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Keywords:

  • sucrose synthase;
  • sucrose metabolism;
  • Arabidopsis;
  • plant mutant;
  • enzyme kinetics;
  • gene family

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The properties and expression patterns of the six isoforms of sucrose synthase in Arabidopsis are described, and their functions are explored through analysis of T-DNA insertion mutants. The isoforms have generally similar kinetic properties. Although there is variation in sensitivity to substrate inhibition by fructose this is unlikely to be of major physiological significance. No two isoforms have the same spatial and temporal expression patterns. Some are highly expressed in specific locations, whereas others are more generally expressed. More than one isoform is expressed in all organs examined. Mutant plants lacking individual isoforms have no obvious growth phenotypes, and are not significantly different from wild-type plants in starch, sugar and cellulose content, seed weight or seed composition under the growth conditions employed. Double mutants lacking the pairs of similar isoforms sus2 and sus3, and sus5 and sus6, are also not significantly different in these respects from wild-type plants. These results are surprising in the light of the marked phenotypes observed when individual isoforms are eliminated in crop plants including pea, maize, potato and cotton. A sus1/sus4 double mutant grows normally in well-aerated conditions, but shows marked growth retardation and accumulation of sugars when roots are subjected to hypoxia. The sucrose synthase activity in roots of this mutant is 3% or less of wild-type activity. Thus under well-aerated conditions sucrose mobilization in the root can proceed almost entirely via invertases without obvious detriment to the plant, but under hypoxia there is a specific requirement for sucrose synthase activity.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Sucrose synthase is widely believed to be the main route of entry of carbon from sucrose into cellular metabolism in plants. There is direct evidence of its importance in several sink organs of crop plants, where reductions in activity have been shown to reduce the availability of carbon for synthesis of storage products and growth. For example, the rugosus4 (rug4) mutation of pea (Craig et al., 1999) and the shrunken1 (sh1) and sucrose synthase1 (sus1) mutations of maize (Chourey et al., 1998) eliminate isoforms of sucrose synthase that are highly expressed in the developing seed, resulting in reductions in seed sucrose synthase activity and reduced accumulation of starch. The rug4 mutation reduces sucrose synthase activity in the Rhizobium nodules of pea roots as well as in the seed, resulting in a loss of nitrogen-fixing capacity and premature senescence of these organs (Craig et al., 1999; Gordon et al., 1999). Tubers of transgenic potato plants with reduced sucrose synthase activity also accumulate less starch than wild-type tubers (Zrenner et al., 1995). In transgenic cotton plants with reduced sucrose synthase activity, fibre cell initiation and elongation and seed development are compromised (Ruan et al., 2003).

In species examined to date, sucrose synthase is encoded by a small multigene family. Analysis of transgenic and mutant crop plants reveals that particular isoforms have specific functions within the plant. For example, the rug4 mutations of pea eliminate the SUS1 isoform but have no effect on SUS2 and SUS3, suggesting that these two isoforms are unable to compensate for the loss of SUS1 in the seed and the root nodule (Barratt et al., 2001). In potato, reductions in SUS1 affect the tuber but reductions in SUS2 have minimal effects on metabolism (Weckwerth et al., 2004; Zrenner et al., 1995). In maize, loss of the SH1 isoform has different effects from loss of the SUS1 isoform. Both isoforms are necessary for wild-type rates of starch synthesis, but SH1 is also necessary for normal cell wall formation during endosperm development (Chourey et al., 1998).

The reasons for this differentiation of function of isoforms of sucrose synthase within the plant are not clear. Different isoforms may fulfil identical functions within the cell but operate in distinct cell types, developmental periods or environmental conditions. However, it is also possible that different isoforms may have distinct, non-overlapping functions within the same cell. Although much of the enzyme is apparently soluble, and presumably mediates the conversion of sucrose to a pool of hexose phosphates available for general cellular metabolism, a significant fraction appears to be membrane-bound in a wide variety of plant organs (Amor et al., 1995; Etxeberria and Gonzalez, 2003; Winter et al., 1997), and some may also be associated with the cytoskeleton (Azama et al., 2003; Winter et al., 1998). Several functions for membrane-associated sucrose synthase have been postulated. First, it may supply uridine diphosphate glucose (UDPglucose) to cellulose and callose synthases on the inner face of the plasma membrane (Amor et al., 1995; Subbaiah and Sachs, 2001). Second, it may perform the same function for xyloglucan synthesis, associated with the membrane of the Golgi apparatus (Buckeridge et al., 1999; Konishi et al., 2004). Third, it may be involved in transport of sucrose across the tonoplast from the vacuole to the cytosol in sucrose-storing organs (Etxeberria and Gonzalez, 2003). There is indirect evidence that different isoforms contribute to the soluble and membrane bound activities of sucrose synthase. Although both the SH1 and SUS1 isoforms of maize are necessary for normal starch synthesis in the developing endosperm, only SH1 appears to be necessary for normal cell wall synthesis (Chourey et al., 1998). However, post-translational modification of the enzyme may also determine whether it is membrane associated. There is both in vitro and in vivo evidence that phosphorylation of the sucrose synthase protein decreases its association with membranes (Hardin et al., 2004; Huber et al., 1996; Subbaiah and Sachs, 2001; Winter et al., 1997; Zhang et al., 1999).

Insufficient information exists about the sucrose synthase gene family in any plant to resolve the questions about the precise roles of its members in metabolism. No systematic analysis of properties of isoforms encoded by the gene family has been carried out, and although the roles of a few individual isoforms have been probed in transgenic and mutant plants there has been no systematic functional analysis for the gene family. The model plant Arabidopsis offers the opportunity for such an analysis. The Arabidopsis genome contains six sucrose synthase genes. The isoforms they encode fall into three distinct pairs based on comparisons of amino acid sequences (Baud et al., 2004). SUS1 and SUS4 are 89% identical to each other, but less than 68% identical to other isoforms. SUS2 and SUS3 are 74% identical to each other but 67% or less identical to other isoforms, and SUS5 and SUS6 are 58% identical to each other but 48% or less identical to other isoforms. SUS5 and SUS6 have C-terminal extensions of 3 and 14 kDa, respectively, relative to other isoforms. At least two of the three pairs of isoforms appear to be represented in other dicotyledonous species. Phylogenetic analysis shows that AtSUS1 and AtSUS4 are closely related to pairs of isoforms from potato (Solanacae), pea (Fabacae) and carrot (Umbelliferae). AtSUS2 and AtSUS3 are closely related to a pair of isoforms from Craterostigma plantagineum (Scrophulariacae) (Barratt et al., 2001; Baud et al., 2004). The third pair, AtSUS5 and AtSUS6, is closely related to a pair of sucrose synthase genes from rice (OsSUS5 and OsSUS6; Harada et al., 2005). Thus the three pairs of isoforms in Arabidopsis are unlikely to be the result of recent gene duplication events. Each isoform may have a specific function conserved in a wide range of plants. Arabidopsis is also typical of plants studied so far in that members of the gene family are strongly differentially expressed in different organs of the plant, through development and in response to environmental stress (Baud et al., 2004).

Our aims were first to discover whether the Arabidopsis isoforms show major differences in properties, second to gather and collate evidence about their patterns of expression in the plant, and third to provide information about their functions through examination of T-DNA insertion lines (referred to as knockout mutants) lacking individual isoforms, or the pairs of isoforms described above. Our examination of knockout mutants focused on three characters that might be expected to be affected by loss of specific isoforms: sugar and starch content in leaves at the end of the night and at the end of the day, which would be affected by loss of sink capacity elsewhere in the plant and by reduced energy supply for phloem loading (Martin et al., 1993); seed weight and composition, which are affected by the loss of specific isoforms in pea and maize (Chourey et al., 1998; Craig et al., 1999); and cellulose content, which would be affected if specific isoforms contribute carbon from sucrose to cellulose synthesis. These experiments uncovered a growth and metabolic phenotype in a line lacking both the sus1 and sus4 isoforms. We report further experiments in which the role of these isoforms in tolerance of hypoxia in the roots was investigated.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Kinetic characteristics of sucrose synthase isoforms

Full-length cDNAs encoding each of the six isoforms of sucrose synthase (SUS1–6) were cloned into pET3d expression vectors to facilitate isopropyl-β-d-thiogalactopyranoside (IPTG)-inducible expression of the complete (untagged) natural polypeptide sequence in Escherichia coli. Five of the six constructs yielded soluble, kinetically active enzyme. The remaining isoform, SUS2, was essentially insoluble under the expression conditions employed.

Based on the phylogenetic relationship between the six SUS genes in Arabidopsis, we selected four of the expressed proteins for detailed study. These were the closely-related isoforms SUS1 and SUS4, and the more distantly related isoforms SUS3 and SUS6. SUS6 is one of the two isoforms possessing novel carboxy-terminal extensions relative to the four other isoforms. The four selected isoforms were purified to near homogeneity by a combination of polyethylene glycol precipitation, anion-exchange chromatography on Q-Sepharose and immobilized metal affinity chromatography on a Ni-NTA (nickel-agarose) column. This procedure typically yielded 0.25–1.5 mg of enzyme from 1.6 l of culture, at a specific activity of 3–12 μmol min−1 (mg protein)−1 (Supplementary Table S1). The specific activity of these isoforms is comparable to that of sucrose synthase recently purified from other plant sources (Barratt et al., 2001; Matic et al., 2004; Römer et al., 2004).

All four isoforms displayed similar pH profiles with maximum activity at pH 6.0–6.5 in the direction of sucrose degradation, and at pH 9.0–9.5 in the direction of sucrose synthesis (data not shown). Accordingly, kinetic characterization was undertaken at pH 6.0 and pH 9.0 in the degradative and synthetic directions, respectively. For SUS1 we confirmed that the response of the enzyme to variation in substrate concentrations measured at pH 7.5 was qualitatively identical to that observed at the pH optima, as described below (data not shown).

Initial studies of the response of SUS1 activity to changes in the concentration of substrates suggested that the enzyme operates by a ternary complex mechanism. However, initial-rate kinetic studies in the absence of product are unable to distinguish the order of binding of substrate and thus provide an incomplete description of the reaction catalysed by the enzyme. Therefore we undertook a comprehensive product-inhibition analysis in which the effect on SUS activity of each product, in isolation, was examined by varying the concentration of one of the substrates at fixed concentrations of the co-substrate. The patterns of inhibition obtained in these studies were entirely consistent with an ordered reaction mechanism in which UDPglucose and UDP are the first substrates to bind, and the last product to be released, depending on the direction of the reaction (Supplementary Table S2). The same conclusion has recently been reached from a more restricted kinetic analysis of a SUS isoform partially purified from sugarcane (Schäfer et al., 2004). Consistent with this proposal, we were able to demonstrate that glucose is an uncompetitive inhibitor of SUS1 activity with respect to both UDPglucose (Kiu = 98.8 ± 5.4 mm) and fructose (Kiu = 82.3 ± 10.1 mm) when operating in the direction of sucrose formation (Supplementary Figure S1). This kinetic response, which is similar to that reported for SUS from maize endosperm (Doehlert, 1987), is most readily explained by glucose binding to the enzyme–UDP complex, probably at the site vacated by sucrose. This mechanism of inhibition would be similar to that resulting in substrate inhibition by high concentrations of fructose (see below).

Having established the reaction mechanism of SUS it was possible to interpret initial-rate kinetic data for each of the four isoforms obtained by varying the concentration of one substrate at each of several fixed concentrations of the co-substrate for the enzyme operating in the direction of sucrose synthesis or sucrose degradation. There are several statistically significant differences in the kinetic constants between the four SUS isoforms (Table 1, see also Supplementary Figure S2 for representative data). However, there is no obvious correlation between the kinetic properties of the different isoforms and their degree of sequence similarity. Despite the presence of a novel carboxy-terminal extension, the kinetic characteristics of SUS6 are within the range exhibited by the other isoforms. The most marked kinetic difference between the isoforms is the variation in sensitivity to substrate inhibition by fructose, even between isoforms exhibiting close sequence similarity (Figure 1). We observed a similar difference in sensitivity to substrate inhibition between a closely related pair of SUS isoforms in our earlier studies on pea (PsSUS1 and PsSUS3; Barratt et al., 2001). This inhibition arises because of the ability of fructose to bind to the enzyme–UDP complex forming a dead-end ternary complex. However, while useful in confirming the order in which substrates bind to the enzyme, it is unlikely that the Kis values observed are sufficiently low for these differences to be physiologically significant.

Table 1.   Comparison of initial rate kinetic constants of Arabidopsis sucrose synthase isoforms expressed in E. coli. Kinetic parameters were determined from rates measured in the direction of sucrose synthesis (synthetic) and degradation (degradative). The concentrations of the reactants were varied over the following ranges: fructose, 0.5–200 mm; UDPglucose, 0.02–5 mm; sucrose, 5–200 mm; UDP, 0.02–5 mm. Data were fitted to the equation describing a freely reversible compulsory ordered ternary complex reaction mechanism including inhibition by the second substrate (Kis) using non-linear regression analysis. Uniform weighting was applied to all data points. Michaelis and inhibition constants are expressed in mm and maximum catalytic activity is expressed as μ mol min−1 (mg protein)−1. Each value is the best-fit estimate ± SE obtained from 192–240 measurements. For each kinetic parameter, values with the same superscript are significantly different from each other as determined by Student's t-test (two-tailed, P < 0.05). The results are representative of the constants obtained from kinetic analysis of at least two independent preparations of each isoform
Kinetic constantSucrose synthase isoform
SUS1SUS3SUS4SUS6
Synthetic
 SynthVmax11.14 ± 0.30a7.32 ± 0.44ab12.01 ± 0.45bc3.27 ± 0.18abc
 FruKm4.37 ± 0.30a41.96 ± 4.01ab4.97 ± 0.35bc12.44 ± 1.29abc
 UDPglcKm0.0439 ± 0.0036a0.1968 ± 0.0172ab0.0687 ± 0.0065ab0.0556 ± 0.0119b
 UDPglcKi0.0173 ± 0.0063a0.1106 ± 0.0170ab0.0470 ± 0.0086ab0.0534 ± 0.0193b
 FruKis107.4 ± 6.58a169.2 ± 20.6ab25.04 ± 1.49abc95.8 ± 9.37bc
Degradative
 DegVmax4.68 ± 0.08a3.03 ± 0.18ab4.99 ± 0.13abc1.43 ± 0.08abc
 SucKm31.58 ± 1.50a108.20 ± 13.04ab67.54 ± 4.39abc31.06 ± 1.86bc
 UDPKm0.0853 ± 0.0053a0.0910 ± 0.0203b0.0735 ± 0.0098c0.2633 ± .0494abc
 UDPKi0.0582 ± 0.01160.0498 ± 0.01970.0938 ± 0.01650.0773 ± 0.0133
 SynthVmax:DegVmax2.382.422.412.29
image

Figure 1.  Substrate inhibition of SUS activity by fructose. (a) The activities of SUS1 (circles), SUS3 (squares), SUS4 (triangles) and SUS6 (inverted triangles) were measured in the direction of sucrose synthesis in the presence of 1 mm UDPglucose. Fructose was varied between 0.5 and 200 mm, as shown. (b) SUS4 activity was measured in the presence of 0.02 (circles), 0.05 (squares), 0.2 (triangles) and 2.0 (inverted triangles) mm UDPglucose; similar curves at 0.1, 0.5, 1.0 and 5 mm UDPglucose are omitted for clarity. Fructose was varied as shown. Each value is the mean ± SE of three measurements and is expressed relative to the maximum activity measured at the concentration of UDPglucose used in the assay.

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Expression patterns of SUS genes in the plant

Quantitative data on transcript levels of the six isoforms of sucrose synthase within the Arabidopsis plant are available from real-time PCR experiments (expressed as a percentage of the transcript level of the EF1A4 α gene; Baud et al., 2004), and from multiple microarrays available via the Gene Chronologer function at http://www.genevestigator.ethz.ch/mm/. These two sources are not directly comparable because different stages of development and organs have been used, and the bases of comparison of expression levels are different. They give different impressions of the transcript distributions for some isoforms. For example, for roots Baud et al. (2004) report that SUS4 transcript levels are almost 100 times higher than those of SUS1, but Gene Chronologer reports that they are about 25% lower than those of SUS1. Baud et al. (2004) report that SUS5 and SUS6 transcript levels are comparable with or higher than those of SUS1 in roots, stems, leaves and flowers, but Gene Chronologer reports that SUS1 transcript levels are four or more times higher than those of either SUS5 or SUS6 in all these organs.

To provide further information on the expression patterns of SUS genes, we performed semi-quantitative RT-PCR analysis using gene-specific primers on total RNA extracted from large number of organs and developmental stages (Figure 2a). We also performed real-time quantitative RT-PCR analysis on a number of organs, in which transcript levels were expressed as a percentage of the expression of the ubiquitin gene (Figure 2b). Again, these two methods do not give identical results. However, if these data sets are considered together with those of Baud et al. (2004) and Gene Chronologer, several important and consistent features emerge. First, no two isoforms have the same distribution pattern. Second, some isoforms are highly expressed in a few organs or developmental stages and expressed at much lower levels elsewhere, while others are expressed across a wide range of organs and stages. At one extreme, high levels of SUS2 transcript are seen only in seeds during a particular phase of development; in contrast, SUS1, SUS5 and SUS6 are much more widely expressed. Third, more than one isoform is expressed in all of the organs examined, but relative expression levels can change dramatically through organ development. For example, levels of SUS2 transcript rise to a peak and then fall dramatically during seed development, whereas SUS3 transcript appears in mid seed development, increases in abundance as the seed matures, and is still present at high levels in the germinating seed. During leaf development, levels of some transcripts are unchanged, others fall during senescence, and one, SUS3, rises dramatically during senescence (data not shown).

image

Figure 2.  Analysis of spatial patterns of SUS transcript abundance. (a) Semi-quantitative RT-PCR. All organs apart from roots, cultivated shoots and germinating seeds were harvested from glasshouse-grown plants at the stage of flowering and setting seed. Pedicels were taken from immature siliques at a range of developmental stages. Silique samples consisted of whole, seed-containing siliques. Siliques and seeds were harvested at the number of days after flowering (Daf) indicated. Roots and cultured shoots were from 19-day-old plants grown in sterile agar containing MS-sucrose, at 20°C and with a 16-h photoperiod. Germinating seedlings were grown for 5 days in water with gentle shaking, at 20°C and with a 16-h photoperiod. (b) Real-time quantitative PCR. Transcript levels are expressed as a percentage of the transcript level of ubiquitin 10 (At4g05320) in the same organ. Values are the means and standard deviation of three or four replicates. Organs were harvested from 7-week-old plants grown in a soil/vermiculite mixture at 20°C with a 16-h photoperiod. Silique samples contained seeds and represented a full range of developmental stages.

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To provide a more detailed picture of expression patterns within organs, we transformed wild-type plants with constructs containing the GUS reporter gene on one of the SUS gene promoter regions. Plants were stained for GUS activity at various developmental stages. On a broad scale, results were consistent with those from measures of transcript abundance described above. Detailed examination revealed considerable, specific localization of expression in particular cell types and tissues. The following observations were made on multiple lines of the relevant transformants. First, expression of several isoforms appeared to be largely in the vasculature. Although SUS1 is widely expressed in the plant, its expression was clearly confined to the vasculature in cotyledons, mature leaves and siliques (Figure 3a). Expression of SUS5 and SUS6 is also vascular in cotyledons, leaves, petals, anthers and roots (Figure 3b), and expression of SUS3 is vascular in cotyledons. Second, detailed patterns of expression within a single organ differ from one isoform to another. For example, in roots prior to flowering SUS5 and SUS6 are expressed in the vasculature but not in the tips, SUS3 is expressed throughout the mature parts of the root (not confined to the vasculature) and also in the tip but not in the expanding zone, and SUS2 is expressed only in the tip (Figure 3c). In mature leaves, SUS3 is specifically expressed in stomatal guard cells (Figure 3d).

image

Figure 3.  Analysis of spatial patterns of SUS gene expression, using promoter–GUS fusions. Seedlings (whole seedling, cotyledon and root pictures) were grown in sterile agar containing MS-sucrose. Mature plants were grown either in a growth chamber or in a glasshouse. The plants shown in (a) were transformed with a construct employing the promoter fragment from Set 1 (see Appendix S2 in Supplementary Material); all other plants were transformed with constructs employing promoter fragments from Set 2. (a) SUS1 promoter: seedling, mature leaf and developing silique. (b) SUS6 promoter: flower and cotyledons. (c) Seedling roots: from left to right SUS2, SUS3 and SUS6 promoters. For SUS2 and SUS6 seedlings were 7 days old; for SUS3 the seedling was 16 days old. (d) SUS3 promoter: surface of mature leaf.

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Selection and validation of knockout mutants

To examine the roles of the SUS isoforms in the plant, we obtained T-DNA insertion lines (referred to as knockout mutants) for each isoform. The authenticity of the knockout mutations was checked in three main ways. First, for each of the isoforms except SUS4, we obtained more than one mutant, and used these multiple lines in our subsequent analyses. Second, we checked for each of the putative knockout lines that transcript from the relevant SUS gene was absent. Third, for SUS1, SUS2, SUS3 and SUS4, we used specific antisera to check that the relevant protein was also absent.

The origins of the lines used in this study are given in Table 2. Where possible, we used mutant lines in a Col background, and selected a congenic wild-type line as a specific control. For SUS4, no suitable knockout lines are available in public collections. Our single line was the kind gift of Sergei Kushnir (University of Gent; see Babiychuk et al., 1997). This mutation was initially in the ecotype C24. We backcrossed this line six times into the ecotype Col, selecting each time for the T-DNA insertion in the SUS4 gene, and used the resulting line and a congenic wild-type line in our analyses. For all of the lines listed with the exception of the sus1 knockout lines SALK-014303 and SAIL-1226-F04, RT-PCR using isoform-specific primers revealed that transcript was absent. For the two sus1 knockout lines, truncated transcripts were still present in the plant (data not shown).

Table 2.   T-DNA insertion lines used in this study. Col is ecotype Colombia; Ws is ecotype Wassilewskija. Evidence about the presence of the protein was gained from the use of specific antisera (see Figure 4)
GeneT-DNA insertion linesLocation of insertionEcotypePCR evidenceProtein evidence
  1. aLines from which double mutant lines were produced.

  2. bLine was originally in ecotype C24, and was backcrossed six times into Col-0 before experiments commenced.

At5g20830 (SUS1)SALK-014303aExonCol-0Truncated transcriptProtein missing
SAIL-1226-F04ExonCol-0Truncated transcriptProtein missing
At5g49190 (SUS2)SALK-076303aIntron/exon boundaryCol-0Transcript missingProtein missing
GABI-377G03ExonCol-0Transcript missingNo data
At4g02280 (SUS3)WISCExonWsTranscript missingProtein missing
SALK-019405aExonCol-0Transcript missingProtein missing
At3g43190 (SUS4)GENT sus4aExonCol-0bTranscript missingProtein missing
At5g37180 (SUS5)SALK-065271IntronCol-0Transcript missingNo data
SALK-102146ExonCol-0Transcript missingNo data
SALK-152944aExonCol-0Transcript missingNo data
GABI-022D04ExonCol-0Transcript missingNo data
At1g73370 (SUS6)WISCExonWsTranscript missingNo data
SAIL-374-D07aExonCol-0Transcript missingNo data
SALK-019129IntronCol-0Transcript missingNo data

To check that the relevant isoform proteins were eliminated in the knockout mutants, we raised and purified antisera to unique peptides for each isoform. The specificity of the antisera was checked on recombinant proteins purified from an E. coli expression system (data not shown). Each of the antisera except that for SUS4 specifically recognized the protein to which it was raised. Antisera raised to two different peptides from SUS4 (the SUS4 and SUS4r peptides: see Supplementary Material, Appendix S2) recognized SUS4, but one also recognized SUS1 (SUS4r peptide) and the other SUS3 (SUS4 peptide). However, use of these antisera in tandem with the specific antisera for SUS1 and SUS3 enables specific recognition of SUS4.

The antisera for SUS1, SUS2, SUS3 and SUS4 recognized bands of the correct molecular mass on blots of crude extracts of Arabidopsis organs from mature plants in soil. SUS1 protein was detected in flowers and roots, SUS4 protein in roots, SUS2 protein in the mid-stage of seed development and SUS3 protein in mature and imbibed seeds (Figure 4). Levels of both SUS1 and SUS4 protein in roots were strongly enhanced by anoxia, consistent with reports that expression of these isoforms is enhanced by anoxia (Baud et al., 2004; Martin et al., 1993). When extracts of knockout mutants for these four isoforms were probed with the corresponding antisera, no proteins of the expected molecular mass were detected (Figure 4). For both of the sus1 lines no proteins were detected that might represent the products of the truncated transcripts. This indicates strongly that the transcripts are either not translated or result in unstable proteins: thus these lines can be considered to have no SUS1 activity.

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Figure 4.  Presence of SUS proteins in wild-type and mutant plants. Panels show immunoblots of SDS-polyacrylamide gels of soluble proteins from the organs and conditions indicated. For each panel, all lanes are from the same gel and blot, and contain the same amount of protein. (a) Blot developed with antiserum to SUS1. Anoxia was imposed by immersing the entire plant in degassed water for 3 days prior to harvest of the root system. Note the following: (i) the antiserum does not recognize the SUS4 protein; (ii) flooding increases the amount of SUS1 protein in wild-type roots, but the protein is not detectable under normal or flooded conditions in the two mutant lines. (b) Blot developed with antiserum to SUS4. Anoxia was imposed as in (a); hypoxia was imposed by flooding the root systems but not the aerial parts of the plant for 2 days in degassed water. Note the following: (i) the antiserum does not recognize the SUS1 protein. It recognizes the SUS3 protein (see text), but SUS3 protein is not detectable in roots using a specific antiserum with a higher affinity for SUS3 (not shown); (ii) hypoxia increases the amount of SUS4 protein in both wild-type roots and roots of a sus1 mutant, but the protein is not detectable under normal, hypoxia or anoxic conditions in roots of the sus4 mutant. (c) Blot developed with antiserum to SUS2. Note that amounts of SUS2 protein peak at 12–14 days after flowering in wild-type seeds, but the protein is not detectable in seeds of a sus2 mutant. (d) Blot developed with antiserum to SUS3. Note that SUS3 protein is present in dry and imbibed wild-type seeds, but is not detectable in seeds of two sus3 mutant lines.

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The antisera for SUS5 and SUS6 failed to detect proteins in soluble extracts of roots, leaves, stems, flowers and fruits of Arabidopsis plants, and in membrane-containing fractions (100 000 g pellet from the soluble extract) from these organs, under normal or anoxic conditions (not shown). This may reflect very low levels of these proteins.

Phenotypic characterization of the knockout mutants

From visual inspection, plants lacking any one of the six isoforms grew at the same rate as wild-type plants, and displayed the same phenology and vegetative and reproductive morphology. This was true in both the growth room for 12 and 16-h days and the glasshouse under various day lengths and temperatures. We examined in more detail three specific characteristics of these plants that might be expected to be affected by loss of sucrose synthase isoforms: sugar and starch content, cellulose content and the weight and lipid content of mature seeds.

For analysis of sugar and starch content, leaves and roots of 5-week-old plants grown in hydroponic culture were harvested at the end of the day and at the end of the night. The amounts of glucose, fructose, sucrose and starch in each of the knockout lines were not statistically significantly different from those of the equivalent wild-type lines grown under the same conditions at the same time (Table 3). The cellulose content of leaves and roots was measured on the same plants: again there were no significant differences between any of the mutant lines and the equivalent wild-type lines (Table 4).

Table 3.   Starch and sucrose content of leaves and roots of wild-type and mutant plants. Plants were grown hydroponically, and harvested when they were approximately 5 weeks old. Samples were taken at the end of the 16-h light period and at the end of the 8-h dark period. Samples consisted of a whole rosette (Leaf) or a whole root system (Root). Values are means ± SD of measurements on five different samples. Col A, Col B and Ws are wild-type controls; Col A and Col B are the controls for batches of plants grown at two different times. For each mutant line, the relevant wild-type control line is indicated in parenthesis
LineCarbohydrate content [μmol glucose equivalents (g fresh weight)−1]
LeafRoot
StarchSucroseStarchSucrose
LightDarkLightDarkLightDarkLightDark
Col A18.5 ± 4.98.5 ± 3.44.2 ± 1.43.2 ± 0.90.03 ± 0.010.05 ± 0.020.9 ± 0.11.0 ± 0.14
Col B8.1 ± 1.85.2 ± 1.65.5 ± 1.64.2 ± 1.20.07 ± 0.050.04 ± 0.021.36 ± 1.651.18 ± 0.45
Ws24.4 ± 4.214.2 ± 4.35.1 ± 0.95.5 ± 1.80.05 ± 0.030.05 ± 0.021.13 ± 0.261.03 ± 0.19
sus1 (SALK; Col A)20.9 ± 5.67.6 ± 2.74.3 ± 1.43.3 ± 0.70.04 ± 0.010.06 ± 0.010.94 ± 0.131.34 ± 0.33
sus2 (SALK; Col A)18.9 ± 2.29.2 ± 2.34.2 ± 1.03.4 ± 0.50.06 ± 0.020.07 ± 0.020.80 ± 0.181.07 ± 0.26
sus2 (GABI; Col A)21.6 ± 1.76.7 ± 0.74.7 ± 0.63.2 ± 0.60.04 ± 0.010.04 ± 0.010.72 ± 0.220.93 ± 0.17
sus3 (SALK; Col A)21.5 ± 2.48.9 ± 2.66.1 ± 1.62.7 ± 0.40.05 ± 0.020.07 ± 0.040.78 ± 0.061.24 ± 0.37
sus3 (WISC; Ws)27.7 ± 7.514.4 ± 5.27.8 ± 2.25.6 ± 2.10.05 ± 0.010.08 ± 0.031.53 ± 0.461.79 ± 0.69
sus4 (GENT; Col B)9.5 ± 1.26.2 ± 0.94.1 ± 1.23.7 ± 0.90.25 ± 0.170.05 ± 0.031.65 ± 0.371.36 ± 0.16
sus5 (SALK- 065271; Col A)22.4 ± 3.39.5 ± 3.75.2 ± 0.92.8 ± 0.60.04 ± 0.010.06 ± 0.000.99 ± 0.241.11 ± 0.06
sus5 (GABI; Col A)20.3 ± 4.411.1 ± 2.95.4 ± 0.83.5 ± 0.30.04 ± 0.020.05 ± 0.020.71 ± 0.101.17 ± 0.17
sus6 (SALK; Col A)20.3 ± 1.79.5 ± 3.64.6 ± 1.22.4 ± 1.50.04 ± 0.010.08 ± 0.040.94 ± 0.081.14 ± 0.17
sus6 (WISC; Ws)25.2 ± 4.316.9 ± 3.66.8 ± 2.14.4 ± 1.00.08 ± 0.060.09 ± 0.021.18 ± 0.291.05 ± 0.06
Table 4.   Cellulose content of leaves and roots of wild-type and mutant plants. Plants were grown hydroponically, and harvested when approximately 5 weeks old. Samples were taken at the end of the 8-h dark period. Samples consisted of a whole rosette (Leaf) or a whole root system (Root). Values are means ± SD of measurements on five different samples. Col A, Col B and Ws are wild-type controls; Col A and Col B are the controls for batches of plants grown at two different times. For each mutant line, the relevant wild-type control line is indicated in parenthesis
LineCellulose content [μmol glucose equivalents (g fresh weight)−1]
LeafRoot
Col A25.3 ± 3.67.5 ± 2.1
Col B8.1 ± 4.75.0 ± 0.7
Ws21.4 ± 5.37.0 ± 1.9
sus1 (SALK; Col A)26.7 ± 3.610.4 ± 2.7
sus2 (SALK; Col A)26.7 ± 3.68.3 ± 2.5
sus2 (GABI; Col A)28.0 ± 6.68.7 ± 2.1
sus3 (SALK; Col A)20.8 ± 4.810.0 ± 2.6
sus3 (WISC; Ws)16.2 ± 14.79.1 ± 2.1
sus4 (GENT; Col B)9.8 ± 3.08.7 ± 2.4
sus5 (SALK- 065271; Col A)14.4 ± 6.69.8 ± 2.9
sus5 (GABI; Col A)23.1 ± 6.88.3 ± 1.9
sus6 (SALK; Col A)17.8 ± 7.09.2 ± 3.0
sus6 (WISC; Ws)21.0 ± 3.510.2 ± 3.3

To minimize extraneous effects on seed weights and compositions, plants were grown from seeds harvested at the same time from the same environment and subjected to the same storage conditions. All of the plants were grown at the same time in the same conditions, and different lines were randomly distributed on the glasshouse bench. The timing of seed harvest and subsequent storage was also the same for all lines. There was a clear distinction in both seed weight and seed lipid content between lines in a Col background and lines in a Ws background (Table 5). However, there were no statistically significant differences in either seed weight or lipid content between any of the lines in the same background.

Table 5.   Seed weight and seed lipid content of wild-type and mutant plants. Values are means ± SD. For seed weight, values are from six samples taken from two independent pools of seeds, each from five plants. For percentage lipid, values are from five samples, each from an independent pool of seeds from five plants
LineSeed weight (mg)Percentage lipid
  1. n.d., not determined.

Ws0.0138 ± 0.000633.5 ± 0.7
Col0.0146 ± 0.000226.2 ± 1.3
sus1 (SALK; Col)0.0138 ± 0.000826.4 ± 1.6
sus1 (SAIL; Col)0.0154 ± 0.000726.8 ± 1.0
sus2 (SALK; Col)0.0140 ± 0.000525.6 ± 1.0
sus3 (SALK; Col)0.0143 ± 0.000626.1 ± 1.4
sus3 (WISC; Ws)0.0144 ± 0.000732.3 ± 1.3
sus4 (GENT; Col)0.0144 ± 0.000427.3 ± 1.5
sus5 (SALK-065271; Col)n.d.26.6 ± 1.1
sus5 (SALK-102146; Col)0.0140 ± 0.000326.5 ± 1.9
sus5 (SALK-152944; Col)0.0141 ± 0.000425.7 ± 1.7
sus6 (WISC; Ws)0.0140 ± 0.000734.0 ± 0.8
sus6 (SAIL; Col)0.0146 ± 0.000425.8 ± 2.0

Phenotypic characterization of double mutants

The lack of obvious phenotypes in the knockout mutants suggests a high degree of functional overlap between them, or between SUS and invertase. To explore this further, we selected three double mutants, sus1/sus4, sus2/sus3 and sus5/sus6. Each of these lacks one of the three pairs of SUS isoforms described in the Introduction. Double mutants were selected from F2 populations of crosses between lines in a Col-0 background (see Table 2) by screening for plants homozygous for T-DNA insertions in both SUS genes. For sus1/sus4 and sus2/sus3 mutants, the loss of the SUS protein products was confirmed using the specific antisera, as described for the single knockout lines above (data not shown). When grown in soil, the double mutant lines were not obviously different from wild-type controls. When grown hydroponically (as for sugar, starch and cellulose measurements), sus2/sus3 mutants and sus5/sus6 mutants again looked like wild-type controls, but sus1/sus4 mutants grew considerably more slowly (not shown).

The three double mutants were subjected to the same analyses as the single knockout lines. For measurements of seed weight and seed lipid content, double mutant lines were compared with a wild-type line selected from the same F2 population. There were minimal differences between the lines and their respective wild-type controls (Table 6). Seed weight of the sus1/sus4 mutant was significantly reduced relative to that of its wild-type control, but not relative to that of Col-0 plants grown in the same randomized batch. For cellulose, starch and sugar content, there were no significant differences between sus2/sus3 or sus5/sus6 and the wild-type control, in roots or leaves (Table 7a and b). The sus1/sus4 double mutant was also essentially the same as its wild-type control with respect to cellulose and starch content, and the sugar content of the roots. However, the sugar content of leaves was substantially higher than that of the wild-type control. This was most pronounced at the end of the night, when glucose and fructose were almost ninefold higher in mutant than wild-type leaves and sucrose was more than twofold higher.

Table 6.   Seed weight and seed lipid content of wild-type and double mutant plants. The control for each line was a wild-type line selected from the segregating F2 population from which the double mutant was selected. Values are means ± SD. For seed weight, values are from six samples taken from two independent pools of seeds, each from five plants. For percentage lipid, values are from six samples, each from an independent pool of seeds from five plants. Seed weight of sus1/sus4 is statistically significantly different from that of its control (P = 5.5 × 10−6, Student's t-test) but not from that of Col-0
LineSeed weight (mg)Percentage lipid
sus1/sus40.0153 ± 0.000532.81 ± 1.06
Control for sus1/sus40.0181 ± 0.000535.09 ± 1.64
sus2/sus30.0162 ± 0.000635.80 ± 1.84
Control for sus2/sus30.0167 ± 0.000435.79 ± 2.10
sus5/sus60.0150 ± 0.000333.79 ± 1.37
Control for sus5/sus60.0167 ± 0.001134.29 ± 2.85
Col00.0165 ± 0.000733.62 ± 1.99
Table 7.   Starch, sugar and cellulose content of leaves and roots of wild-type and double mutant plants. Plants were grown and harvested as described in Table 4. Measurements of cellulose were made on plants harvested at the end of the night. Values are means ± SD of measurements on five different samples. (a) Starch, sugar and cellulose contents of leaves. Values for sus1/sus4 for glucose, fructose, sucrose and starch are all significantly different from those for Col-0 at the end of the night (P values 0.0006, 0.003, 0.001, 0.023, respectively) and values for glucose, fructose and sucrose are different at the end of the day (P values 0.003, 0.012, 0.005, respectively, Student's t-test). (b) Starch, sugar and cellulose contents of roots
LineCarbohydrate content [μmol glucose equivalents (g fresh weight)−1]
StarchSucroseGlucoseFructoseCellulose
DayNightDayNightDayNightDayNight 
(a) Leaves
 Col024.5 ± 3.510.2 ± 1.65.3 ± 1.04.6 ± 0.50.36 ± 0.070.45 ± 0.080.42 ± 0.060.23 ± 0.0428.3 ± 4.2
 sus1/sus421.2 ± 2.17.1 ± 1.810.2 ± 3.110.1 ± 2.52.84 ± 1.313.87 ± 1.391.37 ± 0.721.97 ± 0.9429.8 ± 3.1
 sus2/sus322.7 ± 4.710.4 ± 1.66.2 ± 1.76.1 ± 1.20.41 ± 0.110.34 ± 0.120.40 ± 0.160.35 ± 0.0630.8 ± 3.7
 sus5/sus623.1 ± 4.39.3 ± 2.25.5 ± 1.05.2 ± 1.20.33 ± 0.090.46 ± 0.130.30 ± 0.060.39 ± 0.1130.4 ± 5.3
(b) Roots
 Col-00.14 ± 0.100.20 ± 0.251.44 ± 0.202.41 ± 0.250.26 ± 0.100.12 ± 0.010.07 ± 0.080.09 ± 0.0114.5 ± 2.4
 sus1/sus40.16 ± 0.150.19 ± 0.121.67 ± 0.482.84 ± 0.620.28 ± 0.170.17 ± 0.040.04 ± 0.070.10 ± 0.0316.8 ± 1.0
 sus2/sus30.15 ± 0.090.13 ± 0.091.32 ± 0.292.25 ± 0.130.24 ± 0.080.17 ± 0.040.06 ± 0.070.09 ± 0.0212.8 ± 1.9
 sus5/sus60.14 ± 0.090.17 ± 0.061.26 ± 0.192.47 ± 0.480.18 ± 0.040.17 ± 0.050.04 ± 0.040.12 ± 0.0314.3 ± 1.9

In the light of the reduced growth rate and elevated sugar content of the sus1/sus4 mutant we assayed the activity of SUS and soluble invertase in roots and leaves of this line, and compared them with those of the other double mutants and Col-0 (Supplementary Table S3). The activity of SUS was reduced to 10% of wild-type levels in leaves of the sus1/sus4 mutant, and was undetectable (reduced to less than 10% of wild-type levels) in roots. The activity of invertase was unaffected in roots of this mutant, but in leaves neutral invertase activity was elevated by about 40%. Activity of SUS was reduced by about one-third in roots of the sus2/sus3 double mutant relative to Col-0, but otherwise the sus2/sus3 and sus5/sus6 mutants were indistinguishable from Col-0.

Effects of root hypoxia on sus1, sus4 and sus1/sus4 mutant plants

As a first step in the further characterization of the phenotype of the sus1/sus4 mutant, we examined the effects of root hypoxia on growth and sucrose synthase activity. Our rationale for this investigation was as follows. In wild-type plants, levels of transcripts for the SUS1 and SUS4 genes, but not the genes encoding the other isoforms, increase under hypoxic conditions (Baud et al., 2004; Klok et al., 2002). The amounts of both proteins also increase under these conditions (our results, Figure 4). The reduced growth rate of the sus1/sus4 mutant in hydroponic culture but not soil might thus be a consequence of a requirement for SUS1 and SUS4 for normal growth rates under the more hypoxic conditions likely to prevail in hydroponic media. The elevated sugar content of the leaves of sus1/sus4 mutant plants in hydroponic culture might be a consequence of reduced rates of root metabolism – although it might also be a direct consequence of the low sucrose synthase activity in the leaves.

Plants were grown in a free-draining grit-like rooting medium that allowed the harvest of complete root systems. We compared the fresh weights of the root systems of wild-type plants with those of sus2/sus3, sus1/sus4 and sus5/sus6 mutant lines before flooding, after flooding (immersion of the rooting medium in degassed water) for 4 days, and after a further ‘recovery’ period of 3 days without flooding. As controls, we examined plants from the same batch that had not been flooded. Root weights of wild-type, sus2/sus3 and sus5/sus6 mutant plants responded similarly to flooding (Table 8). After 4 days of flooding, root weight had increased by 72–86% (compare columns A and B in Table 8). In the 3 days following flooding, root weights increased by 63–71% (compare columns B and C in Table 8). At the end of the 7-day period, the overall gain in root weight of plants that had been flooded was 29–33% less than the gain in root weight of plants that had not been flooded (column E in Table 8). The sus1/sus4 mutant line was much more severely affected by flooding. After the 4 days of flooding, root weight had increased by only 44%, and there was no statistically significant gain in root weight in the 3 days following flooding. At the end of the 7-day period, the flooding treatment had reduced the gain in root weight by 71% relative to unflooded control plants. The aerial parts of the plant were also severely affected by the treatment (Figure 5).

Table 8.   Effects of flooding on root growth of double mutant and wild-type plants. Plants were grown for 4 weeks in a free-draining medium. Flooding consisted of immersion of the pots in degassed water to the top of the growth medium. After 4 days of flooding, the medium was allowed to drain completely, then watered when necessary as for control plants. After harvest, root systems were rapidly blotted and weighed immediately. Values are means ± SD of measurements on five plants. The last column is derived from the previous columns: (C – A)/(D – A) × 100. Root weight of sus1/sus4 is statistically significantly different from that of Col-0 in column C (P = 8.4 × 10−6, Student's t-test)
LineWeight at start of experiment (A)Weight at 4-day flooded (B)Weight at 4-day flooded, 3-day recovery (C)Weight at 7-day unflooded (D)% reduction in weight gain due to the flooding treatment (E)
Col-00.078 ± 0.0060.134 ± 0.0090.228 ± 0.0130.303 ± 0.01833
sus2/sus30.084 ± 0.0030.153 ± 0.0080.256 ± 0.0220.302 ± 0.06721
sus5/sus60.075 ± 0.0060.139 ± 0.0180.227 ± 0.0230.291 ± 0.03229
sus1/sus40.086 ± 0.0080.124 ± 0.0080.140 ± 0.0150.274 ± 0.00871
image

Figure 5.  Effects of flooding of the root system of sus1/sus4 mutants on rosette growth and membrane-associated SUS proteins. (a) Rosettes of sus1/sus4 mutant plants (top) and plants of a wild-type line selected from the segregating F2 population from which the double mutant was selected, after 4 days’ flooding and 3 days’ recovery. (b) Presence of SUS proteins in soluble and membrane fractions of roots. Panels show immunoblots of SDS-polyacrylamide gels of soluble and membrane proteins from the lines and conditions indicated. For each panel, all lanes are from the same gel and blot. Fractions were prepared from a homogenate initially subjected to centrifugation at 10 000 g. The supernatant was further centrifuged at 100 000 g to give the soluble fraction and a pellet containing membranes. Membrane and soluble fractions for a given line/condition are from a single homogenate. Equal proportions of the soluble and membrane fractions were loaded onto the gel.

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Activity of SUS was not statistically significantly different in roots of sus4 and control plants in unflooded conditions, but it was 73% lower in roots of sus1 than control plants under these conditions (Table 9). In agreement with the results from hydroponic culture (Supplementary Table S3), activity in the double mutant sus1/sus4 was at the limit of detection. It was reduced by at least 97% relative to control plants. Plants of sus1, sus4 and wild-type lines all had elevated SUS activities after flooding for 4 days, and these activities declined over the 3 days following flooding. Activity in both sus1 and sus4 roots was statistically significantly lower than that in roots of control plants after these treatments. Activity in the sus1/sus4 mutant remained close to the level of detection after these treatments.

Table 9.   Changes in activity of SUS in roots of plants subjected to flooding. Plants were subjected to the regime described in Table 8. Unflooded plants were harvested at the same time as plants subjected to 4 days of flooding and 3 days of recovery. SUS activity was measured in extracts of whole root systems. The control was a wild-type line selected from the segregating F2 population from which the double mutant was selected. Values are means ± SD of measurements on three plants. Values are expressed on a protein (rather than a fresh weight) basis because dead roots on sus1/sus4 mutant plants after 4 days of flooding and 3 days of recovery contributed to the fresh weight. Except for sus4 under unflooded conditions, all mutant values are significantly different from control values under the same conditions: P values (Student's t-test) are given in parentheses
LineEnzyme activity (μmol min−1 mg−1 protein)
UnfloodedFour-day floodedFour-day flooded then 3-day recovery
Control for sus1/sus40.0307 ± 0.01150.0804 ± 0.00790.0584 ± 0.0049
sus10.0083 ± 0.0035 (P = 0.03)0.0353 ± 0.0017 (P = 0.0006)0.0218 ± 0.0007 (P = 0.0002)
sus40.0279 ± 0.00290.0505 ± 0.0010 (P = 0.003)0.0467 ± 0.0029 (P = 0.024)
sus1/sus40.0001 ± 0.0001 (P = 0.01)0.0003 ± 0.0001 (P = 6.3 × 10−5)0.0008 ± 0.0003 (P = 3.6 × 10−5)

Levels of SUS1 and SUS4 proteins in roots were consistent with the picture derived from measurements of activity (Figure 5). Levels of both proteins increased significantly in the soluble fraction in response to flooding (see also the independent experiment in Figure 4). Anoxia has been reported to increase the fraction of total SUS protein associated with membranes in roots (Subbaiah and Sachs, 2001). We found that flooding increased the amounts of both SUS1 and SUS4 associated with the membrane fraction (Figure 5), but because membrane-associated SUS1 and SUS4 were not readily detectable in unflooded roots, our data do not reveal whether flooding caused a change in ratio of soluble to membrane-associated proteins. Neither protein was detectable in the soluble or the membrane fraction of roots of the sus1/sus4 double mutant.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The isoforms of sucrose synthase in Arabidopsis are closely related to each other in both sequence and general kinetic properties, but have radically different spatial and temporal patterns of expression in the plant. This is consistent with the idea that the isoforms are not highly specialized for distinct functions within the cell, but may fulfil similar functions in different cell types or at different developmental stages. Our analysis of single and double knockout mutants also supports this view. The lack of any obvious phenotype in the single knockout mutants, and in the double knockout mutants in well-aerated, complete growth media, indicates that no single isoform, and none of the three pairs of similar isoforms, has a specific and indispensable function at either the cellular or the organ level in Arabidopsis. Thus more than one isoform or pair of isoforms must be able to provide UDPglucose to cellulose synthase, and it is likely that more than one isoform can provide substrates for energy production for phloem loading and carbon for lipid synthesis in developing embryos.

The apparent redundancy displayed by the isoforms of SUS under normal growth conditions might occur because a significant proportion of the sucrose in Arabidopsis is metabolized via invertase rather than SUS. Invertase hydrolyses sucrose to hexoses, which can then provide hexose phosphate and UDPglucose via the actions of hexokinase, phosphoglucomutase and UDPglucose pyrophosphorylase. The SUS activity in Arabidopsis roots in free-draining media is three to four times greater than the likely rate of utilization of sucrose in respiration. Zeeman and ap Rees (1999) reported a rate of carbon dioxide release of 0.028 μmol sucrose equivalents min−1 g−1 fresh weight from Arabidopsis roots: in Table 9 we report a SUS activity of 0.031 μmol min−1 (mg protein)−1, which equates to 0.115 μmol min−1 g−1 fresh weight. However, activity of neutral invertase – assumed to be in the cytosol – is several times higher than that of SUS (Supplementary Table S3); thus both SUS and invertase, individually, have the capacity to account for sucrose mobilization in normal conditions. In roots of the sus1/sus4 double mutant we assume that essentially all of the sucrose mobilization is via invertase since sucrose synthase activity is at the limits of detection and many-fold lower than is required to account for normal rates of respiration. Nevertheless, the pronounced growth phenotype of the sus1/sus4 double mutant under hypoxic/anoxic conditions clearly establishes that SUS makes a significant metabolic contribution under some conditions.

The apparent lack of specialization among the isoforms does not preclude the possibility that sucrose synthase has multiple, distinct functions within a single cell, for example in supplying substrates for structural polymer synthesis at membrane surfaces and for glycolysis and starch synthesis in the soluble fraction, and in metabolizing sucrose passing across the tonoplast. The partitioning of sucrose synthase protein between these functions may be achieved by post-translational modification of the protein. As discussed in the Introduction, there is good evidence that phosphorylation of sucrose synthase affects its association with membranes in vitro and in vivo.

The lack of any obvious phenotypes among the single and double knockout mutants when grown in controlled conditions in well-aerated, complete media is surprising in the light of previous work on sucrose synthase in maize, pea and potato. In these species, mutant and antisense plants deficient in specific isoforms of SUS have strong phenotypes. Seeds of maize mutants lacking either of isoforms SS1 or SS2 have reduced starch content (Chourey et al., 1998). Pea mutants (rug4) lacking an isoform very similar to SUS1 and SUS4 of Arabidopsis have reduced seed mass and starch content, and fail to assimilate N2 in their Rhizobium root nodules (Craig et al., 1999; Gordon et al., 1999). Antisense inhibition of the main isoform in potato tubers drastically reduces starch accumulation (Zrenner et al., 1995). We suggest two possible reasons for the apparent difference between these species and Arabidopsis in the importance of SUS. First, the high activity and importance of sucrose synthase in seeds and tubers may reflect a requirement for sucrose synthase rather than invertase in organs where oxygen tensions are naturally low. Conversion of sucrose to hexose phosphate via sucrose synthase requires less ATP than conversion via invertase, and it has frequently been proposed that sucrose synthase will be the favoured route in metabolically highly active or bulky organs where ATP synthesis may be limited by low oxygen tension (Guglielminetti et al., 1995; Rolletschek et al., 2002). Consistent with these ideas, transgenic potato tubers expressing elevated levels of invertase have reduced internal oxygen tension, and elevated levels of proteins characteristic of anaerobiosis (Bologa et al., 2003). The SUS activity in pea seeds is ten times higher than neutral invertase activity (Edwards and ap Rees, 1986), and in potato tubers it is about 20 times higher than neutral invertase activity (Ross et al., 1994). The ratios of sucrose synthase to invertase activity reported for non-bulky organs such as roots, in aerated conditions, are generally much lower than those reported for seeds and tubers (e.g. potato, Biemelt et al., 1999; wheat, Albrecht and Mustroph, 2003; soybean, Schubert et al., 2003). Large increases in this ratio occur early in the development of potato tubers (Appeldoorn et al., 1997; Ross et al., 1994) and maize and bean seeds (Tsai et al., 1970; Weber et al., 1997), as synthesis of storage product commences and organ bulk increases. Thus the lack of any pronounced phenotypes in Arabidopsis mutants lacking SUS isoforms may reflect the absence of bulky storage organs. Perhaps the most likely organ to be affected is the developing embryo, a dense, highly metabolically active organ with no direct contact with a gas phase. Development of seeds of the closely related species oilseed rape has been shown to be limited by low internal oxygen concentrations (Vigeolas et al., 2003), and SUS activity far exceeds that of invertase through much of seed development (Hill et al., 2003). However, surprisingly, loss of both of the isoforms of SUS most highly expressed in developing Arabidopsis seeds (SUS2 and SUS3) has no effect on final seed weight and lipid content.

A second possible explanation for the difference between Arabidopsis and other well-studied species in the importance of individual isoforms of SUS lies in the fact that these other species are crop plants. It is conceivable that selection during domestication for high-yielding sink organs in crop species has led to a situation in which one particular isoform of SUS accounts for most of the sucrose synthase activity, and most of the total sucrose metabolizing capacity, in particular cell types at critical developmental stages, such that loss of this isoform results in a significant reduction in flux from sucrose. In an undomesticated species such as Arabidopsis, there may be sufficient activity of more than one isoform throughout sink organ development to ensure that loss of a single isoform does not affect flux, or compensatory increases in other isoforms in the sink organs when one is lost, or a flexible partitioning of flux between sucrose synthase and invertase. If correct, this difference between crop and undomesticated species has potentially important implications for our understanding of carbon partitioning and yield in crop plants.

Although none of the sucrose synthases individually appear to be essential for normal growth under our standard, well-aerated conditions, our data show that SUS1 and SUS4 are together necessary for tolerance of hypoxic conditions. Measurements on single and double mutants indicate that almost all of the SUS activity in both unstressed and flooded roots is contributed by SUS1 and SUS4, the former making the larger contribution. Activities of both isoforms increase under hypoxia: after 4 days of flooding, roots of both sus1 and sus4 mutants have more activity than wild-type unstressed roots and at least 40% of the activity of wild-type roots after the same period of flooding. These data are consistent with previous reports of increases in transcript levels of SUS1 and SUS4– but not other isoforms – under hypoxia (Baud et al., 2004; Klok et al., 2002). Consistent with the relatively small reductions in activity, neither the sus1 nor the sus4 mutant shows impaired growth relative to wild-type plants during flooding or a post-flooding recovery period. In contrast, growth of the sus1/sus4 double mutant, which has almost no SUS activity during either unstressed or flooded conditions, is severely impaired.

As in Arabidopsis, SUS activity in maize roots seems to be important during and following exposure to hypoxia/anoxia, but not in normal conditions. Activity increases in response to hypoxia/anoxia, due to increased expression of SUS1 and SH1. SUS1 responds to hypoxia rather than anoxia, and SH1 to anoxia rather than hypoxia (Zeng et al., 1998). Loss of both isoforms has little effect on growth in normal conditions (Chourey, 1988), but impairs the ability of roots to survive periods of hypoxia/anoxia (Ricard et al., 1998). Increases in SUS activity in response to hypoxia/anoxia have been reported for roots of several other species (e.g. potato, Biemelt et al., 1999; wheat, Albrecht and Mustroph, 2003), and it seems likely that a requirement for SUS for tolerance of such conditions may be a widespread phenomenon. The simplest explanation for this requirement is that the higher yield of ATP per sucrose obtained via SUS as opposed to invertase becomes critical in conditions where oxygen is limiting. However, there is some evidence that the requirement for SUS reflects a specific role in structural polysaccharide synthesis rather than in the provision of carbon for glycolysis. Roots of transgenic potatoes with reduced SUS activity are more rapidly damaged by hypoxia, and recover more slowly on return to air than normal roots (Biemelt et al., 1999). Measurements of sugars and glycolytic enzymes and intermediates indicated that the rate of glycolysis in hypoxia and during recovery was not limited by substrate availability in either wild-type or transgenic plants. The authors suggested that the role of SUS in hypoxia might be to channel carbon to cell wall polyglucans. This would both sustain a sink capacity in the roots during hypoxia, and provide a source of carbon for utilization during a recovery period. Consistent with this suggestion, hypoxia/anoxia induces the deposition of large amounts of callose in maize roots (Subbaiah and Sachs, 2001) and cellulose in wheat roots (Albrecht and Mustroph, 2003). Callose deposition in maize is dependent on the presence of the SH1 isoform, which becomes dephosphorylated and increasingly membrane associated during anoxia. However, the functional significance of this callose deposition is unclear: sh1 mutant plants, with reduced callose deposition, actually survived anoxia better than wild-type plants under the conditions of the experiment. In general terms, although SUS activity is clearly important in the tolerance of hypoxia/anoxia, there is little understanding of its precise role. Our Arabidopsis mutants provide valuable tools for further research.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Plant material and growth conditions

For isolation of RNA and protein for semi-quantitative RT-PCR and immunoblot experiments, wild type Arabidopsis thaliana of ecotypes Wassilewskija (Ws) and Columbia (Col) and T-DNA insertion lines in these backgrounds were sown in a peat-based compost and held at 4°C in the dark for 3–5 d, then grown at 20°C, 75% relative humidity in a 12 h-light/12 h-dark photoperiod with a irradiance of 180–200 μmol m−2 s−1. For anoxic/hypoxic (flooding) treatment, plants were grown as above but in a 1:1 mixture of sand and Terragreen medium (Oil-Dri, http://www.oildri.com) supplemented with a slow-release fertilizer. All plant material was immediately frozen in liquid nitrogen after harvesting and stored at −80°C prior to analysis.

For studies of mature seed weight and composition plants were grown in a peat-based compost with five plants per 9-cm pot, in a randomized block design in a glasshouse. Conditions were 18°C during the day, 15°C at night, and supplementary lighting to give a 16-h day. Mature seeds were placed in a desiccator for 2 weeks at room temperature then stored at 4°C with desiccant prior to analysis.

For isolation of RNA for real-time quantitative RT-PCR and for measurement of cellulose, starch and sugars, seeds were surface sterilized, put on half-strength MS medium containing 0.7% agar and 1% sucrose (including 50 μg ml−1 kanamycin in the case of plants for promoter analysis), imbibed in the dark for 48 h at 4°C then grown at 20°C/18°C in a 16-h light/8-h dark photoperiod with a irradiance of 120 μmol m−2 sec−1. Plants for RNA isolation were transferred after 2 weeks to a soil–vermiculite mixture and grown for at least 4 weeks in growth chambers at 20°C/16°C at 60%/75% relative humidity in a 16-h light/8-h dark photoperiod with an irradiance of 120 μmol photons m−2 sec−1. Plants for cellulose, starch and soluble sugar analysis were transferred after 2 weeks into hydroponic culture for 3 weeks, in growth chambers as described above. The hydroponic solution contained: 1 mm KNO3, 2.5 mm Ca(NO3)2, 0.5 mm MgSO4, 0.5 mm KH2PO4/K2HPO4 (pH 5.8), 20 μm Fe-EDTA, 150 μm H3BO3, 35 μm MnSO4, 2.5 μm ZnSO4, 1.5 μm CuSO4, 1 μm NiCl2, 0.75 μm HMoO4 and 50 nm CoCl2. The medium was continuously aerated, and was changed every 4 days. Whole rosettes and whole root systems were harvested, and all plant material was immediately frozen in liquid nitrogen and stored at −80°C prior to analysis.

Isolation of mutant lines

Homozygous mutants were identified by screening T-DNA insertion lines from the Salk collection, the Arabidopsis Knockout Facility at the University of Wisconsin, the Syngenta Arabidopsis Insertion Library (SAIL) lines, the GABI-Kat collection (http://www.gabi-kat.de/) and an insertion library described by Babiychuk et al. (1997), with gene- and T-DNA-specific primers. Genomic DNA was extracted from leaves according to the protocol provided by the Arabidopsis Knockout Facility. The PCR products obtained using T-DNA left border primers and gene-specific primers were sequenced to confirm the location of the inserts. The sus4 knockout, which was in the ecotype C24, was backcrossed six times into the ecotype Col selecting each time for T-DNA insertion in the SUS4 gene.

Semi-quantitative RT-PCR analysis

Total RNA was isolated using Concert Plant RNA Reagent (Invitrogen, http://www.invitrogen.com/) then purified and treated with DNaseI using the RNeasy RNA Cleanup Kit (Qiagen, http://www.qiagen.com/). Gels were stained with ethidium bromide to confirm RNA integrity. First-strand cDNA was synthesized from 5 μg of total RNA and amplified according to Frohman et al. (1988) using the dT(17) adapter as a primer and Superscript II RNase H Reverse Transcriptase (Invitrogen). Primers for the gene encoding ubiquitin 10 were used to verify the quality of the cDNA and to ensure equal loadings of the various cDNA templates. Oligonucleotides used as primers for the six SUS genes are specified in Supplementary Material, Appendix S2.

To confirm that amplification was linear and to establish the number of amplification cycles for detection of cDNA within the linear range, PCRs were carried out for the six SUS genes for 15, 20, 25 and 30 cycles and the products separated by gel electrophoresis then blotted onto Duralon-UV membranes (Stratagene, http://www.stratagene.com/). The filters were probed with the appropriate cDNA, purified from PCR products using the gene-specific primers, then labelled using Rediprime II Random Prime Labelling System (Amersham Biosciences, http://www5.amershambiosciences.com/).This enabled the following conditions to be established: 15 cycles for SUS1 and SUS2 amplification, and 20 cycles for SUS3, SUS4, SUS5 and SUS6.

Real-time quantitative PCR analysis

Total RNA was isolated from frozen plant material using NucleoSpin RNA Plant kit (Macherey-Nagel, http://www.macherey-nagel.com/), then further purified and precipitated (RNA:5 m LiCl:ethanol at 1:1:2 (v:v:v) at 4°C for 30 min, centrifuged for 10 min at 20 000 g and 4°C and then washed with 70% aq. (v.v) ethanol. Two micrograms of total RNA were digested with RNase-free DNaseI (D5307, Sigma-Aldrich, http://www.sigmaaldrich.com/). Absence of genomic DNA contamination was confirmed by PCR, using primers designed on intron sequence of a control gene (At5g65080). Gels were stained with ethidium bromide to confirm RNA integrity.

Reverse transcriptase reactions were performed with SuperScript III Reverse Transcriptase (Invitrogen), according to the manufacturer's instructions. The efficiency of cDNA synthesis was assessed by real-time PCR amplification of control genes encoding ubiquitin 10 and elongation factor 1α. Oligonucleotides used as primers are specified in Supplementary Material, Appendix S2.

Polymerase chain reactions were performed in an optical 96-well plate with an ABI PRISM® 5700 HT Sequence Detection System (Applied Biosystems, http://www.appliedbiosystems.com/), using SYBR® Green to monitor synthesis of double-stranded DNA (dsDNA). Reactions contained 10 μl 2× SYBR® Green Master Mix reagent (Applied Biosystems), 1 μl cDNA and 125 nm of each gene-specific primer in a final volume of 20 μl. The following standard thermal profile was used for all PCRs: 50°C for 2 min, 95°C for 10 min, 40 cycles of 95°C for 15 sec, 60°C for 1 min.

Preparation of full-length cDNAs

Total RNA was isolated from frozen plant material according to Edwards et al. (1995). DNaseI-treated poly (A+) RNA (1 μg), purified on oligo (dT)-cellulose columns (Amersham Biosciences), was converted into first-strand cDNA using SuperScript II RNase H Reverse Transcriptase with a dT(17) adaptor (Frohman et al., 1988). Specific primers sets were designed for each gene from sequences present in the Arabidopsis thaliana database. Full-length cDNAs for each gene were isolated by RT-PCR using PfuTurbo DNA Polymerase (Stratagene). The specificity of each primer set was checked by sequencing the PCR product following cloning into pGEM-T Easy (Promega, http://www.promega.com/).

Production and expression of promoter–GUS constructs

Promoter fragment amplification employed standard protocols and either PfuTurbo DNA Polymerase (Stratagene), Plantinum Pfx (Invitrogen) or Expand (Roche, http://www.roche.com/). Fragments were amplified using A. thaliana Col-0 genomic DNA purified either with the DNeasy Plant Mini kit (Qiagen) or with the CTAB method (Sambrook and Russell, 2001) and subcloned using the pCR-Script® Amp Cloning Kit (Stratagene). All promoter fragments were completely sequenced and no differences from the annotated genes were found.

The promoter fragments consisted of either approximately the first 2000 base pairs of the intergenic region from the start codon of the respective sucrose synthase isoform, or the region from the start codon to the adjacent gene in the 5′-direction if this was fewer than 2000 base pairs. Two different sets of promoter fragments were used: full details are given in Supplementary Material, Appendix S2.

For promoter analysis each fragment was cloned either into the binary vector pGPTV (Becker et al., 1992) or into a derivative of the binary vector pGreen0029 (Hellens et al., 2000), pGVT5 (Thole and Rawsthorne, 2003). Constructs contained a selectable marker gene for kanamycin resistance and included a β-glucuronidase gene cassette (pGPTV-based constructs) or a β-glucuronidase-intron gene cassette (pGVT5-based constructs).

Arabidopsis thaliana Col-0 plants were transformed using standard procedures (Bechtold and Pelletier, 1998; Clough and Bent, 1998). After selection of transformed plants on medium containing kanamycin, the presence of the transgene was verified by PCR using a promoter-specific forward primer and a gusA-specific reverse primer. Expression of the reporter gene was monitored in T0, T1 and T2 and/or T3 transgenic plants, using histochemical staining according to Jefferson (1987) and Jefferson et al. (1987).

Plasmid construction and expression in Escherichia coli

Full-length cDNAs cloned in pGEM-T Easy had an NcoI restriction site and an in-frame ATG initiation codon introduced at the start of each clone and a BamHI site introduced downstream from the stop codon. This was achieved by modification of the primers for the initial RT-PCR or by using the Quikchange Site-Directed Mutagenesis Kit (Stratagene). Polymerase chain reaction mutagenesis was also used to remove an NcoI restriction site and a BamHI restriction site 250 and 2200 bp, respectively, downstream from the start codon in SUS1; a BamHI restriction site 2243 bp downstream from the start codon in SUS4; and a BamHI restriction site 2310 downstream from the start codon in SUS5.

Plasmids for the expression of mature, full-length SUS proteins (named pETArabSUS1 to pETArabSUS6) were constructed by insertion of a NcoI/BamHI fragment between the NcoI and the BamHI sites of the expression vector pET3d (Studier et al., 1990). pETArabSUS1 to pETArabSUS6 were each transformed into E. coli and expressed as described by Barratt et al. (2001). The sequence of each construct was checked following each of the above manipulations. Expression of the recombinant protein was induced by adding IPTG at a final concentration of 1 mm, then shaking at 350 r.p.m. at 20°C overnight prior to harvest.

Purification of recombinant sucrose synthase protein for antiserum testing

Escherichia coli cells were harvested by centrifugation. Cells from three 50-ml cultures were resuspended in 4 ml of 50 mm K-HEPES [4-(2-hydroxyethyl)piperazine-1-ethanesulphonic acid] (pH 7.5), 1 mm EDTA, 50 ml l−1 glycerol and 0.1 mm DTT, then disrupted by two passages through a French pressure cell and clarified by centrifugation for 20 min at 30 000 g. The supernatant was further clarified by centrifugation for 1 h at 40 000 g at 4°C, then subjected to ammonium sulphate precipitation. Proteins that precipitated between 20% and 65% saturation with ammonium sulphate were collected by centrifugation at 30 000 gsuspended in 50 mm K-HEPES (pH 7.5), 10 mm MgCl2, 1 mm EDTA, 50 ml l−1 glycerol and 2 mm DTT (solution A) and desalted on a NAP25 (Amersham Biosciences) column equilibrated with solution A. The eluate was applied to a MonoQ HR5/5 column (Amersham Biosciences) equilibrated with solution A. Proteins were eluted with a 50-ml linear gradient from 0 to 300 mm KCl in solution A and 1-ml fractions were collected. Fractions with the highest SUS activity (assayed in the cleavage direction by an indirect method; Ross and Davies, 1992) were combined and concentrated to 200 μl using a Centricon YM-100 filter (Amicon Bioseparations, Millipore, http://www.millipore.com) then applied to a Superdex 200 HR 10/30 column equilibrated with solution A. Fractions of the highest activity were retained and their purity assessed with SDS-PAGE. Protein concentrations were measured using the Bio-Rad (http://bio-rad.com/) Protein Assay, with bovine serum albumin as the standard.

SUS2, -3 and -5 were of insufficient purity, following the above purification, to use for antiserum testing, therefore the recombinant proteins were purified from inclusion bodies. pETArabSUS2, -3 and -5 were over-expressed by induction with 1 mm IPTG for 2 h at 37°C with shaking at 350 r.p.m. This resulted in aggregation of the recombinant protein in inclusion bodies from which it could be isolated to a high degree of purity (Barratt et al., 2001).

Production of peptide-specific antisera

Synthetic peptides for each isoform were designed on the basis of their specificity, ease of synthesis, hydrophilic and antigenic properties. The amino acid sequences of the peptides are given in Supplementary Material, Appendix S2. The peptides were synthesized, conjugated via cysteine to keyhole limpet haemocyanin and used for the production of antisera by Sigma-Genosys (http://www.sigmaaldrich.com). Unconjugated peptides were linked to AminoLink Plus Coupling Gel (Pierce Biotechnology, http://www.piercenet.com/) according to the manufacturer's instructions, and each peptide antiserum was affinity purified.

Purification of recombinant SUS protein for kinetic analyses

To facilitate kinetic analysis of functionally active untagged recombinant SUS proteins, we developed a rapid and efficient procedure to purify each of the isoforms to near-homogeneity from 1.6 l of bacterial culture by exploiting the ability of the native enzyme to bind to immobilized Ni-affinity resins (Römer et al., 2004). Growth of E. coli was as in Barratt et al. (2001) except that cultures were at 28°C rather than 30°C and contained 50 μg ml−1 ampicillin. Expression of the recombinant protein was induced by adding IPTG at a final concentration of 1 mm, then shaking at 250 r.p.m. at 18°C overnight prior to harvest.

Escherichia coli cells were harvested by centrifugation. Cells from 32 50-ml cultures were each resuspended in 1.5 ml of BugBuster with 1.5 μl Benzonase nuclease (Merck-Novagen, http://splash.emdbiosciences.com/) and incubated at room temperature for 20 min with shaking at 250 r.p.m. The cell extracts were harvested by centrifugation for 20 min at 16 000 g at 4°C. The resulting supernatants were combined, adjusted to 30% saturation with polyethylene glycol at 4°C and incubated for 1 h. Precipitated proteins were collected by centrifugation for 15 min at 30 000 g at 4°C. The pellet was resuspended in 20 ml of cold 50 mm K-HEPES (pH 8.0). The resuspended pellet was clarified by centrifugation for 15 min at 30 000 g at 4°C and the supernatant was applied to a Q-Sepharose Fast Flow column (Amersham Biosciences; 15 × 2.6 cm) equilibrated with 50 mm K-HEPES (pH 8.0). The column was washed with 50 ml equilibration buffer then proteins were eluted with a 450-ml linear gradient from 0 to 300 mm KCl in equilibration buffer and 10-ml fractions were collected. Fractions with the highest sucrose synthase activity (assayed in the synthetic direction) were combined and applied to a Ni-NTA column (Qiagen; 10 × 0.9 cm) equilibrated with 50 mm K-HEPES (pH 8.0) and 100 mm NaCl. The column was washed with 50 ml of the equilibrating buffer containing 1.25 mm imidazole. Proteins were eluted with a 100-ml linear gradient from 1.25 to 25 mm imidazole in 50 mm K-HEPES (pH 8.0) and 100 mm NaCl and 5-ml fractions were collected. Fractions with the highest SUS activity were combined and used for kinetic analysis. Protein concentrations were measured using the Bio-Rad Protein Assay, with bovine serum albumin as the standard.

Kinetic analyses

Continuous, coupled assays were performed at 25°C, monitored at 340 nm in a microtitre plate reader and were initiated by the addition of purified enzyme unless otherwise specified. For both the cleavage and synthetic reactions, the components and the pH of the assay were optimized to give the maximum activity.

Assay of sucrose synthase activity in the cleavage direction

The optimized assay contained, in 250 μl, 50 mm K-HEPES (pH 6.0), 1 mm MgCl2, 0.5 mm ATP, 0.5 mm NAD+, 1 mm UDP, 200 mm sucrose, 1.9 U hexokinase, 1.25 U glucose 6-phosphate dehydrogenase (from Leuconostoc mesenteroides) and 0.9 U phosphoglucose isomerase.

Assay of sucrose synthase activity in the synthetic direction

The optimized assay contained, in 250 μl, 50 mm K-N-(1,1-dimethyl-2-hydroxyethyl)-3-amino-2-hydroxypropanesulphonic acid (K-AMPSO, pH 9.5), 20 mm KCl, 4 mm MgCl2, 2 mm K2HPO4, 1 mm phosphoenolpyruvate, 0.25 mm NADH, 1 mm UDPglucose, 25 mm fructose, 5 U pyruvate kinase and 5 U lactate dehydrogenase.

Further details of the kinetic analyses and derivation of kinetic constants are given in Supplementary Material, Appendix S1.

Assay of sucrose synthase and invertase in root extracts

Enzymes were extracted from 100 mg roots or leaves at 4°C into 600 μl 50 mm Na-HEPES (pH 7.5), 5 mm MgCl2, 1 mm EDTA, 1 mm EGTA, 10% (v.v) glycerol, 0.1% (v.v) TritonX-100, 2 mm DTT, 20 mg polyvinylpolypyrrolidone and 1 mm benzamidine, 5 mmɛ-aminocaproic acid, 10 μm leupeptin, 10 μm antipain, 1 mm phenylmethylsulphonylfluoride. After centrifugation (20 000 g, 10 min, 4°C) the supernatant was desalted by size exclusion chromatography using Sephadex G-25 equilibrated with extraction buffer. In some cases desalted samples were stored at −80°C before assay.

The sucrose synthase assay contained 30 μl of 100 mm K-AMPSO, pH 9.4, 10 mm fructose (or water in control assays), 10 mm UDP[U14C]glucose (Amersham) at 11.2 GBq mol−1, and 25 μl extract. Incubations were for 20 min at 20°C, stopped by boiling for 2 min, and diluted 1:1 with water. A sample of 100 μl was applied to a column of Dowex resin (1 × 8 200–400 mesh), which was centrifuged for 1 min at 300 g then washed twice by centrifugation at 300 g with 100 μl of water. The combined eluates were subjected to liquid scintillation counting.

The invertase assay contained in 100 μl either 25 mm Na acetate (pH 4.9) (acid invertase assay), or 20 mm K-HEPES (pH 7.5) (neutral invertase assay) and 100 mm sucrose. Incubations were for 60 min at 30°C. After neutralization with 1 m 2-amino-2-(hydroxymethyl)-1,3-propanediol (TRIS) (pH 8.0), the reaction was stopped by heating at 95°C for 3 min. Control assays were heated before incubation. Glucose and fructose were measured enzymatically (Jellito et al., 1992).

Immunoblotting

Plant tissues were extracted at 4°C in 50 mm Na-HEPES (pH 7.5), 100 mm NaCl, 2 mm EDTA, 1 mm EGTA, 5 mm ascorbic acid, 2 mm DTT, 10 g l−1 polyvinylpolypyrrolidone and 10 ml l−1 protease inhibitor cocktail (Sigma, http://www.sigmaaldrich.com/). The homogenate was subjected to centrifugation for 10 min at 10 000 g at 4°C and the supernatant precipitated with 10% w/v trichloroacetic acid (TCA), washed repeatedly with 80% v.v acetone at −20°C then treated with SDS sample buffer (Barratt and Clark, 1991) before analysis by SDS-PAGE. Where soluble and membrane fractions were studied separately, the 10 000 g supernatant was further centrifuged for 60 min at 100 000 g and 4°C. The resultant supernatant (constituting the soluble fraction) was precipitated with TCA and treated as above. The 100 000 g pellet (constituting the membrane fraction) was dispersed in SDS sample buffer. For each extract, the soluble and membrane fractions were dispersed in the same volume of SDS sample buffer. Sodium dodecyl sulphate-PAGE and immunoblotting were performed as described by Barratt et al. (2001).

Seed weight and oil content

Approximately 100 seeds were accurately counted then weighed on a Cahn C-30 Microbalance (Scientific and Medical Products, http://www.scimed.co.uk/). Approximately 200 mg of seed was weighed accurately into a 10-mm diameter NMR tube and the oil content was determined using a benchtop QP20+ NMR spectrometer (Oxford Instruments, http://www.oxford-instruments.com/) as described by Hobbs et al. (2004). The standard curve was produced using canola standards supplied by the Canadian Grain Council.

Measurement of cellulose, starch and sucrose

A protocol for cell wall analysis published by Peng et al. (2000) was modified as follows.

For measurement of sucrose, frozen, powdered plant material (200 mg) was extracted twice for 20 min at 80°C successively in 80% (v.v) ethanol in 3 mm K-HEPES (pH 7), 50% (v.v) ethanol in 3 mm K-HEPES (pH 7) and 3 mm K-HEPES (pH 7). Sucrose was assayed on the combined soluble extracts according to Jellito et al. (1992).

For measurement of starch, lipid was extracted from the residue [twice 60 min in chloroform:methanol (1:1; v.v) at 40°C, 30 min in methanol at 40°C followed by a wash with water] then starch was solubilized by treatment for 60 min in 0.2 m KOH at 95°C. After acidification, starch was hydrolysed by overnight incubation in 50 mm Na-acetate (pH 5.5) containing amyloglucosidase and α-amylase at 37°C, and glucose was assayed according to Jellito et al. (1992).

For measurement of cellulose, the residue was further extracted successively by: (i) incubation for 24 h in 50 mm Na2CO3 containing 10 mm NaBH4 with shaking, followed by two washes with water, (ii) incubation for 16 h and 8 h in 1 m KOH containing 10 mm NaBH4 with shaking, followed by a wash with water, (iii) incubation for 16 h and 8 h in 4 m KOH containing 10 mm NaBH4 with shaking, followed by a wash with water, (iv) treatment with Updegraff reagent (30 min in concentrated HNO3:80% acetic acid (1:10; v.v) at 95°C, followed by three washes with water). The resulting insoluble cellulose preparation was subjected to Seaman hydrolysis (60 min in 72% aq. (v.v) sulphuric acid in a sonication bath). Glucose was then determined using the anthrone method (Laurentin and Edwards, 2003).

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Work at the John Innes Centre and in Oxford was supported by a grant from the Biotechnology and Biological Sciences Research Council (BBSRC, UK) to NJK, AMS and CM, and by a Core Strategic Grant from the BBSRC to the John Innes Centre. We thank Doug Hobbs (John Innes Centre) for advice on seed analyses, and Markus Pauly and colleagues (Max Planck Institute) for advice on cellulose measurement.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Figure S1. Inhibition of SUS by glucose. Figure S2. Effect of substrate concentrations on SUS activity. Table S1. Summary of representative purification of sucrose synthase encoded by SUS1. Table S2. Patterns of product inhibition displayed by SUS1. Table S3. Enzyme activities in roots of hydroponically-grown plants. Appendix S1. Kinetic properties: derivation of kinetic constants; product inhibition assays; inhibitor studies; references Appendix S2. Primers and peptides: PCR analyses; Promoter fragments; synthetic peptides for antiserum production

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