9 Springer-Verlag 1984
Measurement of the inorganic pyrophosphate in tissues of Pisum sativum L. Jeffrey Edwards, Tom ap Rees, Patricia M. Wilson and Susan Morrell Botany School, University of Cambridge, Downing Street, Cambridge CB2 3EA, UK
Abstract. Purified pyrophosphate: fructose 6-phosphate 1-phosphotransferase (EC 184.108.40.206) was used to measure the inorganic pyrophosphate in unfractionated extracts of tissues of Pisum sativum L. The fructose 1,6-bisphosphate produced by the above enzyme was measured by coupling to N A D H oxidation via aldolase (EC 220.127.116.11), triosephosphate isomerase (EC 18.104.22.168) and glycerol-3phosphate dehydrogenase (EC 22.214.171.124). Amounts of pyrophosphate as low as 1 nmol could be measured. The contents of pyrophosphate in the developing embryo of pea, and in the apical 2 cm of the roots, were appreciable; 9.4 and 8.9 nmol g-1 fresh weight, respectively. The possibility that pyrophosphate acts in vivo as an energy source for pyrophosphate: fructose 6-phosphate 1-phosphotransferase and for UDPglucose pyrophosphorylase (EC 126.96.36.199) is considered. Key words: Phosphate, pyro-(assay)- Pisum (pyrophosphate content) - Pyrophosphate, inorganic Pyrophosphate: fructose 6-phosphate l-phosphotransferase.
Introduction The widespread occurrence of pyrophosphate: fructose 6-phosphate l-phosphotransferase (EC 188.8.131.52) in higher plants (Carnal and Black 1983; Kruger et al. 1983) has led to implications that it mediates glycolysis (Carnal and Black 1983; Preiss 1984). As the reaction catalysed by this enzyme is readily reversible (O'Brien et al. 1975), the mere presence of the enzyme does not demonstrate a glycolytic role; it could be involved in the conversion of triose phosphate to fructose 6-phosphate. The latter is a distinct possibility since most of
the plant tissues in which pyrophosphate:fructose 6-phosphate 1-phosphotransferase has been demonstrated are either photosynthetic or gluconeogenic. For the enzyme to contribute appreciably to glycolysis, the cytosol, where the enzyme is located (Stitt et al. 1982; Kruger et al. 1983), would have to contain a substantial amount of inorganic pyrophosphate. Apart from a number of specialized micro-organisms, there is little direct evidence that inorganic pyrophosphate serves as a source of cellular energy (Wood et al. 1977). Indeed, there is widespread acceptance of the view that, in general, inorganic pyrophosphate is rapidly hydrolysed by inorganic pyrophosphatase, and that this provides the driving force for the appreciable number of cellular biosyntheses catalysed by pyrophosphorylases (Kornberg 1957). As we could find no authenticated estimates of the amounts of pyrophosphate in plant tissues, we have developed a method for its assay, which we now report, together with evidence that two quite different parts of the pea plant, the developing embryo and the root apex, contain appreciable amounts of inorganic pyrophosphate.
Materials and methods Enzymes, substrates and co-factors were from Boehringer, Lewes, UK except that pyrophosphate:fructose 6-phosphate 1-phosphotransferase, fructose 6-phosphate and fructose 2,6-bisphosphate were from Sigma (London) Chemical Co., Poole, Dorset, UK. Embryos were obtained from plants of Pisum sativum L. cv. Greenshaft that had been grown in Fisons (Loughborough, Leics, UK) Levington's compost at 15~ by night and 18~ by day in an 18-h photoperiod in daylight supplemented by light from fluorescent lamps to give a minimum photon flux of 73 gmol m- 2 s- 1 Pods were harvested when the fresh weight of the embryos reached 300-400mg. Roots were obtained from 5 to 7-d-old seedlings (cv. Kelvedon Wonder) grown as described by Smith and ap Rees (t979) except that the time on floats was 48-96 h. Within 5-10 rain of
J. Edwards et a l . Inorganic pyrophosphate in pea tissues harvesting, samples of 2-3 embryos (fresh weight 1 g, prepared by removing the testa), and of the apical 2 cm of 36 roots (fresh weight 0.65 g) were freeze-clamped as described by ap Rees et al. (1977) and dropped into liquid nitrogen. Variations in this procedure are given in the results. Almost all of the liquid nitrogen was allowed to evaporate at room temperature and then 1.5 ml 1.41 M HC10 4 was added to the frozen tissue which was then broken up with a glass rod. The resulting suspension was kept on ice for 90 min and then centrifuged for 2 rain at 25000 g at 2 ~ C. The sediment was rinsed by resuspension and centrifugation in two 0.5-ml portions of water. The initial supernatant and the washings were neutralized with 5 M KzCO 3 and combined. Further centrifugation removed the resulting KCIO 4 to give a final supernatant, the pH of which was adjusted to pH 7-8. The latter was assayed for inorganic pyrophosphate. The basis of the assay was measurement of the fructose 1,6-bisphosphate formed when the extract was added to pure pyrophosphate:fructose 6-phosphate l-phosphotransferase. The assay mixture contained in 1 ml : 50 mM 2-amino-2-(hydroxymethyl)-l,3-propanediol (adjusted to pH 8.0 with glacial acetic acid), 2 m M magnesium acetate, I mM fructose 6-phosphate, 0.15 mM N A D H , 20 gM fructose 2,6-bisphosphate, 0.45 unit fructose-bisphosphate aldolase (EC 184.108.40.206), 1.7 unit glycerol-3-phosphate dehydrogenase (EC 220.127.116.11), 5 units triosephosphate isomerase (EC 18.104.22.168) and up to 0.87 ml tissue extract. The reaction was started by the addition of 0.1 unit pyrophosphate:fructose 6-phosphate 1-phosphotransferase and was allowed to go to completion. Oxidation of N A D H was measured spectrophotometrically at 340 nm. Blank cuvettes contained all the above reagents except that the extract was replaced with water.
Results and discussion
Two moles of NADH should be oxidized per mole of pyrophosphate assayed. Figure I shows that when the assay was applied to standard solutions of inorganic pyrophosphate, the above expectation was met and there was a close and linear relationship between the amounts of pyrophosphate assayed and NADH oxidized. This relationship was found to extend up to concentrations of pyrophosphate at least as high as 40 nmol per assay (data not shown). The lower limit of the assay was 1 nmol per assay. Treatment of the standard solutions of pyrophosphate with inorganic pyrophosphatase (EC 22.214.171.124) from yeast completely abolished NADH oxidation in the assay. The lability of inorganic pyrophosphate in acid solution and the presence of inorganic pyrophosphatase in plant extracts make it difficult to extract pyrophosphate from plants without loss. Our procedure is a compromise between the need to use HC104 to inactivate the pyrophosphatase and the need to keep the extract at pH 7-8. Losses caused by the use of HC104 were checked by subjecting standard solutions of sodium pyrophosphate (10 nmol/assay) to the killing procedure: 91.4_+2% (mean _ S E of five estimates) of the pyrophosphate survived. We also checked whether
E W N
PPi ASSAYED ( nmol ) Fig. l. Relationship between amount of sodium pyrophosphate assayed and N A D H oxidized during measurement of standard solutions of pyrophosphate. Each value is the mean of three estimates
Table 1. Pyrophosphate content of pea tissues. Values are means • SE of estimates from the number of samples shown in parentheses Tissue
PPx content (nmol g-1 FW)
Recovery of added PPI (%)
Developing cotyledons Root apices
9.4+_1.19 (11) 8.9 _+0.47 (6)
90.2_+3.3 (6) 101.8 _+3.2 (5)
there were losses due to failure to inactivate pyrophosphatase in the tissues. To do this, for each test, we prepared duplicate samples of tissue. For the embryos, one sample was freeze-clamped, killed and extracted in the usual way; the other sample was treated similarly except that 10 nmol sodium pyrophosphate was added to the sample just before it was freeze-clamped. The difference between the amounts of pyrophosphate recovered from the duplicate samples is expressed as a percentage of that added to give an estimate of the recovery of the pyrophosphate. The recoveries were satisfactory (Table 1). Repetition of this type of experiment with roots gave less satisfactory recoveries, 76_+4% (mean _+_SE of five estimates). Subsequently, we found that the amount of pyrophosphate that we could detect in samples of freeze-clamped roots was significantly less, 17%, than in samples that were harvested, put straight into liquid nitrogen and then killed with H C 1 0 4. This difference of 17%, probably a consequence of the difficulty of collecting all of the freezeclamped samples into the small volume necessary
for assay, accounts for the relatively low recoveries we obtained with freeze-clamped roots. For the measurements in roots given in Table 1 we omitted the freeze-clamping. Recovery experiments done under these conditions were satisfactory (Table 1). We suggest that all the measurements in Table 1 were made under conditions in which there was little loss of inorganic pyrophosphate. Treatment of the extracts of both tissues with inorganic pyrophosphatase completely abolished their ability to oxidize N A D H in the assay. Our measurements show that both the embryos and the roots contained appreciable amounts of inorganic pyrophosphate. The above conclusion rests on the specificity of yeast inorganic pyrophosphatase and plant pyrophosphate: fructose 6-phosphate l-phosphotransferase for inorganic pyrophosphate. The former has been demonstrated (Butler 1971). The latter enzyme from plants (Anderson and Sabularse 1982; Kombrink et al. 1984) and A m o e b a (Reeves et al. 1974) has been studied in detail. Although a wide range of compounds that includes ATP, ADP, guanosine 5"-triphosphate, cytidine 5'-triphosphate, uridine 5'-triphosphate, inosine 5'-triphosphate, phosphoenolpyruvate and tripolyphosphate, has been tried, none other than inorganic pyrophosphate acted as a phosphoryl donor. It is possible that the pyrophosphate that we measured arose artefactually from some metabolite during the killing and extraction of the tissue. In quantitative terms ATP is perhaps the most likely source of such pyrophosphate. The following is evidence against such a possibility. Duplicate samples of pea embryos (700 mg fresh weight) were prepared, 180 nmol ATP were added to one sample, and both samples were killed, extracted and assayed. Provided the pyrophosphate present as a contaminant in the ATP was taken into account, no difference in the amount of pyrophosphate extracted from the two samples could be detected. The amounts of inorganic pyrophosphate that we have found in pea tissues are appreciable when compared with those reported for known glycolytic intermediates in plant tissues in general. This is particularly true for fructose 1,6-bisphosphate. For example, in pre-thermogenic clubs of A r u m maculatum (ap Rees 1977) and in potato tubers (Dixon and ap Rees 1980), tissues that represent extremes of glycolytic activity, there are 24 and 5 nmol fructose 1,6-bisphosphate per g fresh weight, respectively. We can not determine the concentration of pyrophosphate in the pea tissues because we do not know the volume of the compartments that contain it. If it is assumed that the pyrophosphate
J. Edwards et al. : Inorganic pyrophosphate in pea tissues
in peas is in the cytosol and that the latter is 10% of the tissue volume, then the concentration of pyrophosphate in both tissues would be close to 90 t~M. This value is appreciably greater than the K m for pyrophosphate reported for plant pyrophosphate: fructose 6-phosphate 1-phosphotransferase (Kombrink et al. 1984). The data in Table 1 are powerful evidence against the view that the amount of inorganic pyrophosphate in plant cells is kept vanishingly small by the action of pyrophosphatase, and imply that the latter is either rigidly controlled or compartmented. Our results are consistent with the view that inorganic pyrophosphate acts as an energy source in plant cells and that pyrophosphate: fructose 6-phosphate 1-phosphotransferase can act as a glycolytic enzyme. The pyrophosphate content may also be high enough to allow uridine 5'-diphosphate (UDP) glucose pyrophosphorylase (EC 126.96.36.199) to convert UDPglucose to glucose l-phosphate in vivo. This would permit sucrose to be metabolized to hexose phosphates via sucrose synthetase (EC188.8.131.52) and UDPglucose pyrophosphorylase as both these enzymes are readily reversible (Goodwin and Mercer 1983). Such a route is consistent with the observations of Echeverria and Humphries (1983) and with the relatively high maximum catalytic activities of UDPglucose pyrophosphorylase in plants (ap Rees et al. 1984). J.E. thanks the Science and Engineering Research Council, and S.M. the Potato Marketing Board, for postgraduate research studentships.
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