional and enzymatic changes associated with the sugar-end defect in Russet Burbank potatoes. Am J Potato Res 77: 47–56 [Google Scholar]
  57. Sowokinos JR. (2001) Biochemical and molecular control of cold-induced sweetening in potatoes. Am J Potato Res 78: 221–236 [Google Scholar]
  58. Suttle JC. (2004) Physiological regulation of potato tuber dormancy. Am J Potato Res 81: 253–262 [Google Scholar]
  59. Takahashi H, Imamura T, Konno N, Takeda T, Fujita K, Konishi T, Nishihara M, Uchimiya H (2014) The gentio-oligosaccharide gentiobiose functions in the modulation of bud dormancy in the herbaceous perennial Gentiana. Plant Cell 26: 3949–3963 [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Teper-Bamnolker P, Buskila Y, Lopesco Y, Ben-Dor S, Saad I, Holdengreber V, Belausov E, Zemach H, Ori N, Lers A, et al. (2012) Release of apical dominance in potato tuber is accompanied by programmed cell death in the apical bud meristem. Plant Physiol 158: 2053–2067 [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Turnbull CGN, Hanke DE (1985) The control of bud dormancy in potato tubers: evidence for the primary role of cytokinins and a seasonal pattern of changing sensitivity to cytokinin. Planta 165: 359–365 [DOI] [PubMed] [Google Scholar]
  62. Van den Ende W. (2014) Sugars take a central position in plant growth, development and, stress responses: a focus on apical dominance. Front Plant Sci 5: 313. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Viola R, Pelloux J, van der Ploeg A, Gillespie T, Marquis N, Roberts AG, Hancock RD (2007) Symplastic connection is required for bud outgrowth following dormancy in potato (Solanum tuberosum L.) tubers. Plant Cell Environ 30: 973–983 [DOI] [PubMed] [Google Scholar]
  64. Visser RGF, Vreugdenhil D, Hendriks T, Jacobsen E (1994) Gene expression and carbohydrate content during stolon to tuber transition in potatoes (Solanum tuberosum). Physiol Plant 90: 285–292 [Google Scholar]
  65. Wang SY, Faust M (1994) Changes in the antioxidant system associated with budbreak in ‘Anna’ apple (Malus domestica Borkh.) buds. J Am Soc Hortic Sci 119: 735–741 [Google Scholar]
  66. Wiltshire JJJ, Cobb AH (1996) A review of the physiology of potato tuber dormancy. Ann Appl Biol 129: 553–569 [Google Scholar]
  67. Wisniewski M, Sauter J, Fuchigami L, Stepien V (1997) Effects of near-lethal heat stress on bud break, heat-shock proteins and ubiquitin in dormant poplar (Populus nigra Charkowiensis × P. nigra incrassata). Tree Physiol 17: 453–460 [DOI] [PubMed] [Google Scholar]
  68. Yadav UP, Ivakov A, Feil R, Duan GY, Walther D, Giavalisco P, Piques M, Carillo P, Hubberten HM, Stitt M, et al. (2014) The sucrose-trehalose 6-phosphate (Tre6P) nexus: specificity and mechanisms of sucrose signalling by Tre6P. J Exp Bot 65: 1051–1068 [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Yue T, Ling L, Chuan-yuan L, Dong-mei L, Xiu-de C, Dong-sheng G (2013) Respiratory response of dormant nectarine vegetative buds to high temperature stress. J Integr Agric 12: 80–86 [Google Scholar]
  70. Zhu X, Gong H, He Q, Zeng Z, Busse JS, Jin W, Bethke PC, Jiang J (2016) Silencing of vacuolar invertase and asparagine synthetase genes and its impact on acrylamide formation of fried potato products. Plant Biotechnol J 14: 709–718 [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Zhu X, Richael C, Chamberlain P, Busse JS, Bussan AJ, Jiang J, Bethke PC (2014) Vacuolar invertase gene silencing in potato (Solanum tuberosum L.) improves processing quality by decreasing the frequency of sugar-end defects. PLoS ONE 9: e93381. [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from Plant Physiology are provided here courtesy of Oxford University Press

RESOURCES