Glycolytic flux in permeabilized freshly isolated vascular smooth muscle cells

Christopher D. Hardin and Dorian R. Finder

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

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

To determine whether channeling of glycolytic intermediates can occur in vascular smooth muscle (VSM), we permeabilized freshly isolated VSM cells from hog carotid arteries with dextran sulfate. The dextran sulfate-treated cells did not exclude trypan blue, a dye with molecular weight of ~1,000. If glycolytic intermediates freely diffuse, plasmalemmal permeabilization would allow intermediates to exit the cell and glycolytic flux should cease. We incubated permeabilized and nonpermeabilized cells with 5 mM [1-13C]glucose at 37°C for 3 h. 13C nuclear magnetic resonance (NMR) was used to determine relative [3-13C]lactate production and to identify any 13C-labeled glycolytic intermediates that exited from the permeabilized cells. [3-13C]lactate production from [1-13C]glucose was decreased by an average of 32% (n = 6) in permeabilized cells compared with intact cells. No 13C-labeled glycolytic intermediates were observed in the bathing solution of permeabilized cells. We conclude that channeling of glycolytic intermediates can occur in VSM cells.

compartmentation; substrate channeling; permeable cells; enzyme-enzyme interactions

    INTRODUCTION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

VASCULAR SMOOTH MUSCLE (VSM) is the major cellular component of the arterial system, and the contractile state of VSM is responsible for the maintenance of, and alterations in, blood pressure. The energy requirements of VSM are met by a combination of oxidative and glycolytic metabolism. It has been previously suggested that the ATP produced by oxidative metabolism is specifically utilized by the ATPases associated with contraction (8, 22), whereas the ATP produced by glycolysis may be specifically used by the ATPases of ion pumps (3, 9, 24). Therefore, a structuring of metabolism in VSM exists that results in independent metabolic regulation of different VSM functions.

Indeed, carbohydrate metabolism exhibits considerable organization within the cytoplasm of VSM. It has been shown that the intermediates of glycogenolysis (6, 18) and of portions of gluconeogenesis (11, 12) do not completely mix with the intermediates of glycolysis. This compartmentation of metabolism must result from a spatial organization of the glycolytic enzymes within the cytoplasm. Glycolytic enzymes have been localized to the contractile apparatus (7) and the plasma membrane (9, 24) in smooth muscle and to the actin cytoskeleton (20, 21), microtubules (15, 19), and outer mitochondrial membrane (2, 17) of other cell types.

Because the intermediates of glycolysis, glycogenolysis, and gluconeogenesis do not completely mix, it may be predicted that the intermediates of these pathways are not free to diffuse within the cytoplasm. To directly examine whether the intermediates of glycolysis can freely diffuse in the cytoplasm, we permeabilized VSM cells by an adaptation of the methods of Clegg and Jackson (4, 5). We provided permeabilized cells a physiological concentration of glucose in the form of [1-13C]glucose and observed any [3-13C]lactate production and any efflux of 13C-labeled intermediates with 13C nuclear magnetic resonance (NMR) spectroscopy.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Tissue handling. Hog carotid arteries were obtained from a local abattoir within ~30 min of slaughter. Arteries were placed in a cold (~5-10°C) physiological saline solution (PSS), pH 7.4, equilibrated by bubbling with a gas mixture of 95% O2 and 5% CO2. PSS was composed of (in mM) 116 NaCl, 4.6 KCl, 1.16 KH2PO4, 25.3 NaHCO3, 2.5 CaCl2, and 1.16 MgSO4. At the laboratory, the arteries were placed into fresh PSS equilibrated with the gas mixture at 37°C. Segments were dissected free of loose fat, connective tissue, and adventitia.

Cell isolation. We employed a modification of the method of Warshaw et al. (26) for the preparation of freshly isolated VSM cell suspensions. Briefly, one end of a hog carotid artery segment (~5-15 cm long) was tied with suture. A needle attached to a syringe was inserted into the other end of the artery, and the artery was pinched around the needle. Low-calcium PSS (0.5 mM CaCl2) was injected to inflate the artery to check for leaks. Segments with leaks were trimmed or the artery was discarded. A second needle and syringe were used to inflate the arteries with enzyme solution consisting of 496.6 U/ml collagenase, 39.6 U/ml elastase, 60 U/ml deoxyribonuclease, 1.5% bovine serum albumin, 0.1% soybean trypsin inhibitor, 3.99 mM ATP, 0.01 mM isoproterenol, and 1.3% amino acid standard, in low-calcium PSS at a pH of 7.4. On inflation of the artery, suture was securely tied around the other end of the artery while the needle was removed from the artery. Approximately 40 arteries were used for a single experiment. The inflated artery segments were placed in an Erlenmeyer flask holding enough PSS in which to immerse the arteries, and the flask was placed in a shaking water bath at 37°C for 90 min. After the incubation, one end of the artery was cut and the enzyme solution was drained from the artery. The artery was reinflated using enzyme solution with a lower elastase concentration (8 U/ml), which was empirically determined to yield more viable cells (fewer arteries with "over-digested" cells). The arteries were again placed in the water bath at 37°C for 60 min.

After the second incubation, one end of the artery was cut and the enzyme solution was gently drained. The open end of the artery was pinched around a glass pipette filled with buffer A, consisting of (in mM) 93 NaCl, 5 KCl, 5 MgCl2, 35 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), and 0.1 ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA), pH 7.4. The lumen of the artery was rinsed with buffer A by gentle pumping of the solution in and out of the artery. The cell suspension from each artery was placed in a test tube, and the cells were screened for cell condition and density. Only those samples exhibiting at least 90% long and spindle-shaped cells were pooled for the study. Average pooled cell density was 1.7 × 106 ± 0.85 × 106 cells/ml (mean ± SE, n = 13).

Permeabilization and incubations. We adapted the permeabilization procedure of Clegg and Jackson (4, 5) for cultured L-929 cells to our freshly isolated VSM cells. An equal volume of dextran sulfate (1 mg/ml in buffer A, 4°C) was combined with cell suspension (also in buffer A) in conical tubes and incubated at 4°C for 12.5 min. After circular mixing, the cells were incubated at 4°C for an additional 12.5 min. Cells were centrifuged at 20°C and 650 g. Cells were resuspended in buffer A with a volume four times that of the volume of the original cell suspension and then centrifuged at 20°C and 650 g. Cells were resuspended in buffer B (in mM: 150 sucrose, 35 potassium acetate, 35 KCl, 5 MgSO4, 5 NaH2PO4, 40 HEPES, and 5 [1-13C]glucose, pH 7.55) or buffer K (in mM: 150 sorbitol, 70 potassium gluconate, 5 MgCl2, 35 HEPES, 0.1 EGTA, and 5 [1-13C]glucose, pH 7.55) at a volume equal to the volume of the original cell suspension.

An aliquot of cells in buffer B or buffer K was exposed to trypan blue (0.2% final concentration) and, after 10 min, total cell number and the fraction of the total cells that were permeabilized (trypan blue staining) were determined using a hemocytometer. The remaining cells were incubated in a shaking water bath for 3 h at 37°C. At the end of the 3-h incubation, cell suspensions were centrifuged at 20°C and 650 g. The supernatant and the pellet were separated and frozen for analysis by 13C NMR.

Cell disruption. To determine whether disrupted cells can produce [3-13C]lactate from [1-13C]glucose, Triton X-100 was added to a cell suspension (20°C) to achieve a final concentration of 1%. The cell suspension was then rapidly transferred to a Transonic digital sonicator and sonicated at maximum speed at 37°C for 12.5 min. After mixing, the cells were sonicated for an additional 12.5 min. Cells were centrifuged at 20°C and 650 g. Cells were resuspended in buffer A with a volume four times that of the volume of the original cell suspension and then centrifuged at 20°C and 650 g. Cells were resuspended in buffer B at a volume equal to the volume of the original cell suspension. Cells were incubated with 5 mM [1-13C]glucose for 3 h at 37°C. These conditions paralleled the permeabilization and incubation conditions described above but utilized Triton X-100 and sonication to completely disrupt cells.

NMR spectroscopy. Frozen supernatants (2- to 2.5-ml aliquots) were lyophilized in a Speed Vac (Savant Instruments). The powder was resuspended in 800 µl 99% 2H2O with 25 mM 2,2-dimethyl-2-silapentane-5-sulfonate [DSS; also called 3-(trimethylsilyl)-1-propane-sulfonic acid or TMSPS] as a chemical shift reference. A 650-µl aliquot was transferred to a 5-mm NMR tube. 13C NMR spectroscopy was performed on a Bruker DRX 500 spectrometer with the following acquisition parameters: 20,480 scans with 16 dummy scans, 30° pulse angle at 125.77 MHz, 33,333-Hz sweep width, 1-s predelay, and total collection time of 11 h and 20 min; 32,768 points were acquired and processed with 1-Hz line broadening before Fourier transform. All spectra were broad-band proton decoupled and referenced with DSS at 0 parts per million (ppm). NMR data were processed using Bruker software for peak magnitude determination. All peaks are expressed relative to the DSS peak at 0 ppm, and no corrections were made for nuclear Overhauser effects, which were assumed to be unchanged for all experiments. 13C NMR signal intensities were scaled for the total cell number in each incubation.

Determination of chemical shifts of glycolytic intermediates. The chemical shifts of the carbons of the glycolytic intermediates glucose-6-phosphate, fructose-6-phosphate, dihydroxyacetone phosphate, and pyruvate were determined individually using a 500 mM solution of each intermediate dissolved in 2H2O with 25 mM DSS. NMR acquisition parameters were the same as for the cell solution samples except that the number of acquisitions was 300.

Statistics. Statistical significance was determined using a two-tailed paired Student's t-test assuming unequal variances. A P value <= 0.05 was taken to be significant.

    RESULTS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

The preparation of freshly isolated VSM cells yielded more than ~90% spindle-shaped cells (Fig. 1A) in ~20-40% of the arteries. Only preparations with more than ~90% spindle-shaped cells were chosen for incubations in these studies. Spindle-shaped cells, before permeabilization, always (>99% of cells) excluded trypan blue. When freshly isolated VSM cells were incubated with dextran sulfate in buffer A, the permeabilized cells did not exclude trypan blue (Fig. 1B), indicating that they were permeable to molecules of molecular weight of at least ~1,000. Therefore, glycolytic intermediates, with a maximum molecular weight of 406, would be expected to diffuse out of dextran sulfate-permeabilized cells. Permeabilized cells exhibited a less elongate shape with a poorly defined cell boundary (Fig. 1B) compared with freshly isolated cells. Thus treatment of VSM cells with dextran sulfate results in a permeable plasma membrane that should allow diffusion of glycolytic intermediates.


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Fig. 1.   Digital images of freshly isolated vascular smooth muscle (VSM) cells (A; ×400 magnification) and freshly isolated VSM cells permeabilized with dextran sulfate in buffer B (B; ×100 magnification, each narrow vertical bar represents 50 µm; see MATERIALS AND METHODS) in presence of 0.1% trypan blue.

Cells incubated with 5 mM [1-13C]glucose in either buffer B or buffer K were able to carry out glycolysis, since [3-13C]lactate production was observed. Shown in Fig. 2, top, is an example of a proton-decoupled 13C NMR spectrum of the incubation medium of permeabilized cells incubated for 3 h in the presence of buffer B containing 5 mM [1-13C]glucose. The peaks that are not marked in Fig. 2, top, correspond to the naturally abundant 13C in compounds in buffer B. The marked peaks correspond to the 13C-labeled substrate ([1-13C]glucose) and the product of its metabolism ([3-13C]lactate), as well as the peaks from the chemical shift reference DSS.


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Fig. 2.   Top: example of 13C nuclear magnetic resonance (NMR) spectrum of suspension solution (buffer B) sampled at end of 3-h incubation of permeabilized cells with [1-13C]glucose. Resonance in spectrum from suspension solution incubated with cells is at 22.7 parts per million (ppm), corresponding to [3-13C]lactate. In spectra in which other new resonances were detected (see RESULTS), resonances were not visually discernible because signal-to-noise ratio of spectrum allowed detection of signals that were too small to be visualized on same scale as major resonances. DSS, 3-(trimethylsilyl)-1-propane-sulfonic acid or TMSPS. Bottom: example of 13C NMR spectrum of suspension solution (buffer B) before add3ition of cells.

The [3-13C]lactate production of permeabilized cells depicted in Fig. 2, top, is the result of metabolism of [1-13C]glucose, since the [3-13C]lactate was not initially present in buffer B. Shown in Fig. 2, bottom, is a 13C NMR spectrum of buffer B that had not been exposed to cells. The only new resonance that is visually discernible, comparing Fig. 2, top, with Fig. 2, bottom, is [3-13C]lactate in Fig. 2, top. In addition, when buffer B was incubated for 3 h at 37°C without cells, no [3-13C]lactate production was observed (data not shown). Therefore, the [3-13C]lactate production from permeabilized cells was a new product of [1-13C]glucose metabolism by the cells and not due to microbial growth.

Shown in Fig. 3 is the mean [3-13C]lactate production from permeabilized cells and unpermeabilized cells incubated with [1-13C]glucose in either buffer B or buffer K. Lactate production occurred in both permeabilized and unpermeabilized cells, although permeabilization resulted in a 45.3% (significant) and 19.3% (insignificant) decrease in normalized lactate production in buffer B and buffer K, respectively.


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Fig. 3.   [3-13C]lactate production per 106 cells for cell suspensions incubated in either buffer B (shaded bars) or buffer K (solid bars) (see MATERIALS AND METHODS). Cells were either permeabilized with dextran sulfate or unpermeabilized. Data are means ± SE (n = 3) for lactate peak intensity at end of 3-h incubation. [3-13C]lactate peak intensity was normalized to peak intensity of DSS peak at 0 ppm to account for sensitivity changes in NMR spectrometer, since DSS is used as internal standard and chemical shift reference. [3-13C]lactate peak intensities are also normalized to total cell number in incubation.

If glycolytic intermediates or cofactors were able to freely leave the permeabilized cells, they would be diluted by a factor of ~500 (see DISCUSSION) and lactate production should virtually cease. To determine whether any of the 13C-labeled glycolytic intermediates produced during the conversion of [1-13C]glucose to [3-13C]lactate were diffusing from the permeabilized cells, we concentrated the incubation media and detected 13C-labeled molecules with high-resolution 13C NMR spectroscopy (for example, see Fig. 2, top). The only significant new peak in the 13C NMR spectra from incubations of permeabilized cells was [3-13C]lactate. Table 1 lists the expected chemical shifts of 1-13C-labeled hexoses and 3-13C-labeled trioses of the glycolytic pathway. The chemical shifts of most of the intermediates were determined in our laboratory with 2H2O as a solvent or with PSS as a solvent. Shown in Fig. 4 is a diagrammatic representation of the expected peak positions of 13C-labeled glycolytic intermediates and the observed positions of all new peaks in concentrated media from incubations of permeabilized cells in either buffer B or buffer K. The only new peaks that were sufficiently close in position (within 0.4 ppm) to the peak positions of glycolytic intermediates were peaks at ~64.9 ppm, which approximately correspond in position to fructose-6-phosphate (65.3 ppm). In addition, because glucose-6-phosphate and glucose have the same peak positions (see Table 1) we are unable to determine whether glucose-6-phosphate left the permeabilized cells. Therefore, within the limits of the signal-to-noise ratio of the high-resolution 13C NMR spectra, we were unable to detect any 13C-labeled glycolytic intermediates in the incubation solution, with the possible exception of fructose-6-phosphate.

                              
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Table 1.   13C NMR shifts for glycolytic intermediates


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Fig. 4.   Diagrammatic representation of positions of resonances in suspension solution with [1-13C]glucose but without incubation with cells (above axis) and of positions of new resonances resulting from incubation of solution with cells (below axis) from 0 to 120 ppm. Expected positions for glucose, lactate, and fructose-6-phosphate are indicated. Resonances from [1-13C]glucose and [3-13C]lactate were observed in all incubations, whereas resonances with chemical shift similar to fructose-6-phosphate (within 0.4 ppm) were observed in 2 incubations (with peak intensity that averaged <<FR><NU>1</NU><DE>7</DE></FR> of [3-13C]lactate peak intensity).

In Fig. 4 we diagramatically represent new resonances that appear in the incubation solutions of permeabilized cells. The peak intensities were so small that on a spectrum plotted full scale none of the new peaks were visually detectable, with the exception of [3-13C]lactate (compare Fig. 2, top and bottom, for example). To determine whether the small new resonances were due to metabolism of [1-13C]glucose or its metabolites or due to metabolism of buffer components (with a natural abundance of 13C of ~1.1%), we incubated permeabilized cells in buffer B with unlabeled glucose instead of [1-13C]glucose (Fig. 5). No [3-13C]lactate production was observed (since naturally abundant 13C in glucose is ~100-fold lower in label than in the [1-13C]glucose), and no new resonances were detected. Therefore, both the [3-13C]lactate and the barely detectable new peaks (depicted diagrammatically in Fig. 4) resulted from the metabolism of [1-13C]glucose.


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Fig. 5.   Example of 13C NMR spectrum of suspension solution (buffer B) sampled at end of 3-h incubation of permeabilized cells with 5 mM unlabeled glucose instead of [1-13C]glucose.

Because 13C-labeled glycolytic intermediates were not detectable in the bathing medium of intact cells (Figs. 2, top, and 4), we examined the 13C NMR spectrum of the pellet of intact cells incubated with [1-13C]glucose in buffer B. Shown in Fig. 6 is the 13C NMR spectrum of the cell pellet with 1.2% Triton X-100 added to release any intermediates into the 800 µl of solution, maximizing detection in the NMR spectrometer. No peaks corresponding to glycolytic intermediates were observed in the spectrum. However, new resonances, likely corresponding to Triton X-100 or cellular lipid constituents, were observed.


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Fig. 6.   13C NMR spectrum of pellet of permeabilized VSM cells sampled at end of 3-h incubation with 5 mM [1-13C]glucose. Pellet was resuspended in 800 µl of 2H2O and treated with 1.2% Triton X-100. No 13C-labeled glycolytic intermediates were detectable (see text for details).

To verify that complete cell disruption should prevent metabolism of 5 mM [1-13C]glucose, we performed one experiment with freshly isolated VSM cells that were treated with 0.1% Triton X-100 and sonicated (see MATERIALS AND METHODS). Shown in Fig. 7 is the 13C NMR spectrum of the medium from the incubation of cells treated with Triton X-100, sonicated, and then incubated with 5 mM [1-13C]glucose. No [3-13C]lactate or 13C-labeled glycolytic intermediates are observed. Therefore, the release of glycolytic intermediates, enzymes, and cofactors from the disrupted cells resulted in no detectable glycolytic flux.


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Fig. 7.   Example of 13C NMR spectrum of suspension solution (buffer B) sampled at end of 3-h incubation with [1-13C]glucose with cells disrupted by Triton X-100 and sonication (see MATERIALS AND METHODS). No resonance was observed from [3-13C]lactate (22.7 ppm) or from other 13C-labeled glycolytic intermediates.

    DISCUSSION
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

The current study was designed to determine whether the intermediates of glycolysis are channeled in the cytoplasm of VSM. VSM has been reported to exhibit a compartmentation of metabolism, with the intermediates of glycolysis and glycogenolysis exhibiting incomplete mixing (6, 18) and the intermediates of glycolysis and portions of gluconeogenesis exhibiting incomplete mixing (11, 12). It has been proposed (8) that the incomplete mixing of the intermediates for the pathways for glycolysis and glycogenolysis may result from a differential localization of the enzymes, since glycolytic enzymes have been localized to the contractile apparatus (7) and the plasma membrane (9, 24) in smooth muscle. In addition, in other cell types, glycolytic enzymes have been localized to the actin cytoskeleton (20, 21), microtubules (15, 19), and outer mitochondrial membrane (2, 17). Therefore, because glycolytic enzymes are localized to a variety of cytoplasmic structures and because there appears to be a lack of mixing of glycolytic intermediates in the cytoplasm, it might be predicted that glycolytic enzymes do not readily leave the glycolytic, glycogenolytic, or gluconeogenic "compartments" within the cytoplasm.

To directly determine whether the intermediates of glycolysis were channeled, as might be inferred by the compartmentation studies previously reported (6, 11, 12, 18), we permeabilized hog carotid artery VSM cells with dextran sulfate according to the methods of Clegg and Jackson (4). This permeabilization procedure has been shown to allow proteins of ~400 kDa to cross the cell membrane (4). However, despite the permeability of the plasma membrane to macromolecules, in the studies of Clegg and Jackson (4) only 10% of the total protein of the cell left the cell during 30 min of incubation. When we adapted the Clegg and Jackson (4) permeabilization treatment to freshly isolated VSM cells, we found that none of the cells in suspension were capable of excluding trypan blue, indicating that a molecule of molecular weight of at least 1,000 was capable of crossing the plasma membrane (Fig. 1B). Molecular weight alone is only a rough indicator of permeability through pores of a given size. Additional factors such as charge density and hydrated radius are more accurate measures of permeability. However, because trypan blue has more than twice the molecular weight of the largest glycolytic intermediate and because dextran sulfate-permeabilized cells have been shown to allow large proteins to readily leave the cell (4, 5), we felt that trypan blue exclusion was a reasonable indicator of cell permeability in a large population of cells.

Because the largest glycolytic intermediate, fructose-1,6-bisphosphate, has a molecular weight of 406 in the trisodium form, it might be expected that all glycolytic intermediates would diffuse from the permeabilized cell if the intermediates were freely diffusible. However, 13C-labeled glycolytic intermediates derived from [1-13C]glucose metabolism were not detectable in the cell suspension solution, as measured by high-resolution 13C NMR spectroscopy (Fig. 4). The single exception is the resonance at ~64.9 ppm, which may correspond to fructose-6-phosphate (65.3 ppm). This resonance was found in two samples and had a peak intensity less than one-seventh of that from [3-13C]lactate. Because the [3-13C]lactate peak intensity had a signal-to-noise ratio of >40:1, a glycolytic intermediate with a production rate of 4% of the lactate production rate should be detectable. Therefore, glycolytic intermediates derived from [1-13C]glucose metabolism did not diffuse from the cell (except for perhaps fructose-6-phosphate), even though the permeabilized cells were carrying out 54.7 and 80.7% of the glycolytic rate of the unpermeabilized cells (in buffer B and buffer K, respectively; Fig. 3). These findings are generally consistent with those of Clegg and Jackson (5), who found that addition of unphysiologically high concentrations of unlabeled glycolytic intermediates to cells incubated with [14C]glucose did not decrease the specific activity of the lactate produced. In the current study, we provide evidence of channeling of intermediates of glycolysis when the glycolytic intermediates are at their physiological concentration, which is similar to the concentration of the enzymes (25). The lactate production significantly declined in permeabilized cells in buffer B compared with intact cells in buffer B. This decline in lactate production in permeabilized cells was not likely due to intermediates leaving the permeabilized cell but to a decrease in glycolytic flux, perhaps due to either diffusion of glycolytic cofactors or a change in pathway signals resulting from the permeabilization. This is consistent with most intermediates being of similar concentration as the enzymes and thus almost entirely bound within the cell (25).

In these studies, no attempt was made to replenish glycolytic cofactors such as NAD, NADH, ADP, and Pi that may have exited the cell following permeabilization. Sufficient pools of cofactors must have remained within the permeabilized cells to allow glycolytic flux. Most likely, the cofactors were continuously associated with the enzymes of glycolysis. Experimentally, a direct transfer resulting in a recycling of NAD/NADH between glyceraldehyde-3-phosphate dehydrogenase and lactate dehydrogenase has been reported (see Ref. 25 for a review). Therefore, retention of cofactors, like retention of glycolytic intermediates, may be due to a direct transfer or recycling and not necessarily due to restricted diffusion of small molecules not associated with enzymes. Although we cannot rule out a severe restriction of small charged molecules in the permeabilized cells, dextran sulfate-treated L-929 cells have been shown to be permeable to large proteins and unbound glycolytic cofactors (4, 5). Indeed, in L-929 cells, dextran-sulfate permeabilized cells required exogenously added glycolytic cofactors for glycolytic flux to occur (4, 5). Therefore, dextran sulfate likely allows diffusion of charged molecules. In addition, the buffers chosen (from Refs. 4 and 5) were designed to have a high pH to stimulate glycolytic flux and a high K+ concentration to help stabilize enzyme activity. That glycolysis substantially occurred in VSM cells in the absence of exogenous glycolytic cofactors indicates that sufficient enzyme-bound pools of the cofactors remained following permeabilization of freshly isolated VSM cells but not following permeabilization of L-929 cells. This may reflect a greater number of binding sites resulting in greater retention of enzymes and thus a greater retention of glycolytic intermediates and cofactors in VSM cells compared with L-929 cells specifically. Because glycolytic enzymes have been shown to bind to the contractile apparatus (7), the plasma membrane (9, 24), the actin cytoskeleton (20, 21), and the microtubules (15, 19) of a variety of cells, the number of glycolytic enzyme-binding sites may vary with cell phenotype, cell cycle stage, or metabolic state. Indeed, this increased retention of glycolytic enzymes may represent a common characteristic of noncultured cells, which retain more physiological characteristics than cultured cells.

It should be noted that, although the newly detected peaks in the solutions of permeabilized cells are not glycolytic intermediates, they do appear to be derived from [1-13C]glucose, since the new peaks are not formed when [1-13C]glucose is excluded from the incubation (Fig. 5). Indeed, two-thirds of the new peaks found in the solutions from incubations of permeabilized cells were also found in the solutions from incubations of unpermeabilized cells (data not shown). Therefore, the very small new peaks may be due to a very slight extent of extracellular glucose metabolism in both intact and permeabilized cells.

To rule out the possibility that the lactate production observed in the permeabilized cell suspensions (Fig. 3) was due to activity of released glycolytic enzymes, we disrupted the cells and the cellular contents by treatment of the cell suspension with 1% Triton X-100 and sonication. Cells treated with detergent and sonication were likely to have been completely disrupted and to have released enzymes and cofactors into the surrounding solution. When sonicated cells were incubated with [1-13C]glucose for 3 h, no lactate production or accumulation of any glycolytic intermediates was observed (Fig. 7). The abolition of glycolytic flux is probably due to the loss of cellular organization and to the consequent large dilution of enzymes, intermediates, and cofactors. Assuming each cell is 7 µm in diameter and 100 µm in length, then the approximate volume of each cell is 0.00385 nl. Typically, ~1.2 × 106 cells were used per experiment in an incubation volume of ~2 ml. Therefore, the total volume of the cells per incubation was ~4.6 µl, whereas the total volume of the suspension was ~2 ml. Therefore, for permeabilized cells, if small molecules were readily diffusible, the small molecules would be diluted ~500-fold. The 500-fold dilution of enzymes, cofactors, and intermediates, as well as the loss of any additional organization these components may have in the cell, likely resulted in the absence of glycolytic flux.

The total [3-13C]lactate production from [1-13C]glucose is similar whether cells were permeabilized or intact. In addition, the rate of glycolysis, normalized to cell density, was similar among preparations in both treatment groups. Also, treatment of a VSM cell suspension with 1% Triton X-100 and sonication abolished all [3-13C]lactate production. Finally, incubation of buffer B or buffer K without cells resulted in no [3-13C]lactate production. Taken together, these results indicate that the [3-13C]lactate production was the result of VSM cell metabolism and not microbial contamination.

The [3-13C]lactate production from the cell suspensions is comparable to that produced by intact hog carotid artery. In one of our previous studies (11), we demonstrated that ~300 mg of hog carotid artery also incubated at 37°C for 3 h produced a [2-13C]lactate peak from [2-13C]glucose that was ~2.6-fold higher than the 0-ppm DSS peak. In our previous work (11) and in the current work, DSS was the same concentration (25 mM) in the NMR tube. If hog carotid artery tissue is assumed to have ~30% of its mass due to cells, then the 4.6 µl of cell volume would correspond to 15.3 mg of tissue. In the current study, permeabilized cells produced a [3-13C]lactate peak over 3 h that was 22% of the intensity of the 0-ppm DSS peak. The comparable tissue mass of the cell suspension (15.3 mg) is 5.1% of the tissue mass of the previously reported study (300 mg) and the lactate peak intensity from cells (scaled to DSS) is 8.5% of that produced from 300 mg of tissue. Although these calculations rely on a number of assumptions, they clearly demonstrate that the glycolytic rate of permeabilized cells is similar to the glycolytic rate in intact tissue.

A direct comparison of the glycolytic rate of intact vs. permeabilized cells is difficult in the current study (see Fig. 3). In the data presented, glycolytic rates are within a factor of two in these preparations, despite the likely potassium-induced depolarization in intact cells and the greater activation of permeabilized cells in buffer B (no EGTA) compared with buffer K (0.1 mM EGTA). This is consistent with the approximately twofold range of glycolytic rate observed in intact hog carotid artery under different activation conditions (see Refs. 8, 23). Although it is difficult to interpret the comparison of glycolytic flux in the permeabilized cells vs. intact cells, the important finding is that permeabilized cells had substantial glycolytic flux compared with intact cells and compared with intact tissue, despite being permeabilized.

The expected concentration of a given glycolytic intermediate within a cell is typically 20-100 µM (25). The typical total volume of cells in an incubation is ~4.6 µl in these studies. Therefore, in the 800 µl of solution used to suspend the cell pellet for 13C NMR measurements, the concentration of a given glycolytic intermediate is diluted ~175-fold and thus is ~0.1-0.6 µM in the NMR tube. Not surprisingly, no 13C-labeled glycolytic intermediates were detected in the cell pellet (Fig. 6), even after 20,480 scans.

We have previously shown that glycolytic enzymes are associated with the plasma membrane of smooth muscle (9). We have also reported that glycolysis and glycogenolysis are compartmentalized, with incomplete mixing of pathway intermediates (6), and that glycolysis and glycogenolysis can be independently regulated (13, 14). The results from the current study using hog carotid artery smooth muscle cells are consistent with one possible mechanism of the compartmentation of glycolysis, namely a strict channeling of glycolytic intermediates so that the intermediates have little or no chance to leave the compartment. However, strict channeling of glycolytic intermediates from glucose to lactate in this tissue does not always occur, since exogenous glucose can be utilized for glycogen synthesis (6) and pyruvate from glucose can be oxidized (see Refs. 8 and 23). Therefore, although speculative, organized glycolysis may exist in smooth muscle in the form of strict channeling and in some other form that allows intermediates to diffuse away from the glycolytic enzymes. Alternatively, the substrate channeling may be modulated by factors such as metabolic state. Distinguishing these mechanisms is not possible with the current experimental approach.

In conclusion, we have adapted a cell permeabilization protocol from Clegg and Jackson (4, 5) to VSM cells. We have demonstrated that these permeabilized cells can continue to carry out glycolysis, as measured by [3-13C]lactate production from [1-13C]glucose. By use of high-resolution 13C NMR spectroscopy, we were able to directly measure which, if any, glycolytic intermediates exit permeabilized cells. We found that the only glycolytic intermediate that may have exited the permeabilized cells was fructose-6-phosphate. Complete disruption of the cells abolished glycolytic flux, consistent with the dilution of the glycolytic enzymes, cofactors, and intermediates. We conclude that VSM cells exhibit channeling of glycolytic intermediates. We have previously shown that glycolytic enzymes are bound to the plasma membrane of VSM (9, 24), and we had proposed that glucose catabolism to lactate may occur via a membrane-associated set of glycolytic enzymes (8). Glycolytic flux can occur even with significant disruption of the plasma membrane, indicating that either sufficient glycolytic enzymes remain bound to the plasma membrane to carry out glycolytic flux or glucose metabolism in permeabilized cells is carried out by enzymes located elsewhere in the cytoplasm. Because glycolytic enzymes are localized to a variety of cytoplasmic structures and because channeling of pathway intermediates has now been demonstrated in VSM, future studies should be aimed at elucidation of the specific cytoplasmic structures responsible for the organization of the separate pathways of glycolysis, glycogenolysis, and gluconeogenesis in VSM.

    ACKNOWLEDGEMENTS

This work was supported by National Heart, Lung, and Blood Institute Grant HL-48783, a Grant-in-Aid, and an Established Investigator Grant from the American Heart Association, all to C. D. Hardin. D. Finder was supported, in part, by the University of Missouri School of Medicine Research Council and by a Hughes Research Internship. The support of National Science Foundation Instrumentation Grant 8908304 is also acknowledged. Hog carotid arteries were graciously provided by Excel, Inc. (Marshall, MO).

    FOOTNOTES

Address for reprint requests: C. D. Hardin, Dept. of Physiology, MA-415 Medical Sciences Bldg., University of Missouri, Columbia, MO 65212.

Received 21 February 1997; accepted in final form 11 September 1997.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

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