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Am J Physiol Cell Physiol 278: C803-C811, 2000;
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Vol. 278, Issue 4, C803-C811, April 2000

Sorting of metabolic pathway flux by the plasma membrane in cerebrovascular smooth muscle cells

Pamela G. Lloyd and Christopher D. Hardin

Department of Physiology, University of Missouri-Columbia, Columbia, Missouri 65212


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We used beta -escin-permeabilized pig cerebral microvessels (PCMV) to study the organization of carbohydrate metabolism in the cytoplasm of vascular smooth muscle (VSM) cells. We have previously demonstrated (Lloyd PG and Hardin CD. Am J Physiol Cell Physiol 277: C1250-C1262, 1999) that intact PCMV metabolize the glycolytic intermediate [1-13C]fructose 1,6-bisphosphate (FBP) to [1-13C]glucose with negligible production of [3-13C]lactate, while simultaneously metabolizing [2-13C]glucose to [2-13C]lactate. Thus gluconeogenic and glycolytic intermediates do not mix freely in intact VSM cells (compartmentation). Permeabilized PCMV retained the ability to metabolize [2-13C]glucose to [2-13C]lactate and to metabolize [1-13C]FBP to [1-13C]glucose. The continued existence of glycolytic and gluconeogenic activity in permeabilized cells suggests that the intermediates of these pathways are channeled (directly transferred) between enzymes. Both glycolytic and gluconeogenic flux in permeabilized PCMV were sensitive to the presence of exogenous ATP and NAD. It was most interesting that a major product of [1-13C]FBP metabolism in permeabilized PCMV was [3-13C]lactate, in direct contrast to our previous findings in intact PCMV. Thus disruption of the plasma membrane altered the distribution of substrates between the glycolytic and gluconeogenic pathways. These data suggest that organization of the plasma membrane into distinct microdomains plays an important role in sorting intermediates between the glycolytic and gluconeogenic pathways in intact cells.

beta -escin; glycolysis; gluconeogenesis; permeabilization; channeling; vascular smooth muscle; caveolae


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

IN THE TEXTBOOK VIEW of the cell, the intermediates and enzymes of carbohydrate metabolism are freely diffusible components of the cytoplasm. However, it is unlikely that this is the case in living cells. Most glycolytic enzymes probably exist in both free and bound states (for example, see Ref. 26). The interactions of glycolytic enzymes with the F-actin cytoskeleton are well-documented (2), and an actin-binding region has been found in the glycolytic enzyme aldolase (23). Associations of glycolytic enzymes with microtubules have also been demonstrated repeatedly (36) and a glycolytic enzyme-binding domain has recently been identified on alpha -tubulin (35). Thus glycolytic enzymes are probably not freely diffusible in intact cells.

The enzymes of the glycolytic pathway may engage in metabolite channeling (31). Metabolite channeling occurs when an intermediate is transferred directly from one enzyme to another, or when an intermediate is present at locally high concentrations that are out of equilibrium with the bulk of the cytoplasm (24). Localization of enzymes to structures such as actin filaments or microtubules would facilitate this process. The intermediates of gluconeogenesis may be similarly channeled.

The concentrations of glycolytic enzymes are similar to the concentrations of glycolytic intermediates within the cell (32). Therefore, if the intermediates of glycolysis and gluconeogenesis are channeled, the access of exogenous substrates to the pathways will be limited because most enzyme active sites will be occupied by substrates (32). Likewise, once a particular substrate molecule has entered a pathway, it is unlikely to diffuse away. Thus each intermediate remains within the pathway, making it unavailable for use by other pathways in which it is also an intermediate (compartmentation). Compartmentation of glycolytic, gluconeogenic, and glycogenolytic intermediates has been shown in previous studies in our laboratory (9, 11-13) and others (1, 15).

We have recently found that vascular smooth muscle of isolated pig cerebral microvessels (PCMV) utilizes [1-13C]fructose 1,6-bisphosphate ([1-13C]FBP; a glycolytic intermediate) for gluconeogenesis, while simultaneously utilizing [2-13C]glucose for glycolysis. Thus exogenous [1-13C]FBP does not mix with the [2-13C]FBP derived from glucose breakdown, and this tissue exhibits a compartmentation of glycolysis and gluconeogenesis (18). Because glycolytic enzymes are known to associate with microtubules, we examined the role of microtubules in compartmentation of glycolysis and gluconeogenesis. Our data suggested that glycolytic rate is partially regulated by the availability of binding sites for glycolytic enzymes on tubulin (18). However, microtubules did not appear to be involved in the regulation of gluconeogenic flux. In addition, associations of glycolytic enzymes with microtubules did not appear to be the basis of the compartmentation of metabolism we observed. Based on these results, we hypothesized that gluconeogenic enzymes are localized elsewhere within the cell, and that a portion of the glycolytic pathway is also localized to structural elements other than microtubules.

Recently, a number of studies have demonstrated that the plasma membrane is organized into microdomains (such as caveolae) in which proteins of related functions are concentrated (22). Glycolytic enzymes are known to associate with the plasma membrane (34) and the enzyme phosphofructokinase has recently been identified in caveolae (29). Thus we hypothesized that the plasma membrane could be the site of gluconeogenic enzyme localization, as well as the location of a portion of the glycolytic pathway. We disrupted the plasma membrane of vascular smooth muscle (VSM) of PCMV using beta -escin to examine the role of the plasma membrane in the regulation and compartmentation of carbohydrate metabolism in this tissue. The results of these studies provide important new information about the organization of metabolism in living cells.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Tissue collection. Pig brains were obtained at a local abattoir within 30 min of slaughter. Brains were placed in ice-cold physiological saline solution (PSS) for transport to the laboratory. PSS consisted of the following (in mM): 116 NaCl, 4.6 KCl, 1.16 KH2PO4, 25.3 NaHCO3, 2.5 CaCl2, 1.16 MgSO4, and 5 glucose, pH 7.4. PSS was oxygen- and pH-equilibrated before use by gassing with 95% O2/5% CO2. To prevent microbial contamination, 303 mg/l penicillin G and 100 mg/L streptomycin sulfate were added to PSS. PSS was also filtered through a 0.22-µm filter before use (Micron Separations, Westboro, MA). Brains were stored in fresh PSS at 4°C until use.

Microvessel isolation. Microvessels were isolated as previously described (18) by a modification of the method of Sussman et al. (33). Microvessels were isolated from three brains for each experiment. The brains were placed in HEPES-buffered PSS (HBPSS). HBPSS contained 118 mM NaCl, 5.4 mM KCl, 1.8 mM CaCl2, 1.0 mM MgSO4, 28 mM HEPES, 1.0 mM NaH2PO4, 0.2% (wt/vol) BSA, 1 U/ml heparin, and 10 µM isoproterenol, pH 7.4. HBPSS was supplemented with antibiotics and filtered as described for PSS. The outer layers of the brain were removed and the cerebral cortex was dispersed by aspiration into a plastic vacuum flask. The aspirated material was homogenized by five strokes in a stainless steel Dounce-type homogenizer (Dura-Grind, Thomas Scientific, Swedesboro, NJ). Microvessels were collected by pouring the brain homogenate over nylon meshes, which trapped the vessels while allowing smaller pieces of tissue to pass through. A 210-µm nylon mesh (Small Parts, Miami Lakes, FL) was used first to remove large vessels from the homogenate. These vessels were discarded. The material that passed through the 210-µm mesh was filtered over a 105-µm mesh, which trapped vessels of intermediate size. Smaller vessels and debris, which passed through this mesh, were discarded. The vessels adhering to the 105-µm mesh were rinsed with HBPSS, then collected by inverting the mesh and rinsing the vessels into a clean container. PCMV isolated in this manner are largely composed of VSM cells (18).

Permeabilization of microvessels. Microvessels were permeabilized by incubation in Buffer B containing 50 µM beta -escin for 30 min at 21°C. Buffer B contained (in mM) 150 sucrose, 35 potassium acetate, 5 MgSO4, 5 NaH2PO4, 40 HEPES, and 35 KCl, pH 7.55 (4). Permeabilization conditions and solutions were adapted from Iizuka et al. (14) and from previous studies (3, 4, 7). After permeabilization, microvessels were rinsed with fresh Buffer B.

Metabolic studies of permeabilized microvessels. Permeabilized microvessels were resuspended in 9 ml of Buffer B containing 5 mM [1-13C]FBP (Omicron Biochemicals, South Bend, IN) and 5 mM [2-13C]glucose (pH 7.55; Cambridge Isotope Laboratories, Andover, MA). Buffer B also contained the cofactors ATP (1 mM) and NAD (1 mM), as well as an ATP-regenerating system composed of phosphocreatine (PCr, 10 mM), creatine (Cr, 10 mM), and creatine phosphokinase (CPK, 2.5 U/mL). The suspension was mixed to ensure even distribution of microvessels, and 8 ml was pipetted into a 25-cm2 polystyrene cell culture flask (Corning Costar, Cambridge, MA). The flask was incubated for 3 h at 37°C in a shaking bath. At the conclusion of the incubation, a 5.5-ml sample of the suspension was withdrawn from the flask. The sample was centrifuged (Marathon 6K, Fisher Scientific) at 1,000 g for 5 min to pellet the microvessels. For NMR analysis, 4 ml of the resulting supernatant were frozen in liquid nitrogen. A 4-ml sample of the starting solution containing labeled substrates was also saved for NMR analysis.

Metabolic studies of intact microvessels. Metabolism in intact microvessels was examined largely as described above, except that microvessels were not permeabilized with beta -escin, the incubations were performed in HBPSS rather than Buffer B, and no additional cofactors were supplied.

Effects of exogenous ATP on metabolism in permeabilized vessels. To determine the effect of exogenous ATP concentration on metabolism in permeabilized vessels, microvessels were isolated from three brains and permeabilized as described above. The vessels were then split into two aliquots. One aliquot was incubated in Buffer B containing labeled substrates and additional cofactors as described above. The second aliquot was incubated in an identical solution, except that no ATP was provided.

Effects of exogenous NAD on metabolism in permeabilized microvessels. Metabolism in permeabilized vessels was examined at several concentrations of exogenous NAD to determine how the presence of this cofactor affected glycolytic and gluconeogenic rate. Microvessels were isolated and permeabilized as described above, then split into two aliquots. One aliquot was incubated as described above, in incubation medium containing 1 mM NAD. The second aliquot was incubated in an identical solution, except that the concentration of NAD was changed to 0, 0.2, 2, or 4 mM.

Effect of variations in phosphorylation potential on metabolism in permeabilized microvessels. We also examined metabolism in permeabilized vessels at varying ATP-to-ADP ratios ([ATP/ADP]) to determine whether phosphorylation potential modified glycolytic rate. Microvessels were isolated and permeabilized as described above, then split into two aliquots. One aliquot was incubated in the standard solution described above containing labeled substrates, 1 mM ATP, 1 mM NAD, 10 mM PCr, 10 mM Cr, and 2.5 U/mL CPK. Because K' = [ATP][Cr]/[ADP][PCr] = 100 (20), [ADP] for this solution = 0.01 mM, and [ATP/ADP] = 100. The second aliquot was incubated in the same solution, except that the concentrations of PCr and Cr were changed to either 10 mM PCr and 1 mM Cr ([ATP/ADP] = 1,000) or 1 mM PCr and 10 mM Cr ([ATP/ADP] = 10). Thus [ATP/ADP] was varied over two orders of magnitude in these experiments. Data obtained at [ATP/ADP] = 10 and [ATP/ADP] = 1,000 were normalized to the values obtained at [ATP/ADP] = 100.

NMR spectroscopy. Supernatant solutions from metabolic experiments (and the starting solution for each experiment) were lyophilized to powder in a Speed Vac (Savant Instruments, Farmingdale, NY). Dry samples were resuspended in 800 µl of 99.9% D2O (Cambridge Isotope Laboratories, Andover, MA) containing 25 mM 3-(trimethylsilyl)-1-propanesulfonic acid (TMSPS) as a chemical shift reference. A 650-µl aliquot of this solution was transferred to a 5-mm NMR tube for NMR spectroscopy.

13C-NMR was performed using a Bruker DRX 500 spectrometer. One thousand two hundred scans were acquired after sixty-four dummy scans using a 30° pulse angle at 125.77 MHz, 33,333-Hz sweep width, and 1-s predelay. A total of 32,768 points were acquired and processed with line broadening of 1 Hz before Fourier transform of the data. All spectra were broad-band proton decoupled. All peak positions and intensities were normalized to the signal of TMSPS, set at 0 ppm. Peak intensity was calculated using Bruker software. No corrections for nuclear Overhauser effects were made because these effects were expected to be the same for all experiments. Supernatants from intact PCMV were examined as above, except that the number of scans was 300.

Statistical analysis. Significant differences in the metabolism of intact and permeabilized microvessels were detected by comparing 13C-NMR peak intensities of interest using two-tailed t-tests for two samples assuming unequal variances. Significant differences between 13C-NMR peak intensities of microvessels incubated at 0 and 1 mM ATP were detected using a two-tailed t-test for paired samples. Significant differences between 13C-NMR peak intensities of microvessels incubated at [ATP/ADP] = 10 and 100 and [ATP/ADP] = 100 and 1,000 were detected using two-tailed t-tests for paired samples. Values of P < 0.05 were considered significant. All statistical calculations were performed using Microsoft Excel 97 software.

Reagents. Na2HPO4 and NaH2PO4 were purchased from Aldrich Chemical, Milwaukee, WI. All other chemicals (except where otherwise stated) were obtained from Sigma Chemical, St. Louis, MO.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

PCMV are capable of both glycolysis and gluconeogenesis. beta -Escin-permeabilized PCMV incubated with labeled substrates in the presence of 1 mM ATP, an ATP-regenerating system, and 1 mM NAD retained their glycolytic ability, metabolizing [2-13C]glucose to [2-13C]lactate and [1-13C]FBP to [3-13C]lactate. Permeabilized PCMV also metabolized [1-13C]FBP to [1-13C]glucose, demonstrating that the gluconeogenic pathway remained active in permeabilized cells (Fig. 1). Thus PCMV retain metabolic activity, despite extensive disruption of the plasma membrane and free access of cytoplasmic components to the extracellular solution. These results suggest that both glycolytic and gluconeogenic intermediates are channeled in VSM of PCMV because these small compounds would otherwise diffuse out of cells and be diluted by the extracellular solution, halting metabolism (see DISCUSSION).


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  		Fig. 1.   beta -Escin-permeabilized pig cerebral microvessels (PCMV) are capable of both glycolysis and gluconeogenesis. Permeabilized PCMV metabolized [2-13C]glucose [beta anomer, 76.7; alpha  anomer, 74.0 parts per million (ppm)] to [2-13C]lactate (70.9 ppm). Permeabilized PCMV metabolized [1-13C]fructose 1,6-bisphosphate ([1-13C]FBP; beta  anomer, 68.4; alpha  anomer, 67.4 ppm) to both [3-13C]lactate (22.7 ppm) and [1-13C]glucose (beta  anomer, 98.6; alpha  anomer, 94.8 ppm, is hidden by a solution peak). Peaks at 0, 17.6, and 21.7 ppm represent 3-(trimethylsilyl)-1-propanesulfonic acid (TMSPS).

Permeabilization alters metabolic flux and the distribution of substrates between glycolysis and gluconeogenesis. As discussed above, both the glycolytic pathway and the gluconeogenic pathway remained active in PCMV after beta -escin treatment. However, considerable alterations in both metabolic pathway flux and the distribution of substrates between the two pathways were observed in permeabilized PCMV relative to intact PCMV (Fig. 2). In the presence of 1 mM NAD, 1 mM ATP, and an ATP-regenerating system, permeabilization significantly (P < 0.0001) reduced flux of [2-13C]glucose to [2-13C]lactate, to 20.8% of the flux measured in intact PCMV. The flux of [1-13C]FBP to [1-13C]glucose in permeabilized PCMV was also significantly (P < 0.0001) reduced, to 27.5% of the flux in intact PCMV.


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  		Fig. 2.   Permeabilization alters metabolic pathway flux and distribution of [1-13C]FBP between the glycolytic and gluconeogenic pathways in PCMV. Production of [2-13C]lactate from [2-13C]glucose is significantly decreased in permeabilized PCMV (right, dark gray bar; n = 18) relative to intact PCMV (left, dark gray bar; n = 10). Formation of [1-13C]glucose from [1-13C]FBP is also significantly decreased in permeabilized PCMV (right, solid bar) relative to intact PCMV (left, solid bar). The major product of [1-13C]FBP metabolism in permeabilized PCMV is [3-13C]lactate (right, light gray bar), in contrast to intact PCMV (left), in which no [3-13C]lactate production was detectable. All values are normalized to signal of TMSPS and represent means ± SE. *Significantly different from intact values (P < 0.0001). Intact PCMV data are taken from Ref. 18.

In contrast, the flux of [1-13C]FBP to [3-13C]lactate was greatly increased after permeabilization, with [3-13C]lactate representing the major product of carbohydrate metabolism. The increased glycolytic flux from [1-13C]FBP, coupled with the decreased glycolytic flux from [2-13C]glucose, resulted in similar total lactate production in permeabilized and intact PCMV (permeabilized = 110.2% of intact).

The increase in [3-13C]lactate production after permeabilization represents a significant shift in the distribution of [1-13C]FBP between the glycolytic and gluconeogenic pathways. In intact PCMV, [1-13C]glucose production accounted for 100% of the [1-13C]FBP utilization, with no detectable [3-13C]lactate production. In contrast, [1-13C]FBP metabolism in permeabilized PCMV was largely glycolytic, with [3-13C]lactate production accounting for 64.1 ± 4.0% (mean ± SE) of the total [1-13C]FBP utilization.

These data show that disruption of the plasma membrane alters the partitioning of [1-13C]FBP between the glycolytic and gluconeogenic pathways. In order for compartmentation to exist, exogenous FBP and exogenous glucose must be kept in separate cytoplasmic pools in the cell, from their entry as substrates to their exit as products. We have shown that [1-13C]FBP can access the glycolytic pathway after permeabilization but not before. Thus the intact plasma membrane is required for compartmentation, and transport of [1-13C]FBP and [2-13C]glucose across the membrane must be the first step in sorting these substrates into the glycolytic or gluconeogenic pathways. Both exogenous FBP and exogenous glucose have equal access to the plasma membrane of intact cells, but appear to be directed into different metabolic pathways by it. Therefore, transport of FBP and glucose must occur in spatially separate microdomains within the plasma membrane (see DISCUSSION).

Exogenous ATP is required for maximal glycolysis in permeabilized PCMV. Permeabilized PCMV that were incubated with labeled substrates in the presence of 1 mM NAD and an ATP-regenerating system (but no ATP) had minimal glycolytic flux (Fig. 3). Production of [3-13C]lactate from [1-13C]FBP was significantly enhanced in the presence of 1 mM ATP (P < 0.001). In the presence of ATP, there was a corresponding decrease in the production of [1-13C]glucose from [1-13C]FBP (P < 0.01). Production of [2-13C]lactate from [2-13C]glucose was almost undetectable in the absence of ATP, and was significantly increased in its presence (P < 0.05). Thus exogenous ATP is required for maximal glycolytic activity in permeabilized cells. These data demonstrate that the plasma membrane is permeable to molecules at least as large as glycolytic intermediates (the largest of which is FBP, molecular weight 406.1) because ATP (molecular weight 551.1) is able to enter cells freely. These data also suggest that exogenous FBP has a higher affinity for the glycolytic pathway than the gluconeogenic pathway because gluconeogenesis from FBP was markedly decreased in the presence of ATP.


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  		Fig. 3.   Metabolism in permeabilized cells is sensitive to exogenous ATP (n = 4). Although some [3-13C]lactate is produced via glycolysis from [1-13C]FBP even in absence of exogenous ATP (left, solid bar), [3-13C]lactate production is significantly increased in presence of 1 mM ATP (left, open bar). In absence of exogenous ATP, rate of gluconeogenesis, as represented by [1-13C]glucose production from [1-13C]FBP, is high (middle, solid bar). When exogenous ATP is added and glycolytic rate increases, production of [1-13C]FBP is reduced (middle, open bar). Glycolysis from [2-13C]glucose, as represented by production of [2-13C]lactate, is almost undetectable in absence of exogenous ATP (right, solid bar) and is increased in presence of 1 mM ATP (right, open bar). Values represent means ± SE. *P < 0.001; **P < 0.01; #P < 0.05.

Exogenous NAD is required for maximal glycolytic rate in permeabilized PCMV. Permeabilized PCMV incubated with labeled substrates, ATP, and an ATP-regenerating system (but no NAD) produced very little [3-13C]lactate from [1-13C]FBP (Fig. 4). When metabolism was examined over a range of NAD concentrations (0.2, 1, 2, and 4 mM) a clear relationship between NAD concentration and [3-13C]lactate production was observed, with half-maximal [3-13C]lactate production at 0.68 mM NAD. [1-13C]glucose production showed an inverse relationship to NAD concentration, declining as [NAD] increased. Thus when appropriate cofactors are available, [1-13C]FBP is preferentially metabolized by the glycolytic pathway.


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  		Fig. 4.   Metabolism of [1-13C]FBP by permeabilized cells is sensitive to exogenous NAD. Glycolysis of [1-13C]FBP to yield [3-13C]lactate () is very low in absence of exogenous NAD. As concentration of exogenous NAD increases, there is a corresponding increase in [3-13C]lactate production (solid line, r2 = 0.88) and a corresponding decrease in [1-13C]glucose production (black-triangle, dotted line, r2 = 0.93). Values at each concentration represent means ± SE and are normalized to peak intensity of PCMV incubated in presence of 1 mM NAD. N = 4 (0 mM), 3 (0.2 mM), 3 (2 mM), and 4 (4 mM).

Production of [2-13C]lactate from [2-13C]glucose was also sensitive to the concentration of exogenous NAD, and increased as [NAD] increased (Fig. 5). Half-maximal production of [2-13C]lactate was observed at 0.44 mM NAD. Thus lactate production from either [1-13C]FBP or [2-13C]glucose was similarly dependent on NAD. As for ATP, these data show that NAD (molecular weight 685.4) has free access to the cytoplasm and confirm that PCMV were adequately permeabilized.


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  		Fig. 5.   Metabolism of [2-13C]glucose by permeabilized PCMV is sensitive to exogenous NAD. Glycolysis of [2-13C]glucose to yield [2-13C]lactate (black-diamond ) is low in absence of exogenous NAD, but increases as concentration of NAD increases (r2 = 0.99). Values at each concentration represent means ± SE and are normalized to peak intensity of PCMV incubated in presence of 1 mM NAD. N = 4 (0 mM), 3 (0.2 mM), 3 (2 mM), and 4 (4 mM).

Glycolysis in permeabilized PCMV is not sensitive to variations in phosphorylation potential. We also examined the role of phosphorylation potential in metabolism in permeabilized PCMV. The ratio of ATP-to-ADP ([ATP/ADP]) in the incubation solution was varied from 10 to 1,000 by varying the concentrations of Cr, PCr, and ATP. All data were normalized to the values obtained at [ATP/ADP] = 100. No significant differences in lactate production from either [1-13C]FBP or [2-13C]glucose were observed at either high ([ATP/ADP] = 100) or low ([ATP/ADP] = 10) phosphorylation potentials, relative to lactate production at [ATP/ADP] = 100 (Fig. 6). Therefore, phosphorylation potential did not affect lactate production from either [2-13C]glucose or [1-13C]FBP over a 100-fold range of [ATP/ADP] values.


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  		Fig. 6.   Variations in [ATP/ADP] from 10 to 1000 have little effect on either [3-13C]lactate production from [1-13C]FBP (open circle ) or [2-13C]lactate production from [2-13C]glucose (). Peak intensities of [3-13C]lactate and [2-13C]lactate at [ATP/ADP] = 10 and [ATP/ADP] = 1,000 were not significantly different from corresponding peak intensities at [ATP/ADP] = 100 (P > 0.05). No trend in either [3-13C]lactate production (dashed line, r2 = 0.58) or [2-13C]lactate production (solid line, r2 = 0.52) was observed over the range of [ATP/ADP] examined. All data represent means ± SE and are normalized to peak intensity of PCMV incubated at [ATP/ADP] = 10. N = 3 for each concentration.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We have previously demonstrated that a compartmentation of carbohydrate metabolism exists in vascular smooth muscle of PCMV (18). Intact PCMV metabolized the glycolytic intermediate [1-13C]FBP almost entirely to [1-13C]glucose (gluconeogenesis), rather than to [3-13C]lactate (glycolysis). Simultaneously, intact PCMV metabolized [2-13C]glucose to [2-13C]lactate via glycolysis. Thus the intermediates of glycolysis and the intermediates of gluconeogenesis do not mix freely in the cytoplasm of VSM cells. We are currently investigating the structural basis of this compartmentation.

The intermediates of glycolysis and gluconeogenesis are channeled. Metabolite channeling may be one aspect of the structural basis of compartmentation. Generally, metabolic intermediates are considered to be channeled if they are "transferred from one enzyme to another without complete equilibration with the surrounding medium" (24). This transfer can occur directly, via enzyme-enzyme associations. Alternatively, locally high concentrations of intermediates that are kept out of equilibrium with the bulk solution of the cell can facilitate the interaction of enzyme with substrate (24). Channeling has been demonstrated in a variety of biochemical pathways, including phosphatidylcholine biosynthesis (6) and DNA replication (27). It has been suggested that the intermediates of glycolysis and other metabolic pathways are channeled as well (for a recent review, see Ref. 25). Glycolytic enzymes and intermediates are present at similar concentrations in the cell, suggesting that most enzyme active sites are occupied by substrates in vivo (32). Thus, if glycolytic intermediates are channeled directly between enzymes, exogenous intermediates will have limited access to the pathway because there will be few free active sites. This model is consistent with our previous results in PCMV showing that access of exogenous FBP to the glycolytic pathway is restricted (18).

In this study, we examined metabolism in VSM cells permeabilized with beta -escin. The effective disruption of membrane integrity by beta -escin is demonstrated both by our current results and by studies in other laboratories. Our data show that both glycolytic and gluconeogenic fluxes are markedly affected by external NAD and ATP (see Figs. 3-5). Because cells are normally impermeable to NAD and ATP, these data suggest that beta -escin treatment effectively permeabilizes the plasma membrane. These results are in agreement with published studies by other laboratories on beta -escin and other detergents. In rabbit portal vein, a 30-min treatment with 50-100 µM beta -escin rendered smooth muscle cells permeable to ~150-kDa proteins. Both primary antibodies to smooth muscle alpha -actin and fluorescently labeled secondary antibodies were able to enter the cells after beta -escin treatment. beta -Escin also resulted in the partial efflux of lactate dehydrogenase, demonstrating that proteins could both enter and exit the cells (14). Similarly, contraction in smooth muscle of guinea pig ileum treated with 20 µM beta -escin for 20 min was sensitive to extracellular Ca2+, calmodulin, inositol 1,4,5-trisphosphate, and MgATP (17). These results demonstrate that beta -escin permeabilization allows molecules ranging in size from small cofactors to large proteins to both enter and exit cells. Most glycolytic enzymes are smaller than 150 kDa (32). Therefore, both glycolytic enzymes and intermediates should be able to cross the plasma membrane freely after beta -escin permeabilization, if they are freely diffusible in the cytoplasm. However, although the plasma membrane is permeable to large molecules and enzymes, specific enzymes may be retained within the cells. In this study, we have shown that both the glycolytic and gluconeogenic pathways continued to operate in permeabilized PCMV. Thus all of the enzymes of these pathways were retained by the cells with sufficient activity for flux to continue. Because these enzymes did not diffuse out of the permeabilized cells, their diffusion must be restricted by associations with intracellular structures. Such localization of enzyme activity is requisite for compartmentation of metabolism to occur.

In contrast to the slow diffusion of structurally associated macromolecules such as enzymes, diffusion of metabolites and cofactors into and out of permeabilized cells is likely to be rapid. In saponin-permeabilized guinea pig ventricular myocytes, ATP-sensitive K+ channels (KATP channels) remain open in the absence of either ATP or substrates for ATP production, demonstrating the loss of ATP from the cells. However, a combination of glycolytic substrates and cofactors (FBP, NAD, ADP, and Pi) closes KATP channels within 1 min of addition to the bathing solution, demonstrating the rapid entry of these compounds into permeabilized cells (37). Although only a portion of the plasma membrane was exposed to saponin, the cells were depleted of ATP within 2 min of washing ATP out of the bath. These results show that metabolic substrates and cofactors are able to pass rapidly across the plasma membrane, even in partially permeabilized cells. Thus all of the evidence from this study and from studies in other laboratories suggests that beta -escin treatment effectively removes barriers to the diffusion of glycolytic intermediates across the plasma membrane in our preparation.

Because permeabilization should allow efflux of glycolytic intermediates, the flux of glycolysis and gluconeogenesis might be expected to cease after beta -escin treatment. When pelleted by centrifugation, the microvessels in the incubation mixture represented only ~100 µl of the total 8-ml incubation solution. Thus if intermediates diffused freely out of the permeabilized cells, they would be diluted ~80× by the incubation medium. Such dilution of metabolic intermediates would effectively halt metabolism, even though the enzymes are retained within the cells. However, we found that both glycolysis and gluconeogenesis remain active in permeabilized cells.

If permeabilization resulted in a large increase in the activities of all the glycolytic enzymes, then it is possible that our results could be explained by continued metabolic flux due to mass action. In our experiments, the effective dilution was ~80-fold. However, we observed a decrease in metabolism ([2-13C]lactate production from [2-13C]glucose and [1-13C]glucose production from [1-13C]FBP) of approximately fivefold. Therefore, to support the level of metabolism that we observed, each enzyme in the glycolytic and gluconeogenic pathways would have to increase its activity by ~16-fold. We feel this is a highly unlikely possibility. Therefore, we conclude that glycolytic and gluconeogenic intermediates are channeled in VSM of PCMV, preventing dilution of the intermediates by the bathing medium and allowing metabolism to continue. These results are in agreement with previous studies in our laboratory (7) and others (3, 21) on glycolysis in cells permeabilized with dextran sulfate or saponin. To our knowledge, this is the first study examining carbohydrate metabolism in cells permeabilized with beta -escin. In addition, this is the first report of channeling in two overlapping pathways operating in opposite directions in the same preparation.

Glycolysis and gluconeogenesis are modulated by cofactors in permeabilized cells. We found that both glycolytic rate and gluconeogenic rate were modulated by ATP and NAD in permeabilized PCMV. Therefore, beta -escin permeabilization effectively removed the ability of the plasma membrane to serve as a diffusive barrier to small molecules. Although restrictions to diffusion of small molecules within the cytoplasm may still exist after plasma membrane permeabilization, these intracellular diffusion barriers are unlikely to be significant over the long (3 h) incubation times used in these experiments. Maximal glycolysis was observed when 1 mM NAD, 1 mM ATP, and an ATP-regenerating system were supplied in addition to the labeled substrates. In the absence of NAD, glycolysis from either [1-13C]FBP or [2-13C]glucose was low, indicating that most of the cytoplasmic NAD had diffused out into the incubation medium. Exogenous ATP also modulated both glycolysis and gluconeogenesis. Lactate production was low in the absence of added ATP, again demonstrating diffusion into the extracellular medium. Addition of 1 mM ATP to the incubation medium increased lactate production from both [1-13C]FBP and [2-13C]glucose, while decreasing [1-13C]glucose production from [1-13C]FBP. Conversion of [1-13C]FBP to [1-13C]glucose was highest when glycolysis was inhibited in the absence of NAD and ATP, and declined as [NAD], [ATP], and glycolytic rate increased. Metabolism of [1-13C]FBP to [3-13C]lactate is energetically more favorable than metabolism of [1-13C]FBP to [1-13C]glucose. Thus in the presence of sufficient cofactors (NAD and ATP) and given free access of [1-13C]FBP to the glycolytic pathway (as is provided by permeabilization), it would be expected that the major product of [1-13C]FBP metabolism would be [3-13C]lactate. The decreased production of [1-13C]glucose that we observed in the presence of ATP and NAD may therefore simply reflect the partitioning of [1-13C]FBP between more and less energetically favorable pathways, with metabolism via glycolysis favored when conditions are appropriate.

Role of plasma membrane in pathway sorting. Our results in permeabilized cells can be accounted for by the existence of metabolite channeling. Either enzyme-enzyme interactions or localized enzyme systems (or both) are necessary for channeling to occur. Because glycolysis and gluconeogenesis appear to occur in separate metabolic compartments in intact cells, we hypothesized that the enzymes of glycolysis must be spatially separated from the enzymes of gluconeogenesis. We have investigated the potential role of enzyme associations to microtubules as one structural basis of this phenomenon. However, although associations of glycolytic enzymes with microtubules appeared to regulate glycolytic pathway flux, microtubules did not appear to be required for compartmentation of metabolism to exist (18).

The beta -escin-permeabilized PCMV model described in this study allowed us to examine the role of the plasma membrane in compartmentation of carbohydrate metabolism. In intact PCMV, glucose and FBP must enter the cell via transporters located in the plasma membrane. Unless there are two separate membrane domains for transport of the substrates, mixing of the pathway intermediates should occur at the cytoplasmic side of the plasmalemma. Thus the compartmentation of glycolysis and gluconeogenesis that we observed is consistent with the existence of two spatially distinct sites for transport of FBP and glucose.

When PCMV were permeabilized with beta -escin, we observed a major alteration in the partitioning of [1-13C]FBP between glycolysis and gluconeogenesis. In permeabilized cells, [1-13C]FBP utilization was divided between glycolysis ([3-13C]lactate production) and gluconeogenesis ([1-13C]glucose production). This is in contrast to similar studies using intact PCMV (see Fig. 2 and Ref. 18), where FBP was used almost exclusively for gluconeogenesis. Therefore, permeabilization of the plasma membrane removed the diffusion barrier that limits access of FBP to glycolytic enzymes in intact cells. Because the selective nature of substrate access to metabolic pathways is no longer evident in permeabilized cells, it appears that the intact plasma membrane is required for such selectivity to exist. However, there are also several alternative explanations for our data, as discussed below.

One possible explanation for our data showing that beta -escin treatment greatly decreased glycolysis from [2-13C]glucose while greatly increasing glycolysis from [1-13C]FBP is that permeabilization selectively inhibited [2-13C]lactate production from [2-13C]glucose, relative to [3-13C]lactate production from [1-13C]FBP. Such an effect could possibly be produced by a selective inhibition or loss of hexokinase, phosphohexose isomerase, and phosphofructokinase (the enzymes preceding FBP in the glycolytic pathway) relative to the rest of the glycolytic enzymes. An effect of this type would presumably be independent of the permeabilizing agent used and would reflect differential associations of the enzymes within the cell. However, this scenario seems unlikely because a study of saponin-permeabilized rat adipocytes demonstrated that only 4-8% of the activity of any of the glycolytic enzymes could be found outside of the permeabilized cells. In addition, no enzymes were preferentially released by the permeabilizing treatment (21). Therefore, preferential loss of enzymes from the top portion of the pathway is unlikely to account for our results.

Another possible explanation for our results is suggested by the fact that glycolysis is functionally linked to plasma membrane ion transport activities, including those mediated by the Ca2+-ATPase (10), the Na+-K+-ATPase (28), and KATP channels (37). Therefore, any alterations in these processes caused by beta -escin could also produce alterations in glycolytic rate. For example, inability of beta -escin-permeabilized cells to maintain ion gradients could cause increased ion transport activity, which could then stimulate increased glycolysis. This mechanism would be consistent with our data if the stimulation of glycolysis occurred only between aldolase and lactate dehydrogenase (thus resulting in a selective stimulation of glycolysis from [1-13C]FBP, but not from [2-13C]glucose). Membrane ATPase activity stimulates glycolysis by producing Pi and ADP from ATP (28). Thus one strategy to address this possibility would be to use specific inhibitors of membrane ATPases (such as ouabain, an inhibitor of Na+-K+-ATPase activity) to inhibit production of Pi and ADP. However, since a wide variety of ATPases have been proposed to be coupled with glycolysis, the inhibition of one ATPase would not be sufficient to investigate this general mechanism. Therefore, to investigate the possibility that increased ion transport could account for our results, we altered the ATP-to-ADP ratio over two orders of magnitude to determine whether changes in [ATP/ADP] altered lactate production (from either [2-13C]glucose or [1-13C]FBP). This strategy should affect all ATPases because of the large range of phosphorylation potentials examined. As shown in Fig. 6, lactate production from either [2-13C]glucose or [1-13C]FBP does not change over a large range of [ATP/ADP]. Therefore, it is unlikely that alterations in [ATP/ADP] produced by increased membrane ATPase turnover could account for the increase in glycolysis from [1-13C]FBP that we observed after permeabilization of PCMV with beta -escin.

Finally, beta -escin permeabilization could have caused increased [3-13C]lactate production by 1) allowing increased entry of [1-13C]FBP into permeabilized cells (relative to intact cells), resulting in increased [3-13C]lactate production by mass action; or 2) by damaging the mitochondria, resulting in impaired oxidation of [3-13C]pyruvate. Because the cells are permeable to small molecules, the rate of transport of [1-13C]FBP should not be limiting for metabolism in permeabilized cells. However, increased uptake of [1-13C]FBP cannot in itself explain the greatly increased rate of [3-13C]lactate production in permeabilized PCMV because the rate of [1-13C]glucose production from [1-13C]FBP is significantly reduced (see Fig. 2). Accumulation of [1-13C]FBP within the cell should result in increased activity of both FBP-metabolizing pathways, rather than selectively stimulating the glycolytic pathway while inhibiting the gluconeogenic pathway. It is also unlikely that damage to the mitochondria can account for the increased [3-13C]lactate production in permeabilized cells because such damage would have also increased [2-13C]lactate production from [2-13C]glucose, and we observed the opposite effect.

Therefore, the most likely explanation for our data is that the intact plasma membrane is required for compartmentation of metabolism. Recent studies have demonstrated the existence of plasma membrane microdomains containing specific protein components in vascular smooth muscle (22). A model demonstrating how plasma membrane organization could contribute to metabolic compartmentation is presented in Fig. 7. The plasma membrane may sort intermediates between competing metabolic pathways by restricting access of substrates to the cytoplasm to specific membrane microdomains containing appropriate transporters, which are colocalized with metabolic enzymes on the inner face of the membrane. Glucose and FBP enter cells by different mechanisms. Glucose enters cells via glucose transporters, whereas FBP most likely enters cells via dicarboxylate transporters (5, 8). Some isoforms of the glucose transporter may be localized to caveolae, although conflicting reports on the localization exist (16, 30). In addition, the glycolytic enzyme phosphofructokinase has recently been localized to caveolae (29). Localization of glucose transporters and glycolytic enzymes to caveolae could allow glucose molecules crossing the membrane to immediately enter the glycolytic pathway. In addition to their membrane localization, glycolytic enzymes are also associated with microtubules and other structures deeper within the cytoplasm. Colocalization of dicarboxylate transporters with gluconeogenic enzymes in a separate membrane microdomain could explain the inability of exogenous FBP to access either membrane- or microtubule-associated glycolytic enzymes in intact cells.


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  		Fig. 7.   Model demonstrating how localization of glycolytic and gluconeogenic enzymes to separate membrane microdomains and cytoplasmic sites could account for compartmentation of glycolysis and gluconeogenesis, as well as for changes in [1-13C]FBP metabolism after permeabilization. FBP enters intact cells (top) via dicarboxylate transporters (filled cylinders), which are localized to noncaveolar domains of plasma membrane. Gluconeogenic enzymes (filled circles) are also localized to noncaveolar domains. Colocalization of dicarboxylate transporter with gluconeogenic enzymes favors preferential utilization of exogenous FBP for gluconeogenesis, rather than glycolysis. Glucose enters intact cells via glucose transporters (open cylinders), which are present in both caveolar and noncaveolar domains. Glycolytic enzymes (open circles) localized in caveolae or deeper within cytoplasm on structures such as microtubules convert glucose to lactate. After permeabilization (bottom), exogenous FBP is no longer constrained to entering cells via dicarboxylate transporters, and is able to access both glycolytic and gluconeogenic enzymes.

Therefore, a colocalization of functionally matched transporters and metabolic enzymes may allow for independent regulation of pathway flux. This is analogous to the spatial organization of biological signaling systems involving cytoarchitecture and specific membrane domains to modulate signaling pathway flux and cross talk (38). Indeed, endothelial nitric oxide synthase and the arginine transporter are colocalized in caveolae, allowing for efficient delivery of arginine to the enzyme (19). This arrangement of a membrane transporter with a functionally related signaling enzyme is analogous to the arrangement of transporters and metabolic enzymes that we propose in our model. Therefore, the spatial organization of metabolism and cell signaling may share many common features.


    ACKNOWLEDGEMENTS

The technical assistance of Tina Roberts is appreciated.


    FOOTNOTES

This work was supported by an Established Investigator Grant from the American Heart Association (C. D. Hardin), National Heart, Lung, and Blood Institute Training Grant HL-07094 (support to P. G. Lloyd), American Heart Association (Heartland Affiliate) Predoctoral Fellowship 9910198Z (P. G. Lloyd), and National Science Foundation instrumentation Grant CHE-89-08304. Pig brains were provided by Excel, Marshall, MO.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: C. D. Hardin, Dept. of Physiology, MA415 Medical Sciences Bldg., Univ. of Missouri-Columbia, Columbia, MO 65212 (E-mail: HardinC{at}health.missouri.edu).

Received 16 June 1999; accepted in final form 29 October 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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