Fructan Metabolism in Cereals: Induction in Leaves and Compartmentation in Protoplasts and Vacuoles

Fructan Metabolism in Cereals: Induction in Leaves and Compartmentation in Protoplasts and Vacuoles W. WAGNER,

F. KELLER and A. WIEMKEN

Swiss Federal Institute of Technology, Department of General Botany, CH-8092 Zurich, Switzerland Received August 20,1983 . Accepted September 1,1983

Summary Leaf blades of Triticum aestivum L. cv Kolibri and Hordeum vulgare L. cv Gerbel were induced to accumulate fructan (polyfructosylsucrose of varying molecular size) in large amounts (~70 % of dwt) by impeding the export of photosynthates. This was achieved by subjecting plants to cold stress (5°C at night) or by continuous illumination of excised leaf blades. Concomitantly with the accumulation of fructan in the leaves the activity of a sucrosesucrose-fructosyltransferase (SST), probably the key-enzyme of fructan anabolism, increased several fold in cell-free extracts. Its pH optimum is 5.7. At 8°C its activity is still half of the activity at 28°C, the temperature optimum. This remarkably anomalous dependence on temperature is interesting with regard to fructan accumulation in grasses during the cold season. Protoplasts obtained from fructan-enriched barley leaves were employed for the isolation of vacuoles. All the fructans (~trisaccharide) as well as the SST activity were found to be associated exclusively with the vacuoles, which therefore appear to play the central role in fructan storage and metabolism.

Key words: Hordeum vulgare, Triticum aestivum, assimilate partitioning, cold stress, fructan, protoplasts, vacuoles.

Introduction Unlike tropical grasses which produce starch as the main non-structural polysaccharide in vegetative organs, grasses from temperate and cool climate zones produce fructan of the phlein-type, a (2 -+ 6)-i1-linked polyfructosylsucrose with a DP of up to 260 (Archbold, 1940; Hegnauer, 1963; Smith, 1973; Kandler and Hopf, 1982; Meier and Reid, 1982). In many perennial forage grasses and cereals fructan may account for more than 30 % of dwt of the leaves, stems, and ears depending on their state of development and on environmental conditions such as light intensity, water supply, temperature, and nutrition (Kiihbauch, 1978; Pollock and Jones, 1979; Labhart et a1., 1983). Numerous studies mainly reported in the agricultural literature suggest that whenever the supply of assimilate by photosynthesis exceeds the immediate demand for growth or grain filling the fixed carbon may be diverted into Abbreviations: DP: degree of polymerization; FFT: fructan-fructan-fructosyltransferase; SST: sucrose-sucrose-fructosyltransferase.

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fructan synthesis. The large fructan reserves thus formed may be remobilized when the demand for assimilate exceeds the actual supply by photosynthesis, for instance during growth in early spring, during regrowth after mowing, and during grain filling. Thus fructan plays a key-role as buffer between supply by photosynthesis and utilization for growth (i.e. crop yield). Despite the early insight into this fundamental homeostatic function of fructan in plants, nothing is known about the enzymology and subcellular compartmentation of fructan metabolism in the economically important grasses such as cereals (Pollock, 1979, 1982 a, 1982 b; Meier and Reid, 1982). Furthermore fructan has received surprisingly little attention in studies on assimilate partitioning. Fructans are presumably stored in the vacuoles (Edelman and Jefford, 1968). As the technique for the isolation of vacuoles from barley mesophyll was available (Martinoia et aI., 1981; Kaiser et aI., 1982) it was tempting to see whether the organization of fructan metabolism could be explored using these leaves.

Materials and Methods Plants: Seeds of barley (Hordeum vulgare L. cv Gerbel) and wheat (Triticum aestivum L. cv Kolibri) were soaked in running tap water for 24 h and sown in 12 cm pots of soil (about 150 seeds per pot) and placed in a growth cabinet. The seedlings were watered daily and illuminated with a photosynthetic photon-flux density of 500-600/Lmol photons m- 2s- l . The day/night temperature regime was 25/10 °C with a daylength of 12 h. RH was 70 %. Conditions for induction 0//ructan synthesis in leafblades: Seedlings grown for 12 d were transferred to a growth cabinet with a temperature regime of 15°C (day)/15 °C-5 °C (midnight) 15°C (night) and a daylength of 10 h. The other growth conditions were as indicated above. Alternatively, fructan synthesis was induced in primary leaf blades excised from 10d old seedlings. The leaf blades, standing in water, were illuminated with a photosynthetic photonflux density of 500-600 /Lmol photons m -2S -I at 22°C. Extraction 0/leaves: Primary or secondary leaf blades (1 g fwt) were immersed in 80 % boiling ethanol (10 ml) in which they were subsequently homogenized by use of a glass tissue grinder. Half of the homogenate was centrifuged at 2000 xg for 15 min and the sediment was used for determination of starch. The other half was used for sugar and fructan determination. It was centrifuged at 2000 xg for 15 min and the supernatant withdrawn. The sediment was resuspended in water (8 ml), incubated at 25°C for 15 min, and centrifuged again as before. The two supernatants (80% ethanol and water extract) were combined and brought to dryness under reduced pressure. Analysis 0/ sugars and /ructans: Sugars and fructans (DP;:!: 3) were separated by silicagel TLC (ready foils F 1500 from Schleicher and Schull, Kassel, GFR). The TLC-foils were developed three times with aceton/water (87/13 or 84/16, v/v). The location of sugars and fructans on the TLC-foils was made visible by use of the ureaHCL spray which stains mainly ketoses (Wise et aI., 1955). For quantification the sugars were extracted from TLC and measured colorimetrically with a fructose specific reagent (Nakamura, 1968). Alternatively, or as a control, the sugars were determined enzymatically (before and after separation by TLC) using the assay kits from Boehringer GmbH, Mannheim, GFR (sucrose, glucose, fructose), by GLC of the trimethylsilyl oxime derivatives (Keller and Wiemken, 1982), or by HPLC (Frehner et aI., 1984). For TLC, GLC, and HPLC of samples from cell fractionation the betaine, when present in amounts disturbing the determination, was removed by passage through a column containing 0.5 ml bed-volume of Dowex 50 H+ 200-400 mesh layered onto 0.5 ml bed-volume of Amberlite MB-3 at 4°C within 5 min (forced through the columns

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by centrifugation, 300 xg, 4 min). Total fructose (free and bound) was measured colorimetrically (Nakamura, 1968). Determination of starch: The pellet obtained from the leaf homogenate after the extraction with 80 % ethanol (see above) was resuspended in 0.5 N NaOH (1 ml per 1 g original fwt of leaves) and incubated at 60°C for 1 h under shaking. Then the suspension was neutralized by adding 0.5 M HCI (1 ml) and 0.5 M acetate (NaOH) buffer, pH 4.8 (2 ml). After centrifugation at 2000 xg for 5 min the supernatant was decanted. Aliquots of it (0.5 ml) were incubated with amyloglucosidase (20 ILl of the crude solution of the enzyme from Aspergillus niger obtained from Boehringer, Mannheim, GFR) at 50°C for 90 min. The amount of glucose released upon incubation was determined using the UV-test from Boehringer, and was assumed to be derived from starch. The procedure was tested by internal standardization. Preparation ofprotoplasts: Primary leaf blades of barley seedlings grown for 12 d were excised from the plants and illuminated for 16 h as described for induction of fructan synthesis. Then the leaves were stripped of their abaxial epidermis and floated with the stripped face downwards on the surface of a medium (2 g fwt of leaves per 10 ml of medium in a petri dish) containing 0.7 M sorbitol, 20 mM citrate (NaOH) buffer, pH 5.5 (Medium-P) supplemented with 2 % cellulase R 10 (supplied by Welding & Co., Hamburg, GFR) and 0.1 % Pectolyase (from Seishin Pharmaceutical Co., Noda, Chiba, Japan). After 2 h at 25°C the protoplasts released were passed through a 200 ILm nylon gauze. The resulting protoplast suspension was mixed with an equal volume of Medium-P supplemented with 50 % Percoll (Pharmacia, Uppsala, Sweden). Onto the resulting suspension two layers of Medium-P, the first supplemented with 20 % Percoli, were loaded in a centrifuge tube (equal volumes of the layers to that of the protoplast suspension). After centrifugation at 350 xg for 5 min the protoplasts were collected from the 20 %/0 % Percoll interphase with a Pasteur pipette. They were diluted with 6 volumes of Medium-P and layered onto an equal volume of Medium-P supplemented with 5 % Percol\. After centrifugation at 150 xg for 5 min a loosely packed sediment of purified protoplasts was obtained. Preparation of vacuoles: The purified protoplasts were lysed by resuspension of the protoplast pellet in a solution containing OAM sorbitol, 5mM EDTA (NaOH), 1mgml- 1 BSA, 25mM Tricine (HCI), pH 8 (about 50 ILl of protoplast sediment per ml). After 1 min under gentle stirring at 25°C most protoplasts were lysed. Then half a volume of 1 M sorbitol, 25 mM Tricine (HCI), pH 8, 90 % Percoll was added quickly. Onto the resulting protoplast lysate (one volume) three layers (one volume each) were loaded, containing: 1) 0.6M sorbitol, 25 mM Tricine (HCI) buffer, pH 8,10% Percoll. 2) 0.6M betaine, same buffer, 2.5% Percoll. 3) 0.6M betaine, same buffer, without Percoll. After centrifugation at 1000 xg for 10 min the vacuoles were recovered from the 2.5 %/0 % Percoll interphase. The suspension was diluted with 10 volumes of 0.6M betaine, Tricine (HCI) buffer, pH 8, and centrifuged at 100 xg for 5 min after which the vacuoles were in the pellet. Enzyme assays: Marker enzymes were assayed by standard methods as described in «Methods of Enzymatic Analysis» (Bergmeyer, 1974): malatedehydrogenase, hexose-6-P-isomerase, catalase. Cytochrome c oxidase was determined as indicated elsewhere (Keller and Wiemken, 1982). The activities of a-mannosidase, acid P-ase, i3-N-acetylglucosaminidase were assayed using the corresponding p-nitrophenyl substrates. The assay mixtures (0.2 ml) contained 0-100 ILl enzyme sample, 0.1 M acetate (NaOH) buffer, pH 4.84, and 1 mgml- 1 of the corresponding p-nitrophenyl substrates. After incubation at 30°C for 10-120 min the reaction was stopped by adding 004 ml 1 M Na 2C0 3• The nitrophenol released was measured by the 400 nm absorption. Extraction and assays offructan-enzymes: Primary leaves were homogenized in 20 mM citrateP buffer, pH 5.5,1 mgml- 1 BSA at 4°C in a glass tissue grinder (1 g fwt/ml). The homogenate was centrifuged at 2000 xg for 10 min and the supernatant was passed through a Bio-Gel P-I0 200-400 mesh column (1 x 6 cm) equilibrated with the same citrate-P buffer. The samples from

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cell fractionation were treated identically with Bio-GeI. For measuring the activities of SST and invertase the assay mixture (0.2 ml) contained enzyme sample (0-0.1 ml), 0.1 M sucrose, 1 mg ml- 1 BSA, 20 mM citrate-P buffer, pH 5.5. After incubation for 1-4 h at 27°C the reaction was stopped by incubation for 5 min in a boiling water bath followed by centrifugation at 12,000 xg for 2 min. Aliquots (5 JLI) of the supernatant were spotted on TLC plates for analyzing sugars as described above. The amount of trisaccharide was taken as a measure for the activity of SST. The amount of fructose formed was used as a measure for invertase.

Results

Induction ojJructan synthesis in wheat and barley leaves A temperature regime with cool nights (5°C) induced a conspicuous accumulation of non-structural carbohydrates in the leaf blades of wheat and barley seedlings (Fig. 1 a, b; 2). Initially, sucrose was the main species formed but later, after about 14 days, the level of sucrose remained constant and rapid synthesis of fructans started (Fig. 2). The trisaccharide appeared first and then, sequentially, higher polymers of fructose (Fig. 1 a, b). This is particularly evident in wheat (Fig. 1 b) whereas in barley certain classes of fructans are synthesized preferentially (Fig. 1 a). A much faster and experimentally more amenable way to induce fructan synthesis was found by excision and subsequent continuous illumination of the leaf blades. Since the export of photosynthate was prevented by the excision of the leaves, enormous quantities of fructans were rapidly synthesized. The same oligosaccharide pattern was observed (Fig. 3) as in the leaf extracts of cold-stressed seedlings (Fig. 1 a). Within a few hours the leaf blades accumulated as much fructan as the leaves of the

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cold-stressed seedlings did within days (Fig. 4). After 72 h fructan accounted for more than 70 % of dwt of the leaves. Starch was quantitatively of little importance. Apparently after about 8 to 16 h the amount, first of sucrose, and then of the trisaccharide reached the threshold levels required for triggering rapid synthesis of higher fructans (Fig. 4). If the leaf blades were put in a solution of EDTA instead of water, sucrose was exuded into the solution and fructan formation was much reduced (Fig. 5). This is in Z. Pjlanzenphysiol. Bd. 112. S. 359-372.1983.

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Fig. 5: Reduction of fructan synthesis in excised, illuminated primary leaf blades of H. vulgare upon EDT A-induced exudation of photosynthates. a: control (extract of leaves standing in water), b: water used in a, c: reference sugars (see Fig. 1), d: extract of leaves standing in 2 mM EDTA (NaOH, pH 6.5), e: solution used in d, containing exudate.

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agreement with the notion that the surplus of photosynthates in the leaves caused by the excision induces the accumulation of fructan. Under similar conditions, wilting flowers of Ipomoea tricolor synthesized starch as an alternative non-structural carbohydrate to fructan, when the export of hexoses was prevented by excision (Wiemken et al., 1976).

Induction and properties of sucrose·sucrose fructosyltransferase (SST) in barley leaves Cell-free extracts of the fructan-enriched barley leaves had a marked activity of SST as shown by the formation of a trisaccharide upon incubating the dialyzed extracts with sucrose (Fig.6). The trisaccharide nature of the oligosaccharide formed was derived from the retention times (GLC) and Rf value (TLC) which are identical with those of the trisaccharide isokestose. Upon acid hydrolysis a glucose to fructose ratio of one to two was obtained. The trisaccharide is therefore most probably kestose or isokestose, a proposed precursor-trisaccharide of the phlein-type fructans (Pollock, 1982 b). Besides this trisaccharide a fructan of higher DP (about 6) is faintly visible on TLC (Fig. 6). Furthermore fructose is formed by the action of invertase present in the leaf extracts. The SST but not the invertase activity increased several fold upon stimulation of the leaves to fructan synthesis (Fig. 7). This excludes the possibility that the trisaccharide is formed by a transferase activity of the invertase (Allen and Bacon, 1956). Moreover, the pH optima of SST (5.7) and invertase (4.8) differ markedly {Fig. 8). The pH optimum of SST suggests that this enzyme may be located in the acid vacuolar compartment. Since grasses accumulate fructans particularly during the cool season (Pollock and Jones, 1979) we decided to test the dependence on temperature of SST activity.

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Indeed, at 8 °C still half of the activity present at the temperature optimum of 28°C was found (Fig. 9). The thermal stability is also quite low: half of the enzyme activity was lost after a 30 min incubation at 38°C (Fig. 10). Dependence on temperature, thermal stability as well as pH optimum are thus clearly distinct for SST and invertase.

Subcellular compartmentation offructan metabolism in barley leaves Since fructans are highly water soluble and may be present in large quantities they have been presumed to be located in vacuoles {Edelman and Jefford, 1968, Meier and Reid, 1982}. This location has, however, never been demonstrated. Therefore we thought it worthwhile to isolate the vacuoles from mesophyll protoplasts obtained from fructan-enriched barley leaves in order to assess the fructan content of the vacuoles directly. As these protoplasts and vacuoles had a much higher specific density than the ones isolated previously from barley leaves {Martinoia et aI., 1981} the isolation procedure had to be modified. Percoll and betaine were used, instead of sorZ. Pjlanzenplrysiol. Rd. 112. S. 359-372.1983.

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50

bitol and sucrose, in order to prevent interference in the assaying of sugars and sugar metabolizing enzymes. Percoll and betaine had been applied successfully in previous studies on the subcellular compartmentation of sugars (Kaiser et aI., 1982; Keller and Wiemken, 1982). The purity of the isolated vacuoles was assessed by phase contrast microscopy and by determining several extravacuolar marker enzymes (Keller and Wiemken, 1982), Table 1: Purity of vacuoles isolated from mesophyll protoplasts of excised, illuminated primary leaves of H. vulgare. Marker enzymes

activity in protoplast lysate (nkat/ml)

Extravacuolar 0.08 catalase malatedehydrogenase (NAD) 28.8 cytochrome c oxidase 0.75 hexose-6-P-isomerase 2.50

%in vacuolar fraction 0.Q1 0.05 0.15 0.12

% recovery %in in gradient vacuoles*) (lysate = 100 %) 77 107 111 95

0.10 0.52 1.57 1.26

Vacuolar a-mannosidase fJ- N-acetylglucosaminidase acid-P-ase

0.05 0.05 1.77

10.7 8.4 7.4

104 115 102

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77

*) calculated assuming that a-mannosidase and fJ-N-acetylglucosaminidase are 100% vacuolar (average).

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Table 2: Subcellular localization of non-structural carbohydrates, SST, and invertase in mesophyll protoplasts of excised, illuminated primary leaves of H. vulgare.

fructose glucose sucrose trisaccharide fructan (DP > 3) SST invertase

activity/amount per ml of protoplast lysate

%in vacuoles*)

29ltg 111ltg 351ltg 84ltg 130 Itg 0.04 nkat 0.06 nkat

107 109 65

87 86

92 81

*) see footnote Table 1. and was found to be satisfactory (Tab. 1). The yield of vacuoles from the protoplasts as determined by vacuolar marker enzymes was about 10 % (Tab. 1). The bulk of the non-structural carbohydrates tested is clearly associated with the vacuoles as are the activities of SST and invertase (Tab. 2). Fructose, glucose, the trisaccharide, and total fructan (DP > 3) are most probably located exclusively in the vacuoles. The values of slightly more than 100 % of glucose and fructose in the vacuoles may be due to some hydrolysis of sucrose and fructans during isolation. The values for the trisaccharide, fructan, SST, and invertase, of slightly less than 100 % may be due to the preferential loss of heavy vacuoles containing more than average amounts of fructan and the two enzymes. Such a loss is quite feasible since light vacuoles are selected by the floatation step during isolation. Sucrose, which was present in large quantities in the protoplasts, is quite distinct in that a lower proportion is found in the vacuoles as compared with the other sugars. Taking into account the vacuolar and extravacuolar volumes of the cells (Kaiser et aI., 1982), it appears that sucrose is the only sugar the concentration of which may be equal to or even higher in the cytosol than in the vacuoles. Such a situation has already been observed in gentian roots (Keller and Wiemken, 1928) and in the tubers of Jerusalem artichokes (Frehner et al., 1984). The preferential loss of sucrose from the vacuoles during isolation can of course not be completely ruled out.

Discussion The results presented above suggest that fructan is rapidly synthesized in large quantities by photosynthesizing leaves of cereals when the demand for photosynthates is restricted. The fructans were found to be deposited in the vacuoles of the mesophyll. Vacuolar fructan, a potential sink and source of photosynthates in the immediate vicinity of the sites of photosynthesis therefore deserves more attention in studies on assimilate partitioning than hitherto. The storage of fructans in vacuoles is an elegant way of overcoming the osmotic constraints of accumulation of low Z. Pf/anzenphysiol. Bd. 112. S. 359-372. 1983.

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molecular weight sugars. As fructans have a much lower osmotic potential than sugars, vacuoles offer an almost unlimited capacity to accomodate excessively produced photosynthates within mesophyll cells. It is evident that deposition of carbon in the form of fructan offers several advantages over deposition as starch in non-dehydrated plant tissue. By filling the vacuoles with the highly water-soluble fructan use is made of the largest compartment of the plant cell. In contrast, a piling up of starch in the chloroplasts may eventually impair photosynthesis. Furthermore the mean molecular size of fructan can be changed easily using only a simple transferase system and thus fructans are also ideally suited for osmoregulation during temperature or water stress. An increase of the osmotic potential by a decrease of the mean molecular size of the fructan stored during the winter may assist forage grasses in the rapid leaf expansion growth in early spring (Pollock and Jones, 1979). Alteration of the osmotic potential may also be used to protect cells against freezing by lowering the freezing point. This role of fructans has been proposed for storage organs such as the tubers of Jerusalem artichokes (Edelman and Jefford, 1968) and for grasses (Pollock et aI., 1979). Even high molecular weight fructans which do not substantially increase the osmotic potential may act as efficient antifreeze agents in a similar manner as the highly polar glycopeptides in cold water fish (DeVries, 1982). Fructan may therefore play other roles besides that of a temporary carbohydrate reserve. The properties of SST are in agreement with the roles of fructans mentioned above. It can rapidly produce sufficient trisaccharide even at a low temperature when phloem transport and growth are restricted but photosynthesis is still functioning. Indeed, autumn and winter are the main seasons for fructan accumulation in forage grasses (Pollock and Jones, 1979). As the first step of fructan synthesis catalyzed by SST occurs in the vacuoles, the next step is likely to be located in the same compartment. However, a demonstration of this must await elucidation of the biochemistry of fructan biosynthesis in grasses. Up to now it is not known how many enzymes are involved in this process, and SST may also have FFT activity leading from the trisaccharide to higher polymers (Pollock, 1979). However, if the situation is analogous to that found in the tubers of Helianthus tuberosus, two enzymes, SST and FFT, exist (Edelman and Jefford, 1968) and both are located in the vacuoles (Frehner et aI., 1984). Acknowledgements We thank Ph. Matile for his valuable suggestions and S. Turler and D. Furrer for their aid in preparing the manuscript. The work was supported by the Swiss National Science Foundation.

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BERGMEYER, H. U. (Ed.): Methods of Enzymatic Analysis, 2nd English ed. Academic Press New York, pp. 485, 501, 675 (1974). DEVRIES, A. L.: Biological antifreeze agents in coldwater fishes. Compo Biochem. Physiol. 73A, 627-640 (1982). EDELMAN, J. and T. G. JEFFORD: The mechanism of fructosan metabolism in higher plants as exemplified in Helianthus tuberosus. New Phytol. 67, 517-531 (1968). FREHNER, M., F. KELLER, A. WIEMKEN, and PH. MATILE: Fructan metabolism in Helianthus tube· rosus: compartmentation in protoplasts and vacuoles isolated from tubers. Z. Pflanzenphysiol. in preparation (1984). HEGNAUER, R.: Chemotaxonomie der Pflanzen, Band II, 197-224. Birkhauser Verlag, Basel (1963). KAISER, G., E. MARTINOIA, and A. WIEMKEN: Rapid appearance of photosynthetic products in the vacuoles isolated from barley mesophyll protoplasts by a new fast method. Z. Pflanzenphysiol.l07, 103-113 (1982). KANDLER, O. and H. HOPF: Oligosaccharides based on sucrose (sucrosyl oligosaccharides). In: Encyclopedia of Plant Physiology, F. A. LOEWUS and W. TANNER (Eds.), New Series, Vol. 13 A, 348-383. Springer-Verlag, Berlin, Heidelberg, 1982. KELLER, F. and A. WIEMKEN: Differential compartmentation of sucrose and gentianose in the cytosol and vacuoles of storage root protoplasts from Gentiana lutea L. Plant Cell Reports 1,274-277 (1982). KUEHBAUCH, W.: Die Nichtstrukturkohlenhydrate in Grasern des gemaBigten Klimabereiches, ihre Variationsmoglichkeiten und mikrobielle Verwertung. Landwirtsch. Forschung 31, 251-268 (1978). LABHART, CH., J. NOESBERGER, and C. J. NELSON: Photosynthesis and degree of polymerization of fructan during reproductive growth of meadow fescue at two temperatures and two photon flux densities. J. Expt. Bot. 34, 1037-1046 (1983). MARTINOIA, E., U. HECK, and A. WIEMKEN: Vacuoles as storage compartments for nitrate in barley leaves. Nature 289, 292-294 (1981). MEIER, H. and J. S. G. REID: Reserve polysaccharides other than starch in higher plants. In: Encyclopedia of Plant Physiology, F. A. LOEWUS and W. TANNER (Eds.), New Series, Vol. 13 A. 418-471. Springer-Verlag, Berlin, Heidelberg, 1982. NAKAMURA, M.: Determination of fructose in the presence of large excess of glucose. Agr. BioI. Chern. 32,701-706 (1968). POLLOCK, C. J.: Pathway of fructosan synthesis in leaf bases of Dactylis glomerata. Phytochemistry 18, 777-779 (1979). - Patterns of turnover of fructans in leaves of Dactylis glomerata L. New Phytol. 90, 645-650 (1982 a). - Oligosaccharide intermediates of fructan synthesis in Lolium temulentum. Phytochemistry 21,2461-2465 (1982 b). POLLOCK, C. J. and T. JONES: Seasonal patterns of fructan metabolism in forage grasses. New Phytol. 83, 9-15 (1979). POLLOCK, C. J., G. J. P. RILEY, J. L. STODDART, and H .THOMAS: The biochemical basis of plant response to temperature limitations. Welsh Plant Breeding Station, annual report, pp. 227-246 (1979). SMITH, D.: The non-structural carbohydrates. In: Chemistry and Biochemistry of Herbage, G. W. BUTLER and R. W. BAILEY (Eds.). pp. 106-151. Academic Press, London and New York, 1973. TEORELL, T. and E. STENHAGEN: Ein Universalpuffer fur den pH-Bereich 2.0-12.0. Biochem. Z. 299, 416-419 (1938).

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