Molecular and Biochemical Parasitology, 44 ( 1991) 91-96 Elsevier
91
MOLBIO 01435
Inorganic pyrophosphatase of Trichomonas vaginalis Stephen M.J. Searle* and Mikl6s Mtiller The Rockefeller University, New York, NY, U.S.A. (Received 30 March 1990; accepted 27 July 1990)
Trichomonas vaginalis homogenates were found to have an acid inorganic pyrophosphatase activity with a specific activity at pH 4.8 of about 7 nmol min -1 (mg protein) - l . This activity was localized predominantly in hydrotase containing particles, showed structure-bound latency and was tightly membrane-bound. The activity showed no magnesium dependence, a Km of about 2 mM inorganic pyrophosphate, a pH optimum of 5.2 and was inhibited by fluoride at millimolar levels. No evidence was obtained for the existence of a cytosolic magnesium-dependent activity but the existence of a low level of magnesium-independent cytosolic activity cannot be excluded. These observations correlate with the importance of cytosolic inorganic pyrophosphate in the carbohydrate catabolism in T. vaginalis. Key words: Trichomonas vaginalis; Inorganic pyrophosphate; Pyrophosphatase; Lysosome
Introduction Inorganic pyrophosphate (PPi) is generally regarded a waste product of various biosynthetic reactions which needs to be efficiently removed [1]. In agreement with this notion is the presence of high levels of magnesium-dependent inorganic pyrophosphatases (PPase) in the cytosol of almost all cells. There is increasing evidence, however, that PPi can be of functional significance in various cells. A notable case is when PPi replaces ATP as a phosphoryl donor in some key steps of glycolysis [2]. This function of PPi was detected in pioneering studies in the bacterium Propionobacterium shermanii [3] and the protist Entamoeba histolytica [4]. Phosphofructokinase of these organisms utilizes PPi, and not ATP, to convert fructose 6-phosphate to Correspondence address: Mikl6s Miiller, The Rockefeller University, 1230 York Avenue, New York, NY 10021, U.S.A. *Present address: School of Biological Sciences, University of Bath, Claverton Down, Bath BA2 7AY, U.K. Abbreviations: PPase, inorganic pyrophosphatase; PPi, inorganic pyrophosphate.
fructose 1,6-bisphosphate. Recently, several additional anaerobic protists, Trichomonas vaginalis, Tritrichomonas foetus, Isotricha prostoma [5] and Giardia lamblia [6] were found to contain only PPi-linked phosphofructokinase, a characteristic that might be correlated with their anaerobic nature. The metabolic significance of PPi in these organisms implies the need for a cytoplasmic concentration of PPi at metabolite level in support of glycolysis. In view of this consideration, high levels of PPase activity are not expected to be present in the cytoplasm. In seeming agreement, high levels (180/zM) of PPi [7] and no cytosolic alkaline PPase [8] were detected in E. histolytica, the only representative of the above group to have been studied in some detail. The discovery of additional protist species containing PPi-dependent phosphofructokinase prompted us to initiate a study of their PPase. In the present communication we show that, similarly to E. histolytica [8], the flagellate, T. vaginalis, contains only an acid PPase, primarily localized in hydrolase-bearing organelles and lacks a cytoplasmic, magnesium-dependent alkaline PPase.
0166-6851/91/$03.50 © 1991 Elsevier Science Publishers B.V. (Biomedical Division)
92 Materials and Methods
Organism and culture. T. vaginalis strain C-1 (ATCC30001) was grown axenically for 22 h in tryptose-yeast extract-maltose medium [9] supplemented with 10% horse serum.
Cellfractionation. The procedures used have been described earlier [10,11]. All steps of the fractionation procedure were performed at 4°C in the presence of 250 mM sucrose. Cells from 2000 ml culture were harvested by centrifugation, washed twice with sucrose, and the cell pellet homogenized with a Potter homogenizer. The suspension was centrifuged for 10 min at 350 × g in a Sorvall SS34 rotor. The supernatant was removed, the pellet rehomogenized and centrifuged under the same conditions again. The supernatants were combined (cell extract). The pellet was resuspended in sucrose using a Dounce homogenizer (nuclear fraction). The extract was centrifuged at 31 500 × g for 30 min in an SS34 rotor. The supernatant was removed (cytosolic fraction) and the pellet resuspended. This suspension was then centrifuged again at 31 500 × g for 30 min in the SS34 rotor. The supernatant was removed (cytosolic wash) and the pellet resuspended in sucrose using a Dounce homogenizer (particle fraction). In one experiment an intermediate centrifugation step of 5500 × g for 10 rain was included to separate the large particle fraction. The pellet obtained in the subsequent step of 31 500 × g for 30 min represented a small particle fraction. In some experiments the extract was centrifuged at 105 000 × g for 60 min at 0°C in a Beckman Ti50 rotor, to sediment all organellar material. In one experiment the particle fraction was further fractionated by centrifugation on an isopycnic sucrose density gradient in a zonal rotor at 39000 rpm for 40 min at 4°C [12].
Enzyme assays and protein determination. PPase was determined by the release of inorganic phosphate from PPi [8]. The standard assay mixture contained 5 mM tetrasodium pyrophosphate, 7.5 mM acetate buffer, pH 5 and 0.2% (w/v) Triton X-100, incubated at 30°C for 1 h. In certain experiments the concentration of the substrate was varied or inhibitors or putative activators were included in the assay mixture.
Acid phosphatase was assayed primarily with p-nitrophenol phosphate as substrate [10]. The released p-nitrophenol was determined spectrophotometrically at 410 nm. In certain experiments the hydrolysis of glycerol 2-phosphate was also determined by replacing PPi by 20 mM glycerol 2phosphate in the standard PPase assay described above. N-acetyl-/%glucosaminidase was assayed using p-nitrophenol N-acetyl-/3-glucosaminide as substrate [10]. Malate dehydrogenase (decarboxylating) was assayed by monitoring the reduction of NADP in the presence of malate at 340 nm [13]. Protein concentrations were determined by both the manual and automated versions of the method of Lowry et al. [14].
Latency of hydrolases. In these experiments the samples were assayed on the day of fractionation with the use of the standard assay mixtures with 250 mM sucrose included to render them isotonic for the organelles. Free activity was measured in the absence of detergent and total activity in the presence of 0.1% Triton X-100. All assays were incubated for 30 min at 30°C.
Determination of the effect of pH. In the PPase and acid phosphatase assay, the pH of the assay reagent without sample was adjusted to the desired pH. The buffers used were acetate (pH 3.645), PIPES (pH 6.5-7) and Tris (pH 7.5-9), all at 7.5 mM concentrations. Results
Specific activity in homogenates. Total cellular activity of PPase in T. vaginalis, calculated as a sum of activities in the nuclear fraction and extract, was 7.3 _+0.9 nmol min-~ (mg protein)-~ at pH 4.8 (mean _+ SD for 10 experiments) and 0.44 nmol min-1 (mg protein)-1 at pH 8.0. Addition of 1 mM Mg 2+ did not affect these activities.
Subcellular localization. After sedimentation of particles by centrifugation of the homogenates, most of the PPase activity was found in the particulate fraction (Table I). Specific activity in the particulate fraction was 13.9 _+ 2.4 nmol min -1 (mg protein) -1 and in the supernatant 2.5 + 0.1 nmol m i n - i (mg protein)-l (means + SD for 5 experiments). Acid phosphatase, formerly shown
93 TABLE I Sedimentability of T. vaginalis phosphohydrolases and protein after centrifugation of cell extracts at 31 500 × g for 30 min Fraction
Pyrophosphatase
Acid phosphatase
Protein
Particulate Supernatant Wash of particles
80.4 + 4.7 17.9 + 3.8 1.6 + 1.0
88.0 + 5.3 9.5 + 4.6 2.5 + 1.4
38.6 + 5.7 49.8 + 4.9 11.6 _+ 4.1
Mean percentage + SD for 5 experiments. Recoveries for all activities were between 90 and 95%.
to be localized in lysosome-like particles, had a similar distribution, although its activity in the cytosol was slightly but consistently lower. Centrifugation of the cell homogenate at higher speed did not increase the proportion of activity found in the pellet. The results of a four-fraction differential centrifugation (Fig. 1) show the presence of PPase in both particulate fractions, with a relative specific activity about equal in the large and small particle fractions. This distribution was similar to those of the lysosomal marker enzymes [11], acid phosphatase and N-acetyl-fl-glucosaminidase. The hydrogenosomal marker, malate dehydrogenase (decarboxylating) is predominantly localized in the large particle fraction. The distribution of particulate PPase was also studied by isopycnic centrifugation of the par3
ticulate fraction in a sucrose gradient (Fig. 2). PPase activity showed a broad distribution with a modal density of 1.18 g ml -~ . This distribution is identical to that of acid phosphatase and similar to that of N-acetyl-/3-glucosaminidase, confirming the lysosomal localization of PPase. The hydrogenosomes, as shown by the marker malate dehydrogenase (decarboxylating), banded at a higher density.
Characterization of PPase activity in the particulate fraction. Since the PPase activity was localized predominantly in the particulate fraction, its properties were studied in such fractions. The activity had a pH optimum of about 5.2 (Fig. 3). The pH dependency of acid phosphatase activity was somewhat different from that of the PPase suggesting the presence of two distinct enzymes.
INORGANIC MALATEDEHYDROGENASE PYROPHOSPHATASE {DECARBOXYLATING)
2 INORGANICPYROPHOSPHATASE 0 0 3
I ACIDPHOSPHATASE
MALATEDEHYDROGENASE ~ 5 0 (DECARBOXYLATING) 40
2
o_
30
1,1. 0 ~O Z
~u
N.ACE1.YL.!3.GLUCOS" AMINIDASE
AQD I~I~PH~ASE
20
z
3
20t --co,L.L0c 50 100 50 PERCENTOF PROTEIN
20 >"
100
Fig. 1. Distribution of enzymes after differential centrifugation of a Trichomonas vaginalis homogenate. Relative specific activity was plotted against cumulative percentage of protein recovered in each fraction. The fractions are (from left to right): nuclear fraction (350 x g for 4 rain), large particle fraction (5500 × g for 10 min), small particle fraction (31 500 × g for 30 min). The right-hand fraction represents the final supernatant. Recoveries for all activities were between 85 and 110%.
1
--,0o,
0
•
.oT,, ~
1.2
1.3 1.1 EQUILIBRIUMDENSITY
10
1.2
1.3
Fig. 2. Distribution of enzymes after isopycnic centrifugation in a sucrose gradient of a large particle fraction of Trichomonas vaginalis. Enzyme frequency is plotted against density [11]. Arrow designates modal density. Recoveries for all activities were between 80 and 110%.
94 TABLE II Latency of hydrolases in T. vaginalis homogenates and particulate fractions
_•IOO
~
4o
ul
20
0
4
5
6
7
8
9
pH
Fig. 3. Effect of pH on phosphatase activities in a particulate fraction of Trichomonas vaginalis. Inorganic pyrophosphatase in the absence (0) and presence of 1 mM Mg2~- (&); acid phosphatase (0).
Reaction velocities obtained by varying the PPi concentration between 0.5 and 8 mM were plotted against [PPi] as half reciprocal plot and direct linear plot [15,16]. These plots showed a Km of about 2 mM PPi. Added magnesium ions (1 mM) had no effect on the PPase activity of the particulate fraction at any pH value tested. No magnesium effect was noted in experiments in which low-molecularweight components were removed from the fraction by gel filtration (passage through a PD12 G25 Sephadex desalting gel column). The activity in the particulate fraction was inhibited by fluoride (Fig. 4), as usual for PPases [17-19], but not by cyanide (5 mM), cysteine (5 mM), tartrate (10 mM) or EDTA (5 mM). Fluoride inhibition was studied with tetrasodium pyrophosphate, p-nitrophenol phosphate and glycerol 2-phosphate as substrates (Fig 4). About 120 100
0 ~ 0
60
~
2o 0 0.001
0.01
0.1
1
lO
100
Enzyme
Cell extract
Particle fraction
Acid phosphatase Pyrophosphatase N-acetyl-J3-glucosaminidase
30 43 42
37 + 7 42 + 3 48 + 4
Free activity as percent of total activity given. Mean values + SD for 2 experiments with particle fraction.
1 mM F - gave a 50% inhibition of PPi hydrolysis. Acid phosphatase measured with either p-nitrophenol phosphate or glycerol 2-phosphate was inhibited to the same extent at 100 #M concentration. This again suggests that the PPi and the substrates of acid phosphatase are being hydrolyzed by distinct enzymes. The activity in tile particulate fraction showed some latency using the standard assay conditions described, but the long incubation at relatively high temperature meant that it was difficult to detect. When the assay time was reduced to 30 min the activity showed structure-bound latency (Table II). The significant free activity noted was probably due to damaged particles, as the other enzymes assayed, which are known to be localized inside particles, showed similar free activities. After exposure of particles to conditions which are known to break open organelles (addition to the particle fraction of 0.1% Triton X- 100 or sonication in the presence or absence of 100 mM KC1) and centrifugation at high speed, all PPase activity was recovered in the sedimentable fraction. In contrast, between 25 and 55% of the proteins became non-sedimentable due to various combinations of these factors. This suggests that the PPase is tightly bound to the membrane of an organelle.
Activity in the nonsedimentable fraction. The low activity in the cytosolic fraction made the determination of its pH optimum impossible but its activity was much lower at pH 8 than at pH 4.8. The activity in the cytosolic fraction showed no magnesium dependence at either pH 4.8 or 8.
I F ' - ] (raM)
Fig. 4. The effect of F - on phosphatase activities in a particulate fraction of Trichomonas vaginalis. Inorganic pyrophosphatase (0); acid phosphatase assayed with p-nitrophenyl phosphate (O) and glycerol 2-phosphate (A) as substrate.
Discussion
The observed overall activity of PPase in T.
95
vaginalis is relatively low. Under optimum conditions T. vaginalis homogenates exhibit a specific activity of about 7 nmol min -1 (mg protein)This can be compared to levels of 130 nmol m i n - l (mg protein) -1 in mammalian liver [20] and 190 nmol min -~ (mg protein) -1 in E. histolytica [8]. Still higher specific activities are observed in certain prokaryotes, 1800 nmol min -~ (mg protein) - l in Streptococcus faecium [18], 1200 nmol m i n - l (mg protein)-I in Escherichia coli [18] and 750 nmol min - l (rag protein) -1 in Ureaplasma urealyticum [21 ]. The results also show that the major PPase activity in this organism is a magnesium independent enzyme with acid pH optimum and a relatively low (millimolar) affinity for PPi. This activity is sedimentable, exhibits considerable structure-bound latency and is bound to membranes. Subcellular fractionation showed that this activity is localized in lysosome-like organelles, also containing acid phosphatase and N-acetyl-/3glucosaminidase [ 11 ]. This distribution is different from that of cytosolic magnesium-dependent glycerol 3-phosphatase of T. vaginalis [22]. The PPase residing inside a membrane-bounded organelle is likely to be isolated from the cytosolic compartment, where glycolysis occurs. Its participation in the regulation of cytosolic PPi levels is questionable. Inorganic PPase of E. histolytica has similar properties and localization [8]. Although, as discussed below, the dominant PPase of most eukaryotes is an enzyme of significantly different properties, PPases with acid pH optimum, and in several cases localized in lysosomes, have been observed in different animal [23,24] and plant [25,26] cells. The specific activity of the acid enzymes is usually lower than that of the cytosolic alkaline one. These enzymes were little studied and their role in cellular economy remains to be elucidated, The latter conclusion also pertains to the enzymes detected in T. vaginalis and E. histolytica. The dominant PPase of most eukaryotic cells is the extensively studied, magnesium-dependent enzyme with an alkaline pH optimum [27]. This enzyme, localized in the cytosol, is assumed to play a leading role in the removal of PPi generated in the biosynthetic processes. In addition, magnesium-dependent neutral PPases have been
detected in intracellular membranes of higher plants [28-30] which were implicated in transmembrane proton translocation [28,31]. We were unable to detect the presence of either type of activity in T. vaginalis. It is of interest to compare the level of PPase observed in the cytoplasm with estimates of the rates of utilization and production of PPi by T. vaginalis. Utilization of endogenous carbohydrate proceeds at a rate of 10 to 20 nmol min -~ (mg protein)-1 and of exogenous glucose at a rate of 50 to 100 nmol min -~ (mg protein) - t [32]. Since at least one mole of PPi is used in the catabolism of one mole glucose [5], the rate of carbohydrate utilization corresponds to the minimal required rate of PPi production. The rate of PPi production by biosynthetic reactions can be estimated. The generation time of T. vaginalis at 37°C is about 5 h [33], and its biomass contains about 60 percent protein [34]. Klemme takes the PPi liberation to be 7.4 #mol per mg biomass synthesized in organisms which utilize preformed precursors for macromolecular synthesis [35]. This gives a rate of approximately 50 nmol min -~ (mg protein)-~ PPi production in T. vaginalis. This value is of the same order as the estimated utilization of PPi; thus it is likely that biosynthetic reactions are the major if not the sole source of PPi production. The total cellular PPase activity observed at pH 8.0 is at. least two orders of magnitude lower than the estimated rates of PPi production and utilization. The ratio of PPase activity to PPi production becomes still lower for the cytosol in view of the compartmentation of the PPase, indicating that this enzyme plays hardly any role in regulating cytoplasmic PPi levels. Among the microorganisms compared by Klemme [35] none had a similarly low ratio. The mechanisms whereby PPi levels are regulated remain to be elucidated in this organism and other anaerobic protists.
Acknowledgements This work was supported by US Public Health Service grant AI 11942 and CA 46157. S.M.J.S. was a guest scientist at The Rockefeller University on a summer placement from the University of Bath, Bath, U.K. The competent technical help
96
of Ms. Deborah Mulready is gratefully acknowledged. References 1 Kornberg, A. (1962) On the metabolic significance of phosphorolytic and pyrophosphorolytic reactions. In: Horizons in Biochemistry (Kasha, M. and Pullman, B., eds.), pp. 251-264, Academic Press, New York. 2 Wood, H.G. (1977) Some reactions in which inorganic pyrophosphate replaces ATP and serves as a source of energy. Fed. Proc. 36, 2197-2205. 3 Wood, H.G., O'Brien, W.E. and Michaels, G. (1977) Properties of carboxytransphosphorylase; pyruvate, phosphate dikinase; pyrophosphate-phosphofructokinase and pyrophosphate-acetate kinase and their roles on the metabolism of inorganic pyrophosphate. Adv. Enzymol. 45, 85-155. 4 Reeves, R.E. (1984) Metabolism of Entamoeba histolytica Schaudinn, 1903. Adv. Parasitol. 23, 105-142. 5 Mertens, E., Van Schaftingen, E. and Mtiller, M. (1989) Presence of a fructose-2,6-bisphosphate-insensitive pyrophosphate: fructose-6-phosphate phosphotransferase in the anaerobic protozoa Tritrichomonasfoetus, Trichomonas vaginalis and lsotricha prostoma. Mol. Biochem. Parasitol. 37, 183-190. 6 Mertens, E. (1990) Occurrence of pyrophosphate:fructose6-phosphate 1-phosphotransferase in Giardia lamblia trophozoites. Mol. Biochem. Parasitol. 40, 147-150. 7 Reeves, R.E., South, D.J., Blytt, H.J. and Warren, L.G. (1974) Pyrophosphate:D-fructose 6-phosphate 1phosphotransferase. A new enzyme with the glycolytic function of 6-phosphofructokinase. J. Biol. Chem. 249, 7737-7741. 8 McLaughlin, J., Lindmark, D.G. and MUller, M. (1978) Inorganic pyrophosphatase and nucleoside diphosphatase in the parasitic protozoon, Entamoeba histolytica. Biochem. Biophys. Res. Commun. 82, 913-920. 9 Diamond, L.S. (1957) The establishment of various trichomonads of animals and man in axenic cultures. J. Parasitol. 43, 488~.90. 10 MUller, M. (1973) Biochemical cytology of trichomonad flagellates. I. Subcellular localization of hydrolases, dehydrogenases, and catalase in Tritrichomonas foetus. J. Cell Biol. 57, 453~,74. 11 Lindmark, D.G., MUller, M. and Shio, H. (1975) Hydrogenosomes in Trichomonas vaginalis. J. Parasitol. 61, 552-554. 12 Leighton, F., Poole, B., Beaufay, H., Baudhuin, P., Coffey, J.W., Fowler, S. and de Duve, C. (1968) The large-scale separation of peroxisomes, mitochondria, and lysosomes from the livers of rats injected with Triton WR-1339. J. Cell Biol. 37, 482-513. 13 Steinbiichel, A. and MUller, M. (1986) Anaerobic pyruvate metabolism of Tritrichomonasfoetus and Trichomonas vaginalis hydrogenosomes. Mol. Biochem. Parasitol. 20, 57~65. 14 Lowry, O.H., Rosebrough, N.J., Farr, A.L. and Randall, R.J. (1951) Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265-275. 15 Hanes, C.S. (1932) Studies on plant amylases, I. The effect of starch concentration upon the velocity of hydrolysis by the amylase of germinated barley. Biochem. J. 26, 1406-1421.
16 Eisenthal, R. and Cornish-Bowden, A. (1974) The direct linear plot. A new graphical procedure for estimating enzyme kinetic parameters. Biochem. J. 139, 715-720. 17 Krishnan, V.A. and Gnanam, A. (1988) Properties and regulation of Mg 2+-dependent chloroplast inorganic pyrophosphatase from Sorghum vulgare leaves. Arch. Biochem. Biophys. 260, 277-284. 18 Start, P.R. and Oginsky, E.L. (1972) Inorganic pyrophosphatase of Streptococcus faecium F24. Can. J. Microbiol. 18, 183-191. 19 Irie, M., Yabuta, A., Kimura, K., Shindo, Y. and Tomita, K. (1970) Distribution and properties of alkaline pyrophosphatases of rat liver. J. Biochem. 67, 47 58. 20 Dianzani, M.U. (1954) Uncoupling of oxidative phosphorylation in mitochondria from fatty livers. Biochim. Biophys. Acta 14, 514-532. 21 Davis, J.W., Jr., Mose~, I.S., Ndubuka, C. and Ortiz, R. (1987) Inorganic pyrophosphatase activity in cell-free extracts of Ureaplasma urealyticum. J. Gen. Microbiol. 133, 1453-1459. 22 Steinbiichel, A. and Miiller, M. (1986) Glycerol, a metabolic end product of Trichomonas vaginalis and Tritrichomonasfoetus. Mol. Biochem. Parasitol. 20, 45-55. 23 Norberg, B. (1950) Isodynamic pyrophosphatases in rat liver. Acta Chem. Scand. 4, 601~509. 24 Brightwell, R. and Tappel, A.L. (1968) Lysosomal acid pyrophosphatase and acid phosphatase. Arch. Biochem. Biophys. 124, 333-343. 25 Naganna, B., Raman, A., Venugopal, B. and Sripathi, C.E. (1955) Potato pyrophosphatases. Biochem. J. 60, 215-223. 26 Naganna, B., Venugopal, B. and Sripathi, C.E. (1955) Occurrence of alkaline pyrophosphatase in vegetable tissues. Biochem. J. 60, 224-225. 27 Butler, L.G. (1971) Yeast and other inorganic pyrophosphatases. In: The Enzymes, Vol. IV, 3rd Ed. (Boyer, P.D., ed.), pp. 529-541, Academic Press, New York. 28 Weiner, H., Stitt, M. and Heldt, H.W. (1987) Subcellular compartmentation of pyrophosphate and alkaline pyrophosphatase in leaves. Biochim. Biophys. Acta 893, 13-21. 29 Rea, P.A. and Poole, R.J. (1986) Chromatographic resolution of H+-translocating pyrophosphatase from H ÷translocating ATPase of higher plant tonoplast. Plant Physiol. 81, 126-129. 30 Maeshima, M. and Yoshida, S. (1989) Purification and properties of vacuolar membrane proton-translocating inorganic pyrophosphatase from mung bean. J. Biol. Chem. 264, 20068-20073. 31 Chanson, A., Fichmann, J., Spear, D. and Taiz, L. (1985) Pyrophosphate-driven proton transport by microsomal membranes of corn coleoptiles. Plant Physiol. 79, 159-164. 32 Mack, S.R. and MUller, M. (1980) End products of carbohydrate metabolism in Trichomonas vaginalis. Comp. Biochem. Physiol. 67B, 213-216. 33 Mack, S.R. and MUller, M. (1978) Effect of oxygen and carbon dioxide on the growth of Trichomonas vaginalis and Tritrichomonas foetus. J. Parasitol. 64, 927-929. 34 Michaels, R.M. and Treick, R.W. (1962) The mode of action of certain 3- and 5-nitropyridines and pyrimidines, III. Biochemical lesions in Trichomonas vaginalis. Exp. Parasitol. 12, 401417. 35 Klemme, J.-H. (1976) Regulation of intracellular pyrophosphatase-activity and conservation of the phosphoanhydride-energy of inorganic pyrophosphate in microbial metabolism. Z. Naturforsch. C 31, 544-550.
91
MOLBIO 01435
Inorganic pyrophosphatase of Trichomonas vaginalis Stephen M.J. Searle* and Mikl6s Mtiller The Rockefeller University, New York, NY, U.S.A. (Received 30 March 1990; accepted 27 July 1990)
Trichomonas vaginalis homogenates were found to have an acid inorganic pyrophosphatase activity with a specific activity at pH 4.8 of about 7 nmol min -1 (mg protein) - l . This activity was localized predominantly in hydrotase containing particles, showed structure-bound latency and was tightly membrane-bound. The activity showed no magnesium dependence, a Km of about 2 mM inorganic pyrophosphate, a pH optimum of 5.2 and was inhibited by fluoride at millimolar levels. No evidence was obtained for the existence of a cytosolic magnesium-dependent activity but the existence of a low level of magnesium-independent cytosolic activity cannot be excluded. These observations correlate with the importance of cytosolic inorganic pyrophosphate in the carbohydrate catabolism in T. vaginalis. Key words: Trichomonas vaginalis; Inorganic pyrophosphate; Pyrophosphatase; Lysosome
Introduction Inorganic pyrophosphate (PPi) is generally regarded a waste product of various biosynthetic reactions which needs to be efficiently removed [1]. In agreement with this notion is the presence of high levels of magnesium-dependent inorganic pyrophosphatases (PPase) in the cytosol of almost all cells. There is increasing evidence, however, that PPi can be of functional significance in various cells. A notable case is when PPi replaces ATP as a phosphoryl donor in some key steps of glycolysis [2]. This function of PPi was detected in pioneering studies in the bacterium Propionobacterium shermanii [3] and the protist Entamoeba histolytica [4]. Phosphofructokinase of these organisms utilizes PPi, and not ATP, to convert fructose 6-phosphate to Correspondence address: Mikl6s Miiller, The Rockefeller University, 1230 York Avenue, New York, NY 10021, U.S.A. *Present address: School of Biological Sciences, University of Bath, Claverton Down, Bath BA2 7AY, U.K. Abbreviations: PPase, inorganic pyrophosphatase; PPi, inorganic pyrophosphate.
fructose 1,6-bisphosphate. Recently, several additional anaerobic protists, Trichomonas vaginalis, Tritrichomonas foetus, Isotricha prostoma [5] and Giardia lamblia [6] were found to contain only PPi-linked phosphofructokinase, a characteristic that might be correlated with their anaerobic nature. The metabolic significance of PPi in these organisms implies the need for a cytoplasmic concentration of PPi at metabolite level in support of glycolysis. In view of this consideration, high levels of PPase activity are not expected to be present in the cytoplasm. In seeming agreement, high levels (180/zM) of PPi [7] and no cytosolic alkaline PPase [8] were detected in E. histolytica, the only representative of the above group to have been studied in some detail. The discovery of additional protist species containing PPi-dependent phosphofructokinase prompted us to initiate a study of their PPase. In the present communication we show that, similarly to E. histolytica [8], the flagellate, T. vaginalis, contains only an acid PPase, primarily localized in hydrolase-bearing organelles and lacks a cytoplasmic, magnesium-dependent alkaline PPase.
0166-6851/91/$03.50 © 1991 Elsevier Science Publishers B.V. (Biomedical Division)
92 Materials and Methods
Organism and culture. T. vaginalis strain C-1 (ATCC30001) was grown axenically for 22 h in tryptose-yeast extract-maltose medium [9] supplemented with 10% horse serum.
Cellfractionation. The procedures used have been described earlier [10,11]. All steps of the fractionation procedure were performed at 4°C in the presence of 250 mM sucrose. Cells from 2000 ml culture were harvested by centrifugation, washed twice with sucrose, and the cell pellet homogenized with a Potter homogenizer. The suspension was centrifuged for 10 min at 350 × g in a Sorvall SS34 rotor. The supernatant was removed, the pellet rehomogenized and centrifuged under the same conditions again. The supernatants were combined (cell extract). The pellet was resuspended in sucrose using a Dounce homogenizer (nuclear fraction). The extract was centrifuged at 31 500 × g for 30 min in an SS34 rotor. The supernatant was removed (cytosolic fraction) and the pellet resuspended. This suspension was then centrifuged again at 31 500 × g for 30 min in the SS34 rotor. The supernatant was removed (cytosolic wash) and the pellet resuspended in sucrose using a Dounce homogenizer (particle fraction). In one experiment an intermediate centrifugation step of 5500 × g for 10 rain was included to separate the large particle fraction. The pellet obtained in the subsequent step of 31 500 × g for 30 min represented a small particle fraction. In some experiments the extract was centrifuged at 105 000 × g for 60 min at 0°C in a Beckman Ti50 rotor, to sediment all organellar material. In one experiment the particle fraction was further fractionated by centrifugation on an isopycnic sucrose density gradient in a zonal rotor at 39000 rpm for 40 min at 4°C [12].
Enzyme assays and protein determination. PPase was determined by the release of inorganic phosphate from PPi [8]. The standard assay mixture contained 5 mM tetrasodium pyrophosphate, 7.5 mM acetate buffer, pH 5 and 0.2% (w/v) Triton X-100, incubated at 30°C for 1 h. In certain experiments the concentration of the substrate was varied or inhibitors or putative activators were included in the assay mixture.
Acid phosphatase was assayed primarily with p-nitrophenol phosphate as substrate [10]. The released p-nitrophenol was determined spectrophotometrically at 410 nm. In certain experiments the hydrolysis of glycerol 2-phosphate was also determined by replacing PPi by 20 mM glycerol 2phosphate in the standard PPase assay described above. N-acetyl-/%glucosaminidase was assayed using p-nitrophenol N-acetyl-/3-glucosaminide as substrate [10]. Malate dehydrogenase (decarboxylating) was assayed by monitoring the reduction of NADP in the presence of malate at 340 nm [13]. Protein concentrations were determined by both the manual and automated versions of the method of Lowry et al. [14].
Latency of hydrolases. In these experiments the samples were assayed on the day of fractionation with the use of the standard assay mixtures with 250 mM sucrose included to render them isotonic for the organelles. Free activity was measured in the absence of detergent and total activity in the presence of 0.1% Triton X-100. All assays were incubated for 30 min at 30°C.
Determination of the effect of pH. In the PPase and acid phosphatase assay, the pH of the assay reagent without sample was adjusted to the desired pH. The buffers used were acetate (pH 3.645), PIPES (pH 6.5-7) and Tris (pH 7.5-9), all at 7.5 mM concentrations. Results
Specific activity in homogenates. Total cellular activity of PPase in T. vaginalis, calculated as a sum of activities in the nuclear fraction and extract, was 7.3 _+0.9 nmol min-~ (mg protein)-~ at pH 4.8 (mean _+ SD for 10 experiments) and 0.44 nmol min-1 (mg protein)-1 at pH 8.0. Addition of 1 mM Mg 2+ did not affect these activities.
Subcellular localization. After sedimentation of particles by centrifugation of the homogenates, most of the PPase activity was found in the particulate fraction (Table I). Specific activity in the particulate fraction was 13.9 _+ 2.4 nmol min -1 (mg protein) -1 and in the supernatant 2.5 + 0.1 nmol m i n - i (mg protein)-l (means + SD for 5 experiments). Acid phosphatase, formerly shown
93 TABLE I Sedimentability of T. vaginalis phosphohydrolases and protein after centrifugation of cell extracts at 31 500 × g for 30 min Fraction
Pyrophosphatase
Acid phosphatase
Protein
Particulate Supernatant Wash of particles
80.4 + 4.7 17.9 + 3.8 1.6 + 1.0
88.0 + 5.3 9.5 + 4.6 2.5 + 1.4
38.6 + 5.7 49.8 + 4.9 11.6 _+ 4.1
Mean percentage + SD for 5 experiments. Recoveries for all activities were between 90 and 95%.
to be localized in lysosome-like particles, had a similar distribution, although its activity in the cytosol was slightly but consistently lower. Centrifugation of the cell homogenate at higher speed did not increase the proportion of activity found in the pellet. The results of a four-fraction differential centrifugation (Fig. 1) show the presence of PPase in both particulate fractions, with a relative specific activity about equal in the large and small particle fractions. This distribution was similar to those of the lysosomal marker enzymes [11], acid phosphatase and N-acetyl-fl-glucosaminidase. The hydrogenosomal marker, malate dehydrogenase (decarboxylating) is predominantly localized in the large particle fraction. The distribution of particulate PPase was also studied by isopycnic centrifugation of the par3
ticulate fraction in a sucrose gradient (Fig. 2). PPase activity showed a broad distribution with a modal density of 1.18 g ml -~ . This distribution is identical to that of acid phosphatase and similar to that of N-acetyl-/3-glucosaminidase, confirming the lysosomal localization of PPase. The hydrogenosomes, as shown by the marker malate dehydrogenase (decarboxylating), banded at a higher density.
Characterization of PPase activity in the particulate fraction. Since the PPase activity was localized predominantly in the particulate fraction, its properties were studied in such fractions. The activity had a pH optimum of about 5.2 (Fig. 3). The pH dependency of acid phosphatase activity was somewhat different from that of the PPase suggesting the presence of two distinct enzymes.
INORGANIC MALATEDEHYDROGENASE PYROPHOSPHATASE {DECARBOXYLATING)
2 INORGANICPYROPHOSPHATASE 0 0 3
I ACIDPHOSPHATASE
MALATEDEHYDROGENASE ~ 5 0 (DECARBOXYLATING) 40
2
o_
30
1,1. 0 ~O Z
~u
N.ACE1.YL.!3.GLUCOS" AMINIDASE
AQD I~I~PH~ASE
20
z
3
20t --co,L.L0c 50 100 50 PERCENTOF PROTEIN
20 >"
100
Fig. 1. Distribution of enzymes after differential centrifugation of a Trichomonas vaginalis homogenate. Relative specific activity was plotted against cumulative percentage of protein recovered in each fraction. The fractions are (from left to right): nuclear fraction (350 x g for 4 rain), large particle fraction (5500 × g for 10 min), small particle fraction (31 500 × g for 30 min). The right-hand fraction represents the final supernatant. Recoveries for all activities were between 85 and 110%.
1
--,0o,
0
•
.oT,, ~
1.2
1.3 1.1 EQUILIBRIUMDENSITY
10
1.2
1.3
Fig. 2. Distribution of enzymes after isopycnic centrifugation in a sucrose gradient of a large particle fraction of Trichomonas vaginalis. Enzyme frequency is plotted against density [11]. Arrow designates modal density. Recoveries for all activities were between 80 and 110%.
94 TABLE II Latency of hydrolases in T. vaginalis homogenates and particulate fractions
_•IOO
~
4o
ul
20
0
4
5
6
7
8
9
pH
Fig. 3. Effect of pH on phosphatase activities in a particulate fraction of Trichomonas vaginalis. Inorganic pyrophosphatase in the absence (0) and presence of 1 mM Mg2~- (&); acid phosphatase (0).
Reaction velocities obtained by varying the PPi concentration between 0.5 and 8 mM were plotted against [PPi] as half reciprocal plot and direct linear plot [15,16]. These plots showed a Km of about 2 mM PPi. Added magnesium ions (1 mM) had no effect on the PPase activity of the particulate fraction at any pH value tested. No magnesium effect was noted in experiments in which low-molecularweight components were removed from the fraction by gel filtration (passage through a PD12 G25 Sephadex desalting gel column). The activity in the particulate fraction was inhibited by fluoride (Fig. 4), as usual for PPases [17-19], but not by cyanide (5 mM), cysteine (5 mM), tartrate (10 mM) or EDTA (5 mM). Fluoride inhibition was studied with tetrasodium pyrophosphate, p-nitrophenol phosphate and glycerol 2-phosphate as substrates (Fig 4). About 120 100
0 ~ 0
60
~
2o 0 0.001
0.01
0.1
1
lO
100
Enzyme
Cell extract
Particle fraction
Acid phosphatase Pyrophosphatase N-acetyl-J3-glucosaminidase
30 43 42
37 + 7 42 + 3 48 + 4
Free activity as percent of total activity given. Mean values + SD for 2 experiments with particle fraction.
1 mM F - gave a 50% inhibition of PPi hydrolysis. Acid phosphatase measured with either p-nitrophenol phosphate or glycerol 2-phosphate was inhibited to the same extent at 100 #M concentration. This again suggests that the PPi and the substrates of acid phosphatase are being hydrolyzed by distinct enzymes. The activity in tile particulate fraction showed some latency using the standard assay conditions described, but the long incubation at relatively high temperature meant that it was difficult to detect. When the assay time was reduced to 30 min the activity showed structure-bound latency (Table II). The significant free activity noted was probably due to damaged particles, as the other enzymes assayed, which are known to be localized inside particles, showed similar free activities. After exposure of particles to conditions which are known to break open organelles (addition to the particle fraction of 0.1% Triton X- 100 or sonication in the presence or absence of 100 mM KC1) and centrifugation at high speed, all PPase activity was recovered in the sedimentable fraction. In contrast, between 25 and 55% of the proteins became non-sedimentable due to various combinations of these factors. This suggests that the PPase is tightly bound to the membrane of an organelle.
Activity in the nonsedimentable fraction. The low activity in the cytosolic fraction made the determination of its pH optimum impossible but its activity was much lower at pH 8 than at pH 4.8. The activity in the cytosolic fraction showed no magnesium dependence at either pH 4.8 or 8.
I F ' - ] (raM)
Fig. 4. The effect of F - on phosphatase activities in a particulate fraction of Trichomonas vaginalis. Inorganic pyrophosphatase (0); acid phosphatase assayed with p-nitrophenyl phosphate (O) and glycerol 2-phosphate (A) as substrate.
Discussion
The observed overall activity of PPase in T.
95
vaginalis is relatively low. Under optimum conditions T. vaginalis homogenates exhibit a specific activity of about 7 nmol min -1 (mg protein)This can be compared to levels of 130 nmol m i n - l (mg protein) -1 in mammalian liver [20] and 190 nmol min -~ (mg protein) -1 in E. histolytica [8]. Still higher specific activities are observed in certain prokaryotes, 1800 nmol min -~ (mg protein) - l in Streptococcus faecium [18], 1200 nmol m i n - l (mg protein)-I in Escherichia coli [18] and 750 nmol min - l (rag protein) -1 in Ureaplasma urealyticum [21 ]. The results also show that the major PPase activity in this organism is a magnesium independent enzyme with acid pH optimum and a relatively low (millimolar) affinity for PPi. This activity is sedimentable, exhibits considerable structure-bound latency and is bound to membranes. Subcellular fractionation showed that this activity is localized in lysosome-like organelles, also containing acid phosphatase and N-acetyl-/3glucosaminidase [ 11 ]. This distribution is different from that of cytosolic magnesium-dependent glycerol 3-phosphatase of T. vaginalis [22]. The PPase residing inside a membrane-bounded organelle is likely to be isolated from the cytosolic compartment, where glycolysis occurs. Its participation in the regulation of cytosolic PPi levels is questionable. Inorganic PPase of E. histolytica has similar properties and localization [8]. Although, as discussed below, the dominant PPase of most eukaryotes is an enzyme of significantly different properties, PPases with acid pH optimum, and in several cases localized in lysosomes, have been observed in different animal [23,24] and plant [25,26] cells. The specific activity of the acid enzymes is usually lower than that of the cytosolic alkaline one. These enzymes were little studied and their role in cellular economy remains to be elucidated, The latter conclusion also pertains to the enzymes detected in T. vaginalis and E. histolytica. The dominant PPase of most eukaryotic cells is the extensively studied, magnesium-dependent enzyme with an alkaline pH optimum [27]. This enzyme, localized in the cytosol, is assumed to play a leading role in the removal of PPi generated in the biosynthetic processes. In addition, magnesium-dependent neutral PPases have been
detected in intracellular membranes of higher plants [28-30] which were implicated in transmembrane proton translocation [28,31]. We were unable to detect the presence of either type of activity in T. vaginalis. It is of interest to compare the level of PPase observed in the cytoplasm with estimates of the rates of utilization and production of PPi by T. vaginalis. Utilization of endogenous carbohydrate proceeds at a rate of 10 to 20 nmol min -~ (mg protein)-1 and of exogenous glucose at a rate of 50 to 100 nmol min -~ (mg protein) - t [32]. Since at least one mole of PPi is used in the catabolism of one mole glucose [5], the rate of carbohydrate utilization corresponds to the minimal required rate of PPi production. The rate of PPi production by biosynthetic reactions can be estimated. The generation time of T. vaginalis at 37°C is about 5 h [33], and its biomass contains about 60 percent protein [34]. Klemme takes the PPi liberation to be 7.4 #mol per mg biomass synthesized in organisms which utilize preformed precursors for macromolecular synthesis [35]. This gives a rate of approximately 50 nmol min -~ (mg protein)-~ PPi production in T. vaginalis. This value is of the same order as the estimated utilization of PPi; thus it is likely that biosynthetic reactions are the major if not the sole source of PPi production. The total cellular PPase activity observed at pH 8.0 is at. least two orders of magnitude lower than the estimated rates of PPi production and utilization. The ratio of PPase activity to PPi production becomes still lower for the cytosol in view of the compartmentation of the PPase, indicating that this enzyme plays hardly any role in regulating cytoplasmic PPi levels. Among the microorganisms compared by Klemme [35] none had a similarly low ratio. The mechanisms whereby PPi levels are regulated remain to be elucidated in this organism and other anaerobic protists.
Acknowledgements This work was supported by US Public Health Service grant AI 11942 and CA 46157. S.M.J.S. was a guest scientist at The Rockefeller University on a summer placement from the University of Bath, Bath, U.K. The competent technical help
96
of Ms. Deborah Mulready is gratefully acknowledged. References 1 Kornberg, A. (1962) On the metabolic significance of phosphorolytic and pyrophosphorolytic reactions. In: Horizons in Biochemistry (Kasha, M. and Pullman, B., eds.), pp. 251-264, Academic Press, New York. 2 Wood, H.G. (1977) Some reactions in which inorganic pyrophosphate replaces ATP and serves as a source of energy. Fed. Proc. 36, 2197-2205. 3 Wood, H.G., O'Brien, W.E. and Michaels, G. (1977) Properties of carboxytransphosphorylase; pyruvate, phosphate dikinase; pyrophosphate-phosphofructokinase and pyrophosphate-acetate kinase and their roles on the metabolism of inorganic pyrophosphate. Adv. Enzymol. 45, 85-155. 4 Reeves, R.E. (1984) Metabolism of Entamoeba histolytica Schaudinn, 1903. Adv. Parasitol. 23, 105-142. 5 Mertens, E., Van Schaftingen, E. and Mtiller, M. (1989) Presence of a fructose-2,6-bisphosphate-insensitive pyrophosphate: fructose-6-phosphate phosphotransferase in the anaerobic protozoa Tritrichomonasfoetus, Trichomonas vaginalis and lsotricha prostoma. Mol. Biochem. Parasitol. 37, 183-190. 6 Mertens, E. (1990) Occurrence of pyrophosphate:fructose6-phosphate 1-phosphotransferase in Giardia lamblia trophozoites. Mol. Biochem. Parasitol. 40, 147-150. 7 Reeves, R.E., South, D.J., Blytt, H.J. and Warren, L.G. (1974) Pyrophosphate:D-fructose 6-phosphate 1phosphotransferase. A new enzyme with the glycolytic function of 6-phosphofructokinase. J. Biol. Chem. 249, 7737-7741. 8 McLaughlin, J., Lindmark, D.G. and MUller, M. (1978) Inorganic pyrophosphatase and nucleoside diphosphatase in the parasitic protozoon, Entamoeba histolytica. Biochem. Biophys. Res. Commun. 82, 913-920. 9 Diamond, L.S. (1957) The establishment of various trichomonads of animals and man in axenic cultures. J. Parasitol. 43, 488~.90. 10 MUller, M. (1973) Biochemical cytology of trichomonad flagellates. I. Subcellular localization of hydrolases, dehydrogenases, and catalase in Tritrichomonas foetus. J. Cell Biol. 57, 453~,74. 11 Lindmark, D.G., MUller, M. and Shio, H. (1975) Hydrogenosomes in Trichomonas vaginalis. J. Parasitol. 61, 552-554. 12 Leighton, F., Poole, B., Beaufay, H., Baudhuin, P., Coffey, J.W., Fowler, S. and de Duve, C. (1968) The large-scale separation of peroxisomes, mitochondria, and lysosomes from the livers of rats injected with Triton WR-1339. J. Cell Biol. 37, 482-513. 13 Steinbiichel, A. and MUller, M. (1986) Anaerobic pyruvate metabolism of Tritrichomonasfoetus and Trichomonas vaginalis hydrogenosomes. Mol. Biochem. Parasitol. 20, 57~65. 14 Lowry, O.H., Rosebrough, N.J., Farr, A.L. and Randall, R.J. (1951) Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265-275. 15 Hanes, C.S. (1932) Studies on plant amylases, I. The effect of starch concentration upon the velocity of hydrolysis by the amylase of germinated barley. Biochem. J. 26, 1406-1421.
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