Planta

, Volume 237, Issue 3, pp 681–691 | Cite as

Expression of CsSEF1 gene encoding putative CCCH zinc finger protein is induced by defoliation and prolonged darkness in cucumber fruit

Original Article

Abstract

To find a marker gene for photoassimilate limitation in cucumber fruit, genes induced in young fruit by total defoliation were cloned using the subtraction method. Almost every clone matched perfectly to a member of cucumber unigene ver. 3 of the Cucurbit Genomics Database. From the clones obtained, six genes were selected and the effect of defoliation on their expression was analyzed. In particular, expression of a gene that is highly homologous to the cucumber gene CsSEF1 (CAI30889) encoding putative CCCH zinc finger protein, which is reported to be induced at somatic embryogenesis in suspension culture, was enhanced by the treatment by about 50 times. The sequencing of the full-length cDNA and BLAST search in the Cucurbit Genomics Database indicated that our cloned gene is identical to CsSEF1. In control fruit, the expression of CsSEF1 did not change markedly in terms of development. By contrast, the expression of CsSEF1 was enhanced by prolonged darkness at the transcript level. This increase in the expression of CsSEF1 was temporally correlated with the decline in the fruit respiration rate. In mature leaves under prolonged darkness, enhanced expression was observed in the asparagine synthetase gene, but not in CsSEF1. These results suggest that the asparagine synthetase gene can be a good marker for sugar starvation and that CsSEF1 might be involved in the signal transduction pathway from photoassimilate limitation to growth cessation in cucumber fruit.

Keywords

CsSEF1 Cucumber Fruit growth Photoassimilate Respiration Tandem CCCH zinc finger 

Abbreviations

CCCH

Cysteine-cysteine-cysteine-histidine

DAA

Days after anthesis

GA

Gibberellic acid

qRT-PCR

Quantitative reverse transcriptase–polymerase chain reaction

RACE

Rapid amplification of cDNA ends

TZF

Tandem CCCH zinc finger

UTR

Untranslated region

Introduction

In cucumber, the production of malformed fruit should be avoided. Although it has long been assumed that the production of malformed fruit is related to limited photoassimilate supply (Kato and Oda 1977), this has not been strictly proven. The regulatory mechanism of photoassimilate partitioning to sink organs is poorly understood (Gifford and Evans 1981; Giaquinta 1983; Thorne 1985; Ho 1988; Frommer and Ninnemann 1995; Lalonde et al. 2004; Marcelis et al. 2004; Turgeon and Wolf 2009; Wubs et al. 2009; Zhou et al. 2009; Nunes-Nesi et al. 2010; Ruan et al. 2010; Chen and Thelen 2011). One possible strategy for studying the effect of limited photoassimilate supply at the cellular level may be to find a marker gene that responds to such a limitation in the fruit. Extensive studies on the effect of sugar status in Arabidopsis transcriptomes have been performed (Price et al. 2004; Blasing et al. 2005). Whereas interest in gene expression induced by sugar or sugar starvation has increased (Rolland et al. 2006), only a few published studies have investigated the relationship between photoassimilate supply and gene expression in rapidly growing fruits such as cucurbits (Craft and Lorentz 1944). Defoliation (Tamura et al. 2011) and prolonged darkness (Baena-González et al. 2007) have been widely used as methods to limit photoassimilate supply. Here, we used complete defoliation as a treatment and the subtraction method to clone genes whose expression is enhanced in young fruit by defoliation. We obtained the full-length cDNA from a clone whose expression was markedly enhanced by defoliation, and identified it as the CsSEF1 gene, which is reported to be induced during somatic embryogenesis in cucumber (Grabowska et al. 2009). The response of CsSEF1 expression differed from that of the asparagine synthetase gene, which is known to be a good marker of sugar starvation. The decline in the fruit respiration rate, which is almost proportional to the fruit growth rate (Tazuke and Sakiyama 1991), temporally coincided with the marked enhancement of CsSEF1 expression. Together, these results suggest that CsSEF1 gene action is located somewhere in the signal transduction pathway from photoassimilate starvation to the growth cessation of sink tissues, but is independent of the local sugar-sensing pathway.

Materials and methods

Cucumber (Cucumis sativus L.) cv. Tokiwa (supplied by Dr. Yoshiteru Sakata of the Institute of Vegetable and Tea Science, National Agriculture and Food Research Organization (NARO), Kusawa, Ano, Tsu, Mie, Japan) was used. The plants for subtraction cloning were grown in autumn 2009. Plants were grown in a glasshouse in 15-L pots filled with vermiculite and irrigated with half-strength Hoagland No. 2 solution. The plants were pinched, leaving 12 leaves. Fruits on the primary node of the lateral shoots were used. Female flowers were pollinated with pollen from male flowers of the same plant in the morning of the day of anthesis. Seven days after anthesis (DAA) at 11 a.m., when fruits were about 9 cm long, a control fruit was harvested. At the same time, the plant was completely defoliated for the fruit treatment. At 11 a.m. the next day, the treated fruit was harvested. The whole fruit (about 5 g) was frozen with liquid N2 and powdered with a mortar and pestle. Total RNA was extracted with ISOGEN (Nippon Gene, Tokyo, Japan). Messenger RNA was purified with the Oligotex™ −dT 30 <Super> mRNA Purification Kit (TaKaRa, Ohtsu, Japan). The extract was concentrated as follows. To the 600-μL extract, 600 μL isopropanol and 120 μL 4 M LiCl were added; the mix was incubated at −20 °C for 10 min and centrifuged at 12,000×g. Water (20 μL) was then added to the pellet, which was incubated at 55 °C. The extracted total mRNA was used for subtraction cloning using the PCR-Select cDNA Subtraction Kit (Clontech, Mountain View, CA, USA). Double-stranded cDNA was synthesized from the mRNA and digested with RsaI. To the two aliquots of the treatment digest (tester), different oligonucleotides (adaptor) were ligated. Each ligated tester was hybridized with the control digest (driver), combined, hybridized with the driver again, and then amplified by polymerase chain reaction (PCR) using a thermal cycler (Dice mini, TaKaRa) using primers specific to the adaptors. Thus, fragments of cDNAs specific to the treatment were amplified. The PCR product was subcloned with a TA vector (pGEM-T Easy, Promega, Tokyo, Japan). About 100 colonies (white and pale blue) were subcultured overnight, centrifuged, and stored at −80 °C. Plasmids were extracted from 78 samples and sequenced using the BigDye Terminator v3.1 Cycle Sequencing Kit (Life Technologies Japan, Tokyo, Japan) as the reagent and the Applied Biosystems 3130xl Genetic Analyzer (Life Technologies). Matches to unigenes (cucumber ver. 3) of the Cucurbit Genomics Database (http://www.icugi.org/) were examined.

For the sequencing of full-length cDNA, plants were grown similarly in autumn 2011. Four DAA, three plants were completely defoliated at 11 a.m. Their fruits were harvested at 11 a.m. the next day, and portions of fruit tissues in the proximal and distal halves were frozen with liquid N2 and powdered with a mortar and pestle. One hundred milligrams of the aliquot was taken and the total RNA was extracted using the RNeasy Plant Mini Kit (Qiagen, Hilden, Germany). 5′-Rapid amplification of cDNA ends (RACE) was conducted using the 5′-Full RACE Core Set (TaKaRa) with LA Taq (TaKaRa). The primers were designed from the known sequence obtained above. The PCR product was subcloned and sequenced as above.

Plants for the expression analysis were grown similarly in autumn 2011 and spring 2012. When fruit weight was about 5 g, three control fruits were harvested at 11 a.m. Plants selected for fruit treatment were completely defoliated at the same time. At 11 a.m. the next day, three treated fruits were harvested. Extraction of total RNA was conducted as above using the RNeasy Plant Mini Kit (Qiagen). First-strand cDNA was synthesized from the total RNA extract using the PrimeScript RT Reagent Kit (TaKaRa) and stored at −20 °C. Expression of the six selected genes was analyzed by quantitative reverse transcriptase–polymerase chain reaction (qRT-PCR) using SYBR Premix Ex Taq™ II as reagent and the Thermal Cycler Dice Real Time System (TaKaRa). For the expression analysis in prolonged darkness, plants were placed in a darkened growth chamber at 25 °C at 10 a.m. when fruit weight was about 5 g. Three fruits were harvested, at 10 a.m. (control) and 4 p.m. that day, and at 1 p.m. the next day. The effect of prolonged darkness on the expression of the two selected genes was examined similarly. For the analysis of the effect of prolonged darkness on the expression of the two selected genes in plant parts other than fruit, plants were placed in a darkened growth chamber at 25 °C at 10 a.m. Mature leaves, the apical region of lateral shoots, and roots were harvested at 1 p.m. the next day. Plants grown hydroponically in Hoagland No. 2 solution were used to sample roots. Expression of the two selected genes was analyzed as above. To analyze developmental change in the expression of the two selected genes in control fruit, three fruits were harvested, at 10 a.m. that day (5 DAA), the next day (6 DAA), and 2 days later (7 DAA), when fruit weight was about 5 g. Expression of the two selected genes was analyzed as above.

A fruit weighing about 5 g still on the vine was put into a double cylindrical fruit chamber (outer diameter 15 cm, inner diameter 5 cm, length 30 cm) that was wrapped in urethane foam and aluminum foil for heat insulation and light shielding to analyze the fruit respiration rate. Openings in the cylinder were sealed with silicon stoppers. To avoid air leak, the fruit peduncle was sealed with soft rubber. The cylinder temperature was maintained at 25 °C by circulating water controlled by a thermostatic machine (ZL-100, Taitec, Koshigaya, Japan). Room air was drawn by a compressor (Hitachi, Tokyo, Japan) and passed through a column of soda lime. The flow rate was maintained at 2.0 L min−1 using a mass flow controller (SEC-E40, Horiba, Kyoto, Japan), and brought into the inner cylinder, where 0.5 L min−1 of the air flow from the opposite opening was drawn and its CO2 concentration was measured with an infrared gas analyzer (VA-3000, Horiba). Analyzer readings were recorded every minute by a data logger (CR10X, Campbell Scientific, Logan, UT, USA). For the defoliation treatment, plants were supplied with light from 6 a.m. to 6 p.m. by two high pressure sodium lamps (IAN-361FL, GS Yuasa, Kyoto, Japan) with a photosynthetic photon flux density 1,400 μ mol m−2 s−1. After a fruit was placed in the fruit chamber, the plant was defoliated completely at 1 p.m. For the prolonged dark treatment, no light was supplied after a fruit was placed in the fruit chamber. The respiration rate was monitored several times.

For the analysis of sugar and starch, harvested tissue was frozen by liquid N2, powdered with a mortar and pestle, and homogenized with 80 % ethanol. The homogenate was extracted by incubating for 15 min at 80 °C, filtrated through a glass filter (GF/F, Whatman, Kent, UK), and made up to 100 mL. The residue was dried at 55 °C for 1 week. The starch content of the ethanol-insoluble residue was analyzed with F-kit (Roche Diagnostics, Basel, Switzerland). Aliquots of the ethanol extract were dried with a rotary evaporator and dissolved in water. Concentrations of glucose, fructose, and sucrose, which are the major sugars in cucumber fruit (Pharr et al. 1977), were then determined by F-kit. Hereafter, the sum of glucose and fructose concentrations is referred to as the hexose concentration.

Statistical analyses of the significance of treatment effects on the transcript level were conducted using R (R Development Core Team 2011).

Results

Clones obtained by subtraction cloning

After defoliation, fruit grew slightly for about 1 day and increased in volume by about 40 %. They then stopped growing, but stayed green without signs of withering. Fruit harvested a few days after defoliation remained fleshy.

A total of 108 colonies consisting of 78 white colonies and 30 pale-blue colonies were collected and cultured. From these, 78 clones were sequenced, 74 of which matched unigenes (cucumber ver. 3) in the Cucurbit Genomics Database, as shown in Table 1. The remaining four clones may correspond to failures in sequencing. Twenty-one unigenes were detected overall. Of the 74 clones that were sequenced successfully, 54 clones corresponded to five unigenes that matched more than five clones. The remaining 20 clones corresponded to unigenes that matched fewer than three clones. Among the matched unigenes, CU100471 was the most abundant, being highly homologous to 24 clones. This unigene is highly homologous to aquaporin PIP1-3 of Arabidopsis thaliana. The next most abundant unigene was CU107225, which was highly homologous to 12 clones but had no match in the nucleotide and protein databases. The next most abundant unigene was CU105387, which was highly homologous to seven clones. This unigene is highly homologous to the ubiquitin gene of asparagus. The next most abundant unigene was CU114757, which was highly homologous to six clones. According to cucumber unigene ver. 3 in the Cucurbit Genomics Database, this unigene is highly homologous to the asparagine synthetase gene of asparagus. The BLAST search in GenBank found the closest match to C. sativus clone CU29H5 asparagine synthetase mRNA, partial cds (E-value = 4E−129). Therefore, this unigene is hereafter referred to as the asparagine synthetase gene. The next most abundant unigene was CU107833, which was highly homologous to five clones. This unigene is highly homologous to probable aquaporin TIP2-2 of A. thaliana. Other unigenes matched to two clones or fewer. Of these, CU092580, which matched two clones (clones No. 4 and No. 14), was of interest. A search in SwissProt at the amino acid level revealed it to be highly homologous (E-value = 1E−38) to A. thaliana zinc finger cysteine–cysteine–cysteine–histidine (CCCH) domain-containing protein 20 (O82199, Wang et al. 2008). On the other hand, a search in GenBank at the nucleotide level revealed it to be highly homologous (E-value = 2e−80) to a cucumber full-length cDNA CsSEF1 (CAI30889, 924 bp; Grabowska et al. 2009) that encodes putative CCCH-type zinc finger protein. Our clones No. 4 (694 bp) and No. 14 (704 bp) were 100 % identical (694/694) at the nucleotide level. No. 14 was 10 bases longer in the 5′ direction. Clone No. 14 was 99 % identical (529/532) to CsSEF1.
Table 1

Description of clones obtained

Unigene

E-value

Most closely matched homolog in SwissProt

Organism

E-value

Number of clones

CU100471

0.0

Aquaporin PIP1-3

A. thaliana

6E−144

24

CU107225

0.0

No matches

12

CU105387

0.0

Ubiquitin

Asparagus

2E−35

7

CU114757

1E−160

Asparagine synthetase

Asparagus

1E−64

6

CU107833

0.0

Probable aquaporin TIP2-2

A. thaliana

3E−84

5

CU092580

0.0

Zinc finger CCCH domain-containing protein 20

A. thaliana

1E−38

2

CU100456

0.0

Chlorophyll a/b-binding protein of LHCII type 1, chloroplastic

Cucumber

6E−68

2

CU105533

0.0

Phenazine biosynthesis-like domain-containing protein 2

Mouse

8E−22

2

CU111456

0.0

GDSL esterase/lipase 5

A. thaliana

2E−32

2

CU107857

0.0

Tublin alpha-3/alpha-5 chain

A. thaliana

2E−236

1

CU094539

0.0

Glycerate dehydrogenase

Cucumber

9E−208

1

CU100891

0.0

Aquaporin PIP1-3

A. thaliana

7E−106

1

CU155640

3E−7

Lipoxygenase 1

Potato

9E−31

1

CU100696

1E−131

ATP synthase gamma chain, chloroplastic

Spinach

6E−20

1

CU108064

0.0

Photosystem II reaction center W protein, chloroplastic

A. thaliana

1E−18

1

CU105437

1E−108

Chlorophyll a/b-binding protein 3C, chloroplastic

Tomato

1E−12

1

CU085560

1E−141

No matches

1

CU115603

0.0

No matches

1

CU122562

7E−39

No matches

1

CU137825

2E−79

No matches

1

CU149415

4E–35

No matches

1

Not read

4

Most clones were highly homologous to cucumber unigene ver. 3 of the Cucurbit Genomics Database

5′-RACE of clone No. 14

We performed a 5′-RACE analysis and obtained the 5′ upstream sequence (Fig. 1). The sequence, together with clone No. 14, covers the full-length cDNA, which turned out to be identical to CsSEF1. The BLAST search of the cucumber genome in the Cucurbit Genomics Database showed only a single gene that is highly homologous to CsSEF1 (E-value = 0, the next value is E-value = 10−16). These results indicate that our clones No. 4 and No. 14 are identical to CsSEF1. Multiple alignment by ClustalW was conducted for the top 10 matches by BLAST search in UniProt (Fig. 2). Except for one, all showed the Cx7Cx5Cx3H-x16-Cx5Cx4Cx3H motif. From this multiple alignment, a phylogenic tree of these genes was drawn (Fig. 3).
Fig. 1

Sequence obtained by 5′-RACE analysis of clone No. 14. a Relative positions of the sequence obtained by 5′-RACE and clone No. 14 (homologous to unigene CU092580 in Table 1) in relation to the CsSEF1 gene. The length of the CsSEF1 cDNA (beginning from the 5′ UTR to the end of the 3′ UTR) is 1,096 bp. The coding sequence begins from the 85th base. The sequence read by 5′-RACE begins from the beginning of 5′ UTR and ends after the first CCCH motif. b The sequence obtained by 5′-RACE

Fig. 2

Multiple alignment by ClustalW of the 10 closest proteins retrieved by BLAST in the UniProt database. The reversed characters on the black background indicates the tandem CCCH motif

Fig. 3

Phylogenic tree constructed by the multiple alignment shown in Fig. 2. HUA1 was included as an outlier. The scale bar represents 0.1 % sequence divergence. Bootstrap values based on 1,000 replications at branch nodes

Expression of the six selected unigenes after defoliation treatment

As subtraction cloning can yield false-positive results, expression of the cloned genes was examined. Because of the sample limitations, six unigenes were selected for expression analysis. The four most abundant unigenes were included and the remaining two were selected arbitrarily, CU092580 and CU100456, the latter of which was homologous to one clone that is highly homologous to the chlorophyll a/b-binding protein of LHCII type 1 (chloroplastic). The BLAST search of cucumber genome ver. 2 revealed two genes highly homologous to the asparagine synthetase gene. The primers for qRT-PCR were designed to be specific to unigene CU114757.

Results of the qRT-PCR analyses are presented in Fig. 4. The fruit respiration rate declined immediately after defoliation and became very low at the time of harvest the next day (Fig. 4a). Remarkably, the treatment enhanced expression of CsSEF1 by about 50 times. In addition, the treatment enhanced expression of the asparagine synthetase gene by about 10 times. Other unigenes showed no enhanced expression from the treatment, so they were regarded as artifacts of subtraction cloning (Fig. 4b).
Fig. 4

Effect of defoliation on the respiration rate (a) and expression (b) of the six selected unigenes (Cucurbit Genomics Database cucumber unigene ver. 3) in young cucumber fruits as examined by qRT-PCR. Respiration was measured in a growth chamber at 25 °C. Fruits for qRT-PCR were grown in a glasshouse. CU114757, asparagine synthetase; CU100471, plasma membrane aquaporin; CU107225, no match found; CU105387, ubiquitin; CU100453, chlorophyll a/b-binding protein of LHCII; CU092580, zinc finger CCCH domain-containing protein CsSEF1. The expression of each unigene was expressed as relative to that of the actin gene. The ratio of average expression of three replications in treated fruit to that in control fruit is shown. Vertical bars are SE of means (n = 3). Significance levels shown by Welch’s test

Detailed expression analysis of CsSEF1 and the asparagine synthetase gene

The complete defoliation treatment can cause not only photoassimilate starvation, but also water or mineral stresses. To exclude these possibilities, the prolonged darkness treatment was undertaken (Fig. 5). Changes in the fruit respiration rate under prolonged darkness are shown in Fig. 5a. In the prolonged darkness treatment, the timing of marked enhancement of CsSEF1 gene expression seemed to coincide with the decline in respiration rate (Fig. 5a, b). The expression of CsSEF1 was markedly enhanced, whereas the expression of the asparagine synthetase gene was only slightly, although significantly, enhanced by the treatment.
Fig. 5

Effect of prolonged darkness on the respiration rate (a) and expression (b) of CsSEF1 (circles) and the asparagine synthetase gene (triangles) as examined by qRT-PCR of young fruits grown in a growth chamber at 25 °C. Plants were transferred to a darkened growth chamber at 25 °C in the morning. The expression of each gene was expressed as relative to that of the actin gene. The ratio of average expression of three replications in treated fruit to that in control fruit is shown. Vertical bars are SE of means (n = 3). Significance levels shown by Welch’s test

To exclude the possibility that the enhancement in expression of CsSEF1 and the asparagine synthetase gene was merely due to developmental change, fruits on 5, 6, and 7 DAA were analyzed. The developmental change in expression of CsSEF1 was significant, but the value remained very low compared with the treated fruits (Fig. 6a). The developmental change in expression of the asparagine synthetase gene was not significant, and remained very low compared with the treated fruits (Fig. 6b).
Fig. 6

Developmental change in the expression of CsSEF1 (a) and the asparagine synthetase gene (b) in young cucumber fruit as examined by qRT-PCR. The expression of each gene was expressed as relative to that of the actin gene. The ratio of average expression of three replications in treated fruit to that in defoliation treated fruit is shown. Vertical bars are SE of means (n = 3). Different letters indicate significant difference at 5 % level examined by Tukey’s test

The effects of prolonged darkness on expression of CsSEF1and the asparagine synthetase gene of mature leaves, the apical region of lateral shoots, and roots are shown in Fig. 7. Expression of CsSEF1 was enhanced in the apical region and roots, but not in mature leaves. However, expression of the asparagine synthetase gene was markedly enhanced in all three tissues.
Fig. 7

Effect of prolonged darkness on the expression of CsSEF1 (a) and the asparagine synthetase gene (b) in mature leaves, the apical region of lateral shoots, and roots as examined by qRT-PCR. Plants were transferred to a darkened growth chamber at 25 °C in the morning. For the sampling of roots, hydroponically grown plants were used. All tissues were harvested 27 h after the darkness treatment began. The expression of CsSEF1 was expressed as relative to that of the actin gene. The ratio of average expression of three replications in treated fruit to that in control fruit is shown. Vertical bars are SE of means (n = 3). Significance levels shown by Welch’s test

Carbohydrate concentration of leaves and fruits

In mature leaves, hexose and sucrose concentrations were very low, consistent with a report by Goldschmidt and Huber (1992). The starch concentration of mature leaves became close to zero after 6 h in the darkness (Fig. 8a). Fruits contained small amounts of starch, but their concentrations became close to zero after 6 h in the darkness and 1 day after defoliation. The sucrose concentration of fruits was very low and their hexose concentration declined slightly in the darkness and 1 day after defoliation (Fig. 8b).
Fig. 8

Effects of prolonged darkness and complete defoliation on the starch, hexose, and sucrose concentrations of mature leaves (a) and young fruits (b) of cucumber. For the darkness treatment, plants were transferred to a darkened growth chamber at 25 °C in the morning. For the defoliation treatment, plants grown in a glasshouse were completely defoliated and young fruits were harvested the next day. Vertical bars are SE of means (n = 3)

Discussion

It has been suggested that water and minerals enter fruit mainly via phloem (Ho et al. 1987) although this view was recently criticized based on a nuclear magnetic resonance imaging study (Windt et al. 2009). Xylem flow is mainly driven by leaf transpiration (Nobel 2009). These facts suggest that defoliation treatment must have a drastic influence on both phloem and xylem flows, and it is possible that the effects of defoliation can be caused by water and/or ion stresses rather than by photoassimilate starvation. To rule out the possibility that water and/or ion stresses caused the effects of defoliation, prolonged darkness treatments were conducted. Although this kind of treatment also affects phloem and xylem flows, it is much milder than defoliation because no severing of these conduits occurs.

As seen, the transcript level of CsSEF1 increased markedly after prolonged darkness. This strongly suggests that the initial cause of the upregulation of CsSEF1 by defoliation is photoassimilate starvation. Grabowska et al. (2009) reported that the expression of CsSEF1 is correlated with cucumber somatic embryogenesis. To ensure fruit set, we used pollinated fruit as the material. Because the fruits treated with defoliation and prolonged darkness were 1 day older than the control fruits, the increase in the transcript level of CsSEF1 might have been due to developmental changes in the embryo. However, the transcript level of CsSEF1 in the control fruits was very low compared with that in the treated fruits, irrespective of developmental stage, which indicates that the enhancement effect was due to the defoliation or prolonged darkness treatment.

An aspect requiring further examination is whether the site of marked CsSEF1 induction in fruit tissue is concentrated in the embryo. In this regard, it is noteworthy that enhanced expression of CsSEF1 was observed under prolonged darkness not only in fruit, but also in the apical region of lateral shoots and in the roots. However, it was not observed in mature leaves. This suggests that CsSEF1 expression might be confined to rapidly growing sink tissue. It is unclear how to reconcile this view with the induction of CsSEF1 during somatic embryogenesis.

Enhancement of expression of the asparagine synthetase gene has been widely studied, and it is regarded as a particularly good reporter gene for sugar starvation (Lam et al. 1998; Contento et al. 2004; Price et al. 2004; Thum et al. 2004; Blasing et al. 2005). Our results are consistent with this view. It is noteworthy that expression of the asparagine synthetase gene was markedly enhanced under prolonged darkness in leaves, but to a much lesser extent in fruits, although the minor increase was statistically significant. This may indicate that there was marked sugar starvation in the leaves and that the sugar starvation in fruits was much less severe. The patterns of expression of CsSEF1 and the asparagine synthetase gene suggest that enhanced CsSEF1 expression is related to the growth cessation of rapidly growing sink tissue and is not directly related to local sugar starvation. It is possible that CsSEF1 is located somewhere between photoassimilate limitation and growth cessation of the rapidly growing sink (Fig. 9). The temporal coincidence of enhanced CsSEF1 expression and the decline in fruit respiration rate under prolonged darkness also supports a correlation between CsSEF1 induction and growth cessation. In the defoliation treatment, both CsSEF1 and the asparagine synthetase gene were induced. The immediate decline in fruit respiration rate after defoliation suggests that a rapid reduction in photoassimilate supply was occurring, which in turn may have caused the induction of CsSEF1. A more detailed analysis of the time course of the induction of CsSEF1 and the asparagine synthetase gene could be informative in examining such possibilities.
Fig. 9

Schematic drawing of the hypothetical function of CsSEF1 in cucumber plants treated with complete defoliation or prolonged darkness

The observed decline in leaf starch concentration is consistent with photoassimilate depletion, but the absence of a difference between 6 and 27 h in darkness and the lack of a marked decline in hexose concentration are in contrast to the response of CsSEF1 expression and the fruit respiration rate. The relationship between fruit abortion and sugar concentration is controversial (Reed and Singletary 1989; Zinselmeier et al. 1999; Marcelis et al. 2004). Despite years of research, the control mechanism for photoassimilate partitioning remains poorly understood (Gifford and Evans 1981; Giaquinta 1983; Thorne 1985; Ho 1988; Frommer and Ninnemann 1995; Lalonde et al. 2004; Marcelis et al. 2004; Turgeon and Wolf 2009; Wubs et al. 2009; Zhou et al. 2009; Nunes-Nesi et al. 2010; Ruan et al. 2010; Chen and Thelen 2011). It is possible that CsSEF1 is related to the cessation of sink growth, i.e., the reduction in sink activity. Examination of CsSEF1 function in the signal transduction pathway could provide some insight into the control mechanism of photoassimilate partitioning.

Arabidopsis and rice genomes have 68 and 67 CCCH zinc finger genes, respectively (Wang et al. 2008). CCCH proteins are found in a range of organisms, from humans to yeast (Nie et al. 1995; Mello et al. 1996; Thompson et al. 1996; De et al. 1999), and are suggested to be RNA-binding proteins functioning in RNA processing (Lai et al. 2000, 2003). CCCH proteins in plants are poorly characterized compared with those in animals. In Arabidopsis, a CCCH zinc finger gene, HUA1, is reported to participate in the regulation of flower development (Li et al. 2001). Another CCCH zinc finger gene, PE11, is involved in embryogenesis (Li and Thomas 1998). Other reported plant CCCHs include ArabidopsisAtCPSF30 (Delaney et al. 2006), FES1 (Schmitz et al. 2005), and rice OsDOS (Kong et al. 2006). Specifically, CsSEF1 has a tandem CCCH zinc finger (TZF) motif. In animals, TZF proteins are characterized by two identical Cx8Cx5Cx3H motifs separated by 18 amino acids (Blackshear et al. 2005). Each CCCH zinc finger is capable of binding to specific RNA motifs (Carrick et al. 2004; Hudson et al. 2004; Barreau et al. 2005). In humans, the TZF family consists of three genes: TTP, BRF1, and BRF2. The functions of these TZFs, especially TTP, have been well studied, and include mRNA degradation (Carballo et al. 1998; Lai et al. 2006). Arabidopsis, rice, and soybean have 11, 9, and 23 TZF genes, respectively (Pomeranz et al. 2011). However, the function of TZF genes in plants is poorly understood, compared with that in animals (Pomeranz et al. 2011). In plants, TZF genes with a plant-specific motif of Cx7-8Cx5Cx3H-x16-Cx5Cx4Cx3H are found. The CsSEF1 gene has this motif. Unlike other plant CCCH families, there has been no report on the specific RNA that binds to plant TZFs (Pomeranz et al. 2011). In Arabidopsis, a TZF gene, AtTZ1 (AtC3H23), was identified in a transcriptome analysis as a glucose-responsive gene (Price et al. 2004). AtTZ1 binds both DNA and RNA in vitro, and traffics between the nucleus and cytoplasm (Pomeranz et al. 2010). Lin et al. (2011) indicated that the expression of AtTZ1 is reduced by glucose in a hexokinase-dependent manner and suggested that AtTZF1 serves as a regulator connecting sugar, abscisic acid, gibberellic acid (GA), and peptide hormone responses.

Thus, the response of CsSEF1 in the present study seems to be different from that of AtTZF1. As shown in Fig. 3, AtTZF1 (AtC3H23) is moderately close to CsSEF1. It will be interesting to see if the expression of more closely related homologs such as AtC3H49 and AtC3H20 is enhanced by prolonged darkness. If so, it will be interesting to see if such enhancement is influenced in hexokinase knockout plants. Lee et al. (2012) studied AtC3H49 and AtC3H20 in relation to GA and jasmonic acid responses.

In conclusion, our results confirm that the asparagine synthetase gene is a good marker gene of sugar starvation in cucumber fruit. The CsSEF1 gene may be involved in the signal transduction pathway that leads to growth cessation of sink organs, but dependence of CsSEF1 expression enhancement on the hexokinase pathway should be further examined.

Notes

Acknowledgments

We thank Dr. Yoshiteru Sakata of the Institute of Vegetable and Tea Science, National Agriculture and Food Research Organization, for kindly offering the seeds of the cucumber cultivar ‘Tokiwa’. This work was supported by Grant-in-Aid for Scientific Research (C) (22580285) on Priority Areas from the Ministry of Education, Culture, Sports, Science and Technology of Japan to TA.

References

  1. Baena-González E, Rolland F, Thevelein JM, Sheen J (2007) A central integrator of transcription networks in plant stress and energy signaling. Nature 448:938–942PubMedCrossRefGoogle Scholar
  2. Barreau C, Paillard L, Osborne HB (2005) AU-rich elements and associated factors: are there unifying principles? Nucleic Acids Res 33:7138–7150PubMedCrossRefGoogle Scholar
  3. Blackshear PJ, Phillips RS, Lai WS (2005) Tandem CCCH zinc finger proteins in mRNA binding. In: Luchi S, Kuldell N (eds) Zinc finger proteins: from atomic contact to cellular function. Kluwer Academic/Plenum Publishers, Austin, Texas, pp 80–90CrossRefGoogle Scholar
  4. Blasing OE, Gibson Y, Grunther M, Hohne M, Morcuende R, Osuna D, Thimm O, Usadel B, Scheible W-R, Stitt M (2005) Sugars and circadian regulation make major contributions to the global regulation of diurnal gene expression in Arabidopsis. Plant Cell 17:3257–3281PubMedCrossRefGoogle Scholar
  5. Carballo E, Lai WS, Blackshear PJ (1998) Feedback inhibition of macrophage tumor necrosis factor-∝ production by tristetrapolin. Science 281:1001–1005PubMedCrossRefGoogle Scholar
  6. Carrick DM, Lai WS, Blackshear PJ (2004) The tandem CCCH zinc finger protein tristetraprolin and its relevance to cytokine mRNA turnover and arthritis. Arthritis Res Ther 6:248–264PubMedCrossRefGoogle Scholar
  7. Chen M, Thelen JJ (2011) Plastid uridine salvage activity is required for photoassimilate allocation and partitioning in Arabidopsis. Plant Cell 23:2991–3006PubMedCrossRefGoogle Scholar
  8. Contento AL, Kim SJ, Bassham DC (2004) Transcriptome profiling of the response of Arabidopsis suspension culture cells to Suc starvation. Plant Physiol 135:2330–2347PubMedCrossRefGoogle Scholar
  9. Craft AS, Lorentz OA (1944) Fruit growth and food transport in cucurbits. Plant Physiol 19:131–138CrossRefGoogle Scholar
  10. De J, Lai WS, Thorn JM, Goldsworthy SM, Liu X, Blackwell TK, Blackshear PJ (1999) Identification of four CCCH zinc finger proteins in Xenopus, including a novel vertebrate protein with four zinc fingers and severely restricted expression. Gene 228:133–145PubMedCrossRefGoogle Scholar
  11. Delaney KJ, Xu R, Zhang J, Li QQ, Yun KY, Falcone DL, Hunt AG (2006) Calmodulin interacts with and regulates the RNA-binding activity of an Arabidopsis polyadenylation factor subunit. Plant Physiol 140:1507–1521PubMedCrossRefGoogle Scholar
  12. Frommer WB, Ninnemann O (1995) Heterologous expression of genes in bacterial, fungal, animal and plant cells. Annu Rev Plant Physiol Plant Mol Biol 46:419–444CrossRefGoogle Scholar
  13. Giaquinta RT (1983) Phloem loading of sucrose. Annu Rev Plant Physiol 34:347–387CrossRefGoogle Scholar
  14. Gifford RM, Evans LT (1981) Photosynthesis, carbon partitioning, and yield. Annu Rev Plant Physiol 32:485–509CrossRefGoogle Scholar
  15. Goldschmidt EF, Huber SC (1992) Regulation of photosynthesis by end-product accumulation in leaves of plants storeing starch, sucrose, and hexose sugars. Plant Physiol 99:1443–1448PubMedCrossRefGoogle Scholar
  16. Grabowska A, Winsniewska A, Tagashira N, Malepszy S, Filipecki M (2009) Characterization of CsSEF1 gene encoding putative CCCH-type zinc finger protein expressed during cucumber somatic embryogenesis. J Plant Physiol 166:310–323PubMedCrossRefGoogle Scholar
  17. Ho LC (1988) Metabolism and compartmentation of imported sugars in sink organs in relation to sink strength. Annu Rev Plant Physiol Plant Mol Biol 39:355–378CrossRefGoogle Scholar
  18. Ho LC, Grange RI, Picken AJ (1987) An analysis of the accumulation of water and dry matter in tomato fruit. Plant Cell Environ 10:157–162Google Scholar
  19. Hudson BP, Martinez-Yamout MA, Dyson HJ, Wright PE (2004) Recognition of the mRNA AU-rich element by the zinc finger domain of TIS11d. Nat Struct Mol Biol 11:257–264PubMedCrossRefGoogle Scholar
  20. Kato T, Oda H (1977) Studies on the control of physiological disorders in fruit vegetable crops under plastic films. VIII. On the occurrence of abnormal fruits in cucumber plants. (II) On the development of carrot type and bottle gourd type fruits, so-called sakibosori and shiributo fruits in Japan. (Japanese text with English abstract) Res Rep Kochi Univ 26 (Agric Sci):175–182Google Scholar
  21. Kong Z, Li M, Yang W, Xu W, Xue Y (2006) A novel nuclear-localized CCCH-type zinc finger protein, OsDOs, is involved in delaying leaf senescence in rice. Plant Physiol 141:1376–1388PubMedCrossRefGoogle Scholar
  22. Lai WS, Carballo E, Thorn JM, Kennington EA, Blackshear PJ (2000) Interactions of CCCH zinc finger proteins with mRNA. Binding of tristeraprolin-related zinc finger proteins to AU-rich elements and destabilization of mRNA. J Biol Chem 275:17827–17837PubMedCrossRefGoogle Scholar
  23. Lai WS, Kennington EA, Blackshear PJ (2003) Tristeraprolin and its family members can promote the cell-free deadenylation of AU-rich element-containing mRNA by poly(A) ribonuclease. Mol Cell Biol 23:3798–3812PubMedCrossRefGoogle Scholar
  24. Lai WS, Parker JS, Grissom SF, Stumpo DJ, Blackshear PC (2006) Novel mRNA targets for tristetraprolin (TTP) identified by global analysis of stabilized transcripts in TTP-deficient fibroblasts. Mol Cell Biol 26:9196–9208PubMedCrossRefGoogle Scholar
  25. Lalonde S, Wipf D, Frommer WB (2004) Transport mechanisms for organic forms of carbon and nitrogen between source and sink. Annu Rev Plant Biol 55:341–372PubMedCrossRefGoogle Scholar
  26. Lam HM, Hsieh MH, Coruzzi G (1998) Reciprocal regulation of distinct asparagine synthetase genes by light and metabolites in A. thaliana. Plant J 16:345–353PubMedCrossRefGoogle Scholar
  27. Lee S-J, Jung HJ, Kang H, Kim SY (2012) Arabidopsis zinc finger proteins AtC3H49/AtTZF3 and AtC3H20/AtTZF2 are involved in ABA and JA responses. Plant Cell Physiol 53:673–686PubMedCrossRefGoogle Scholar
  28. Li Z, Thomas TL (1998) PEI1, an embryo-specific zinc finger protein gene required for heart-stage embryo formation in Arabidopsis. Plant Cell 10:383–398PubMedGoogle Scholar
  29. Li J, Jia D, Chen X (2001) HUA1 a regulator of stamen and carpel identities in Arabidopsis, code for a nuclear RNA-binding protein. Plant Cell 13:2269–2281PubMedGoogle Scholar
  30. Lin P-C, Pomeranz MC, Jikumaru Y, Kang SG, Hah C, Fujioka S, Kamiya Y, Jang J-C (2011) The Arabidopsis tandem zinc finger protein AtTZF1 affects ABA- and GA-mediated growth, stress and gene expression responses. Plant J 65:253–268PubMedCrossRefGoogle Scholar
  31. Marcelis LFM, Heuvelink E, Baan Hofman-Eijer LR, Den Bakker J, Xue LB (2004) Flower and fruit abortion in sweet pepper in relation to source and sink strength. J Exp Bot 406:2261–2268CrossRefGoogle Scholar
  32. Mello CC, Schubert C, Draper B, Zhang W, Lobel R, Priess JR (1996) The PIE-1 protein and germline specification in C. elegans. Nature 382:710–712PubMedCrossRefGoogle Scholar
  33. Nie XF, Maclean KN, Kumar V, McKay IA, Bustin SA (1995) ERF-2, the human homologue of murine Tis11d early response gene. Gene 152:285–286PubMedCrossRefGoogle Scholar
  34. Nobel PS (2009) Physicochemical and environmental plant physiology. Academic Press, London, pp 439–505Google Scholar
  35. Nunes-Nesi A, Fernie AR, Stitt M (2010) Metabolic and signaling aspects underpinning the regulation of plant carbon nitrogen interactions. Mol Plant 3:973–996PubMedCrossRefGoogle Scholar
  36. Pharr DM, Sox HN, Smart EL, Lower RL (1977) Identification and distribution of soluble saccharides in pickling cucumber plants and their fate in fermentation. J Amer Soc Hort Sci 102:406–409Google Scholar
  37. Pomeranz M, Hah C, Lin P-C, Kang SG, Finer JJ, Blackshear PJ, Jang J-C (2010) The Arabidopsis tandem zinc finger protein AtTZF1 traffics between nucleus and cythplasmic foci and binds both DNA and RNA. Plant Physiol 152:151–165PubMedCrossRefGoogle Scholar
  38. Pomeranz M, Finer J, Jang J-C (2011) Putative molecular mechanism underlying tandem CCCH zinc finger protein mediated plant growth, stress and gene expression responses. Plant Signal Behav 6:647–651PubMedCrossRefGoogle Scholar
  39. Price J, Laxmi A, St Martin SK, Jang JC (2004) Global transcription profiling reveals multiple sugar signal transduction mechanisms in Arabidopsis. Plant Cell 16:2128–2150PubMedCrossRefGoogle Scholar
  40. R Development Core Team (2011) R: a language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. http://www.R-project.org/
  41. Reed AJ, Singletary GW (1989) Roles of carbohydrate supply and phytohormones in maize kernel abortion. Plant Physiol 91:986–992PubMedCrossRefGoogle Scholar
  42. Rolland F, Baena-Gonzales E, Sheen J (2006) Sugar sensing and signaling in plants: conserved and novel mechanisms. Annu Rev Plant Biol 57:675–709PubMedCrossRefGoogle Scholar
  43. Ruan Y-L, Jin Y, Yang Y-J, Li G-J, Boyer JS (2010) Sugar input, metabolism, and signaling mediated by invertase: roles in development, yield potential, and response to drought and heat. Mol Plant 3:942–955PubMedCrossRefGoogle Scholar
  44. Schmitz RJ, Hong L, Michaels S, Amasino RM (2005) FRIGIDA-ESSENTIAL 1 interacts genetically with FRIGIDA AND FRIGIDA-LIKE 1 to promote the winter-annual habit of A. thaliana. Development 132:5471–5478PubMedCrossRefGoogle Scholar
  45. Tamura K, Sanada Y, Tase K, Komatsu T, Yoshida M (2011) Pp6-FEH1 encodes an enzyme for degradation of highly polymerized levan and is transcriptionally induced by defoliation in timothy (Phleum pratense L.). J Exp Bot 62:3421–3431PubMedCrossRefGoogle Scholar
  46. Tazuke A, Sakiyama R (1991) Relationships between growth in volume and respiration of cucumber fruit attached on the vine. J Japan Soc Hort Sci 59:745–750Google Scholar
  47. Thompson MJ, Lai WS, Talor GA, Blackshear PJ (1996) Cloning and characterization of two yeast genes encoding members of the CCCH class zinc finger proteins: zinc finger-mediated impairment of cell growth. Gene 174:225–233PubMedCrossRefGoogle Scholar
  48. Thorne JH (1985) Phloem unloading of C and N assimilates in developing seeds. Annu Rev Plant Physiol 36:317–343CrossRefGoogle Scholar
  49. Thum KE, Shin MJ, Palenchar PM, Kouranov A, Coruzzi GM (2004) Genome-wide investigation of light and carbon signaling in Arabidopsis. Genome Biol 5:R10PubMedCrossRefGoogle Scholar
  50. Turgeon R, Wolf S (2009) Phloem transport: cellular pathways and molecular trafficking. Annu Rev Plant Biol 60:207–221PubMedCrossRefGoogle Scholar
  51. Wang D, Guo Y, Wu C, Yang G, Li Y, Zheng C (2008) Genome-wide analysis of CCCH zinc finger family in Arabidopsis and rice. BMC Genomics 9:44. doi:10.1186/1471-2164-9-44 PubMedCrossRefGoogle Scholar
  52. Windt CW, Gerkema E, Van As H (2009) Most water in the tomato truss is imported through the xylem, not the phloem: a nuclear magnetic resonance flow imaging study. Plant Physiol 151:830–842PubMedCrossRefGoogle Scholar
  53. Wubs AM, Ma Y, Heuvelink E, Marcelis LFM (2009) Genetic differences in fruit-set patterns are determined by differences in fruit sink strength and a source : sink threshold for fruit set. Ann Bot 104:957–964PubMedCrossRefGoogle Scholar
  54. Zhou Y, Chan K, Wang TL, Hedley CL, Offler CE, Patrick JW (2009) Intracellular sucrose communicates metabolic demand to sucrose transporters in developing pea cotyledons. J Exp Bot 60:71–85PubMedCrossRefGoogle Scholar
  55. Zinselmeier C, Jeong B-R, Boyer JS (1999) Starch and the control of kernel number in maize at low water potentials. Plant Physiol 121:25–35PubMedCrossRefGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2012

Authors and Affiliations

  1. 1.College of AgricultureIbaraki UniversityIbarakiJapan

Personalised recommendations