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Sucrose synthase is a key enzyme in sucrose metabolism. Sucrose metabolism is required by the plant to form carbon required for various processes in the plant such as respiration, starch and cell wall formation. The enzyme is encoded by a small multigene family where most plants have at least two isoforms of the enzyme. The kinetics of sucrose synthase show that different Km values and ratios of sucrose breakdown exist for the enzyme. The methods of extracting, assaying and purifying the enzyme are shown in the enzyme characteristics. Factors such as pH, addition of different buffers, metal ions, fungal volatiles as well as environmental factors such as anoxia have all been shown to affect sucrose synthase activity. The enzymes protein sequences have been phylogenetically divided up into six main groups using clustalw. Sucrose synthase is normally present in the cytoplasm but the availability of sucrose in the chloroplast and its ability to use ADP as a substrate would indicate that the enzyme may be able to act in the chloroplast as well as the cytoplasm.
Sucrose synthase is an important enzyme in sucrose metabolism in plants cells. (Persia et al., 2008) The main route of entry of carbon from sucrose is commonly known to be sucrose synthase. (Bieniawska et al., 2007) This carbon is used for respiration and in the synthesis of cell wall polymers and starch. (Persia et al., 2008) The main form of reduced carbon in plants is sucrose. It is used to support growth and synthesis of reserve materials e.g. starch in heterotrophic sink tissues. (Matic et al., 2004) The UDP-glucose supplied by sucrose synthase is used for cell wall biosynthesis while working with the cellulose synthase complex. (Baud, Vaultier and Rochat, 2004) In most fruit tissues, an increase in sucrose synthase activity is alongside with sucrose accumulation. This would suggest that sucrose synthase plays a physiologically important role. (Islam, Matsui and Yoshida, 1996) Carbohydrates are transported from photosynthetic source tissues to sink tissues in the form of sucrose. The consequent cleavage of sucrose in the sink tissues is the first step for its use in various metabolic pathways. The sugar is cleaved in vivo by either sucrose synthase (Sus) or by invertase. Invertase catalyses an irreversible reaction where sucrose is cleaved into glucose and fructose (Matic et al., 2004) while sucrose synthase catalyses the reversible conversion of sucrose and uridine-diphosphate (UDP) into uridine-diphosphoglucose and fructose. (Hirose, Scofield and Terao, 2008) (Hardin and Huber, 2004) These enzymes play a crucial role in plant growth and development. (Abid et al., 2009) Sucrose Synthase is cytosolic (Šebková et al., 1995) and has been characterized in many different plant species such as maize (Hardin and Huber, 2004), rice (Odegard, Liu and Lumen, 1996) and sugarcane (Schäfer, Rohwer and Botha (2005)). Its activity has been studied in many plant organs such as roots, leaves and seeds. (Šebková et al., 1995)
For trees, cellulose biosynthesis is a highly regulated process in which carbon is permanently placed in their primary and secondary cell walls. Sucrose is the main carbon source for cellulose synthesis. The stem is made up of extremely active sink cells which utilise sucrose for cellulose synthesis. Sucrose synthase is the main sucrolytic enzyme in these cells that catalyzes the reversible conversion of sucrose into fructose and UDP-glucose which is needed for cellulose biosynthesis. (Joshi, Bhandari and Ranjan, 2004) It also plays an important role providing adequate sugar supply during anoxic stress. It has been shown that during anoxic germination of rice, sucrose synthase activity was enhanced whereas the activity of invertase was depressed. This would indicate that sucrose synthase is the enzyme predominantly responsible for sucrose breakdown during anoxia. (Joshi, Bhandari and Ranjan, 2004)
Fig 1: Diagram of the cleavage and synthesis reaction of sucrose synthase (Römer et al., 2004)
Different isoforms of the gene are present in most plants. In the case of maize, two non-allelic genes were discovered for sucrose synthase but more investigation lead to the discovery of a third. At least three genes for sucrose synthase have been discovered in rice where the genes show differences in expression between tissues. RSus1 is expressed in root phloem while RSus2 is expressed in leaf phloem. (Schäfer, Rohwer and Botha, 2005) When examining the different isoforms at an amino acid level it is appears that there is less homology between different sucrose synthase genes in a species than when the gene is compared to its corresponding gene in another species. In the case of maize, there is 75% homology between the SS1 gene and SS2 gene of maize but there is 90% homology between rice RSus1 and maize SS2 genes. In sugarcane, the SS1 gene is 97% identical at the amino acid level to maize SS1 gene. (Lingle and Dyer, 2001)
Nolte and Koch (1993) undertook a study to determine whether sucrose synthase was localized to certain part of the vascular strand. It is well known that sucrose synthase is present in vascular bundles for example in transgenic tobacco plants phloem specific expression of a maize sucrose synthase gene has been observed. Their study, using immunohistochemistry, found that sucrose synthase was restricted to the cytoplasm of companion cells of the phloem and did not appear to be present in other organelles of the plant. (Nolte and Koch, 1993)
The molecular mass of sucrose synthase can be determined by gel filtration. Sucrose synthase elutes from the column with a Kav value of 0.17844 which when using a calibration curve correlates to a molecular mass of 362kDa. Using SDS-PAGE gradient gel the molecular mass of each subunit can be estimated at 92kDa. This can conclude that sucrose synthase is a tetrameric enzyme with a molecular mass of 360kDa and four identical subunits of 90kDa. (Hardin and Huber, 2004) (Elling and Kula, 1993) It can associate with membranes and the actin cytoskeleton where its activity is known to be involved with cellulose synthesis. It does this by channelling uridine-diphosglucose to the growing glucan chain by the enzyme cellulose synthase. (Hardin and Huber, 2004)
Analysis of Sucrose Synthase Gene Family:
From the results of species examined to date, it is shown that sucrose synthase is encoded by a small multigene family. (Bieniawska et al, 2007) Most species of plants have at least two isoforms of sucrose synthase. These isoforms usually have comparable biochemical properties and highly homologous amino acid sequences. (Wen et al., 2010) Further analysis of transgenic and mutant crop plants show certain isoforms of sucrose synthase have specific functions in the plant. The rug4 mutation of pea removes the SUS1 isoform but has no effect on SUS2 or SUS3. This would indicate that these two isoforms are not able to make up for the loss of SUS1 in the seed or root nodule. It is clear that the loss of different isoforms affect the plant in certain ways. Loss of the SH1 isoform in maize has different outcomes from the loss of SUS1 isoform. SH1 is required for normal cell wall formation during endosperm development while both isoforms are needed for wild-type rates of starch synthesis. Why different isoforms have different functions is unclear. The same functions can be carried out in the cell by different isoforms but can occur in distinct cell types, developmental periods or environmental conditions. It is likely that different isoforms could have non-overlapping, particular functions in the same cell. (Bieniawska et al., 2007)
It is difficult to decide on the precise roles of the genes in sucrose synthase gene family when there is not enough information in existence. Although there is some information available on some of the isoforms and they're functions in the plant, no analysis of the functions of the gene family has been carried out. The model plant Arabidopsis is ideal for carrying out such an analysis. Six sucrose synthase genes are in the Arabidopsis genome. Based on comparisons of the amino acid sequences the isoforms they encode can be divided into three distinct pair groups. The isoforms SUS1 and SUS4 are 89% identical to each other but have less than 68% similar amino acid sequences to other isoforms. Similarly, SUS2 and SUS3 are 74% identical to other isoforms and are 67% less identical to the other forms of enzyme. SUS5 and SUS6 are 585 identical to each other but have less 48% similarity to the other isoforms. When examining other dicotyledonous species it appears that at least two of the three pairs of isoforms are present. When phylogenetic analysis was carried out, it showed that the isoforms AtSUS1 and AtSUS4 are related to pairs of isoforms from pea (Fabacae), carrot (Umbelliferae) and potato (Solanacae). A pair of isoforms from Craterostigma plantagineum (Scrophulariacae) is closely related to the pair of isoforms AtSUS2 and AtSUS3 in the Arabidopsis. The pair AtSUS5 and AtSUS6 is related strongly to a pair of genes from rice. This evidence shows that it is unlikely that the three pairs of isoforms in Arabidopsis are as a result of gene duplication events. It is possible that each isoform has an exact function preserved in a wide range of plants. The members of Arabidopsis gene family are strongly differentially expressed in different organs of the plant through its development and in response to external stimuli e.g. environmental stress. This is seen in gene families of other plants studied. (Bieniawska et al., 2007)
Fruit quality is determined by the type and quality of sugars present. A study of the sucrose synthase-encoding gene from the muskmelon fruit was carried out to evaluate how to genetically improve the quality of the fruit. This is done by finding the sugar components in fruit, to identify the enzymes involved in sugar metabolism and distinguish the relationship between sugar accumulation and the activities of related enzymes. It is thought that sucrose synthase is the enzyme involved in metabolising sucrose in developing muskmelon fruit. To examine this, a full length cDNA strand encoding sucrose synthase was extracted from a muskmelon fruit by RT-PCR and RACE and identified as CmSS1. Real time PCR analysis showed that CmSS1 expression changed in among different tissues of the plant e.g. root, stem, leaf. It showed that the mRNA levels are highest in the root and lowest in mature fruit.
Fig 2: The patterns of CmSS1 transcript abundance in the different tissues of the muskmelon plant. These results were found using quantitative real-time PCR analysis of total RNA prepared from the root, stem, leaf, flower and mature fruit of muskmelon.
During fruit development and ripening it was shown that CmSS1 mRNA was at its maximum level at five days after pollination and decreased steadily during fruit development until it reached its minimum level of maturity. This was discovered using again real-time RT-PCR analysis of mesocarp tissues from five days of pollination to ripening.
Fig 3: This graph depicts the patterns of CmSS1 transcript abundance in developing muskmelon fruits found by using quantitative real-time PCR analysis of total RNA prepared from muskmelon. (Wen et al., 2010)
The sugar content and SS activity were analysed to show the functions of CmSS1 in regulating fruit quality. It showed that very low concentrations of sucrose are present in young and unripe muskmelons. Between 20 and 30 days after pollination there is a massive rise in the amount of sucrose in the fruit. Sucrose synthase activity increased in the direction of sucrose synthesis and decreased in the direction of sucrose cleavage through fruit development. (Wen et al., 2010)
Fig 4: The depiction of sucrose content and sucrose synthase activity during muskmelon fruit development. The first chart shows sucrose content during fruit development. The second shows sucrose activity in the sucrose synthesis direction and the third shows sucrose cleavage direction during muskmelon fruit development. (Wen et al., 2010)
Enzyme Kinetics of Sucrose Synthase:
An investigation was carried out by Schöfer et al. to the find the properties of three sucrose synthase isoforms present in sugarcane. Kinetic analysis indicated that the three sucrose synthase genes in sugarcane are different isoforms, with major differences in Km values and the ratios of sucrose breakdown synthesis. The kinetic characteristics of the SuSyA and SuSyB isoforms, both expressed in the leaf roll, differ greatly. It was found that SuSyA has almost three times higher affinity for sucrose than the SuSyB isoform whereas SuSyB has a much greater affinity for UDP than SuSyA. Based on the differences in their kinetic properties it can be concluded that SuSyB and SuSyC are different isoforms of sucrose synthase. SuSyC has roughly ten times higher affinity for UDP compared to the other two isoforms. (Schäfer et al., 2005)
Fig 5: The graph shows the Lineweaver-Burk plot of 1/v against 1/S for the isoforms SuSys A, B and c where UDP was the variable substrate. The concentration of sucrose was kept constant at 320nM. The Km values were determined from the non-linear fit of the data to the Michaelis-Menten equation. (Schäfer et al., 2005)
When examining sucrose synthase in soybean nodules Morell and Copeland (1985) found the kinetic constants of UDP, UDPglucose, sucrose and fructose by fitting the data to the following two equations:
1. v = VA/KiaKh + KhA + KhB + AB 2. v = VA/Ka + A + A/Ki
The kinetic constants for ADP, CDP and ADPglucose were found using non linear regression analysis of initial velocity data.
Fig 6: Graph showing the effect of sucrose concentration on the cleavage activity of sucrose synthase in soybean nodule. The lines show the fit of data to equation 1. The reaction mixture were composed of 20µmol Hepes-KOH buffer (pH 7.5) 2µmol UDP, 1.5µmol NAD, 25µg UDPglucose dehydrogenase. Each symbol represents a different concentration of sucrose. The dark circle shows 3.2µM, the clear circle shows 4µM, the dark triangle shows 6.25µM, 10µM is shown by the clear triangle and the dark square depicts 20µM.
In the cleavage and synthesis direction standard Michaelis-Menten kinetics are observed. The variation of concentration of sucrose at different concentrations of UDP gave an intersecting pattern of linear double reciprocal plots. (Morrell and Copeland, 1985)
V (U/mg protein)
Km sucrose (mM)
Ki sucrose (mM)
Km UDP (mM)
Ki UDP (mM)
Fig 7: Table showing the kinetic parameters for the cleavage reaction of sucrose synthase in soybean nodules. (Morrell and Copeland, 1985)
Fig 8: The graph depicting the effect of UDPglucose concentration on the synthesis reaction of sucrose synthase activity in soybean nodules. The reaction mixtures contained 20µmol Hepes-KOH buffer, 15 µmol fructose, 5µmol MgCl2, 0.4 µmol P-enolpyruvate, 0.15 µmol NADH, 20µmol KCl, 25µg pyruvate kinase 25µg lactate dehydrogenase and the required amount of enzyme. As in the previous graph, the amount of UDPglucose was varied in the presence of 2.5mM (dark circle), 3.2mM (clear circle), 4mM (dark triangle), 5mM (clear triangle) and 8mM (dark square) fructose. The results on the graph are representing the fit of data to equation 1.
When the concentration of UDPglucose was varied at the concentrations of fructose in the graph, an intersecting pattern of linear double reciprocal plots was seen. From fitting the data from the graph to equation 1, it is noted that substrate inhibition would have occurred at a concentration greater than 15mM fructose.
V (U/mg protein)
Km fructose (mM)
Ki fructose (mM)
Km UDPglucose (mM)
Ki UDPglucose (mM)
Fig 9: table showing the kinetic results by fitting the figures from the graph to equation 1.
When partially purified SuSyA, SuSyB and SuSyC were blotted to a nitrocellulose filter the results showed that all three isoforms are approximately 94kDa. (Schäfer et al., 2005) The would correlate to the findings of Hardin et al and Lothar et al who stated that sucrose synthase is tetrameric enzyme made up of four 90kDa subunits.
Fig 10: Immunoblot of sugarcane SuSy. A crude extract of protein from leaf roll was loaded into lane 2 while partially purified isoforms of SuSyA, SuSyB and SuSyC were loaded to lane 3, 4 and 5. The molecular weight ladder was used to identify the bands see in each lane. (Schäfer et al., 2005)
Characteristics of Sucrose Synthase:
Extraction of Protein:
The method for extracting protein from the leaves of maize (Zea mays), rice (Oryza sativa) and tobacco was done as follows: 1-3g of leaves was ground in liquid nitrogen and the powder was mixed in the ratio 1:2 with extraction buffer. The buffer was made up of 0.1M tris-HCl, pH 8, 10mM DTT and 1% polyvinylpolypyrrolidone. The samples were then incubated on ice for 15 minutes and then centrifuged at 1,000g for 10 minutes at 4oC. The pellet was then removed and the supernatant was re centrifuged at 100,000g for one hour at 4oC. After this final centrifugation, the pellet and supernatant which contained the soluble proteins was resuspended in sample buffer for electrophoresis. (Persia et al., 2008) When extracting protein from rice seeds, a similar procedure is followed. Seeds weighing roughly 50-100mg at various stages of growth were homogenized in 400µl of extraction buffer and kept at 4oC. The buffer was made up of 50mM Tris-HCl, pH7.5, 1.0mM DTT, 1.0mM EDTA and 2mM PMSF. Ammonium sulphate fractions (30-50% w/v) were precipitated and then resuspended in dialysis buffer made up of 50mM Tris-HCl, pH 8.0, 5mM MgSO4, 5mM 2-mercaptoethanol. This was then dialyzed overnight at 4oC. (Odegard, Liu and De Lumen., 1996) The method for extracting protein from tobacco pollen tubes is slightly different to those mentioned previously. The pollen first was slowly thawed from storage at -20oC and hydrated in a humid chamber overnight. It was then germinated in BK medium and allowed to germinate at 25oC for three hours. After this period had elapsed, the pollen was collected by centrifugation at 1,000g for 5 minutes at 25oC. It was then washed twice with BRB25 buffer which is made up of 25mM HEPES, pH 7.5, 2mM EGTA and 2mM MgCl2 and 15% Suc. After washing, the pollen was resuspended in lysis buffer and lysed on ice using a motor-driven Potter-Elvehk-jem homogenizer. The lysis buffer used was made up of BRB25 buffer along with 2mM dithiothreitol, 1mM phenylmethylsulfonyl fluoride (PMSF), 10µL/mL protease inhibitors, 1mM NaN3 and 10% mannitol. After lysis was carried out, the samples were centrifuged at 1,000g for 10 minutes at 4oC. The supernatant was centrifuged again at 4oC for 45 minutes at 100,000g over a 20% (w/v) Suc cushion. The supernatant was then collected as it contained the soluble protein fraction. (Persia et al., 2008)
After extracting protein, the sucrose synthase activity in sugarbeets was found using a spectrophotometric end point assay. The activity of the enzyme was monitored as fructose formed at 35oC. This was carried out in a solution that contained 250mM sucrose, 2mM UDP and 100mM MES. The control was carried out by assaying for activity in the absence of UDP. The total protein concentration was determined using the Bradford method where bovine serum albumin was the standard. (Klotz and Haagenson., 2008) When assaying for protein from rice, the Bradford method was followed to determine protein concentration as was done in Klotz et al. 40mg of protein was used per assay. The assay was carried out in 20mM MES pH 6.4, 200mM sucrose and 4mM UDP for 15 minutes at 30oC. The reaction was stopped by boiling for 2 minutes and the fructose levels were measured. The control tubes did not contain UDP. (Odegard, Liu and De Lumen., 1996) When examining the effect of sucrose synthase on carbon partitioning a similar method was followed for assaying the protein. Sucrose synthase was assayed in the direction of sucrose breakdown using 50µl poplar plant extract. The tetrazolium blue assay was followed to determine the amount of free fructose. As in previously mentioned assays, the absence of UDP in the assay acted as a control. The total protein content was found by employing the Bradford (Bio-Rad) protein assay. (Coleman, Yan and Mansfield., 2009) A similar method was followed for carrying out an assay for the enzyme on tomato tissue. The reaction mixtures contained 50mM Hepes-NaOH buffer, 15mM MgCl2, 25mM fructose and 25mM UDP glucose. This was incubated at 37oC for 30 minutes and was terminated with the addition of 70µl of 30% KOH. The enzyme blanks were terminated with the addition of KOH at 0 minutes. The tubes were then kept at 100oC for 10 minutes to destroy any fructose. The soluble protein content was determined using the Lowry method whereby bovine serum albumin was the standard. (Islam, Matsui and Yoshida., 1996) Alkaline copper solution is added to each tube and allowed to stand at room temperature for roughly 30 minutes. Dilute folate reagent is then added to each tube rapidly and after 30 minutes the absorbance is read at 750nm. (Lowry et al., 1951) The results were measured as µmole of sucrose per minute per mg protein. (Islam et al., 1996) When assaying for sucrose synthase in the cleavage direction Römer et al used recombinant SuSy1 gene from potato. In a volume of 100µl HEPES buffer with a concentration of 200mM and pH 7.6 recombinant sucrose synthase was incubated along with 2mM UDP and 500mM sucrose for ten minutes at 30oC. HPLC analysis was used for the formulation of UDP-glucose. The Bradford assay was used to determine protein concentrations as was carried out by Klotz et al and Coleman et al. The activity of the enzyme was also tested with the nucleoside diphosphates dTDP, CDP, ADP and GDP at 2mM. For assaying recombinant enzyme in the synthesis direction a similar method was followed as when assaying for standard enzyme. Recombinant sucrose synthase was incubated in a total volume of 100µl HEPES buffer where this time the pH was 8.0 and the concentration was as in cleavage direction of 200mM. 1mM UDP-Glc and 20mM D-fructose was also added to the mixture and it was incubated for five minutes at 30oC. The reaction was heated to 95oC for five minutes and HPLC analysis was used to establish the formation of UDP. The sucrose synthase activity was also tested using dTDP-Glc, CDP-Glc and ADP-Glc. (Römer et al., 2004)
Purification of Protein:
After extraction of the protein from the crude extract, purification can be carried out. This can be done in a number of ways such as Batch adsorption with Sephadex A50, Anion exchange chromatography and Gelfiltration. SDS-PAGE can be carried out after purification to check the purity of the protein sample. The Sephadex A50 gel is loaded into a glass funnel and washed twice with deionised water. The gel was then washed twice with 300ml standard buffer. The protein sample was loaded to the gel and slowly sucked through the gel for 30 minutes. The gel bed was then washed with 300ml standard buffer and then with 300ml standard buffer containing 100mM KCl. The last washing step contained 300mM KCl. 200ml of the first salt preparation was concentrated to 40-50ml by using a cross-flow ultrafiltration module with YM 30 ultrafiltration membrane that had been pretreated with 55 PEG 4000 solution. This was done to prevent the enzyme sticking to the membrane. In anion exchange chromatography a Sepharose Q column was first equilibrated with 300ml Hepes buffer. This was made up of 200mM pH 8 with 50mM KCl. 70-80mg of protein sample was loaded and the elution was started using two different salt gradients. To prevent enzyme inactivation after elution all the fractions were titrated back to pH 7.2. All fractions that contained enzyme activity were pooled and concentrated by using ultrafiltration. Gelfiltration experiments are carried out on a prepacked HiLoad 16/60 Superdex 200 prep grade column that was connected to FPLC equipment. Four samples containing 2mg of protein were loaded and eluted with a flow rate of 1 ml min-1. The fractions were then pooled and stored at -20oC in 500µl aliquots. (Elling and Kula., 1993) To determine the purity of the protein, SDS-PAGE is carried out. This is done by loading 100µg of protein samples to a 125 SDS-polyacrylamide slab gel that was overlaid with stacking gel. The electrophoresis was carried out at 4oC and at 40V for 16 hours and followed by 200V for one hour. Coomassie blue R 250 was used to stain the gel followed by destaining. (Kumutha et al., 2008)
Factors that affect Sucrose Synthase Activity:
Šebková et al (1995) stated that sucrose synthase has two different pHs for optimal activity. In the cleavage direction it was found that most enzyme activity was observed between pH 6.0 and 8.5 at temperatures between 50 to 55oC. In the synthesis direction, a pH between 8.5 to 9.5 and a temperature of 35oC was optimal for enzyme activity. (Šebková et al., 1995) This would correlate with the findings of Morell and Copeland (1985) who found that optimal activity of the enzyme in soybean was at pH 6 in the cleavage direction and at a pH of 9.5, sucrose synthase activity in the synthesis direction was at its highest. It was also found that at a pH of 7.5 the cleavage and synthesis activities were their highest. (Morrell and Copeland., 1985) Elling and Kula (1995) examined the effect of buffers TES-NaOH, MOPS-NaOH, TEA-NaOH and Tris-HCl on the pH optimum of sucrose synthase activity. These were determined using UDP and TDP as substrates for the reaction. They found that the enzyme had its highest activity in Hepes-NaOH buffer. When MOPS-NaOH and TES-NaOH buffer was used, only 60-80% activity was noted. (Elling and Kula 1995) It was also found that the velocity of the reaction could be increased by increasing the temperature where optimal activity was seen between 50 and 60oC. Xu at al (1989) reported that potato and bean are also able to withstand these high temperatures. However once the temperature goes above 60oC enzyme activity starts to decreased rapidly and was destroyed once the temperature reached 70oC. (Xu et al., 1989) The cleavage of sucrose by the sucrose synthase enzyme was investigated to find the rate of cleavage reaction using different nucleosidediphosphates as cosubstrates. They found that the rate of reaction was UDP>TDP>ADP>CDP>GDP. Echt and Chourey (1985) found similar results when examining nucleotide specificity. They found that substrate specificity for SS1 and SS2 were UDP>TDP>ADP>CDP>UTP where each substrate was at a concentration of 4mM. (Echt and Chourey 1985) Low levels of heavy metal ions such as mercurate inhibited cleavage activity of the enzyme. This would lead to the assumption that sulfhydryl groups are involved in the catalytic process. It is also inhibited by Tris-HCl and by small concentrations of MgCl2 and MnCl2. (Šebková et al., 1995) Cations were shown by Elling and Kula (1995) to have a slight influence on enzyme activity. The activity was lessened slightly (10%) by the presence of 1mM Mn2+ and Mg2+ ions with UDP. The enzyme is completely inactivated in the presence of 1mM Cu2+ or Fe2+. (Elling and Kula., 1993)
A recent study was undertaken to examine the effects of volatile emissions on carbohydrate metabolism. Studies on this area have taken place before but it is usually examining the results of physical contact between the host plant and the microbe. No work has taken place until now on the effect on the plant in the absence of physical contact. Many microbes such as Pseudomonas spp, Strepomyces spp, Penicillin spp and a selection of truffles produce ethylene. This gaseous plant hormone plays an important role in many aspects of plant growth and development such as seed germination, root hair initiation, fruit ripening and starch accumulation. In the work of Ezquer et al (2010), the possible effects of volatiles released from gram-negative bacteria, gram-positive bacteria and fungi on starch metabolism was studied. The results showed that the volatile compounds released by microbes promoted high levels of starch accumulation in mono- and dicotyledonous plants. It also revealed fungal volatiles (FVs) promoted massive changes in expression of genes involved in many important processes in plant such as metabolism of carbohydrates, amino acids, sulphur and lipids, energy production, protein translation and stability, cell wall biosynthesis and photosynthesis. However no changes were noted in the expression in some of the genes that coded for proteins involved in starch and sucrose metabolism such as plastidial hexokinase, plastidial phosphoglucose isomerase, plastidial adenylate kinase, alkaline invertase and UDPglucose (UDPG) pyrophosphorylase.
It was found in the study that FVs strongly upregulate the expression of Sucrose Synthase in potato leaves. The plants were cultured in the presence and absence of FVs emitted by A. Alternata. This caused a massive enhancement of expression of Sus4 isoform. A 29.4- and 31.63-fold increase was observed in expression when the plants were cultured in the presence and absence of sucrose. This isoform of the enzyme controls the accumulation of ADPG, UDPG and starch in potato source leaves and tubers. Analyses of the intracellular amounts of starch and nucleotide-sugars in the leaves of the plant show a positive correlation between patterns of enzyme activity and starch, UDPG and ADPG amounts. This was noted when the leaves were cultured in the presence and absence of FVs. Western blot analyses and quantitative RT-PCR confirmed also the increase in expression. (Xu et al., 1989)
Environmental Factors affecting Sucrose Synthase Activity:
Waterlogging is where oxygen supply is blocked to root leading a severe decrease in the amount of oxygen available to the plant. This leads to inhibition of root respiration that causes a major decline in energy of root cells affecting vital metabolic processes of the plant. This is restriction of oxygen supply is known as anoxia. The presence of glucose in an anoxic incubation medium drastically decreases meristem death and studies have shown that sucrose synthase is the enzyme mainly responsible for sucrose breakdown under anoxia. (Kumutha et al., 2008) The increase in glycolytic demands caused by these demands is the cause of increased sucrose synthase expression. This has been demonstrated in many plant species e.g. sucrose synthase gene is induced in wheat and in rice when oxygen levels are low. (Ricard et al., 1998) Harada et al (2005) also found an increase in sucrose synthase activity in pondweed turins while under anoxia. (Harada et al., 2005) Klotz and Haagenson (2008) found that sugarbeet contained two genes for sucrose synthase activity-SBSS1 and SBSS2. They demonstrated that anaerobic conditions caused a large increase in the transcription levels of SBSS1 and a quick increase and succeeding decline in SBSS2 transcription levels. However this did not correlate with a significant increase in sucrose synthase enzyme activity. A 23% increase in sucrose synthase activity was noted after initiation of anaerobic conditions but otherwise the activity of the enzyme did not differ greatly to that of the controls. (Klotz and Haagenson., 2008)
Fig 11: The graph outlines the different rates of sucrose synthase activity in the control and anaerobic treated plant. The asterisk shows the days when there was major difference, in this case 23% difference, in the control and anaerobic roots. (Klotz and Haagenson, 2008)
Experiments carried out by Kumutha et al indicate that the ability to utilise sucrose synthase while under oxygen deprivation may be due to different genotypes of the plant being more capable of using the enzyme when stressed. When examining the effect of water logging on carbohydrate metabolism in pigeon pea, they used four different genotypes: two tolerant to water logging stress (ICPL 84023 and ICP 301) and two susceptible (ICP 7035 and Pusa 207). ICPL 84023 and ICP 301 had an increase in enzyme activity until the fourth day of treatment whereas ICP 7035 and Pusa 207 had a continuous decline in enzyme activity while waterlogged. When the plants were removed from waterlogging treatment, the sucrose synthase activity in ICP 7035 and Pusa 207 increased slightly while the activity in ICPL 84023 and ICP 301 decreased. This would point to a carbohydrate-based tolerance mechanism in ICPL 84023 and ICP 301 that gives them a greater concentration of total, non reducing and reducing sugars along with greater enzyme activity of sucrose synthase over the susceptible genotypes. (Kumutha et al., 2008)
Fig 12: Bar chart which depicts the effect of waterlogging on sucrose synthase activity (A) and alcohol dehydrogenase activity (B) in root tissues of the pigeon pea genotypes. ICPL 84023 and ICP 301 showed a steady increase in enzyme activity until the fourth day of treatment where the 2.4 and 2.5 times higher than the pre-stress levels. ICP 7035 and Pusa 207 showed a continuous decline in activity. (Kumutha et al., 2008)
Decrease in Temperature:
Klotz and Haagenson (2008) noted that exposure to cold temperature for 24 hours increased SBSS1 and SBSS2 transcript levels in roots of plants when compared to controls. When plants were exposed to 4oC for 24 hours the transcription levels of SBSS1 and SBSS2 were 30-60% higher than in the controls. These differences were not observed however in sucrose synthase activity or in protein levels. The levels of SBSS1 and SBSS2 protein were very in the controls as in the cold treated roots. The activity of sucrose synthase was similarly unaffected by cold treatment. Roots were stored at 2oC and 20oC. No significant differences were noted in activity between the two different temperatures apart from a 19% decrease in activity in the roots treated at 2oC at 0.3h after the initial cold treatment.
Fig 13: Graph showing the sucrose synthase activity in roots after being treated with cold temperatures. The asterisk indicates the times at which enzyme activity was notably different between 2 and 20oC treated roots. (Klotz and Haagenson et al., 2008)
Phylogenetic Analysis of Sucrose Synthase:
The entire sucrose synthase (Sus) protein sequences were aligned using ClustalW to investigate the molecular evolution and phylogenetic relationship between sucrose synthase in plants and the tree was calculated by neighbour-joining algorithms. The distance method produced one tree by using the full length of the alignment of all plant Sus selected.
From the tree it is clear that Sus can be broken up into six main groups: A, B, C, D, E and F. Of the 52 plant genes analysed, 17 were classified into group A. Each of the main groups was further sub divided into smaller groups of related proteins. In the D and E group the massive expansion of dicotyledonous and monocotyledonous Sus genes shows that these two groups of genes should have extended both before and after the monocot-dicot split. The monocotyledonous Sus genes are clustered together in one specific group (group C). This shows a considerable split between monocotyledonous and dicotyledonous Sus. The evolution of dicotyledonous plant groups A and B probably occurred due to two duplication events: group B and the common ancestor group A were produced as a result of the first duplication and the second duplication produced the last group. Some putatively orthologous and paralogous pairs can also be noted from the tree such as Ms_SUS1/MT_SUS1, Hv_SUS1/Ta_SUS1, Mt_SUS3/Ps_SUS3 and Os_SUS/Os_SUS6. The most homogenous group on the tree is group B. Sus genes are known to be well conserved over a long period of evolutionary time and genes that provide comparable functions belong to the same group. This would mean that the identification of orthologs and paralogs would aid in a massive way the annotation of uncharacterised Sus. (Abid et al., 2009)
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Figure 14: The phylogentic tree from the analysis of aligned amino acid sequences from many Sus genes. The amino acid sequences were aligned using the programme ClustalX-2.0.3-win. The programme Protdist was used by distance analyses and the tree was calculated by neighbour-joining algorithms. (Abid et al., 2009)
Sucrose Synthase and the Chloroplast:
Sucrose formed in leaves is done so by using photosynthesis. After sucrose formation, it is transported other parts of the plant such as roots, seeds and tubers where it is converted to starch. Carbohydrate metabolism is an example of integrated control between the cytosol and plastid. Gluconeogensis is carried out in both compartments although starch is restricted to the plastids and sucrose is metabolised in the cytosol. The formation of both of these is biochemically regulated by metabolites and enzymes that exist in the cytosol and chloroplast.
Sucrose synthase activity is usually highest in sink tissues that require sucrose for starch synthesis. Young leaves require more starch than mature leaves and therefore have higher sucrose synthase activity. Sucrose synthase produces UDPG in heterotrophic tissues which can be converted to ADPG by UGPase and AGPase. ADPG can then be used as a substrate for starch biosynthesis. UDP is generally recognised as the main nucleoside diphosphate for sucrose synthase although studies by Curatti et al., (2000) have shown that ADP can act as a substrate for sucrose synthase to produce ADPG.
Fig 15: The model scheme showing starch biosynthesis in green leaves. The scheme shows the coordinated actions of sucrose synthase and ADPG. The enzymes are numbered as follows: 1. Sucrose synthase, 2. Starch synthase. 3. Amylase, 4. Hexokinase, 5. cpPGM and 6. AGPase.
Experiments using isolated chloroplasts show that chloroplasts can both synthesize and mobilise starch and that cpPGM and AGPase have a role to play in the sourcing of glucose molecules from starch breakdown. As sucrose synthase can produce ADPG from sucrose and ADP and the availability of starch in the chloroplast, it has been suggested that sucrose synthase can catalyse the de novo synthesis of ADPG in the cytosol. (Romero, Ardila and Akazawa., (1991)). This implies that a form of C6 molecule enters the chloroplast and is used as a precursor for starch biosynthesis. The rate of starch biosynthesis could be determined by the rate of import of ADPG synthesised by the enzyme in the cytosol. When there is high light intensity, sucrose will accumulate and sucrose synthase will produce all the ADPG necessary for starch biosynthesis and when the light intensity decreases, the production of ADPG will be decreased and starch biosynthesis will be decreased. (Baroja-Fernández et al., 2001)
Sucrose metabolism is a very important process in the plant. The carbon formed from sucrose is required for many processes such as respiration, cell wall formation and starch synthesis. Sucrose synthase is a key enzyme in sucrose metabolism. This enzyme is encoded by a small multigene family where most plant species have at least two isoforms of sucrose synthase. Kinetic analysis of the enzyme show that Km values and different ratios of sucrose breakdown occur for different isoforms of the enzyme. When characterising the enzyme, methods of extraction, assaying and purification are examined. It was shown that pH, buffers, temperature, metal ions, fungal volatiles and anoxia all influence sucrose synthase activity. Sucrose synthase is normally present in the cytoplasm but the availability of sucrose in the chloroplast and the enzymes ability to use ADP indicates that the enzyme could potentially be used in the chloroplast.