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M. H. Dixon, S. A. Hill, M. B. Jackson, R. G. Ratcliffe, L. J. Sweetlove, Physiological and Metabolic Adaptations of Potamogeton pectinatus L. Tubers Support Rapid Elongation of Stem Tissue in the Absence of Oxygen, Plant and Cell Physiology, Volume 47, Issue 1, January 2006, Pages 128–140, https://doi.org/10.1093/pcp/pci229
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Abstract
Tubers of Potamogeton pectinatus L., an aquatic pondweed, over-winter in the anoxic sediments of rivers, lakes and marshes. Growth of the pre-formed shoot that emerges from the tuber is remarkably tolerant to anoxia, with elongation of the stem occurring faster when oxygen is absent. This response, which allows the shoot to reach oxygenated waters, occurs despite a 69–81% reduction in the rate of ATP production, and it is underpinned by several physiological and metabolic adaptations that contribute to efficient energy usage. First, extension of the pre-formed shoot is the result of cell expansion, without the accumulation of new cellular material. Secondly, after over-wintering, the tuber and pre-formed shoot have the enzymes necessary for a rapid fermentative response at the onset of growth under anoxia. Thirdly, the incorporation of [35S]methionine into protein is greatly reduced under anoxia. The majority of the anoxically synthesized proteins differ from those in aerobically grown tissue, implying an extensive redirection of protein synthesis under anoxia. Finally, anoxia-induced cytoplasmic acidosis is prevented to an unprecedented degree. The adaptations of this anoxia-tolerant plant tissue emphasize the importance of the mechanisms that balance ATP production and consumption in the absence of oxygen.
Introduction
Higher plants rely on a continuous supply of oxygen to provide the ATP that is required for normal growth and development. However, it is not uncommon for plants to experience periods of oxygen shortage, for example when soil becomes waterlogged due to heavy rainfall or flooding (Drew 1997). Prolonged periods of oxygen deficiency inevitably lead to cell death, and the extent to which plant tissues can tolerate the absence of oxygen varies widely between species and during development. For example, maize roots cannot survive for much longer than 48 h in the absence of oxygen (Subbaiah et al. 2000) and waterlogging has a severe impact on the growth and yield of this and many other crop plants. In contrast, the rhizomes of the semi-aquatic Acorus calamus can survive many months under anoxia (Braendle and Crawford 1987).
Many wetland, marsh and aquatic plants thrive in permanently waterlogged soils. Physiological adaptations allow these plants to avoid the adverse consequences of submergence, and the development of aerenchyma, which enables submerged tissues to avoid a potential oxygen shortage by receiving oxygen transported from the aerial, oxygenated tissues, is commonly observed (Jackson and Armstrong 1999). While this mechanism is effective when at least part of a plant is in contact with oxygen, there are times during the life cycle of aquatic and semi-aquatic plants when this is not the case. For instance, regenerative organs such as the tubers of Potamogeton pectinatus, turions of Potamogeton distinctus and rhizomes of Iris pseudacorus over-winter in the sediments at the bottom of lakes, rivers and marshes. These sediments are known to be anoxic a few millimetres below the surface (Wetzel 1983) and, for these plants to regenerate, the tubers or turions have to support the growth of new tissues for a sufficient period for them to reach oxygenated water. Metabolic adaptations must therefore have evolved to support such growth in the absence of oxygen.
The absence of oxygen inhibits oxidative phosphorylation, and so anoxic tissues are forced to rely on fermentation for the production of ATP. Fermentation only produces a small fraction of the ATP that would be obtainable from oxidative phosphorylation (Brand 1994), and it can also impair pH regulation, leading to a potentially lethal acidification of the cytoplasm (Roberts et al. 1984, Xia and Roberts 1994). It is therefore remarkable that some plant tissues are able not only to maintain cellular functions, but also to support the elongation of new tissue under these extreme conditions. A number of anoxia-tolerant tissues from wetland and aquatic species have been studied, and they demonstrate several common features that appear to be essential for anoxia tolerance. For example, all such tissues contain substantial carbohydrate reserves as well as the enzymatic machinery to mobilize these reserves for fermentation (Rumpho and Kennedy 1981, Ishizawa et al. 1999, Harada et al. 2005). In addition, ethanolic fermentation, which reduces the accumulation of protons that can contribute to cytoplasmic acidosis, is predominant and lactate fermentation is minimal or absent (Monk et al. 1984, Rivoal et al. 1991). These studies have revealed the basic mechanisms of anoxia tolerance, but the particular features that allow growth of new tissue under anoxic conditions remain unknown. However, it seems likely that these features will be species specific, suggesting that detailed investigations of the anoxia-tolerant tissues of individual species, rather than comparative studies of anoxia-tolerant plants with a range of physiological adaptations, could be informative.
Accordingly, the aim of this study was to carry out a detailed biochemical investigation of Potamogeton pectinatus L., a plant with the ability to grow under anoxia. Potamogeton pectinatus is an aquatic monocot that reproduces vegetatively by means of small tubers with a pre-formed shoot. After over-wintering, new plants are initiated by elongation of the pre-formed shoot and remarkably the rate of elongation of the shoot is faster in the absence of oxygen than under aerobic conditions. The physiological response of the pre-formed shoots to anoxia and its hormonal regulation have been examined in detail (Summers and Jackson 1994, Summers and Jackson 1996, Summers et al. 1996, Summers and Jackson 1998) and preliminary consideration has been given to the underlying metabolic adaptation of the plant (Summers et al. 2000). In this study, the metabolic analysis is extended with detailed measurements of respiration, ethanolic fermentation, starch utilization, protein synthesis and cytoplasmic pH regulation in P. pectinatus stem tissues derived from the pre-formed shoot. Plants were grown under aerobic and anoxic conditions, and the results compared.
Results
Growth in the presence and absence of oxygen
Fig. 1A shows the difference in shoot morphology of plants grown from tubers under aerobic and anoxic conditions for 7 d. As expected (Summers and Jackson 1994), the elongation of growing shoots was significantly enhanced by anoxia, with the first and second internodes reaching a combined length of 104 ± 13 mm (SE; n = 12) in the absence of oxygen, and only 26 ± 3 mm (SE; n = 11) under aerobic conditions (Fig. 1B). Shoots grown under anoxic conditions were mainly composed of elongated first and second internode (stem) tissue, with only short, first leaves above the second node. In contrast, shoots grown under aerobic conditions were mainly composed of leaf tissue above the second node, with both first and second leaves visible, and there was very little extension of the internode tissue. Roots emerged at the first and second nodes under aerobic conditions, but no root growth was observed under anoxia (Dixon 2001).
Comparison of the fresh and dry weights of the internodes from shoots grown for 7 d under aerobic and anoxic conditions revealed that the increased length of the anoxically grown stems was associated with a significant increase in fresh weight (Fig. 1C). In contrast, the dry weight of the stems grown under the two conditions was not significantly different (Fig. 1D). This suggests that increased shoot elongation under anoxic conditions was largely driven by the uptake of water.
Respiration and ATP synthesis in stem tissue
Ethanol was the principal fermentation product under anoxia (Summers et al. 2000, Dixon 2001). Anoxic growth increased the levels of malate and alanine in stem tissues, but the amounts produced were small in comparison with ethanol. Moreover, lactate was undetectable using an assay with a detection limit of 10 nmol (g FW)–1. Assays of the fermentative enzymes in tissue extracts expressed as g DW–1 (Table 1) showed that lactate dehydrogenase (LDH) was absent from pre-formed shoot tissue taken directly from cold storage, as well as from stem tissue grown for 7 d under aerobic conditions or anoxia. Pyruvate decarboxylase (PDC) and alcohol dehydrogenase (ADH) were present in all tissues, and while there was no difference in the maximum catalytic activity of PDC in any of the tissues, ADH increased significantly in stem tissue grown under anoxia (Table 1).
Ethanol production occurred under both aerobic (Fig. 2A–C) and anoxic conditions (Fig. 2D–F) in the sealed flask experiment, and detectable quantities of ethanol, typically 1.36 ± 0.14 µmol (g DW)–1, were present in aerobically grown stem tissue at the end of 7 d, even though the incubation medium was continuously sparged with air. Including the ethanol that diffused from the tissue into the medium, the results in Fig. 2 indicate an ethanol production rate of 323 nmol min–1 (g DW)–1 in the aerobic experiment (Fig. 2C; r = 0.99, P < 0.05) and 857 nmol min–1 (g DW)–1 under anoxia (Fig. 2F; r = 0.85, P < 0.05).
The respiration rate of stem tissue grown under aerobic conditions was 807 ± 57 nmol O2 min–1 (g DW)–1. The underlying rates of hexose consumption in these experiments, assuming a negligible contribution to respiration from fatty acid and amino acid oxidation, can be calculated from the stoichiometric relationships between glucose and oxygen during respiration (1 : 6), and glucose and ethanol during fermentation (1 : 2). These rates were 135 nmol hexose min–1 (g DW)–1 under aerobic conditions ignoring ethanol production, 296 nmol hexose min–1 (g DW)–1 under aerobic conditions in the sealed flask experiment, and 429 nmol hexose min–1 (g DW)–1 under anoxia. Assuming that 31 mol of ATP are generated per mol of hexose via oxidative phosphorylation, and 2 mol via ethanolic fermentation, then these hexose consumption rates correspond to maximal rates of ATP production of 4,185 nmol ATP min–1 (g DW)–1 under aerobic conditions ignoring ethanol production, 4,493 nmol ATP min–1 (g DW)–1 under aerobic conditions in the sealed flask experiment, and 857 nmol ATP min–1 (g DW)–1 under anoxia. Thus, the stimulation of growth under anoxia (Fig. 1) occurred even though the availability of ATP was reduced to only 19–20.5% of the supply under aerobic conditions.
The predicted rate of ATP synthesis under anoxia increases if it is assumed that sufficient PPi is available to allow pyrophosphate:fructose-6-phosphate 1-phosphotransferase (PFP), rather than phosphofructokinase (PFK), to catalyse the phosphorylation of fructose 6-phosphate. Both enzymes were present in stem tissue, and growth under anoxia increased the activity of PFP and decreased the activity of PFK (Table 1), although there was still sufficient PFK activity to support the measured flux to ethanol. Using PFP rather than PFK under anoxia would increase the glycolytic yield of ATP to 3 mol per mol of hexose respired, and thus increase the rate of anoxic ATP production to 1,286 nmol ATP min–1 (g DW)–1 or 29–31% of the maximum aerobic value.
Starch breakdown in tubers and stem tissue
The starch content of P. pectinatus tubers and stem tissue declined steadily during growth over 11 d (Fig. 3). The starch content of dormant tubers was 160 ± 30 µmol hexose (g FW)–1 (SE; n = 8), and the rate of degradation during growth, calculated from lines fitted to the data by linear regression, was 8.2 nmol hexose min–1 (g FW)–1 (r = 0.97, P < 0.05) under aerobic conditions and 7.2 nmol hexose min–1 (g FW)–1 (r = 0.95, P < 0.05) under anoxic conditions (Fig. 3A). The starch content of pre-formed shoot tissue was 13.2 ± 1.9 µmol hexose (stem)–1 (SE; n = 5) and the rate of degradation during growth, again calculated by linear regression, was 0.57 nmol hexose min–1 (stem)–1 (r = 0.92, P < 0.05) under aerobic conditions and 0.70 nmol hexose min–1 (stem)–1 (r = 0.97, P < 0.05) under anoxic conditions (Fig. 3B). The tuber, with a typical FW of 0.8 g, stored much more starch than the pre-formed shoot, but the availability of oxygen had little effect on the rate of starch degradation in either tissue.
The pathway of starch degradation was investigated by measuring both the total amylolytic activity and the activity of starch phosphorylase, in tubers and shoot tissue (Tables 1, 2). There were no significant differences in these activities between dormant tuber tissue and tuber tissue from plants grown under aerobic or anoxic conditions for 7 d (Table 2). Moreover, the maximum catalytic activity of starch phosphorylase was found to be similar to that of the total amylolytic activity in all tuber tissue types. In stem tissues, the total amylolytic activity was higher in pre-formed shoots than in stem tissue from P. pectinatus plants grown under aerobic or anoxic conditions for 7 d (Table 1). The activity of starch phosphorylase was not significantly different between pre-formed shoot tissue and stem tissue grown under aerobic conditions for 7 d, but was significantly reduced in tissue grown under anoxic conditions for 7 d (Table 1).
Protein profile of stem tissue
The two-dimensional (2D) gels of the most abundant soluble proteins from the stem tissue of P. pectinatus plants grown for 7 d under aerobic (Fig. 4A) and anoxic (Fig. 4B) conditions were very similar, with 474 and 453 proteins visible on the respective gels. The relative positions and abundances of the proteins were analysed using the sophisticated pattern recognition algorithms in the Z3 software package (Compugen, Tel Aviv, Israel). This procedure provides a powerful tool for matching common spots across 2D gels (Smilansky 2001), and it showed that 27 proteins were only found in aerobically grown tissue, and that eight proteins were only found in plants grown under anoxia. Moreover, under anoxic conditions, 17 proteins were enhanced >5-fold and seven proteins were reduced >5-fold relative to aerobic conditions.
De novo protein synthesis in stem tissue
Potamogeton pectinatus stem tissue from plants grown for 7 d under aerobic conditions incorporated 258 ± 67 kBq (stem section)–1 (SE; n = 4) of [35S]methionine into protein during a 4 h incubation, whereas anaerobically grown tissue under anoxic conditions incorporated only 33 ± 11 kBq (stem section)–1 (SE; n = 4) over the same period. The rate of uptake of [35S]methionine was 5-fold higher under anoxic conditions, presumably reflecting a difference in the morphology or permeability of the anoxically grown tissue, and so the reduced rate of incorporation of [35S]methionine into protein under anoxia must have been caused by a slower rate of protein synthesis.
Autoradiographs of 2D gels loaded with protein containing equal amounts of [35S]methionine showed that the pattern of the most abundant soluble proteins synthesized de novo in shoot tissue grown and incubated under aerobic conditions (Fig. 4C) was very different from the pattern obtained for tissue grown and incubated under anoxia (Fig. 4D). The gels from the aerobic and anoxic shoot tissue revealed 97 and 105 proteins, respectively, but with relatively few spots in common. Thus, apart from a cluster of major spots (shown boxed; Fig. 4C, D) and two other major proteins (shown circled; Fig. 4C, D), no other proteins migrated to a common position, indicating that the majority of the newly synthesized proteins were different under the two conditions.
Response of cytoplasmic pH to oxygen deprivation
Measurements of cytoplasmic pH (pHcyt) were obtained from the chemical shift of the cytoplasmic inorganic phosphate (Pi) signal observed in the in vivo 31P nuclear magnetic resonance (NMR) spectra of P. pectinatus stem tissue (Fig. 5A, B). Aerobically grown tissue responded to the imposition of anoxia with a small fall in pHcyt from 7.41 ± 0.01 (SE; n = 3) to 7.08 ± 0.04 (SE; n = 3) (Fig. 5C). Acidification occurred over the first hour of anoxia, and the pH recovered to its initial value after oxygenation was re-established. There was also a steady decline in the vacuolar pH (pHvac) from 5.73 ± 0.02 (SE; n = 3) to 5.53 ± 0.03 (SE; n = 3) over the 8 h time course, but the reliability of these values was reduced by the presence of unresolved phytate signals under the vacuolar Pi signal. The 31P NMR spectrum of stem tissue grown and recorded under anoxic conditions (Fig. 5B) was similar to the spectrum obtained from aerobically grown tissue under aerobic conditions (Fig. 5A). The mean pHcyt values recorded over a 9 h time course (Fig. 5D) were 7.47 ± 0.02 (SE, n = 3) for the aerobically grown stem tissue under oxygenated conditions and 7.41 ± 0.01 (SE, n = 3) for the stem tissue grown and recorded under anoxic conditions. The relatively high pHvac of the tissue grown and recorded under anoxic conditions, and the gradual acidification of the vacuole over the 9 h time course (Fig. 5D), were shown to be the result of the prior accumulation of an unidentified compound that was released during the time course, since the drift in pHvac was abolished when the same experiment was performed without the continuous replacement of the suspending medium in the NMR tube (Dixon 2001). It should be noted that the release of the unknown compound, thought to be a freely permeable weak base, had no effect on the pHcyt time course (Fig. 5D), making it highly unlikely that the high pHcyt value under anoxia could be attributable to the presence of the compound.
Discussion
The stimulating effect of anoxia on stem elongation in P. pectinatus is a short-term response to oxygen deprivation that appears to support an escape from the anoxic environment surrounding the tubers. This growth response is a remarkable example of anoxia tolerance, and a detailed biochemical investigation of P. pectinatus tubers and shoots under anoxia was undertaken to gain some insight into the metabolic mechanisms that underpin the effect. In the experiments reported here, measurements of respiration and starch breakdown were used to estimate the impact of the complete absence of oxygen on the rate of ATP production, while two energy-consuming processes, protein synthesis and pH regulation, were investigated to provide insight into the adaptation of the plant to the reduced ATP supply.
Production of ATP during anoxia
A quantitative assessment of the impact of anoxia on ATP production can be made by measuring the rate of respiration under aerobic conditions and the rate of fermentation under anoxic conditions. Although these two fluxes are often estimated from the rate of CO2 production, the consumption or production of CO2 by pathways other than respiration and fermentation reduces the reliability of such measurements (Gibbs and Greenway 2003). For example, there was an appreciable flux through the oxidative pentose phosphate pathway in P. pectinatus shoot tissue under anoxia (Dixon 2001). In order to avoid this problem, the respiratory flux under aerobic conditions and the fermentative flux under anoxia were measured from the rates of oxygen consumption and ethanol production, respectively.
To be completely accurate, this approach needs to allow for the appreciable rate of ethanol production under aerobic conditions. It was noted in an earlier study (Summers et al. 2000) that tissue from P. pectinatus plants that had been grown for 7 d in a constantly aerated solution contained measurable amounts of ethanol, and in the experiments reported here the ethanol content of aerobically grown stem tissue was typically 1.36 ± 0.14 µmol (g DW)–1 at the end of 7 d. Since sparging will tend to displace the ethanol produced under aerobic conditions, an attempt was made to estimate aerobic ethanol production in a sealed flask experiment (Fig. 2). It is possible that sealing the flask led to the development of hypoxia, since depletion of the endogenous oxygen level in metabolically active tissues is a significant risk (Berry and Norris 1949, Armstrong and Beckett 1987, Gibbs and Greenway 2003). This possibility cannot be ruled out, but the PDC and ADH levels (Table 1) were sufficient to support the rate of ethanol production observed in the aerobic sealed flask, and there was no lag phase in the production of ethanol (Fig. 2C), which argues against a gradual imposition of anoxia. Fortunately, any uncertainty in the interpretation of this experiment has little bearing on the estimate of ATP production, since the ATP yield from glycolysis is much lower than that from respiration, but it does affect estimates of hexose consumption.
Ignoring aerobic ethanol production, the rate of hexose consumption increased by a factor of 3.18 in P. pectinatus stem tissue under anoxia; while, if the ethanol production observed in the aerobic sealed flask experiment is accepted as genuine, then hexose consumption increased by a factor of 1.45. This result is broadly in line with measurements on a range of tissues (Gibbs and Greenway 2003), but it is lower than the 6-fold increase deduced for P. pectinatus from measurements of CO2 production (Summers et al. 2000). However, the latter measurements were made on unsubmerged P. pectinatus shoots, and uncontrolled physiological changes, for example rapid dehydration due to lack of a waxy cuticle, may have significantly altered the respiratory response in comparison with the submerged plant. Moreover, the aerobic respiration rate was underestimated by ignoring aerobic ethanol production, and the use of CO2 outputs could have led to incorrect glycolytic fluxes because of the potential contribution to the CO2 balance from pathways other than respiration or fermentation. On this basis, the new measurements are a more accurate reflection of the glycolytic capacity of P. pectinatus, and it can be concluded that the stems respond to anoxia with a modest increase.
The hexose consumption rates reported here imply that the rate of ATP synthesis under anoxia is only 19–20.5% of that under aerobic conditions, rising to 29–31% in the unlikely event that the glycolytic flux is facilitated exclusively by PFP rather than PFK under anoxia. A 16-fold increase in the glycolytic flux would be required to compensate fully for the inhibition of oxidative phosphorylation under anoxia, but increases in glycolytic flux of this magnitude have not been found in anoxia-tolerant plant species. For example, Echinochloa phyllopogon showed no major change in glycolytic rate under anoxia (Kennedy et al. 1991), and a maximum increase in rate of only 3- to 4-fold has been observed in other anoxia-tolerant species (Mayne and Kende 1986, Small et al. 1989). Two conclusions can be drawn: first, an increase in the rate of glycolysis is not essential for anoxia tolerance (Kennedy et al. 1991, Kennedy et al. 1992); and, secondly, even when it occurs, a faster glycolytic flux cannot compensate for the lack of oxidative phosphorylation. Some caution is required, because the precise shortfall in ATP is difficult to estimate given the variable contribution of the alternative oxidase to respiration, for example a recent in vivo study provided estimates of between 11 and 49% in five grasses (Millenaar et al. 2001), and the diversion of energy stored in the proton electrochemical gradient into processes other than ATP synthesis, for example mitochondrial transport processes and uncoupling. However, all anoxia-tolerant plants appear to be ATP compromised under anoxic conditions, and their survival presumably depends on avoiding unnecessary ATP consumption.
How does P. pectinatus support an enhanced rate of growth under anoxia despite the reduced availability of ATP?
The allocation of scarce ATP under anoxia is likely to be particularly important in P. pectinatus, since the increased growth rate (Fig. 1) can only be sustained by a supply of ATP for the biosynthesis of macromolecules and the generation of turgor pressure. At 857–1,286 nmol ATP min–1 (g DW)–1, the rate of ATP synthesis under anoxia in P. pectinatus stem tissue corresponds to 3.8–5.7 µmol ATP h–1 (g FW)–1 or 0.54–0.81 µmol ATP h–1 mg–1 protein. As noted elsewhere (Greenway and Gibbs 2003), this rate is relatively high for anaerobic ATP synthesis in an anoxia-tolerant plant tissue, and it is noteworthy that this is achieved without a massive acceleration of glycolysis. Moreover, the detection of ATP signals in the 31P NMR spectrum of the anoxically grown P. pectinatus stems under anoxia (Fig. 5B) comparable in intensity with those observed under aerobic conditions in the spectrum of the aerobically grown tissue (Fig. 5A) provides further evidence that a balance is maintained between ATP supply and demand under anoxia. Similarly, the negligible effect of anoxia on the ATP content and adenylate energy charge of P. distinctus shoots (Ishizawa et al. 1999) testifies to a remarkable ability to sustain the energy status of the tissue under anoxia.
From a physiological standpoint, P. pectinatus has several adaptations that maximize the chance of surviving prolonged anoxia while promoting the elongation of the shoot. First, P. pectinatus over-winters as a tuber with a pre-formed shoot, and both the tuber and shoot are rich in starch, providing ample substrate for prolonged fermentation (Fig. 3). A similar trait has been observed in P. distinctus turions where stem growth is also stimulated under anoxia (Harada and Ishizawa 2003). Secondly, the presence of a pre-formed shoot means that growth can be supported by expansion of existing cells rather than cell division (Fig. 1). The extent to which cell division needs to occur during anaerobic growth in anoxia-tolerant plants is still unclear. Cell counts and measurements of cell length suggested that there could be some cell division in the second internode in P. pectinatus during stem elongation under anoxia (Summers and Jackson 1994), whereas similar measurements on arrowhead (Sagittaria pygmaea) shoots showed that stimulated growth under anoxia was mainly due to cell elongation (Tamura et al. 2001). Cell expansion is a considerably less energetic process than cell division, and this growth mechanism has a clear advantage when ATP is a limited commodity. Furthermore, cell expansion and elongation allow rapid extension of the stem tissue, which can be viewed as a physiological adaptation to escape the anoxic environment. In other words, the survival strategy of P. pectinatus is to prioritize energy expenditure for the rapid elongation of stem tissue in an ‘all or nothing’ bid to reach oxygenated waters.
Metabolically, the presence of ADH and PDC in pre-formed shoots ensures a rapid fermentative response at the onset of anoxia, but it also reduces the ATP requirement for de novo synthesis of new enzymes. Similarly, P. pectinatus maintains high activities of starch-degrading enzymes and is capable of the same rate of starch breakdown under both anoxia and aerobic conditions (Fig. 3). This contrasts with rice, for example, where the induction of amylase activity under anoxia is insufficient to maintain the aerobic rate of starch breakdown (Perata et al. 1993, Perata et al. 1998). Furthermore, unlike rice, P. pectinatus has the capacity to degrade starch via starch phosphorylase, which avoids the need to use ATP for the subsequent phosphorylation of glucose. As starch is the main source of carbohydrate for fermentation, the ability to maintain a high rate of starch breakdown under anoxia, and to do so via an ATP-efficient pathway, may be a key factor in the survival of P. pectinatus under anoxia. In fact, the rate of starch breakdown is unchanged during anoxia (Fig. 3), implying that the increased flux through glycolysis is supported by redirecting carbohydrate from other potential sinks, such as cell wall synthesis.
Despite the use of ATP-saving steps in biochemical pathways and the adoption of a less energy-demanding growth strategy, it is evident that some biosynthetic processes must be down-regulated to save ATP under anoxia (Geigenberger 2003). For example, protein synthesis represents a considerable drain on the energy resources of a cell, and a reduction in the rate of protein synthesis is always observed in tissues under anoxia (Ricard and Pradet 1989, Chang et al. 2000). The rate of protein synthesis in P. pectinatus under anoxia decreased to 13% of that under aerobic conditions, a decrease that is comparable with that observed in maize root tips (10–15%; Chang et al. 2000) and rice coleoptiles (18%; Ricard and Pradet 1989). Importantly, most of the newly synthesized proteins had different isoelectric points and molecular weights from those synthesized under aerobic conditions, indicating that despite a general down-regulation, the synthesis of proteins that are presumably essential for growth and survival under anoxia continued (Fig. 4). A predominantly different set of proteins is also synthesized under anoxia in rice seedlings (Ricard and Pradet 1989) and S. pygmaea shoots (Ishizawa et al. 1999). It is likely that these proteins include not only those involved in fermentative metabolism, but also those involved in signal transduction, regulation of gene expression and novel pathways associated with anoxia tolerance (Klok et al. 2002). The down-regulation and redirection of protein synthesis under anoxia is thus a controlled process that can be presumed to favour survival, and it is likely that the identification of these proteins (Chang et al. 2000) will eventually provide significant insight into the mechanism of anoxia tolerance.
While some processes can be down-regulated to save ATP, the unstinting expenditure of energy on other processes is essential for survival under anoxia. The regulation of pHcyt appears to fall into this category, and reducing the acidosis that accompanies anoxia in less anoxia-tolerant tissues is seen as a crucial factor for surviving oxygen deprivation (Roberts et al. 1984, Xia and Roberts 1994). Aerobically grown P. pectinatus stems responded to anoxia with a smaller acidification of the cytoplasm [Fig. 5C; see also Summers et al. (2000) for observations on plants grown aerobically under unsparged conditions] than the pH 0.5–0.6 observed for non-acclimated maize root tips (Roberts et al. 1984, Fox et al. 1995) or pea internodes (Summers et al. 2000). Moreover, in an unprecedented observation, the steady-state pHcyt of anoxically grown P. pectinatus stem tissue recorded under anoxia was essentially the same as the pH obtained for aerobically grown tissue under aerobic conditions (Fig. 5D). This suggests that energy limitations imposed by anoxia do not affect the ability of P. pectinatus to regulate pHcyt effectively and this almost certainly contributes to the anoxia tolerance of the species. Since the interplay between lactate and ethanol production appears to be a key factor in achieving satisfactory pH regulation under anoxia (Ratcliffe 1999), it is interesting to note, first, that both lactate and LDH were undetectable in P. pectinatus stem tissue, and, secondly, that there was a detectable level of aerobic ethanol production (Fig. 2). In a variety of less anoxia-tolerant tissues, the usual sequence of events is for a period of lactate production to be followed by a switch to ethanol production after the associated fall in pHcyt has activated PDC (Roberts et al. 1984, Fox et al. 1995, Ratcliffe 1999). In contrast, the occurrence of aerobic ethanol production in P. pectinatus implies that there is no need for a H+-producing phase of glycolysis under anoxia, thus improving the regulation of pHcyt and minimizing the acidification of the cytoplasm under both short-term (Fig. 5C) and long-term (Fig. 5D) oxygen deprivation. The maintenance of the ATP level in anoxic P. pectinatus stems (Fig. 5A, B) and P. distinctus shoots (Ishizawa et al. 1999) is likely to be another factor favouring cytoplasmic pH regulation, since: (i) it minimizes the acidifying effect of the net hydrolysis of ATP that occurs during the onset of anoxia; and (ii) it ensures the availability of ATP for the H+ pumping ATPases in the tonoplast and plasma membranes (Ratcliffe 1999, Gout et al. 2001).
Common mechanisms of anoxia tolerance across kingdoms?
Although the growth response of P. pectinatus under anoxia is unusual among plants, the underpinning metabolic events are not. Thus P. pectinatus relies on a switch to ethanolic fermentation, fuelled by a substantial store of carbohydrate at or near to the site of utilization, and mechanisms as yet unknown for balancing the continuing requirement for energy with the reduced supply of ATP. There are differences in metabolic response in comparison with less well adapted plants, for example the remarkably efficient pH regulation and the implied lack of a role for pHcyt in the regulation of PDC activity, but ultimately it is probably the mechanisms for achieving an appropriate balance between ATP supply and consumption, allowing non-essential ATP-consuming processes to be down-regulated in favour of essential processes, that lie at the heart of the adaptive process.
In the animal kingdom, a number of species have been identified with a remarkable tolerance to hypoxic or anoxic conditions (Storey 1996, Jackson 2000), and a comparison with the metabolic response observed in P. pectinatus may be instructive. For example, one of the best characterized anoxia-tolerant animal species is the painted turtle, Chrysemys picta, which can survive anoxic submergence for up to 5 months at 3°C. Extensive studies of the biochemistry and physiology of the painted turtle have identified two key features for its survival (Jackson 2000): regulation of acid–base status; and reducing the demand for ATP to match the reduction in the rate of ATP synthesis. Under aerobic conditions, the demand for ATP is distributed between protein synthesis, which accounts for approximately 38% of available ATP, Na+/K+ pumping (25%), glucose biosynthesis (18%), protein degradation (18%) and urea biosynthesis (1%) (Jackson 1968). Under anoxia, protein synthesis is reduced to <10% of the aerobic rate while glucose and urea biosynthesis cease. The ATP demand for Na+/K+ pumping is also reduced, but less than other processes, with the result that it represents the dominant energy sink under anoxia. There seem to be no equivalent energy budgets for plant tissues susceptible to periods of oxygen deprivation, and this may offer a pointer for future investigations of plant metabolism under anoxia. For example, a clearer appreciation of the energy requirements for ion transport under aerobic and anoxic conditions would be desirable.
The energy status in animal cells is sensed by an AMP-activated protein kinase (AMPK) (Hardie et al. 2003). This protein responds to a fall in the ATP : AMP ratio by up-regulating processes that produce ATP and down-regulating processes that consume ATP. AMPK homologues are found in all eukaryotic cells, and it is presumed that a similar energy management scheme will operate in plants (Sugden et al. 1999, Geigenberger 2003). However, because the ATP : ADP ratio falls in response to low oxygen before any detectable effect on the NADH : NAD+ ratio, it appears that the activity of the putative AMPK system must itself be regulated by an oxygen sensor that operates above the Km of cytochrome oxidase (Geigenberger et al. 2000, Geigenberger 2003). This conclusion assumes that a fall in the ATP : ADP ratio will necessarily reflect changes in the NADH : NAD+ ratio—a link that could be weakened in the presence of alternative oxidase activity—but as yet the identity of the sensor and the role of the AMPK homologues in plants is unknown. It is also unclear whether such a system, which appears to be required to explain the metabolic responses of plant tissues to a declining oxygen concentration, will also be relevant to the prioritization of ATP utilization under anoxia itself.
In summary, we have identified a number of physiological and metabolic adaptations that allow P. pectinatus to execute an ‘anoxia escape’ strategy. This involves the rapid elongation of a pre-formed shoot by cell expansion and it is achieved despite a marked reduction in the ATP supply. Ethanolic fermentation fuelled by extensive starch reserves provides the necessary energy but, due to the lower efficiency of fermentation, a number of ATP-saving by-passes are utilized and there is a down-regulation of protein synthesis to conserve ATP. Despite the restriction in ATP availability, pH homeostasis is maintained to an unprecedented degree. These adaptations are broadly similar to those present in some anoxia-tolerant animals, and they suggest the existence of a common anoxia survival strategy that has evolved across kingdoms.
Materials and Methods
Enzymes and chemicals
Enzymes and substrates were supplied by Roche Molecular Biochemicals (Lewes, UK) or Sigma Chemical Company (St Louis, MO, USA). All other chemicals were from BDH Chemicals Ltd (Poole, UK). Radiochemicals were supplied by Amersham International plc (Amersham, UK).
Plant material
Over-wintering tubers of P. pectinatus (L.) were harvested each year from stands of adult plants growing in the River Evenlode, Oxfordshire, UK. Harvesting was completed during late September and early October. The dormant tubers were cleaned, graded and surface sterilized before being stored for at least 3 months in damp gravel at 2°C in air-tight jars.
Plant growth conditions
Dormant tubers were removed from cold storage and placed either individually or in pairs at the bottom of perspex tubes stoppered at one end with a rubber bung. The tubers were weighted down with 6 g of washed gravel. Plants were grown in the dark at 20°C under aerated or anoxic conditions for 7–10 d. Aerobic conditions were created by submerging tubers in 0.1 mM CaSO4 solution and bubbling the solution continuously with air (flow rate of 25–30 ml min–1). Anoxic conditions were created by submerging tubers in de-oxygenated CaSO4 solution inside an anaerobic work-station (Don Whitely Scientific, Shipley, UK). Oxygen was eliminated in the anaerobic chamber by circulating a gas mixture containing 90% nitrogen, 10% hydrogen (BOC Gases, Murray Hill, NJ, USA) over a palladium catalyst. These growth conditions, described in earlier work on P. pectinatus as aerobic sparged and anaerobic unsparged (Summers and Jackson 1994), elicit the greatest effect on the growth response to anoxia (Summers and Jackson 1994, Dixon 2001).
Enzyme assays
Freshly cut sections of stem and tuber tissue were snap-frozen and ground to a fine powder in liquid nitrogen with a pestle and mortar. The frozen powder was stored at –80°C until required. Aliquots of between 0.2 and 1 g of frozen powder were ground at 4°C in 1–3 ml of an extraction buffer containing 100 mM Tris (pH 8.0), 10 mM EDTA, 10 mM MgCl2, 5 mM dithiothreitol, 0.5% (w/v) bovine serum albumin (BSA), 0.1% (w/v) sodium metabisulfite and 0.1% (w/v) polyvinylpyrrolidone (PVP-40; Sigma). After 5 min, the suspension was centrifuged at 20,000×g for 10 min at 4°C and the supernatant was desalted by passage through a Sephadex G-25 M PD-10 column (Pharmacia LKB, Uppsala, Sweden) equilibrated with extraction medium.
Unless otherwise stated, enzymes were assayed at 25°C in 1 ml of the following optimized reaction mixtures by measuring the change in absorbance at 340 nm: ADH (EC 1.1.1.1), 47 mM Tris (pH 8.9), 1.26 mM NAD+, 100 mM ethanol (Smith and ap Rees 1979a); LDH (EC 1.1.1.27), 45 mM MES (pH 6.5), 0.24 mM NADH, 1 mM MgCl2, 6 mM pyruvate (Smith and ap Rees 1979a); PFK (EC 2.7.1.11), 43 mM MOPS (pH 7.8), 0.24 mM NADH, 2 mM fructose-6-phosphate, 1 mM MgCl2, 0.2 U of adolase, 9 U of triosephosphate isomerase, 3 U of glycerol phosphate dehydrogenase, 0.5 mM ATP (Smith and ap Rees 1979b); PDC (EC 4.1.1.1), 100 mM MES (pH 6.0), 0.24 mM NADH, 6 mM pyruvate, 1 U of ADH (added after recording the background activity of LDH) (Bucher et al. 1994); PFP (EC 2.7.1.90), 100 mM MOPS (pH 7.4), 0.24 mM NADH, 2 mM fructose-6-phosphate, 1 mM MgCl2, 1 U of aldolase, 1 U of triosephosphate isomerase, 3 U of glycerol phosphate dehydrogenase, 2 µM fructose 2,6-bisphosphate, 0.5 mM pyrophosphate (Tobias et al. 1992); starch phosphorylase (EC 2.4.1.1), 100 mM Tris (pH 7.5), 5 mM MgCl2, 0.4 mM NAD+, 0.024 mM glucose 1,6-bisphosphate, 0.0025% (w/v) amylopectin (from potato), 2 U of of phosphoglucomutase, 1.4 U of glucose 6-phosphate dehydrogenase (from Leuconostoc mesenteroides), 4.5 mM Na2HPO4 (Sweetlove et al. 1996). Total amylolytic activity was determined according to Bernfeld (1955). Assays were optimized with respect to pH and substrate concentration, and for each enzyme it was shown that the assay gave a linear response to varying amounts of enzyme.
Respiration measurements
Several freshly cut 5 mm sections of stem tissue were placed in 1 ml of 100 mM glucose, 0.1 mM CaSO4, 10 mM MES (pH 6.5), and oxygen consumption was measured at 25°C using a Clark oxygen electrode (Hansatech, Kings Lynn, UK).
Metabolite assays
Ethanol production was measured over a time course using three replicate sealed flasks for each time point. A single 1 cm section of P. pectinatus stem tissue was placed in 10 ml of 0.3 mM glucose, 100 mM sucrose and 10 mM Tris (pH 7.4) in a sealed 150 ml flask at 25°C. The flasks were swirled every 30 min to keep the medium mixed. Tissue sections and samples of the medium taken at each time point were snap-frozen in liquid N2. Ethanol production under anoxic conditions was assayed every hour for the first 8 h, and after 20 h, whilst ethanol production under aerobic conditions was recorded every hour for 8 h. The volume of the incubation medium was calculated to contain sufficient oxygen to support aerobic respiration for 9 h. Frozen tissue was extracted using trichloroacetic acid according to the method of Mohanty and ap Rees (1992). Tissue extracts and samples of the medium were assayed at 25°C in an assay mixture containing 1 ml of Sigma glycine reagent (pH 9.0) (Sigma product reference 332–9), 0.6 mM NAD+ and 10–50 µl of extract. The ethanol content in the medium before incubation under aerobic and anoxic conditions was determined and the value subtracted from all subsequent medium measurements. Starch was assayed by digestion to glucose (Zrenner et al. 1996) and lactate was assayed according to Maurer and Poppendiek (1974).
Phenol extraction of proteins
Aliquots of frozen P. pectinatus stem tissue powder from plants grown for 7 d under aerobic and anoxic conditions were ground in 1 ml of ice-cold extraction buffer containing 0.5 M Tris (pH 7.5), 10 mM EDTA, 1% (v/v) Triton X-100 and 2% (v/v) β-mercaptoethanol. Extracts were clarified by centrifugation at 20,000×g for 10 min. An aliquot of supernatant containing 400 µg of protein was added to an equal volume of ice-cold 0.5 M Tris (pH 8.0)-saturated phenol and mixed thoroughly. The sample was then centrifuged (20,000×g for 1 min). Eighty percent of the upper aqueous phase was removed and discarded. The lower phenol phase was re-extracted with an equal volume of extraction buffer as before. Five volumes of 1 M ammonium acetate in methanol were added and incubated at –20°C overnight to precipitate protein from the phenol. Precipitated proteins were pelleted by centrifugation at 20,000×g for 5 min, and the supernatant was discarded. The pellet was washed with 1 M ammonium acetate in methanol and then with 80% (v/v) acetone. The pellet was allowed to dry thoroughly before being re-suspended in 330 µl of sample buffer [6 M urea, 2 M thiourea, 2% (w/v) CHAPS, 0.5% (v/v) IPG buffer (3–10NL; Amersham Pharmacia Biotech, Uppsala, Sweden) and 0.002% (w/v) bromophenol blue]. Any remaining insoluble material was pelleted by centrifugation (20,000×g for 5 min).
Separation of proteins using two-dimensional gel electrophoresis
Proteins (400 µg) were separated in the first dimension by isoelectric focusing (IEF) using an immobilized, non-linear pH gradient of 3–10, and in the second dimension by SDS–PAGE as described by Millar et al. (2001). Protein spots were visualized by silver staining (Blum et al. 1987) and the stained gel imaged using a CCD-based imaging system. Protein spots in the image were detected and quantified automatically using Z3 automated 2D gel analysis software (Compugen, Tel Aviv, Israel). Detected spots were filtered statistically to remove false spots and background artefacts. Pairs of gels were matched by automatic image warping to account for local differences in the way the gels had run. Differences in normalized matched spot intensity, determined as total pixel intensity in each spot expressed relative to the total spot intensity of all detected spots, were compared using a cut-off of either 5-fold greater intensity or 5-fold less.
Uptake of [35S]methionine by stem tissue and incorporation into protein
Freshly cut 1 cm stem segments from plants grown for 7 d under aerobic and anoxic conditions were incubated in 5 ml volumes of sterile 0.1 mM CaSO4 containing 1,850 kBq of [35S]methionine (9.25 GBq mmol–1). After 4 h, the suspending medium was removed and the tissue was washed three times with sterile distilled water. The uptake of [35S]methionine was determined by comparing the amount of [35S]methionine in the incubation buffer before and after the incubation. Aliquots (100 µl) of the incubation buffer were added to 4 ml of Optiphase scintillation fluid and the amount of [35S]methionine was determined by scintillation counting. To determine the amount of [35S]methionine incorporated into protein, frozen tissue sections were ground to a fine powder under liquid nitrogen and the proteins were extracted in phenol as described above. A 200 µg aliquot of protein was re-suspended in 400 µl of buffer containing 63 mM MES (pH 6.8), 2% (w/v) SDS, 10% (v/v) glycerol, 10% (v/v) β-mercaptoethanol. After heating to 100°C for 3 min, 200 µl was added to 4 ml of Optiphase scintillation fluid and the amount of 35S was determined by scintillation counting.
Separation of 35S-labelled proteins using two-dimensional IEF-SDS–PAGE
Proteins were extracted in phenol and separated by 2D gel electrophoresis as described above. After the proteins had been separated in the second dimension, the gels were dried between two sheets of cellophane. Once dry, the upper sheet was peeled off and the exposed gel surface laid against X-ray film (Kodak Biomax ML, Sigma Chemical Company, St Louis, MO, USA) for 2 weeks in a light-proof cassette and then developed. The autoradiograph was scanned and the protein pattern analysed using Z3 software (Compugen, Tel Aviv, Israel).
Sample preparation for NMR spectroscopy
Potamogeton pectinatus stem tissue was analysed by 31P NMR spectroscopy using procedures similar to those described by Fox et al. (1995). After 7 d growth under aerobic or anaerobic conditions, 1.5 cm long stem sections were cut into a continuously aerated or nitrogenated medium containing 0.1 mM CaSO4, 10 mM MES (pH 6.0). The stem sections were vacuum infiltrated for 3 min in the same medium to eliminate the intercellular air spaces and to improve the quality of the NMR spectra. All manipulations of tissue grown under anoxic conditions were completed in the anaerobic work-station or under an atmosphere of nitrogen gas. After vacuum infiltration, 10–15 stem sections were transferred to 6 ml of medium in a 10 mm diameter NMR tube where they were maintained under oxygenated or anoxic conditions using an air-lift system operating with a gas flow rate of 100% O2 or N2 at 50 ml min–1 (Fox et al. 1989). In addition, oxygenated or de-oxygenated medium was circulated through the NMR tube at 6 ml min–1 to maintain a fixed external pH. The tissue was allowed to stabilize in the NMR tube for 30 min at 21°C before starting the NMR experiment.
In vivo 31P NMR spectroscopy
31P NMR spectra were recorded at 121.49 MHz on a Bruker CXP 300 NMR spectroscope using a double-tuned 13C/31P 10 mm diameter probehead. 1H-decoupled 31P NMR spectra were accumulated with a 45° pulse angle, a recycle time of 0.5 s and a total acquisition time of 30 or 60 min. Chemical shifts were measured in resolution-enhanced spectra relative to the signal from a capillary containing a 2% (v/v) aqueous solution of the tetraethyl ester of methylene diphosphonic acid, but they are quoted on the scale that puts the resonance from 85% orthophosphoric acid at 0 ppm Cytoplasmic and vacuolar pH values were determined using calibration curves similar to those described by Spickett et al. (1993). The identification of phytate as a contributor to the P. pectinatus spectra was confirmed by spiking a trichloroacetic acid extract with an authentic sample of myo-inositol hexakisphosphate.
Acknowledgments
The authors thank Mr. P. Symington, Manor Farm, Long Hanborough and The Estates office of Blenheim Palace, Woodstock, Oxfordshire for access to stretches of the River Evenlode. We also thank Professor C. J. Leaver (University of Oxford) for access to equipment, and Dr. J. E. Summers (Long Ashton Research Station, University of Bristol) for advice and assistance. M.H.D. thanks the UK Biotechnology and Biological Sciences Research Council and Long Ashton Research Station for the funding of a CASE studentship.
Enzyme | Pre-formed shoot | Aerobic growth | Anoxic growth |
PDC | 1.3 ± 0.1 a | 0.9 ± 0.3 a | 1.5 ± 0.4 a |
ADH | 43.5 ± 0.7 a | 36.7 ± 3.9 a | 60.1 ± 7.5 b |
LDH | ND | ND | ND |
PFK | NM | 2.1 ± 0.1 a | 1.7 ± 0.1 b |
PFP | NM | 7.4 ± 0.5 a | 17.1 ± 0.8 b |
Total amylolytic activity | 5.0 ± 0.2 a | 1.8 ± 0.4 b | 1.9 ± 0.3 b |
Starch phosphorylase | 0.6 ± 0.2 a | 0.6 ± 0.1 a | 0.3 ± 0.0 b |
Enzyme | Pre-formed shoot | Aerobic growth | Anoxic growth |
PDC | 1.3 ± 0.1 a | 0.9 ± 0.3 a | 1.5 ± 0.4 a |
ADH | 43.5 ± 0.7 a | 36.7 ± 3.9 a | 60.1 ± 7.5 b |
LDH | ND | ND | ND |
PFK | NM | 2.1 ± 0.1 a | 1.7 ± 0.1 b |
PFP | NM | 7.4 ± 0.5 a | 17.1 ± 0.8 b |
Total amylolytic activity | 5.0 ± 0.2 a | 1.8 ± 0.4 b | 1.9 ± 0.3 b |
Starch phosphorylase | 0.6 ± 0.2 a | 0.6 ± 0.1 a | 0.3 ± 0.0 b |
Maximum catalytic activities were determined for pre-formed shoots, and for stem tissue from plants that had been grown for 7 d under either aerobic or anoxic conditions. Each value, expressed in µmol min–1 (g DW)–1, is the mean of the measurements from three extracts ±SE. In each row, means with different letters are significantly different (P < 0.05) as determined by ANOVA and t-test (ND, not detected; NM, not measured).
Enzyme | Pre-formed shoot | Aerobic growth | Anoxic growth |
PDC | 1.3 ± 0.1 a | 0.9 ± 0.3 a | 1.5 ± 0.4 a |
ADH | 43.5 ± 0.7 a | 36.7 ± 3.9 a | 60.1 ± 7.5 b |
LDH | ND | ND | ND |
PFK | NM | 2.1 ± 0.1 a | 1.7 ± 0.1 b |
PFP | NM | 7.4 ± 0.5 a | 17.1 ± 0.8 b |
Total amylolytic activity | 5.0 ± 0.2 a | 1.8 ± 0.4 b | 1.9 ± 0.3 b |
Starch phosphorylase | 0.6 ± 0.2 a | 0.6 ± 0.1 a | 0.3 ± 0.0 b |
Enzyme | Pre-formed shoot | Aerobic growth | Anoxic growth |
PDC | 1.3 ± 0.1 a | 0.9 ± 0.3 a | 1.5 ± 0.4 a |
ADH | 43.5 ± 0.7 a | 36.7 ± 3.9 a | 60.1 ± 7.5 b |
LDH | ND | ND | ND |
PFK | NM | 2.1 ± 0.1 a | 1.7 ± 0.1 b |
PFP | NM | 7.4 ± 0.5 a | 17.1 ± 0.8 b |
Total amylolytic activity | 5.0 ± 0.2 a | 1.8 ± 0.4 b | 1.9 ± 0.3 b |
Starch phosphorylase | 0.6 ± 0.2 a | 0.6 ± 0.1 a | 0.3 ± 0.0 b |
Maximum catalytic activities were determined for pre-formed shoots, and for stem tissue from plants that had been grown for 7 d under either aerobic or anoxic conditions. Each value, expressed in µmol min–1 (g DW)–1, is the mean of the measurements from three extracts ±SE. In each row, means with different letters are significantly different (P < 0.05) as determined by ANOVA and t-test (ND, not detected; NM, not measured).
Enzyme | Dormant | Aerobic growth | Anoxic growth |
Total amylolytic activity | 195 ± 67 a | 220 ± 58 a | 155 ± 14 a |
Starch phosphorylase | 175 ± 5 a | 279 ± 57 a | 248 ± 63 a |
Enzyme | Dormant | Aerobic growth | Anoxic growth |
Total amylolytic activity | 195 ± 67 a | 220 ± 58 a | 155 ± 14 a |
Starch phosphorylase | 175 ± 5 a | 279 ± 57 a | 248 ± 63 a |
Maximum catalytic activities were determined for dormant tubers, and for tubers from plants that had been grown for 7 d under either aerobic or anoxic conditions. Each value, expressed in nmol min–1 (g FW)–1, is the mean of the measurements from three extracts ±SE. In each row, means with different letters are significantly different (P < 0.05) as determined by ANOVA and t-test.
Enzyme | Dormant | Aerobic growth | Anoxic growth |
Total amylolytic activity | 195 ± 67 a | 220 ± 58 a | 155 ± 14 a |
Starch phosphorylase | 175 ± 5 a | 279 ± 57 a | 248 ± 63 a |
Enzyme | Dormant | Aerobic growth | Anoxic growth |
Total amylolytic activity | 195 ± 67 a | 220 ± 58 a | 155 ± 14 a |
Starch phosphorylase | 175 ± 5 a | 279 ± 57 a | 248 ± 63 a |
Maximum catalytic activities were determined for dormant tubers, and for tubers from plants that had been grown for 7 d under either aerobic or anoxic conditions. Each value, expressed in nmol min–1 (g FW)–1, is the mean of the measurements from three extracts ±SE. In each row, means with different letters are significantly different (P < 0.05) as determined by ANOVA and t-test.
Abbreviations
- ADH
alcohol dehydrogenase
- AMPK
AMP-activated protein kinase
- IEF
isoelectric focusing
- LDH
lactate dehydrogenase
- NMR
nuclear magnetic resonance
- PDC
pyruvate decarboxylase
- PFK
phosphofructokinase
- PFP
pyrophosphate:fructose-6-phosphate 1-phosphotransferase
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