Cadmium induced changes in carbohydrate status and enzymes of carbohydrate metabolism, glycolysis and pentose phosphate pathway in pea

Environmental and Experimental Botany 61 (2007) 167–174

Cadmium induced changes in carbohydrate status and enzymes of carbohydrate metabolism, glycolysis and pentose phosphate pathway in pea Rachana Devi, Nidhi Munjral, Anil K. Gupta, Narinder Kaur ∗ Department of Biochemistry and Chemistry, Punjab Agricultural University, Ludhiana 141004, India Received 9 February 2007; accepted 20 May 2007

Abstract With 50 ␮mol/l CdCl2 , reduction of about 40% in shoot and 70% in root lengths was observed in pea seedlings. The reduction in seedling growth was accompanied by decreased water content of shoots and roots. Increasing trend of sucrose was observed in shoots of cadmium-stressed seedlings. Cotyledons of stressed seedlings had lower ␣-and ␤-amylase activities resulting in their higher starch content. Reducing sugar and sucrose contents were not significantly affected in roots of cadmium-stressed seedlings but starch content was low in comparison with normal seedlings which appeared to be due to lesser conversion of sucrose into starch rather than its rapid hydrolysis. In general activities of acid and alkaline invertases were low in stressed seedlings with exception to cotyledons where alkaline invertase was more under stress. Cadmium-stressed seedlings had lower sucrose synthase and sucrose phosphate synthase (SPS) activities in roots and shoots but cotyledons had 10–15% higher SPS activity. Phospho glucoisomerase activity was maximum in cotyledons showing its involvement in conversion of glucose-1-phosphate to glucose-6phosphate whereas the activity of this enzyme was almost negligible in roots. Exogenous cadmium up regulated the activity of glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase in cotyledons whereas activities of these enzymes were down regulated by 40–50% in shoots. Higher activities of hexokinase (10–15%) and phospho glucoisomerase in roots and shoots of stressed seedlings indicated that hexoses are preferably channeled towards glycolysis than pentose phosphate pathway under cadmium toxicity. © 2007 Elsevier B.V. All rights reserved. Keywords: Cadmium toxicity; Pisum sativum; Carbohydrate metabolism; Glycolysis; Pentose phosphate pathway

1. Introduction The increased tempo of industrial activity has resulted in the discharge of high volume of liquid effluents. The sewage water contains high amounts of toxic elements like lead, cadmium, nickel, mercury, arsenic and other organic and inorganic chemicals. Cadmium, a common pollutant from various industrial productions, to a great extent is accumulated in the soil. The main sources of cadmium pollution are metal mining, manufacture and disposal as well as applying of cadmium containing phosphate fertilizers, sewage sludge and pesticides (Van Bruwaene et al., 1984). It is readily taken up by plants and can interfere with many physiological processes associated with normal growth and development (Artetxe et al., 2002; Maksymiec and Krupa, 2002). Mercury, cadmium and lead all show a strong affinity for

∗

Corresponding author. E-mail address: [email protected] (N. Kaur).

0098-8472/$ – see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.envexpbot.2007.05.006

ligands such as phosphates, cysteinyl and histidyl side chains of proteins, purines, pteridines and porphyrins. Hence, these elements can act on a large number of enzymes having functional –SH groups, bind to and affect the conformation of nucleic acids and disrupt pathways of oxidative phosphorylation, although in each instance the precise response depends upon the individual properties of the metal (Fodor, 2002). Unlike heavy metals, the other abiotic stresses like drought, salt and temperature do not have significant effect on sulphydryl groups of proteins. The toxicity of heavy metals for plant metabolism, including photosynthesis, which is one of the most metal sensitive processes, is well known (Krupa and Baszynski, 1995 and Fodor, 2002). One of the major consequences of exposure of plants to heavy metals is the enhanced accumulation of oxygen free radicals. Peroxidases can be used as stress markers in heavy metal poisoning (Mocquot et al., 1996). However, increase in their activity is thought to be a common response and can protect plants from various stress factors (Gaspar et al., 1985) as all abiotic stresses result in oxidative stress.

168

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

Degradation of starch and synthesis of sucrose in cotyledons/endosperms is one of the essential features for germination. Sucrose synthesized in such source tissues is translocated to growing tissues (embryonic axis and then shoot and root) wherein it is metabolized by invertase and sucrose synthase. The hexoses thus formed are partitioned into glycolytic cycle and pentose phosphate pathway. The effect of heavy metals on these metabolic events is not clear. Environmental stresses like drought, cold and salinity lead to major alterations in carbohydrate metabolism (Gupta et al., 1993; Wanner and Junttila, 1999; Kaur et al., 2000; Gupta and Kaur, 2005) and up regulation of many genes corresponding to carbohydrate metabolism (Seki et al., 2002). High carbohydrate accumulation during early seedling development may reflect undesirable growth conditions (Lopez-Molina et al., 2001). Sugars regulate growth activities by modulations of gene expression and enzyme activities in both carbohydrate exporting and importing tissues (Coruzzi and Bush, 2001). The up regulation of extracellular invertase was suggested to be a common response to various biotic and abiotic stresses (Roitsch et al., 2003). In many plants, the genes for sucrose synthase and invertase are subjected to sugar regulation (Ehness et al., 1997). However, with exception to recently published report on carbohydrate metabolism in growing rice seedlings under arsenic toxicity (Jha and Dubey, 2004), there is hardly any literature on this aspect. Pea is an important vegetable crop that is grown all over the world. The present study was undertaken to understand the effect of cadmium toxicity on carbohydrate status and important enzymes of starch and sucrose metabolism along with pentose phosphate and glycolytic pathways in pea seedlings. 2. Materials and methods 2.1. Plant material and seedling growth Pea seeds (Pisum sativum L) cv PB 88 were dipped in 0.1% HgCl2 for 5–10 min and washed thoroughly with sterilized water under aseptic conditions. Seeds were germinated in absence and presence (0.01–1 mmol/l) of CdCl2 on 0.9% agar in conical flasks in an incubator at 25 ◦ C in the dark. Growth data were taken after 7 days of growth. Samples in triplicate were taken randomly from different flasks. 2.2. Extraction and determination of soluble sugars and starch Soluble sugars were extracted from 500 mg of each tissue twice with 80% and then with 70% ethanol. Ethanol extracts were combined and concentrated by evaporation at 50 ◦ C under vacuum (Chopra et al., 1998). Reducing sugars were determined colorimetrically using reaction with arsenomolybdate (Nelson, 1944). Sucrose was determined after its complete hydrolysis with invertase and then estimating glucose by glucose oxidase (Kaur et al., 2003). Starch was estimated in the residue left after the extraction of soluble sugars by hydrolysing it by excess amyloglucosidase and then estimating glucose as reducing sugars

(Kaur et al., 2003). Total sugars were estimated colorimetrically using phenol-sulphuric acid method of Dubois et al. (1956). 2.3. Extraction and determination of enzyme activities Enzymes were extracted at 4 ◦ C. Invertases were extracted by crushing the tissues (500 mg) with 3–4 ml of chilled 0.1 M NaCl (Dey, 1986). The extract was centrifuged at 10,000 × g for 10 min. The supernatant was passed through a sephadex G-25 column to remove reducing sugars. Acid invertase (EC 3.2.1.26) activity was measured by incubating 160 mmol/l of sodium acetate buffer (pH 5.0), 100 mmol/l of sucrose and 0.1 ml of enzyme in total volume of 1 ml at 37 ◦ C for 30 min. Reducing sugars formed were estimated (Nelson, 1944). The assay system for alkaline invertase (EC 3.2.1.27) was same as that described for acid invertase except that sodium acetate buffer was replaced by sodium phosphate buffer (pH 8.0). Sucrose phsophate synthase (SPS) (EC 2.4.1.14) and sucrose synthase (EC 2.4.1.13) were extracted from plant tissues (100–500 mg) with buffer (3–4 ml pH 8.2) containing 100 mmol/l HEPES, 10 mmol/l EDTA, 15 mmol/l KCl, 5 mmol/l MgCl2 , 2 mmol/l sodium diethyl dithiocarbamate and 5 mmol/l ␤-mercaptoethanol. Polyvinylpyrollidone (100 mg/g tissue) was also added. The extracted material was centrifuged at 10,000 × g for 15 min. The supernatant was made sugar free by passing through sephadex G-25 column using 10 mM HEPES buffer (pH 7.0). Assay system for SPS consisted of 1.1 ␮mole of UDP-glucose, 4.4 ␮mole of fructose-6-phosphate, 100 ␮mole of HEPES (pH 8.2) containing 3.5 ␮mole of MgCl2 and 2 ␮mole of NaF in a total volume of 40 ␮l. To this 100 ␮l of enzyme was added and contents incubated at 37 ◦ C for 15 min. The sucrose formed was determined using anthrone reagent (Kaur et al., 2000). The assay system for sucrose synthase was similar to SPS except that fructose-6-phosphate was replaced with fructose and NaF was not added in the assay system. Amylase was extracted with 50 mmol/l sodium acetate buffer (pH 5.0) containing 1 mmol/l CaCl2 and contents centrifuged at 10,000 × g for 10 min. ␣-Amylase (EC 3.2.1.1) activity was determined after destroying the ␤-amylase by heating the enzyme at 70 ◦ C for 20 min and estimating reducing sugars formed from 2% starch in 50 mmol/l sodium acetate buffer (pH 5.0) in presence of 1 mM CaCl2 at 37 ◦ C. ␤-Amylase (EC 3.2.1.2) was extracted with 100 mmol/l sodium acetate buffer (pH 3.6) containing 1 mmol/l EDTA. Activity of ␤-amylase was determined by estimating the reducing sugars formed after the enzyme action on 1% starch prepared in 50 mmol/l sodium acetate buffer pH 5.0 containing 1 mmol/l EDTA (Miyagi et al., 1990). Enzymes for assaying various glycolytic enzymes were extracted essentially by the procedure used by Copeland et al. (1989). The required tissue (100–500 mg) was homogenized in a chilled pestle and mortar with extraction buffer consisting of 3–5 ml 20 mmol/l HEPES buffer (pH 8.0) containing 1 mmol/l EDTA sodium salt, 5 mmol/l MgCl2 , and 5 mmol/l 2-mercaptoethanol. Polyvinylpyrrolidone (100 mg g−1 tissue) was also added during extractions. The homogenate was filtered through four layers of cheese cloth and centrifuged (1000 × g,

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

169

15 min). The supernatant was then centrifuged at 10,000 × g for 15 min and activities of glycolytic enzymes were determined in the supernatant. Assay system for glycolytic and pentose phosphate pathway enzymes were based on coupled reactions leading to reduction of NADP and increase in absorbance at 340 nm was monitored. Assay system for hexokinase (EC 2.7.1.1) consisted of 50 mmol/l HEPES buffer (pH 8.0), 5 mmol/l KCl, 5 mmol/l glucose, 5 mmol/l ATP, 7 mmol/l MgCl2 , 0.5 mmol/l NADP+ , 2 units of glucose-6-phosphate dehydrogenase and 0.1 ml of enzyme in total volume of 1 ml. Assay system of fructokinase (EC 2.7.1.4) consisted of 50 mmol/l HEPES buffer (pH 8.0), 5 mmol/l KCl, 0.4 mmol/l fructose, 1 mmol/l ATP, 1.5 mmol/l MgCl2 , 0.5 mmol/l NADP+ , 2 units of glucose-6phosphate dehydrogenase, 2 units of phospho glucoisomease and required amount of enzyme (0.1 ml) in total volume of 1 ml. Phospho glucomutase (EC 2.7.5.1) was assayed by incubating 50 mmol/l HEPES buffer (pH 8.0), 10 mmol/l MgCl2 , 0.5 mmol/l NADP+ , 2.5 mmol/l glucose-1-phosphate, 2 units of glucose-6-phosphate dehydrogenase and enzyme in total volume of 1 ml. Phospho hexose isomerase (EC 5.3.1.9) was assayed by incubating 50 mmol/l HEPES buffer (pH 8.0), 10 mmol/l MgCl2 . 0.5 mmol/l NADP+ , 2.5 mmol/l fructose-6phosphate, 2 units of glucose-6-phosphate dehydrogenase and enzyme in total volume of 1 ml. Fructose 1,6 bisphosphatase (EC 3.1.3.11) was assayed by incubating 50 mmol/l HEPES buffer (pH 8.0), 5 mmol/l MgCl2 , 4 mmol/l fructose-1,6-bisphosphate, 0.5 mmol/l NADP+ , 2 units of glucose-6-phosphate dehydrogenase, 2 units of phospho glucoisomerase and enzyme in total volume of 1 ml. Glucose-6-phosphate dehydrogenase (1.1.149) was assayed by the method of Yuan and Anderson (1987). The reaction mixture contained 50 mmol/l HEPES buffer (pH 7.5), 0.2 mmol/l NADP, 5 mmol/l MgCl2 , 10 mmol/l KCl, 1 mmol/l EDTA and 0.03% triton X-100 and enzyme. The reaction was started by adding of 2.5 mmol/l glucose-6-phosphate. The method for assaying 6-phosphogluconate dehydrogenase (1.1.1.44) was same as that of glucose-6-phosphate dehydrogenase except that instead of glucose-6-phosphate 6-phosphogluconate was added as a substrate. Protein content was estimated by the procedure of Lowry et al. (1951). Statistical analysis was done using Student’s t-test. 3. Results and discussion The data for the lengths and biomass of different tissues were taken at 7th day of seedling growth. Cadmium ions inhibited seedling growth and inhibition became more pronounced with increase in concentration of exogenous cadmium. With 50 ␮mol/l cadmium chloride about 40% reduction in shoot length and 70% reduction in root length was observed (Fig. 1). The reduction of seedling growth was accompanied by higher percent dry weight of cotyledons, shoots and roots (Fig. 1). Cadmium could possibly interfere with the water uptake capacity of the seedlings that might have resulted in reduced seedling growth. In Arabidopsis, cadmium is reported to induce stomatal closing leading to wilting, however, exogenous cadmium

Fig. 1. Effect of increasing concentration of CdCl2 on %dry biomass and seedling growth at 7 days of growth. Growth data are mean ± S.D. of three samples of 10 seedlings each.

did not cause significant differences in relative water content of leaf tissues (Perfus-Barbeoch et al., 2002). Nevertheless, strong decrease in plant growth, photosynthesis and leaf conductance in leaves of cadmium supplied Arabidopsis suggested perturbation of plant–water relationship (Perfus-Barbeoch et al., 2002). Since, the present study is with etiolated seedlings; the disturbance in plant–water relationship on cadmium application could lead to lower water content in different tissues.

170

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

Table 1 Effect of cadmium chloride (50 ␮mol/l) on the carbohydrate composition (mg g−1 FW) of different tissues of pea seedlings at different days of growth Tissue

Days

Total sugars

Reducing sugars

Control

Cd-stressed

Control

Sucrose Cd-stressed

Starch

Control

Cd-stressed

Control

Cd-stressed

Root

3 5 7

14.8 ± 1.1 9.7 ± 0.4 12.9 ± 0.3

14.4 ± 1.0 8.5 ± 0.3 9.5 ± 0.5

10.0 ± 0.4 9.4 ± 0.6 7.2 ± 0.7

9.2 ± 0.1 9.0 ± 1.1 9.8 ± 0.5

3.1 ± 0.3 3.3 ± 0.2 3.1 ± 0.3

3.4 ± 0.3 3.8 ± 0.3 2.8 ± 0.6

4.1 ± 0.6 3.9 ± .0.7 3.0 ± 0.4

3.5 ± 0.3 1.9* ± 0.2 1.1* ± 0.2

Shoot

5 7

9.8 ± 0.3 10.2 ± 0.3

10.6 ± 0.3 10.7 ± 0.4

6.9 ± 0.0 5.5 ± 0.1

5.3 ± 0.1 5.0 ± 0.4

4.0 ± 0.5 3.6 ± 0.5

4.8 ± 0.2 4.0 ± 0.3

2.0 ± 0.1 1.4 ± 0.1

2.1 ± 0.5 1.8 ± 0.6

Cotyledons 1 3 5 7

59 34 25 13

± ± ± ±

1 1 1 1

55 32 32* 26*

± ± ± ±

1 2 2 3

3.6 2.1 2.4 1.9

± ± ± ±

0.1 0.1 0.1 0.2

3.1 1.7 1.0* 0.7*

± ± ± ±

0.3 0.1 0.0 0.1

34 26 22 11

± ± ± ±

3 3 3 1

34 24 31* 23*

± ± ± ±

2 5 1 2

87 66 44 47

± ± ± ±

13 19 3 8

94 70 50 56

± ± ± ±

4 14 13 9

Values are mean ± S.D. of data obtained from triplicate samples. *Differences significant in comparison with respective control at ≤0.01 (Student’s t-test).

Shukla et al. (2003) observed that the root, shoot-leaf lengths and the root, shoot-leaf biomass decreased with the increasing concentrations of cadmium in the nutrient medium. Since with 50 ␮mol/l CdCl2 , a reasonable rate of growth and biomass partitioning into roots and shoots was observed, therefore, this concentration of CdCl2 was selected for studying carbohydrate metabolizing enzymes under cadmium toxicity. Exogenous cadmium did not cause any significant change in total sugars, reducing sugars and sucrose content of roots, however starch content was significantly less in the roots of cadmium stressed seedlings at days 5 and 7 (Table 1). In shoots an increasing trend of sucrose concentration in cadmium grown seedlings was observed. In cotyledons of control seedlings, total sugars and sucrose content decreased with progress of seedling growth indicating that sucrose is rapidly transported to growing shoots and roots and is not accumulated in the cotyledons. However, in stressed cotyledons decrease in sucrose content with seedling growth was arrested indicating slowing down of the transport of sucrose from cotyledons to the growing tissues. With progress of seedling growth, reduction in starch content of cotyledons was also slowed down in stressed seedlings which led to an increased starch content in cotyledons of stressed seedlings (Table 1). Treatment with cadmium inhibited rice growth and stimulated carbohydrate accumulation especially in seeds from which seedlings were developing (Moya et al., 1995).

In comparison with control, in cadmium-stressed seedlings activity of ␣-amylase was less in all the tissues (Table 2). It appears that low starch content in the roots of stressed seedlings is due to low rate of its synthesis because of less availability of carbon due to decreased rate of starch hydrolysis in the cotyledons. Metabolism of starch is adversely affected in cotyledons of growing seedlings under stressed conditions (Gupta et al., 1993; Kaur et al., 2000; Chugh and Sawhney, 1996). Displacement of calcium by cadmium in calmodulin, important in cell signaling, has been reported (Rivetta et al., 1997). ␣-Amylase requires calcium ions for its activity (Kaur et al., 2001). Decrease in amylase activity could be due to reduced availability of calcium in presence of cadmium. Acid invertase activity was high in growing tissues like roots and shoots and low in cotyledons (Table 3). Acid invertase activity was more in comparison with alkaline invertase in roots and shoots but in cotyledons activity of alkaline invertase was more (Table 3). In general activities of acid and alkaline invertases were less in stressed seedlings with exception to cotyledons where alkaline invertase was more under stress (Table 3). This could lead to wasteful hydrolysis of sucrose formed from the products of starch hydrolysis in the cotyledons of the stressed seedlings. Under arsenic toxicity, a reduced acid invertase activity in rice seedlings has been reported (Jha and Dubey, 2004).

Table 2 Effect of CdCl2 (50 ␮mol/l) on ␣ and ␤-amylase activity (nmoles of reducing sugars formed min−1 mg−1 protein) during seedling growth Tissue

Days

␣-Amylase Control

␤-Amylase Cd-stressed

Control

Cd-stressed

Root

3 5 7

11.9 ± 2.3 27.8 ± 4.4 37.9 ± 1.6

5.9* ± 0.2 25.5 ± 3.6 28.9 ± 2.3

285 ± 35 514 ± 118 613 ± 62

158* ± 26 403 ± 22 552 ± 23

Shoot

5 7

59.5 ± 6.8 25.5 ± 5.0

14.9* ± 2.2 16.2 ± 1.6

499 ± 53 633 ± 73

415 ± 17 517 ± 36

Cotyledons

1 3 5 7

0.8 2.9 9.4 5.2

± ± ± ±

0.2 0.6 3.2 0.8

0.6 2.3 4.6 4.9

± ± ± ±

0.2 0.2 0.6 1.3

4.0 7.5 13.3 45.2

± ± ± ±

0.4 1.0 2.1 4.0

3.8 6.5 5.5* 11.1*

Values are mean ± S.D. of data obtained from triplicate samples. *Differences significant in comparison with respective control at ≤0.01 (Student’s t-test).

± ± ± ±

0.2 0.2 0.4 1.3

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

171

Table 3 Effect of CdCl2 (50 ␮mol/l) on acid and alkaline invertase activity (nmoles of sucrose hydrolysed min−1 mg−1 protein) during seedling growth Tissue

Days

Acid invertase

Alkaline invertase

Control

Cd-stressed

Control

Cd-stressed

Root

3 5 7

22.2 ± 2.5 41.5 ± 1.6 106 ± 14

22.1 ± 1.0 28.2* ± 1.7 51* ± 10

27.4 ± 2.6 28.0 ± 1.0 33.1 ± 6.1

13.7* ± 1.8 14.9* ± 1.7 23.2 ± 3.0

Shoot

5 7

25.8 ± 3.8 94 ± 16

20.8 ± 3.6 54* ± 6

26.4 ± 1.3 35.5 ± 1.2

12.2* ± 0.9 26.9* ± 0.1

Cotyledons

1 3 5 7

0.36 0.30 0.40 0.76

± ± ± ±

0.02 0.01 0.05 0.07

0.34 0.32 0.38 0.44

± ± ± ±

0.03 0.02 0.05 0.05

1.8 1.3 1.5 1.3

± ± ± ±

0.2 0.1 0.2 0.2

2.1 2.0 3.6 3.5*

± ± ± ±

0.4 0.3 0.9 0.3

Values are mean ± S.D. of data obtained from triplicate samples. *Differences significant in comparison with respective control at ≤0.01 (Student’s t-test).

With progress of seedling growth, cadmium caused reduction in activities of sucrose synthase and sucrose phosphate synthase (SPS) in roots and shoots but in cotyledons it did not cause significant effect on sucrose synthase activity where as SPS activity was 10–15% more in stressed seedlings (Table 4). These results showed that sucrose synthesizing capacity is not affected in cotyledons upon cadmium toxicity but in shoots and roots the sucrose synthesis is down regulated which is expected because of inhibition of sucrose transport from cotyledons to the growing tissues under conditions of stress (Gupta et al., 1993). An enhanced activity of SPS under arsenic toxicity has also been reported in rice seedlings (Jha and Dubey, 2004). For synthesis of sucrose the hydrolytic products of starch in the cotyledons need to be converted to fructose-6-phosphate (F-6-P). Similarly glucose and fructose in roots or shoots are utilized by glycolytic cycle or pentose phosphate pathway or they may be converted back to sucrose. The sucrose in the cotyledons is synthesized from glucose and glucose1-phosphate formed from starch. This sucrose is transported to the growing tissues where it is converted to glucose and fructose by hydrolytic and non-hydrolytic cleavage. The glucose formed from sucrose cleavage is phosphorylated to glucose-6-phosphate (G-6-P). G-6-P could be metabolized either through glycolytic sequence or it could be channeled

into pentose phosphate pathway which is primarily responsible for meeting the NADPH requirement for various biosynthetic reactions. Addition of cadmium increased the activity of glucose-6phosphate dehydrogenase (G6PDH) and 6-phosphogluconate dehydrogenase (6PGDH) in cotyledons (Table 5). However, significance of such an increase is still not clear. The roots of cadmium-stressed seedlings had about 10–15% lower activity of G6PDH and 6PGDH. However, in shoots the activity of these enzymes was down regulated by about 40–50% (Table 5). Both these enzymes determine the operation of pentose phsophate pathway. These data showed a lower rate of NADPH production in roots and shoots of stressed seedlings thus affecting the overall biosynthesizing capacity of these tissues where NADPH is required. Hexokinase activity was about 10–15% more in the cotyledons, roots and shoots of stressed seedlings. Fructokinase activity was also not significantly affected in cadmium-stressed seedlings. These data showed that phosphorylation of hexoses is not affected under cadmium toxicity. Phospho glucoisomerase (PGI) activity was high in all the tissues showing that G-6-P to F-6-P conversion is also not affected under cadmium toxicity. It appears that in spite of low starch hydrolysis in cotyledons of stressed seedlings, the hexoses thus formed are rapidly phos-

Table 4 Effect of CdCl2 (50 ␮mol/l) on sucrose synthase (SS) and sucrose phosphate synthase (SPS) activity (nmoles of product formed min−1 mg−1 protein) during seedling growth Tissue

Days

SPS

SS

Control

Cd-stressed

Control

Cd-stressed

Root

3 5 7

7.1 ± 0.5 11.3 ± 1.8 8.3 ± 0.7

10.2 ± 0.2 6.4* ± 0.1 4.3* ± 0.3

4.3 ± 0.2 4.0 ± 0.1 4.3 ± 0.2

5.7* ± 0.2 2.6* ± 0.2 3.3 ± 0.3

Shoot

5 7

3.4 ± 0.3 2.2 ± 0.3

3.5 ± 0.7 1.6 ± 0.4

2.3 ± 0.3 4.6 ± 0.3

2.8 ± 0.1 2.4* ± 0.1

Cotyledons

1 3 5 7

12.3 25.6 32.4 27.0

± ± ± ±

1.4 0.6 2.6 1.1

15.3 28.4 38.4 33.2

± ± ± ±

1.9 1.2 3.4 1.8

11.5 20.3 29.3 24.7

± ± ± ±

1.0 2.2 2.8 3.1

12.0 26.4 23.4 20.4

Values are mean ± S.D. of data obtained from triplicate samples. *Differences significant in comparison with respective control at ≤0.01 (Student’s t-test).

± ± ± ±

1.1 2.0 3.1 2.1

172

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

Table 5 Effect of CdCl2 (50 ␮mol/l) on activity of enzymes of pentose phosphate pathway and glycolytic cycle at 7 days of growth Tissue

Root

Shoot

Control G6PDH 6PGDH Hexokinase Fructokinase PGM PGI F1,6BP

1.5 5.3 3.8 7.2 2.6 116 18.9

± ± ± ± ± ± ±

Cd-stressed 0.2 0.8 0.2 0.8 0.4 4 1.5

1.2 4.4 4.3 7.6 4.9 114 19.3

± ± ± ± ± ± ±

0.1 0.4 0.2 0.1 0.8 2 1.7

Cotyledons

Control 3.3 17.0 3.0 7.1 9.3 97 19.9

± ± ± ± ± ± ±

Cd-stressed 0.7 0.1 0.2 0.2 1.4 2 0.7

1.7 9.6* 4.0 7.3 13.3 95 18.7

± ± ± ± ± ± ±

0.3 1.5 0.7 0.5 2.3 3 1.0

Control 0.8 13.6 7.7 7.8 48.7 73 19.6

± ± ± ± ± ± ±

Cd-stressed 0.1 1.1 1.7 0.9 1.7 5 1.6

2.8* 16.6 9.1 9.0 50.2 70 19.5

± ± ± ± ± ± ±

0.5 3.0 1.3 0.8 2.1 3 1.1

Enzyme activity is expressed as nmole of product formed min−1 mg−1 protein. Values are mean ± S.D. of data obtained from triplicate samples. *Differences significant in comparison with respective control at ≤0.01 (Student’s t-test).

phorylated. This is also reflected by higher activity of phospho glucose isomerase in cotyledons which rapidly transforms G-6P to F-6-P for its conversion into sucrose-6-phosphate by SPS. PGI activity was high in roots and shoots showing the importance of this enzyme in converting G-6-P into F-6-P. These data indicated that hexoses are preferably utilized for glycolysis than pentose phosphate pathway under cadmium toxicity.

In comparison with PGI, the activity of phospho glucomutase (PGM) was almost negligible in roots and low in shoots (Table 5). However, activity of this enzyme was maximum in cotyledons showing that G-1-P formed by starch phsophorylase in cotyledons is rapidly converted to G-6-P and then to F-6-P for synthesis of sucrose. Based on the results of the present investigation, effect of cadmium on various metabolic

Fig. 2. Effect of CdCl2 on various metabolic processes in pea seedlings. Downregulation and upregulation. AMY, amylases; HXK, hexokinase; PGI, phospho glucoisomerase; PGM, phospho glucomutase; SPS, sucrose-6-phosphate synthase; SUC, sucrose; F, fructose; G, glucose; G-6-P, glucose-6-phosphate; F-6-P, fructose6-phosphate; G-1-P, glucose-1-phosphate; INV, invertase; G6PDH, glucose-6-phosphate dehydrogenase; 6PGDH, 6-phosphogluconate dehydrogenase; PGI, phospho glucoisomerase.

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

steps has been summarized in Fig. 2. The inhibition of enzyme activities by cadmium could either be due to its interaction with sulphydryl groups of cysteine or indirectly disturbing the cation balance at the subcellular level (Van and Clijisters, 1990). The present studies are unable to pinpoint the precise reason for the effects of cadmium on various metabolic events. The observed effects of cadmium could be due to reduced availability of water (Fig. 1). Cadmium may also replace calcium in calmodulins or it may react with sulphydryl groups of enzymes/proteins. The presence of thiol groups has been shown in the active site of yeast hexokinase (Gray et al., 1983) and 6-PGDH (Dallocchio et al., 1983). However, activities of these enzymes are not significantly down regulated in cadmium stressed seedlings. On the contrary, invertase which contains cysteine residue in its catalytic domain (Sturm, 1999) is down regulated in shoots and roots of cadmium stressed seedlings. These results indicated that effect of cadmium is not solely based on its reaction with sulphydryl groups. Genes encoding stress related proteins (heat shock proteins, chitinase, chalcone isomerase), metallothionein, ␥-glutamyl cysteine synthase and glutathione were up regulated after 1 week exposure to cadmium (Rivera-Becerril et al., 2005). However, no reports are available on the effect of cadmium on expression of genes related with carbohydrate metabolism. Acknowledgement Thanks are due to University Grants Commission, New Delhi for funding this work. References Artetxe, U., Garcias-Plazoala, J.I., Hernandez, A., Becerril, J.M., 2002. Low light grown duckweed plants are more protected against the toxicity induced by Zn and Cd. Plant Physiol. Biochem. 40, 859–863. Copeland, L., Vella, J., Hong, A., 1989. Enzymes of carbohydrate metabolism in soybean nodules. Phytochemistry 28, 57–61. Chopra, J., Kaur, N., Gupta, A.K., 1998. Carbohydrate status and sucrose metabolism in mung bean roots and nodules. Phytochemistry 49, 1891–1895. Chugh, L.K., Sawhney, S.K., 1996. Effect of cadmium on germination, amylases and rate of respiration of germinating pea seeds. Environ. Pollut. 92, 1–5. Coruzzi, G.M., Bush, D.R., 2001. Nitrogen and carbon nutrient and metabolite signaling in plants. Plant Physiol. 125, 61–64. Dallocchio, F., Matteuzzi, M., Bellini, T., 1983. Evidence for the proximity of cysteine and a lysine residue in the active site of 6-phosphogluconate dehydrogenase. Ital. J. Biochem. 32, 124–130. Dey, P.M., 1986. Change in the forms of invertases during germination of mungbean seeds. Phytochemistry 25, 51–53. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, R.A., Smith, F., 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28, 350–356. Ehness, R., Ecker, M., Godt, D.E., Roitsch, T.H., 1997. Glucose and stress independently regulate source and sink metabolism and defence mechanisms via signal transduction pathways involving protein phosphorylation. Plant Cell. 9, 1825–1841. Fodor, F., 2002. Physiological responses of vascular plants to heavy metals. In: Prasad, M.N.V., Strzalka, K. (Eds.), Physiology and Biochemistry of Metal Toxicity and Tolerance in Plants. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp. 149–177.

173

Gaspar, T., Penel, C., Castillo, F.J., Greppin, H., 1985. A two step control of basic and acidic peroxidases and its significance for growth and development. Physiol. Plant 64, 418–423. Gray, A.J.W., Barron, N.M., Barnard, E.A., 1983. Location in the yeast hexokinase structure of residues related to the enzyme activity. Biosci. Rep. 3, 963–971. Gupta, A.K., Kaur, N., 2005. Sugar signalling and gene expression in relation to carbohydrate metabolism under abiotic stresses. J. Biosci. 30, 761– 776. Gupta, A.K., Singh, J., Kaur, N., Singh, R., 1993. Effect of polyethylene glycolinduced water stress on uptake, interconversion and transport of sugars in chickpea seedlings. Plant Physiol. Biochem. 31, 743–747. Jha, A.B., Dubey, R.S., 2004. Carbohydrate metabolism in growing rice seedlings under arsenic toxicity. J. Plant Physiol. 161, 867–872. Kaur, N., Kaur, P., Gupta, A.K., 2001. Inter organ comparison of amylases and starch content in mungbean seedlings. Acta Physiol. Plant. 23, 429– 435. Kaur, S., Gupta, A.K., Kaur, N., 2000. Effect of GA3 , kinetin and indole acetic acid on carbohydrate metabolism in chickpea seedlings germinating under water stress. Plant Growth Regul. 30, 61–70. Kaur, S., Gupta, A.K., Kaur, N., 2003. Indole acetic acid mimics the effect of salt stress in relation to enzymes of carbohydrate metabolism in chickpea seedlings. Plant Growth Regul. 39, 91–98. Krupa, Z., Baszynski, T., 1995. Some aspects of heavy metal toxicity towards photosynthetic apparatus-direct and indirect effect on light and dark reactions. Acta Physiol. Plant 17, 177–190. Lopez-Molina, L., Mongrand, S., Chua, N.H., 2001. A post-germination developmental arrest checkpoint is mediated by abscisic acid and requires the AB 15 transcription factor in Arabidopsis. Proc. Natl. Acad. Sci. U.S.A. 98, 4782–4787. Lowry, O.H., Rosebrough, N.J., Farr, A.L., Randall, R.J., 1951. Protein measurement with the folin phenol reagent. J. Biol. Chem. 193, 265–275. Maksymiec, W., Krupa, Z., 2002. Jasmonic acid and heavy metals in Arabidopsis plants- a similar physiological response to both stressors? J. Plant Physiol. 159, 509–515. Miyagi, M., Oku, H., Chinen, I., 1990. Purification and action pattern on soluble starch of ␣-amylase from sugarcane leaves. Agric. Biol. Chem. 54, 849– 855. Mocquot, B., Vangronsveld, J., Clijsters, H., Mench, M., 1996. Copper toxicity in young maize (Zea mays L.) plants: effects on growth, mineral and chlorophyll contents, and enzyme activities. Plant Soil 128, 287–300. Moya, J.L., Ros, R., Picazo, I., 1995. Heavy metal-harmone interactions in rice plants: effects on growth, net photosynthesis and carbohydrate distribution. J. Plant Growth Regul. 14, 61–67. Nelson, N., 1944. A photometric adaptation of Somogyi method of determination of glucose. J. Biol. Chem. 153, 375–380. Perfus-Barbeoch, L., Leonhardt, N., Vavasseur, A., Forestier, C., 2002. Heavy metal toxicity: cadmium permeates through calcium channels and disturbs plant water status. Plant J. 32, 539–548. Rivera-Becerril, F., Metwally, A., Laurent, M.F., Tuinen, D.V., Dietz, K.-J., Gianinazzi, S., Gianinazzi-Pearson, V., 2005. Molecular responses to cadmium in roots of Pisum sativum L. Water Air Soil Pollut. 168, 171– 186. Rivetta, A., Negrini, N., Cocucci, M., 1997. Involvement of Ca2+ -calmodulin in cadmium toxicity during early phase of radish (Raphanus sativus) seed germination. Plant Cell Environ. 20, 600–608. Roitsch, T., Balibrea, H.E., Hofmann, M., Proels, R., Sinha, A.K., 2003. The upregulation of extracellular invertase was suggested to be a common response to various biotic and abiotic stress related stimuli like pathogen infection and salt stress. J. Exp. Bot. 54, 513–524. Seki, M., Narusaki, M., Ishida, J., Nanjo, T., Fujita, M., Oone, Y., Kamiya, A., Nakajima, M., Enju, A., Sakurai, T., Satou, M., Akiyama, K., Taji, T., Shinozaki, K.Y., Carninci, P., Kawai, J., Hayashizaki, K.Y., Shinozaki, K., 2002. Monitoring the expression profiles of 7000 Arabidopsis genes under drought, cold and high salinity stresses using a full length cDNA micro array. Plant J. 31, 279–292. Shukla, V.C., Singh, J., Joshi, P.C., Kakkar, P., 2003. Effect of bioaccumulation of cadmium on biomass productivity, essential trace elements, chlorophyll

174

R. Devi et al. / Environmental and Experimental Botany 61 (2007) 167–174

biosynthesis and macromolecules of wheat seedlings. Biol. Trace Element Res. 92, 257–273. Sturm, A., 1999. Invertases: primary structures, functions and roles in plant development and sucrose partitioning. Plant Physiol. 121, 1– 7. Van, A.F., Clijisters, H., 1990. Effects of metals on enzyme activity in plants. Plant Cell Environ. 13, 195–206.

Van Bruwaene, R., Kichmann, R., Impens, R., 1984. Cadmium contamination in agricultural and zoo technology. Experientia 40, 43–52. Wanner, L.A., Junttila, O., 1999. Cold-induced freezing tolerance in Arabidopsis. Plant Physiol. 120, 391–399. Yuan, X.H., Anderson, I.E., 1987. Changing activity of glucose-6-phosphate dehydrogenase from pea chloroplasts during photosynthetic induction. Plant Physiol. 85, 598–600.