The transport of glucuronides synthesized in the luminal compartment of the endoplasmic reticulum by UDP-glucuronosyltransferase isoenzymes was studied in rat liver microsomal vesicles. Microsomal vesicles were loaded with p-nitrophenol glucuronide (5 mM), phenolphthalein glucuronide or UDP-glucuronic acid, by a freeze–thawing method. It was shown that: (i) the loading procedure resulted in millimolar intravesicular concentrations of the different loading compounds; (ii) addition of UDP-glucuronic acid (5 mM) to the vesicles released both intravesicular glucuronides within 1 min; (iii) glucuronides stimulated the release of UDP-glucuronic acid from UDP-glucuronic acid-loaded microsomal vesicles; (iv) trans-stimulation of UDP-glucuronic acid entry by loading of microsomal vesicles with p-nitrophenol glucuronide, phenolphthalein glucuronide, UDP-glucuronic acid and UDP-N-acetylglucosamine almost completely abolished the latency of UDP-glucuronosyltransferase, although mannose 6-phosphatase latency remained unaltered; (v) the loading compounds by themselves did not stimulate UDP-glucuronosyltransferase activity. This study indicates that glucuronides synthesized in the lumen of endoplasmic reticulum can leave by an antiport, which concurrently transports UDP-glucuronic acid into the lumen of the endoplasmic reticulum.
The mechanisms by which the in vivo intoxication with BrCCl3 inhibits the calcium sequestration activity of liver microsomes were studied. The initial rate of Ca2+ transport is inhibited by nearly 50% in the intoxicated rats as compared to the controls; this indicates that the active transport of Ca2+ is markedly affected by the intoxication. The microsomal ATPase activities both in the presence and in the absence of Ca2+ were not decreased at all in the intoxicated animals. However, the Ca2+-dependent extra ATP-hydrolysis shows a different kinetics in the BrCCl3-poisoned rats with respect to the controls. The release of Ca2+ from Ca2+-loaded liver microsomes is higher in the intoxicated animals. It seems therefore that the increased permeability of the membrane to Ca2+ contributes to some extent to the haloalkane-induced inhibition of the calcium sequestration activity of liver microsomes.
Glucose-6-phosphate transport was investigated in rat or human liver microsomal vesicles using rapid filtration and light-scattering methods. Upon addition of glucose-6-phosphate, rat liver microsomes accumulated the radioactive tracer, reaching a steady-state level of uptake. In this phase, the majority of the accumulated tracer was glucose, but a significant intraluminal glucose-6-phosphate pool could also be observed. The extent of the intravesicular glucose pool was proportional with glucose-6-phosphatase activity. The relative size of the intravesicular glucose-6-phosphate pool (irrespective of the concentration of the extravesicular concentration of added glucose-6-phosphate) expressed as the apparent intravesicular space of the hexose phosphate was inversely dependent on glucose-6-phosphatase activity. The increase of hydrolysis by elevating the extravesicular glucose-6-phosphate concentration or temperature resulted in lower apparent intravesicular glucose-6-phosphate spaces and, thus, in a higher transmembrane gradient of glucose-6-phosphate concentrations. In contrast, inhibition of glucose-6-phosphate hydrolysis by vanadate, inactivation of glucose-6-phosphatase by acidic pH, or genetically determined low or absent glucose-6-phosphatase activity in human hepatic microsomes of patients suffering from glycogen storage disease type 1a led to relatively high intravesicular glucose-6-phosphate levels. Glucose-6-phosphate transport investigated by light-scattering technique resulted in similar traces in control and vanadate-treated rat microsomes as well as in microsomes from human patients with glycogen storage disease type 1a. It is concluded that liver microsomes take up glucose-6-phosphate, constituting a pool directly accessible to intraluminal glucose-6-phosphatase activity. In addition, normal glucose-6-phosphate uptake can take place in the absence of the glucose-6-phosphatase enzyme protein, confirming the existence of separate transport proteins. Glucose-6-phosphate transport was investigated in rat or human liver microsomal vesicles using rapid filtration and light-scattering methods. Upon addition of glucose-6-phosphate, rat liver microsomes accumulated the radioactive tracer, reaching a steady-state level of uptake. In this phase, the majority of the accumulated tracer was glucose, but a significant intraluminal glucose-6-phosphate pool could also be observed. The extent of the intravesicular glucose pool was proportional with glucose-6-phosphatase activity. The relative size of the intravesicular glucose-6-phosphate pool (irrespective of the concentration of the extravesicular concentration of added glucose-6-phosphate) expressed as the apparent intravesicular space of the hexose phosphate was inversely dependent on glucose-6-phosphatase activity. The increase of hydrolysis by elevating the extravesicular glucose-6-phosphate concentration or temperature resulted in lower apparent intravesicular glucose-6-phosphate spaces and, thus, in a higher transmembrane gradient of glucose-6-phosphate concentrations. In contrast, inhibition of glucose-6-phosphate hydrolysis by vanadate, inactivation of glucose-6-phosphatase by acidic pH, or genetically determined low or absent glucose-6-phosphatase activity in human hepatic microsomes of patients suffering from glycogen storage disease type 1a led to relatively high intravesicular glucose-6-phosphate levels. Glucose-6-phosphate transport investigated by light-scattering technique resulted in similar traces in control and vanadate-treated rat microsomes as well as in microsomes from human patients with glycogen storage disease type 1a. It is concluded that liver microsomes take up glucose-6-phosphate, constituting a pool directly accessible to intraluminal glucose-6-phosphatase activity. In addition, normal glucose-6-phosphate uptake can take place in the absence of the glucose-6-phosphatase enzyme protein, confirming the existence of separate transport proteins. Liver plays a major role in regulation of blood glucose levels. In response to stress or low blood glucose levels, it releases glucose for use by other tissues. The terminal step of both glycogenolysis and gluconeogenesis, the two glucose producing pathways, is catalyzed by glucose-6-phosphatase (EC 3.1.3.9) (1Nordlie R.C. Sukalski K.A. Martonosi A.N. The Enzymes of Biological Membranes. 2. Plenum Press, New York1985: 349-398Google Scholar). The importance of this enzyme in the regulation of blood glucose levels is clear from the debilitating effects in the absence of enzyme activity in glycogen storage disease (GSD) 1The abbreviations used are: GSD, glycogen storage disease; ER, endoplasmic reticulum; Mops, 4-morpholinepropanesulfonic acid. type 1 (2Chen Y.T. Burchell A. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill, New York1995: 935-965Google Scholar). The enzyme activity was originally recovered with the microsomal subcellular fraction in the 1950s, e.g. de Duve et al. (3de Duve C. Pressman R. Gianetto R. Wattiaux R. Appelmans F. Biochem. J. 1955; 60: 604-617Google Scholar), which mainly derives from the endoplasmic reticulum (ER) membranes (4de Duve C. Beaufay H. Bull. Soc. Chim. Biol. 1954; 36: 1525-1526Google Scholar). The latency of its activity, discovered in early studies (5Ashmore J. Weber G. Vitam. Horm. 1959; 17: 91-132Google Scholar, 6Nordlie R.C. Boyer P.D. The Enzymes. Academic Press, New York1971: 543-609Google Scholar), together with histochemical studies (7Leskes A. Siekevitz P. Palade G.E. J. Cell Biol. 1971; 49: 264-287Google Scholar) indicated the compartmentation of the enzyme in the ER lumen. Consistent with this, more recent sequence information (8Lei K.-J. Shelly L.L. Pan C.-J. Sidbury J.B. Chou J.Y. Science. 1993; 262: 580-583Google Scholar, 9Shelly L.L. Lei K.-J. Pan C.-J. Sakata S.F. Ruppert S. Schutz G. Chou J.Y. J. Biol. Chem. 1993; 268: 21482-21485Google Scholar) revealed that mammalian glucose-6-phosphatases contain the carboxyl-terminal two-lysine retention motif by which transmembrane proteins are retained in the ER by retrieval from the Golgi (10Jackson M.R. Nilsson T. Peterson P.A. EMBO J. 1990; 9: 3153-3162Google Scholar, 11Jackson M.R. Nilsson T. Peterson P.A. J. Cell Biol. 1993; 121: 317-333Google Scholar). They are also very hydrophobic proteins (12Burchell A. Allan B.B. Hume R. Mol. Membr. Biol. 1994; 11: 217-227Google Scholar), and a variety of topological studies indicate that the active site of the enzyme is located in the lumen of microsomes (13Nilsson O.S. Arion W.J. Depierre J.W. Dallner G. Ernster L. Eur. J. Biochem. 1978; 82: 627-634Google Scholar,14Waddell I.D. Burchell A. Biochem. J. 1991; 275: 133-137Google Scholar). There is no consensus of opinion to date, however, concerning the catalytic mechanism of the enzyme. Presently there are essentially two models to explain it. According to the "translocase-catalytic unit" or "substrate transport" model (15Arion W.J. Wallin B.K. Lange A.J. Ballas L.M. Mol. Cell. Biochem. 1975; 6: 75-83Google Scholar, 16Arion W.J. Lange A.J. Walls H.E. Ballas L.M. J. Biol. Chem. 1980; 255: 10396-10406Google Scholar, 17Burchell A. FASEB J. 1990; 4: 2978-2988Google Scholar, 18Waddell I.D. Scott H. Grant A. Burchell A. Biochem. J. 1991; 275: 363-367Google Scholar), the catalytic site of the glucose-6-phosphatase enzyme, situated inside the lumen of the ER (16Arion W.J. Lange A.J. Walls H.E. Ballas L.M. J. Biol. Chem. 1980; 255: 10396-10406Google Scholar), acts in concert with at least three putative ER transport proteins for the substrate glucose-6-phosphate and for the products phosphate and glucose, which have been named T1, T2, and T3 (2Chen Y.T. Burchell A. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill, New York1995: 935-965Google Scholar, 12Burchell A. Allan B.B. Hume R. Mol. Membr. Biol. 1994; 11: 217-227Google Scholar, 16Arion W.J. Lange A.J. Walls H.E. Ballas L.M. J. Biol. Chem. 1980; 255: 10396-10406Google Scholar, 18Waddell I.D. Scott H. Grant A. Burchell A. Biochem. J. 1991; 275: 363-367Google Scholar), respectively. In the "combined conformational flexibility-substrate transport model" (19Schulze H.-U. Nolte B. Kannler R. J. Biol. Chem. 1986; 261: 16571-16578Google Scholar, 20St-Denis J.-F. Berteloot A. Vidal H. Annabi B. van de Werve G. J. Biol. Chem. 1995; 270: 21092-21097Google Scholar, 21Berteloot A. St-Denis J.-F. van de Werve G. J. Biol. Chem. 1995; 270: 21098-21102Google Scholar), there is no T1 transport protein. Instead, glucose-6-phosphatase enzyme traverses the microsomal membrane forming a water-filled space around the catalytic site in the ER membranes. The catalytic site is thus accessible from the cytosol, and the latency would be caused by the interactions between the enzyme and its membrane environment. The substrate transport model was suggested more than two decades ago after comparing the enzyme kinetic behavior of native microsomes, which exhibit enzyme latency, and that of detergent-disrupted microsomes,i.e. after removal of the microsomal membrane barrier (15Arion W.J. Wallin B.K. Lange A.J. Ballas L.M. Mol. Cell. Biochem. 1975; 6: 75-83Google Scholar,16Arion W.J. Lange A.J. Walls H.E. Ballas L.M. J. Biol. Chem. 1980; 255: 10396-10406Google Scholar). It was subsequently observed that various GSD 1 patients lack the enzyme activity in native but not in disrupted microsomes (2Chen Y.T. Burchell A. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill, New York1995: 935-965Google Scholar) and, most importantly, that the enzyme activities of these patients behave as predicted by the original substrate transport model (22Lange A.L. Arion W.J. Beaudet A.L. J. Biol. Chem. 1980; 255: 8381-8384Google Scholar, 23Nordlie R.C. Sukalski K.A. Munoz J.M. Baldwin J.J. J. Biol. Chem. 1983; 258: 9739-9744Google Scholar, 24Nordlie R.C. Scott H.M. Waddell I.D. Hume R. Burchell A. Biochem. J. 1992; 281: 859-863Google Scholar). In other words, certain cases of GSD can be explained assuming the inherited deficiency of the putative transporter T1 (GSD 1b), T2 (GSD 1c), or T3 (GSD 1d) (2Chen Y.T. Burchell A. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill, New York1995: 935-965Google Scholar). Liver glucose-6-phosphatase enzyme cDNAs have been cloned in humans, rats, and mice (8Lei K.-J. Shelly L.L. Pan C.-J. Sidbury J.B. Chou J.Y. Science. 1993; 262: 580-583Google Scholar, 9Shelly L.L. Lei K.-J. Pan C.-J. Sakata S.F. Ruppert S. Schutz G. Chou J.Y. J. Biol. Chem. 1993; 268: 21482-21485Google Scholar, 25Ruppert S. Boshart M. Bosch F.X. Schmid W. Fournier R.E.K. Schutz G. Cell. 1990; 61: 895-904Google Scholar, 26Mohn K.L. Melby A.E. Tewari D.S. Laz T.M. Taub R. Mol. Cell. Biol. 1991; 11: 381-390Google Scholar), and a number of point mutations of the gene have been shown to underlie GSD 1a (27Lei K.-J. Chen Y.-T. Chen H. Wong L.-J.C. Lui J.-L. McConkie-Rosell A. Van Hove J.L.K. Ou H.C.-Y. Yen N.J. Pan L.Y. Chou J.Y. Am. J. Hum. Genet. 1995; 57: 766-771Google Scholar). In contrast, no mutations of this gene have been found in patients with deficiencies of microsomal glucose-6-phosphate transport (28Lei K.-J. Shelly L.L. Lin B. Sidbury J.B. Chen Y.-T. Nordlie R.C. Chou J.Y. J. Clin. Invest. 1995; 95: 234-240Google Scholar). This indicates that the enzyme protein does not have transport function and that other loci (and proteins) are also needed for glucose-6-phosphate hydrolysis in native (intact) microsomes (2Chen Y.T. Burchell A. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill, New York1995: 935-965Google Scholar, 12Burchell A. Allan B.B. Hume R. Mol. Membr. Biol. 1994; 11: 217-227Google Scholar, 16Arion W.J. Lange A.J. Walls H.E. Ballas L.M. J. Biol. Chem. 1980; 255: 10396-10406Google Scholar). Very recent evidence also indicates that the human chromosome 17 (which contains the glucose-6-phosphatase enzyme gene) is not the site of the defect in GSD 1b (29Lee P. Fenske C. Jeffery S. Weber J. Leonard J. J. Inherited Metab. Dis. 1996; 19: 133Google Scholar). Light-scattering transport experiments showed that liver microsomal vesicles are permeable to glucose-6-phosphate, but not to its isomers mannose-6-phosphate and glucose-1-phosphate (30Fulceri R. Bellomo G. Gamberucci A. Scott H.M. Burchell A. Benedetti A. Biochem. J. 1992; 286: 813-817Google Scholar), which supports a selective ER transport for the substrate of the glucose-6-phosphatase enzyme. Despite the increasing genetic and other evidence of the existence of T1, there is still controversy. The main recent argument for the nonexistence of a T1 transport protein in the conformational model is based on the putative absence of accumulation of glucose-6-phosphate in liver microsomes incubated with the 14C-labeled hexose phosphate (6Nordlie R.C. Boyer P.D. The Enzymes. Academic Press, New York1971: 543-609Google Scholar, 31Berteloot A. Vidal H. van de Werve G. J. Biol. Chem. 1991; 266: 5497-5507Google Scholar). In fact, it was observed that liver microsomes accumulate more 14C than could be explained by the facilitative transport/passive equilibrium of added extravesicular [14C]glucose-6-phosphate, and this was interpreted as a microsomal accumulation of the enzyme product 14C-labeled glucose, assuming that the latter exits the microsomal lumen slowly (6Nordlie R.C. Boyer P.D. The Enzymes. Academic Press, New York1971: 543-609Google Scholar,31Berteloot A. Vidal H. van de Werve G. J. Biol. Chem. 1991; 266: 5497-5507Google Scholar). In other kinetic studies (31Berteloot A. Vidal H. van de Werve G. J. Biol. Chem. 1991; 266: 5497-5507Google Scholar, 32Ajzannay A. Mithieux G. Arch. Biochem. Biophys. 1996; 326: 238-242Google Scholar), the enzyme activity showed an initial burst phase, which was attributed to a tight coupling between glucose-6-phosphate transport and hydrolysis and thus to an extravesicular substrate pool. In addition, neither glucose-6-phosphate nor glucose uptake and accumulation have been observed in liver microsomes obtained from a GSD 1a patient (33St-Denis J.-F. Comte B. Nguyen D.K. Seidman E. Paradis K. Lévy E. van de Werve G. J. Clin. Endocrinol. & Metab. 1994; 79: 955-959Google Scholar). Very recently, knockout mice for the glucose-6-phosphatase enzyme gene have been produced (34Lei K.-J. Chen H. Pan C.-J. Ward J.M. Mosinger Jr., B. Lee E.J. Westphal H. Mansfield B.C. Chou J.Y. Nat. Genet. 1996; 13: 203-209Google Scholar), and it was observed that the addition of [14C]glucose-6-phosphate resulted in lower uptake of14C in microsomes isolated from livers of knockout mice compared with the control, which was discussed as glucose-6-phosphatase-dependent substrate transport (34Lei K.-J. Chen H. Pan C.-J. Ward J.M. Mosinger Jr., B. Lee E.J. Westphal H. Mansfield B.C. Chou J.Y. Nat. Genet. 1996; 13: 203-209Google Scholar). In contrast to this background, we have investigated the nature of the hepatic microsomal intravesicular pools deriving from the transport of glucose-6-phosphate. To this end, we have measured both [14C]glucose-6-phosphate uptake and [14C]glucose accumulation in rat hepatic microsomal vesicles as well as in liver microsomal preparations derived from two GSD 1a cases. Here we show that glucose-6-phosphate crosses the microsomal membrane and forms an intraluminal metabolically active pool allowing the formation of an intraluminal glucose pool whose extents were directly dependent on glucose-6-phosphatase activity. The present results confirm previous genetic evidence that the glucose-6-phosphatase enzyme is not responsible for endoplasmic reticulum glucose-6-phosphate transport and that a different T1 protein/gene is responsible for the ER transport of glucose-6-phosphate. In addition, the results provide an alternative explanation for kinetic data that were previously considered to support the conformational model. 24 h-fasted male Sprague-Dawley rats (180–230 g) were used. Liver microsomes were prepared as reported (35Henne V. Söling H.D. FEBS Lett. 1986; 202: 267-273Google Scholar). Microsomal fractions were resuspended in a buffer (buffer A) containing (in mM): KCl, 100; NaCl, 20; MgCl2, 1; and Mops, 20, pH 7.2. The suspensions were rapidly frozen and maintained under liquid N2 until used. Intactness of microsomal vesicles checked by measuring the latency of mannose-6-phosphatase activity (36Burchell A. Hume R. Burchell B. Clin. Chim. Acta. 1988; 173: 183-192Google Scholar) was greater than 90% in all the preparations employed. Microsomal protein concentrations were determined by biuret reaction using bovine serum albumin as a standard. In some experiments, microsomal glucose-6-phosphatase was inactivated by mild acidic treatment according to (37Arion W.J. Lange A.J. Ballas L.M. J. Biol. Chem. 1976; 251: 6784-6790Google Scholar). To measure microsomal water space, microsomes were diluted (10 mg protein/ml) in buffer A containing [3H]H2O (0.2 μCi/ml) or [3H(C)]inulin (0.17 μCi/ml) and centrifuged (100,000 × g, 60 min), and the radioactivity associated with pellets was measured to enable calculation of extravesicular and intravesicular water spaces (38Marcolongo P. Fulceri R. Giunti R. Burchell A. Benedetti A. Biochem. Biophys. Res. Commun. 1996; 219: 916-922Google Scholar, 39Fulceri R. Bellomo G. Gamberucci A. Romani A. Benedetti A. Biochem. J. 1993; 289: 299-306Google Scholar). Both GSD type 1a patients were initially diagnosed by kinetic analysis of the glucose-6-phosphatase system in microsomes isolated from liver biopsy samples. Case 1 had mild symptoms until adulthood, elevated hepatic glycogen levels, abnormally low glucose-6-phosphatase enzyme activity levels (20% of age-matched control levels), and low levels of abnormally sized glucose-6-phosphatase enzyme protein as judged by immunoblot analysis. 2C. Hinds, and A. Burchell, manuscript in preparation. Case 2 had a much more severe form of type 1a GSD with virtually all the signs and symptoms described for the disorder (2Chen Y.T. Burchell A. Scriver C.R. Beaudet A.L. Sly W.S. Valle D. The Metabolic Basis of Inherited Disease. McGraw-Hill, New York1995: 935-965Google Scholar), elevated hepatic glycogen levels, virtually no glucose-6-phosphatase enzyme activity (∼1% of age-matched control values), and no immunodetectable glucose-6-phosphatase enzyme protein by immunoblot analysis. Portions of liver biopsy samples from the two patients were histologically examined to check for the presence of adenoma or hepatoma cells. The need to do this is illustrated by case 2. Histology demonstrated that a first needle biopsy sample from the patient was partially adenoma and was not used for the present study (or diagnosis). In contrast, a second liver sample was found to contain only (nontransformed) liver cells and thus was used to prepare the microsomes investigated here. Human liver microsomes were prepared in 0.25 m sucrose containing 5 mm Hepes (pH 7.4) by differential centrifugation as described previously (40Burchell A. Jung R.T. Lang C.C. Shepard A.N. Lancet. 1987; i: 1059-1062Google Scholar). The intactness of the two type 1 GSD microsomal preparations, based on the latency ofp-nitrophenol UDP-glucuronosyltransferase activity (41Fulceri R. Bánhegyi G. Gamberucci A. Giunti R. Mandl J. Benedetti A. Arch. Biochem. Biophys. 1994; 309: 43-46Google Scholar), was greater than 90%. Protein concentrations were estimated by the method of Lowry as modified by Peterson (42Peterson G.L. Anal. Biochem. 1977; 83: 346-356Google Scholar). The study of the glucose-6-phosphatase system in human liver samples was approved by the Ethics Committee of Tayside Health Board. Liver microsomes (1 mg protein/ml) were incubated in buffer A containing 0.2, 0.5, 1, 5, 10, or 30 mm glucose-6-phosphate plusd-[14C(U)]glucose-6-phosphate (2–3 μCi/ml) at 22 °C. At the indicated time intervals, samples (0.1 ml) were rapidly filtered through cellulose acetate/nitrate filter membranes (pore size 0.22 μm), and filters were washed with 4 ml of Hepes (20 mm) buffer (pH 7.2) containing 250 mm sucrose and 0.5 mm 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid. This latter compound was added to reduce eventual efflux of vesicular glucose-6-phosphate during the washing procedure (30Fulceri R. Bellomo G. Gamberucci A. Scott H.M. Burchell A. Benedetti A. Biochem. J. 1992; 286: 813-817Google Scholar). The time required to execute filtration and washing was 15–20 s. Filtration of samples of media containing no microsomes, and washing of filters as above, resulted in negligible amounts of radioactivity retained by filters. The total 14C associated to microsomes retained by filters was measured by liquid scintillation counting. Parallel filters were treated with ZnSO4-Ba(OH)2 to separate [14C]glucose from [14C]glucose-6-phosphate, and labeled glucose-6-phosphate and glucose were recovered, respectively, in the pellet and supernatant, after centrifugation (43Fulceri R. Bellomo G. Gamberucci A. Benedetti A. Biochem. J. 1991; 275: 805-807Google Scholar). Briefly, washed filters were transferred into tubes containing 0.3 ml of 0.2 m ZnSO4 and were pushed to the bottom. After mixing, 0.6 ml of a saturated solution of Ba(OH)2 was added. Tubes were centrifuged to remove the white precipitate and filters. A 0.45-ml portion of the clear supernatant was used to measure [14C]glucose produced from [14C]glucose-6-phosphate by liquid scintillation spectroscopy. Routinely, the amount of [14C]glucose-6-phosphate was calculated by subtracting [14C]glucose from the total 14C associated to microsomes measured in parallel samples (see above). In preliminary experiments, we verified i) that more than 97% of standard [14C]glucose and of standard [14C]glucose-6-phosphate applied on filters were recovered in the clear supernatant and in the precipitate, respectively, and ii) that direct measurements of precipitated [14C]glucose-6-phosphate (after resuspending barium precipitates and filters with ZnSO4-Ba(OH)2solutions and centrifuging to remove [14C]glucose carry-over) gave essentially similar results. In each experiment, alamethicin (0.05 mg/ml) was added to the parallel incubates to distinguish the intravesicular and the bound radioactivity (41Fulceri R. Bánhegyi G. Gamberucci A. Giunti R. Mandl J. Benedetti A. Arch. Biochem. Biophys. 1994; 309: 43-46Google Scholar, 44Ojcius D.M. Young J.D.-E. Trends Biochem. Sci. 1991; 16: 225-229Google Scholar). The alamethicin-permeabilized microsomes were recovered on filters and washed as above. More than 95% of the microsomal protein was retained by filters, indicating that the alamethicin treatment did not affect the vesicular structure of microsomes as already reported (41Fulceri R. Bánhegyi G. Gamberucci A. Giunti R. Mandl J. Benedetti A. Arch. Biochem. Biophys. 1994; 309: 43-46Google Scholar). The alamethicin-permeabilized microsomes retained amounts of radioactivity ≤ 20% of that associated to untreated microsomes. Intravesicular radioactive compounds were bona fide lost during the washing procedure since the alamethicin nonreleasable portion did not further decrease even after extensive washing of filters (and microsomes). This allowed us to regard the alamethicin releasable portion of radioactivity as intravesicular. To unequivocally identify the intraluminal material precipitated with barium as glucose-6-phosphate, some microsomal samples were incubated for 5 min in the presence of (30 mm) glucose-6-phosphate and in the presence or in the absence of alamethicin as described above. Samples (1 mg of protein) were filtered, and, after washing, the filters were treated with perchloric acid (3%, 1 ml). Samples were neutralized with KHCO3, tubes were centrifuged to remove the precipitate and filters, and the glucose-6-phosphate content of the neutralized supernatants was measured enzymatically. To this end, 0.5 ml of the supernatant was mixed with an equal volume of buffer A containing 2 mm NADP+, and NADPH formation was detected fluorimetrically (excitation and emission wavelengths were 360 and 470 nm, respectively) upon the addition of glucose-6-phosphate dehydrogenase (0.7 IU/ml). Pulse additions of standard glucose-6-phosphate (1 to 5 nmol) to the reaction mixture allowed the quantitation of microsomal glucose-6-phosphate. Where indicated, the microsomal passive equilibration of intra- and extravesicular glucose-6-phosphate was calculated according to the formula: apparent intravesicular glucose-6-phosphate space (μl/mg of protein) = glucose-6-phosphate accumulated by microsomes (nmol/mg of protein)/concentration of added glucose-6-phosphate (nmol/μl) (30Fulceri R. Bellomo G. Gamberucci A. Scott H.M. Burchell A. Benedetti A. Biochem. J. 1992; 286: 813-817Google Scholar,45Meissner G. Methods Enzymol. 1988; 157: 417-437Google Scholar). Glucose-6-phosphatase activity was measured after 5 min of incubation in buffer A at 22 °C on the basis of d-[14C(U)]glucose production from d-[14C(U)]glucose-6-phosphate according to (43Fulceri R. Bellomo G. Gamberucci A. Benedetti A. Biochem. J. 1991; 275: 805-807Google Scholar). At high substrate concentrations (30 mm), the enzyme activity was also evaluated by measuring glucose production with a glucose (Trinder) kit (Sigma). Osmotically induced changes in microsomal vesicle size and shape were monitored at 400 nm at right angles to the incoming light beam using a fluorimeter (Perkin-Elmer model 650–10S) equipped with a temperature-controlled cuvette holder (22 °C) and magnetic stirrer as described elsewhere (30Fulceri R. Bellomo G. Gamberucci A. Scott H.M. Burchell A. Benedetti A. Biochem. J. 1992; 286: 813-817Google Scholar, 45Meissner G. Methods Enzymol. 1988; 157: 417-437Google Scholar). The mV output signals were acquired at 0.25 s intervals, using MacLabTM hardware (AD Instruments) equipped with Chart Ver. 3.2.5. software. Glucose-6-phosphate (monosodium salt), mannose-6-phosphate (disodium salt), alamethicin, NADP+, and 4,4′-diisothiocyanostilbene-2,2′-disulfonic acid were obtained from Sigma. Na3VO4 was from Fisher Science Co.d-[14C(U)]glucose-6-phosphate (300 mCi/mMol) was from American Radiolabeled Chemicals Inc., St. Louis, MO. [3H]H2O (1 mCi/g) and [3H(C)]inulin (500 mCi/g) were from DuPont NEN, Dreieich, Germany. Glucose-6-phosphate dehydrogenase (from yeast, 350 IU/ml) was from Boehringer Mannheim, Germany. Cellulose acetate/nitrate filter membranes (pore size 0.22 μm) were from Millipore. All other chemicals were of analytical grade. In a first set of experiments, rat liver microsomal vesicles were incubated in the presence of various concentrations of glucose-6-phosphate (plus [14C]glucose-6-phosphate as a tracer). The radioactivity associated with microsomes was measured in vesicles incubated both in the presence and absence of the pore-forming antibiotic alamethicin (41Fulceri R. Bánhegyi G. Gamberucci A. Giunti R. Mandl J. Benedetti A. Arch. Biochem. Biophys. 1994; 309: 43-46Google Scholar, 44Ojcius D.M. Young J.D.-E. Trends Biochem. Sci. 1991; 16: 225-229Google Scholar) to determine net intravesicular accumulation. Because, in addition to [14C]glucose-6-phosphate, [14C]glucose (produced by glucose-6-phosphatase activity) can contribute to the intravesicular measured radioactivity (as it was indeed the case, see below), we expressed as "glucose-6-phosphate + glucose" the 14C-radioactivity accumulated (see Fig.1). Glucose-6-phosphate did not likely undergo major reactions other than dephosphorylation in our incubation system, therefore glucose-6-phosphate + glucose concentrations could be calculated on the basis of the concentrations of glucose-6-phosphate added. Microsomes rapidly accumulated glucose-6-phosphate + glucose until a steady-state level was reached over a 2-min period of incubation (Fig. 1 A). The steady-state intraluminal accumulation of glucose-6-phosphate + glucose increased by increasing the extravesicular concentration of glucose-6-phosphate although not in a linearly proportional fashion (Fig. 1 B). Based on the measured intravesicular water space of rat liver microsomes (3.6 ± 1.1 μl/mg protein, mean ± SD, n = 6) the mm intravesicular concentrations of glucose-6-phosphate + glucose were also calculated (Fig. 1 B). Relatively low extravesicular concentrations (≤1 mm) of glucose-6-phosphate resulted in intravesicular steady-state concentrations of glucose-6-phosphate + glucose higher than those of the extravesicularly added glucose-6-phosphate (Fig. 1 B,inset; concentrations of added glucose-6-phosphate are indicated by the dotted line). On the other hand, at glucose-6-phosphate concentrations ≥5 mm, steady-state intravesicular concentrations of glucose-6-phosphate + glucose were lower than the added glucose-6-phosphate concentrations (Fig.1 B). In theory, intravesicular concentrations of glucose-6-phosphate can maximally equal the extravesicular ones since no energy, or ion gradients, were present in the system to allow microsomal inward transport of glucose-6-phosphate over the passive equilibrium. Therefore, the higher intravesicular concentrations of glucose-6-phosphate + glucose, in the presence of glucose-6-phosphate concentrations ≤1 mm, were logically contributed to by glucose accumulation. Separate measurements of glucose and glucose-6-phosphate present in the lumen of microsomal vesicles revealed that accumulation of glucose was responsible for the apparent increase in intravesicular concentration of 14C-labeled compounds over that of extravesicular glucose-6-phosphate. However, besides the intravesicular glucose pool, a glucose-6-phosphate pool could also be demonstrated by two different methods. The intravesicular steady-state glucose-6-phosphate content of the microsomes at 30 mm glucose-6-pho
Carbonyl compounds released during the NADPH-Fe dependent lipid peroxidation and identified as 4-hydroxyalkenals (almost entirely as 4-hydroxynonenal), when incubated with isolated hepatocytes, produce loss of viability in 95% of the cells, as measured by the trypan blue exclusion test. They also produce an almost complete permeabilization of the plasma-membrane, as measured by the test of the permeability to NADH. concomitantly with the permeabilization of the plasma-membrane, a marked release of enzymes (lactate dehydrogenase and glutamate-pyruvate transaminase) from the hepatocytes occurs. These and other activities of the above mentioned carbonyl compounds suggest the possibility that these products represent some of the effective mediators of the liver injury produced by those toxins, such as CCL4, which promote the peroxidation of membrane lipids.
The inactivation of liver microsomal glucose 6 phosphatase induced either by Fe2+ or by haloalkanes (CCl4, CBrCl3) was investigated in NADPH-microsomes systems. In the case of haloalkanes, EDTA was included in the incubation mixtures, so to exclude participation of free Fe2+ in the ensuing lipid peroxidation. Microsomal glucose 6 phosphatase activity was measured along with the release of malonic dialdehyde and the appearance of carbonyl products bound to microsomal protein, taken as indices of the peroxidative process. Fe2+ was added to NADPH-microsomes at different concentrations, one (6 microM) resulting in an extent of lipid peroxidation comparable with that induced by haloalkanes, the other (60 microM) representing a situation of excess Fe2+, leading to massive lipid peroxidation. Inhibition of glucose 6 phosphatase caused by 6 microM Fe2+ was comparable to that induced by haloalkanes in EDTA-microsomes systems, which supports the view that lipid peroxidation--rather than covalent binding of free radical metabolites--represents the main event leading to the inactivation of glucose 6 phosphatase caused by haloalkanes. The production of 4-hydroxynonenal--the known toxic product of lipid peroxidation--was also studied. A remarkable accumulation of 4-hydroxynonenal was observed in the microsomal membranes after peroxidation induced by 6 microM Fe2+ or haloalkanes, as compared to the incubation medium. In addition, experiments carried out with CCl4 and CBrCl3 in vivo suggested the possible existence of a cytosolic detoxification system able to remove lipid-derived carbonyls bound to microsomal protein.
Loss‐of‐function mutations in the gene encoding GLUT10 are responsible for arterial tortuosity syndrome (ATS), a rare connective tissue disorder. In this study GLUT10‐mediated dehydroascorbic acid (DAA) transport was investigated, supposing its involvement in the pathomechanism. GLUT10 protein produced by in vitro translation and incorporated into liposomes efficiently transported DAA. Silencing of GLUT10 decreased DAA transport in immortalized human fibroblasts whose plasma membrane was selectively permeabilized. Similarly, the transport of DAA through endomembranes was markedly reduced in fibroblasts from ATS patients. Re‐expression of GLUT10 in patients’ fibroblasts restored DAA transport activity. The present results demonstrate that GLUT10 is a DAA transporter and DAA transport is diminished in the endomembranes of fibroblasts from ATS patients.
1. MgATP-dependent Ca2+ uptake by rat liver microsomal preparations and permeabilized hepatocytes was measured in the presence or absence of Pi. 2. Monitoring of free Ca2+ in incubation systems with a Ca2+ electrode in the presence of Pi (2-7 mM) revealed a biphasic Ca2+ uptake, with the onset of a second, Pi-dependent, Ca2+ accumulation. 3. Increasing Pi concentrations (up to 10 mM) caused a progressive enlargement of 45Ca2(+)-loading capacity of microsomal fractions. 4. As a result of Pi stimulation of active Ca2+ uptake, [32P]Pi and 45Ca2+ were co-accumulated. 5. Experiments with permeabilized hepatocytes revealed that the amount of Ca2+ releasable by myo-inositol 1,4,5-trisphosphate is unaffected by Pi.
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