Department of Physiology, University of Missouri-Columbia, Columbia, Missouri 65212
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ABSTRACT |
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We used 13C-labeled substrates and nuclear magnetic resonance spectroscopy to examine carbohydrate metabolism in vascular smooth muscle of freshly isolated pig cerebral microvessels (PCMV). PCMV utilized [2-13C]glucose mainly for glycolysis, producing [2-13C]lactate. Simultaneously, PCMV utilized the glycolytic intermediate [1-13C]fructose 1,6-bisphosphate (FBP) mainly for gluconeogenesis, producing [1-13C]glucose with only minor [3-13C]lactate production. The dissimilarity in metabolism of [2-13C]FBP derived from [2-13C]glucose breakdown and metabolism of exogenous [1-13C]FBP demonstrates that carbohydrate metabolism is compartmented in PCMV. Because glycolytic enzymes interact with microtubules, we disrupted microtubules with vinblastine. Vinblastine treatment significantly decreased [2-13C]lactate peak intensity (87.8 ± 3.7% of control). The microtubule-stabilizing agent taxol also reduced [2-13C]lactate peak intensity (90.0 ± 2.4% of control). Treatment with both agents further decreased [2-13C]lactate production (73.3 ± 4.0% of control). Neither vinblastine, taxol, or the combined drugs affected [1-13C]glucose peak intensity (gluconeogenesis) or disrupted the compartmentation of carbohydrate metabolism. The similar effects of taxol and vinblastine, drugs that have opposite effects on microtubule assembly, suggest that they produce their effects on glycolytic rate by competing with glycolytic enzymes for binding, not by affecting the overall assembly state of the microtubule network. Glycolysis, but not gluconeogenesis, may be regulated in part by glycolytic enzyme-microtubule interactions.
vascular smooth muscle; carbohydrate metabolism; tubulin; glycolytic enzymes; carbon-13 nuclear magnetic resonance; taxol; vinblastine; cytoarchitecture; gluconeogenesis
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INTRODUCTION |
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IN THE PAST, the glycolytic enzymes and their substrates were thought to be freely diffusible within the aqueous phase of the cell, due to their presence in high-speed supernatants ("cytosol"; see Ref. 26). It is becoming apparent that the cytoplasm of intact cells is considerably more organized than was previously thought (for a review, see Ref. 8). This model of the organized cytoplasm is consistent with evidence demonstrating that many of the glycolytic enzymes associate specifically with cellular structures. Glycolytic enzymes interact with membranes (for a review, see Ref. 44) and membrane proteins (band 3; see Refs. 28 and 29). Glycolytic enzymes also bind to F-actin (2-4, 25, 32) and microtubules (23, 27, 31, 33, 51).
Vascular smooth muscle (VSM) is a useful model for the study of carbohydrate metabolism, since it maintains a high rate of glycolysis even when well supplied with oxygen (15). In addition, gluconeogenesis (18, 19) and glycogenolysis (13, 14) occur in this tissue. These three carbohydrate metabolizing pathways (glycolysis, glycogenolysis, gluconeogenesis) contain many enzymes and intermediates that are identical, yet incomplete mixing of the intermediates of glycolysis and glycogenolysis (13, 30) and glycolysis and gluconeogenesis (18) have been observed in VSM. Thus a compartmentation of carbohydrate metabolism exists in VSM. Localization of the enzymes specific to each pathway on spatially separate cellular structures may be the mechanism by which carbohydrate metabolism is organized in this tissue. One functional consequence of this organization may be that energy is produced near energy-consuming sites. For example, membrane-associated glycolytic enzymes appear to produce ATP that is utilized preferentially by the plasma membrane Ca2+-ATPase of pig stomach (16). Similarly, plasma membrane ATP-sensitive K+ channels in guinea pig ventricular myocytes are closed more effectively by glycolytic ATP than ATP generated by oxidative metabolism (52).
Although the association of the enzymes of carbohydrate metabolism to cellular structures appears likely to have physiological significance, it remains unknown whether these associations are involved in producing compartmentation of carbohydrate metabolism or in regulating pathway flux. Thus the present study was undertaken to determine whether there is a structural basis for compartmentation and whether the association of enzymes with microtubules alters pathway flux. We hypothesized that interactions between glycolytic enzymes and microtubules represent one structural basis for compartmentation of carbohydrate metabolism in VSM of pig cerebral microvessels (PCMV). To test this hypothesis, we examined the effects of both microtubule disruption and stabilization on glycolysis and gluconeogenesis in VSM of PCMV. Our results suggest that the binding of glycolytic enzymes to tubulin, rather than the integrity of the microtubule network, plays a significant role in the regulation of glycolytic flux. However, binding of gluconeogenic enzymes to tubulin does not contribute to the regulation of gluconeogenic flux. Thus these pathways may have spatially separate locations within the cytoplasm. The association of glycolytic enzymes with microtubules appears to have functional significance in terms of the regulation of pathway flux.
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MATERIALS AND METHODS |
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Tissue Collection
Pig brains were obtained at a local abbatoir within 30 min of slaughter. Brains were placed in ice-cold physiological saline solution (PSS) for transport to the laboratory. PSS consisted of (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. In addition, PSS was filtered through a 0.22-µm nylon filter before use (Micron Separations, Westboro, MA). At the laboratory, brains were stored in fresh PSS at 4°C until use.Microvessel Isolation
Microvessels were isolated by a modification of the method of Sussman et al. (43). Brains were kept wet at all times with 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 two halves of the brain were gently separated. The outer layers of the brain, containing large blood vessels, were carefully peeled away to expose the cerebral cortex. The gray matter was dispersed by aspiration into a plastic vacuum flask. Aspirated material was further dissociated by five strokes in a stainless steel Dounce-type homogenizer (Dura-Grind; Thomas Scientific, Swedesboro, NJ) and then poured over a 210-µm nylon mesh (Small Parts, Miami Lakes, FL) placed above a plastic container. Large vessels were trapped on the mesh while small vessels and debris passed through into a container placed below the mesh. Material passing through the 210-µm mesh was saved for the next isolation step. The large vessels trapped on the mesh were rinsed several times with HBPSS. Next, the mesh was inverted, and the adherent vessels were washed off into a clean container using HBPSS. These vessels represented the largest size fraction.
The material that was collected after passing through the 210-µm mesh contained vessels too small to be trapped by the mesh. This material was now passed over a 105-µm mesh, which trapped vessels of intermediate size while allowing smaller vessels and debris to pass through into a container placed below the mesh. Material passing through the 105-µm mesh was saved for the next isolation step. The vessels adhering to the mesh were rinsed with HBPSS and then collected by inverting the mesh and rinsing the vessels off into a clean container. These vessels represented an intermediate size fraction.
The brain material remaining after filtration with the 105-µm mesh contained vessels too small to be trapped by either the 210- or 105-µm mesh and debris. This material was filtered a final time using a 35-µm mesh to collect these small vessels. Vessels trapped on the mesh were rinsed with HBPSS and then were collected by inverting the mesh and washing the vessels off. Material passing through the 35-µm mesh was discarded.
Microvessels collected from the 210-, 105-, and 35-µm meshes were used for experiments on microvessel size. All other experiments used only vessels from the 105-µm mesh. When examined microscopically, the material collected from each mesh size consisted of microvessels, with virtually all debris having been removed by the rinsing process.
Microvessel Size Experiments
Previous work in our laboratory utilizing 13C-labeled substrates demonstrated that VSM of hog carotid artery metabolize fructose 1,6-bisphosphate (FBP) largely to glucose (gluconeogenesis) while simultaneously metabolizing glucose largely to lactate (glycolysis; see Ref. 18). Thus glycolysis and gluconeogenesis are compartmentalized in hog carotid artery smooth muscle. Although a compartmentation of glycolysis and gluconeogenesis might also be expected in PCMV, this tissue has both smooth muscle cells (the major cellular component) and endothelial cells, and thus any observed compartmentation may be intercellular and not intracellular. Because microvessels of different sizes will have markedly different contributions of endothelial cell mass, microvessels of three size classes (and therefore varying percentages of endothelium) were studied to determine if endothelial cell metabolism contributes significantly to overall observed metabolism.Microvessels were isolated from 3.5 brains for each microvessel size experiment. Vessels adhering to the 210-, 105-, and 35-µm meshes were collected, poured over meshes (210, 105, or 35 µm, as appropriate), and rinsed with HBPSS. The meshes were inverted, and the microvessels were flushed into clean plastic beakers using 9 ml of HBPSS containing 5 mM [1-13C]FBP (Omicron Biochemicals, South Bend, IN) and 5 mM [2-13C]glucose (pH 7.4; Cambridge Isotope Laboratories, Andover, MA). The suspensions were mixed to ensure even distribution of microvessels, and 8 ml were transferred to 25-cm2 polystyrene cell culture flasks (Corning Costar, Cambridge, MA). The flasks were placed in a shaking bath and were incubated for 3 h at 37°C. At the conclusion of the incubation, the vessel suspension was mixed to ensure homogeneity. A 5.5-ml sample of the vessel suspension was removed from each flask. The samples were placed in 15-ml centrifuge tubes and centrifuged (Marathon 6K; Fisher Scientific) at 1,000 g for 5 min to pellet the microvessels. The resulting supernatant (4 ml) was frozen in liquid nitrogen and was saved for nuclear magnetic resonance (NMR) analysis. A 4-ml sample of the starting solution containing labeled substrates was also saved for NMR analysis.
Metabolic Experiments
Vinblastine. Microvessels were isolated from three brains for each metabolic experiment. For these studies, microvessels were collected from the 105-µm mesh only. The microvessel suspension was brought to a total volume of 100 ml using HBPSS and was mixed thoroughly. The suspension was then split into two 50-ml aliquots before any treatment was administered, allowing for a self-paired design. One aliquot was poured over a 105-µm mesh and rinsed lightly with HBPSS. The mesh was then inverted over a plastic beaker, and the vessels were rinsed off using 50 ml of HBPSS containing 4 mM FBP (Esafosfina, Biomedica Foscama; gift from Dr. Guiseppe Lazzarino) and 5 mM glucose, pH 7.4. This suspension was poured in a 50-ml centrifuge tube and was placed in a shaking water bath at 37°C for 1.5 h. This group of microvessels served as the control.The second aliquot of microvessels was treated as above except that HBPSS contained vinblastine sulfate (5 µM) to disrupt microtubules, in addition to FBP and glucose. Vinblastine was prepared as a concentrated stock solution (10 mM). In some experiments, the order of preparation was reversed so that the first aliquot received vinblastine treatment, whereas the second served as the control.
After the initial 1.5-h incubation, each of the vessel suspensions was mixed and again poured over a 105-µm mesh. The microvessels on the meshes were rinsed with substrate-free HBPSS to remove unlabeled FBP and glucose. Next, the meshes were inverted. The microvessels were flushed into clean plastic beakers using 9 ml of HBPSS containing 5 mM [1-13C]FBP and 5 mM [2-13C]glucose (pH 7.4). The resulting suspensions (8 ml) were incubated in culture flasks for 3 h at 37°C. The flask containing vinblastine-treated vessels also contained 4 µl of vinblastine stock solution, resulting in a concentration of 5 µM.
Excess vessel suspension from both the control and vinblastine-treated groups (~1 ml) was used for indirect immunofluorescence to determine the level of microtubule disruption induced by vinblastine (see Indirect Immunofluorescence). Examination of control and treated vessels using transmitted light did not reveal any differences in appearance between groups.
At the conclusion of the 3-h incubation, the material in each flask was thoroughly mixed to ensure homogeneity of the suspension. A 3.5-ml sample of the suspension was removed from each flask. The samples were placed in 15-ml centrifuge tubes and centrifuged (Marathon 6K; Fisher Scientific) at 1,000 g for 5 min to pellet the microvessels. A 3-ml aliquot of each supernatant was placed in a cryovial and was frozen in liquid nitrogen for NMR analysis. In addition, a 3-ml sample of the starting solution containing labeled substrates was saved for NMR analysis. Excess vessel suspension remaining in the flasks after removal of the 3.5-ml sample was used for indirect immunofluorescence to verify the disruption of microtubules by vinblastine.
Metabolic Experiments
Taxol and vinblastine. Experiments to examine the effects of taxol alone and in the presence of vinblastine were performed as described above, except the initial microvessel suspension was divided into three equal parts. Two of the aliquots were treated with the microtubule-stabilizing agent taxol (10 µM) for 30 min at 37°C in the presence of unlabeled FBP and glucose as described above. Taxol was prepared as a concentrated stock solution (10 mM, in DMSO). After this initial drug treatment, vinblastine sulfate (5 µM) was added to one of the taxol-treated aliquots, and all three aliquots were incubated for an additional 1.5 h at 37°C. Metabolic incubations (3 h) with labeled substrates were then performed as described above, with appropriate additions of taxol or taxol plus vinblastine to the incubation medium. One of the three aliquots received additions of DMSO alone (control).NMR Spectroscopy
The supernatant solutions from microvessel size experiments and metabolic experiments (and the starting solution for each experiment) were lyophilized to powder in a Speed Vac (Savant Instruments, Farmingdale, NY). Immediately before NMR analysis, each sample was suspended in 800 µl of 99.9% D2O (Cambridge Isotope Laboratories) containing 25 mM 3-(trimethylsilyl)-1-propanesulfonic acid (TMSPS) as a chemical shift reference, and 650 µl of this solution were transferred to a 5-mm NMR tube.13C-NMR was performed using a Bruker DRX 500 spectrometer. For microvessel size experiments and vinblastine experiments, data were acquired with the following parameters: 300 scans plus 64 dummy scans, 30° pulse angle at 125.77 MHz, 33,333 Hz sweep width, and 1 s predelay. For experiments in which both taxol and vinblastine were used, samples were subjected to 600 scans plus 64 dummy scans to increase the signal-to-noise ratio (which was decreased by splitting the vessel suspension into 3 rather than 2 parts). Points (32,768) were acquired and processed with line broadening of 1 Hz before Fourier transform of the data. Spectra were broad-band proton decoupled, and peaks were referenced to the signal of TMSPS, set at 0 ppm. Data were processed for determination of peak intensity using Bruker software. No corrections for nuclear Overhauser effects were made, as these were assumed to be the same for all experiments. All peak intensities were normalized to the intensity of TMSPS before analysis. Significant differences in the metabolism of the three vessel size classes were determined by calculating the ratio of [2-13C]- and [3-13C]lactate to [1-13C]glucose and comparing the values using two-tailed t-tests for two samples, assuming equal variances. Significant differences between 13C-NMR peak intensities of control and vinblastine-treated microvessels and between control, taxol, and taxol plus vinblastine-treated microvessels 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.
Criteria for Data Inclusion
In 3 of the 13 vinblastine experiments, the signal-to-noise ratio of the [1-13C]glucose peak at 94.8 ppm was less than three. The data from these experiments supported our overall conclusions but were excluded from the analysis due to the difficulty of precise quantification of such small peaks.Indirect Immunofluorescence
Microvessels from metabolic experiments were fixed and stained for tubulin to determine the effects of vinblastine and taxol on microtubule organization in this tissue. Samples of control, vinblastine-treated, taxol-treated, and taxol plus vinblastine-treated microvessels were collected after the 1.5-h initial incubation with vinblastine and the 3-h metabolic incubation (1.5- and 4.5-h total exposure to vinblastine). Samples were collected from 5 of the 10 vinblastine experiments and 2 of the 5 taxol and vinblastine experiments.The microvessels were poured onto a 105-µM mesh and were washed with
PBS to remove excess albumin before being transferred to glass
coverslips coated with Cell-Tak (Collaborative Biomedical Products,
Bedford, MA). Microvessels remained attached to the coated coverslips
throughout the fixation procedure. The protocol for the visualization
of tubulin using indirect immunofluorescence was performed essentially
as described by Zhai et al. (55). Microvessels were extracted for 2 min
at room temperature in microtubule stabilizing buffer [60 mM
PIPES, 25 mM HEPES, 10 mM EGTA, and 2 mM
MgCl2, pH 6.9 (PHEM)]
containing 0.2% Triton X-100. After three rinses with PBS,
microvessels were fixed in PHEM containing 3.7% formaldehyde (30 min,
37°C). Microvessels were rinsed three times in PBS and then were
incubated in PBS containing 0.2% Triton X-100 (4 min, 21°C). The
rinsing procedure was repeated. Coverslips were next incubated in PBS
containing 10% normal goat serum to block nonspecific binding of the
secondary antibody (30 min, 37°C), followed by incubation in PBS
containing 1% normal goat serum and mouse monoclonal anti-bovine
-tubulin (1:200 dilution; overnight, 4°C; Molecular Probes,
Eugene, OR). After incubation with the primary antibody, coverslips
were rinsed three times in PBS and then were incubated in PBS
containing 1% normal goat serum and goat anti-mouse IgG conjugated to
Alexa 488 (1:200 dilution; 45 min, 37°C; Molecular Probes). After
incubation with the secondary antibody, coverslips were rinsed four
times in PBS and then mounted on glass slides with Mowiol 4-88
mounting medium (Calbiochem, La Jolla, CA).
Quantification of the Effects of Vinblastine on Microtubule Organization
Fluorescence images of the microtubule network were collected from 108 vessels in the following groups: 1.5-h control (n = 27), 4.5-h control (n = 28), 1.5-h vinblastine (n = 25), and 4.5-h vinblastine (n = 28). The images were displayed in random order to three unbiased observers who were unaware of the treatment each vessel had received. The observers were required to classify the staining pattern of each image as "filamentous" or "not filamentous." Significant differences between control and vinblastine-treated vessels at each time point were detected with a one-tailed t-test for two samples assuming equal variances. Values of P < 0.05 were considered significant. Statistical calculations were performed using Microsoft Excel 97 software.Determination of Cellular Components of Microvessels
Because microvessels contain a mixture of VSM cells and endothelial cells, the percentage of each cell type in this tissue was determined by immunofluorescence. Immunofluorescent labeling of smooth muscleDigital Imaging Microscopy
Immunofluorescence microscopy was performed on a wide-field epifluorescence microscope (Nikon Diaphot; Nikon, Garden City, NY) using a ×40 oil immersion objective (numerical aperture 1.3; Nikon). Images were captured by a liquid-cooled charge-coupled device camera (model ATC200L) equipped with a personal computer interface board and liquid recirculation unit (all from Photometrics, Tucson, AZ). Z-scans through the tissue utilized a z-stepper motor (Ludl Electronic Products, Hawthorne, NY). A 150-W xenon arc lamp (Oriel Instruments, Stratford, CT) was used as the light source. The wavelength of the excitation beam was selected using a Ludl 6-position filter wheel and controller (Ludl Electronic Products). Software components of the system included the programs MicroTome and Volume Scan (VayTek, Fairfield, IA) and ImagePro Plus (Media Cybernetics, Silver Spring, MD).All images were acquired using a triple-dye filter cube (XF56; Omega Optical, Brattleboro, VT). The cube allowed for band-pass excitation of 375 ± 20, 485 ± 15, and 575 ± 20 nm, band-pass emission of 452 ± 20, 528 ± 24, and 630 ± 44 nm, and contained 455, 530, and 620 nm dichroic mirrors. Thus DAPI (excitation 358 nm, emission 461 nm), FITC (excitation 494 nm, emission 519 nm) or Alexa 488 (excitation 491 nm, emission 515 nm), and Alexa 568 (excitation 573, emission 596 nm) could be detected in the same tissue sample.
Fluorescence images intended for deconvolution analysis were collected at three to five focal planes separated by 0.5 µm (vinblastine experiments) or 0.25 µM (taxol and vinblastine experiments). The corresponding transmitted light images were acquired at the middle of the z-series. Deconvolution analysis was performed using MicroTome and ImagePro Plus software and utilized the nearest-neighbor algorithm. Adobe Photoshop software was used to color and combine the fluorescence images.
Reagents
Na2HPO4 and NaH2PO4 were obtained from Aldrich Chemical (Milwaukee, WI). Except where stated, all other chemicals were purchased from Sigma Chemical. ![]() |
RESULTS |
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Glycolysis and Gluconeogenesis Occur Simultaneously in PCMV
PCMV that were incubated for 3 h with 5 mM [1-13C]FBP and 5 mM [2-13C]glucose (n = 3) consistently metabolized [1-13C]FBP mainly to [1-13C]glucose (gluconeogenesis) while simultaneously metabolizing [2-13C]glucose to [2-13C]lactate (glycolysis; Fig. 1). The production of [2-13C]lactate demonstrates that glycolysis is occurring in this tissue. However, PCMV produced only minor amounts of [3-13C]lactate from [1-13C]FBP while utilizing [1-13C]FBP for extensive production of [1-13C]glucose. Thus [1-13C]FBP is a much better substrate for gluconeogenesis than for glycolysis in PCMV, despite the fact that it is an intermediate of both pathways.
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Cellular Components of PCMV
Microvessels were immunostained for smooth muscle
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Metabolism Is Similar in Vessels of Three Different Sizes
Microvessels isolated from 210-, 105-, and 35-µm meshes were very similar in overall metabolism. Endothelium represents a substantially (~2-fold) greater percentage of total cell volume in smaller vessels than in larger vessels (see DISCUSSION). Thus endothelial metabolism would be expected to be substantially more evident in smaller vessels than in larger vessels. For example, if gluconeogenesis occurred in endothelium, while glycolysis occurred in VSM, then as vessel size decreased, the ratio of gluconeogenesis to glycolysis would increase by at least a factor of two. When the rate of gluconeogenesis relative to glycolysis is measured in PCMV of differing sizes, there is no significant difference in the rate of gluconeogenesis relative to glycolysis among the three size classes (Fig. 3). Thus the measured metabolism appears to occur in the predominant cell type, VSM.
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Vinblastine Treatment Disrupts Microtubule Organization in PCMV
Because glycolytic enzymes bind to microtubules (see DISCUSSION), we disrupted microtubules of PCMV with vinblastine and examined their metabolism to determine whether this association plays a role in compartmentation or regulation of glycolysis and gluconeogenesis. Microvessels were treated with 5 µM vinblastine for a total of 4.5 h (1.5 h before metabolic incubation and 3 h during metabolic incubation). At low concentrations, vinblastine depolymerizes microtubules as minus ends; at higher concentrations, it depolymerizes microtubules at both ends (38). To verify that vinblastine caused dissociation of microtubules in PCMV, samples of vinblastine-treated and control microvessels were collected at the 1.5-h (Fig. 4) and 4.5-h (Fig. 5) time points during five metabolic experiments. Microvessels were attached to coverslips, then fixed and stained for
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Vinblastine Treatment Differentially Affects Glycolysis and Gluconeogenesis
Metabolic incubations with 13C-labeled substrates were performed both in the presence and absence of 5 µM vinblastine to determine whether microtubules play a role in the compartmentation or regulation of glycolysis and gluconeogenesis observed in VSM of PCMV (n = 10). After microtubule disruption, PCMV continued to metabolize [1-13C]FBP mainly to [1-13C]glucose while simultaneously metabolizing [2-13C]glucose to [2-13C]lactate. Thus dissolution of the microtubule network did not result in a breakdown in compartmentation in this tissue. However, microtubule disruption differentially affected glycolysis and gluconeogenesis. Microtubule disruption had no effect on the rate of gluconeogenesis; the [1-13C]glucose peak intensity in treated vessels was 100.9 ± 5.7% of the peak intensity in self-paired controls (mean ± SE). In contrast, vinblastine treatment significantly (P = 0.04) reduced the intensity of the [2-13C]lactate peak to 87.8 ± 3.7% of self-paired controls (mean ± SE; Fig. 7).
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Taxol Treatment Inhibits Vinblastine-Induced Disruption of Microtubules
Because it is well known that taxol induces microtubule assembly and inhibits microtubule depolymerization (22), we treated PCMV with taxol (10 µM) to determine whether stabilizing the microtubule network would produce effects opposite to those of vinblastine. In addition, we treated PCMV with taxol followed by vinblastine to determine whether taxol-induced microtubule stabilization could inhibit microtubule disruption by vinblastine and block the effect of vinblastine on metabolism. PCMV were treated either with taxol alone (30 min) or were treated with taxol followed by vinblastine (1.5 h before metabolic incubation and 3 h during metabolic incubation) as described above. Control vessels received vehicle (DMSO) alone. Vessels from two experiments were collected at the 1.5-h and 4.5-h time points and were prepared for immunofluorescence microscopy as described above. Taxol-treated vessels qualitatively appeared brighter than control vessels, and camera shutter times were thus reduced, demonstrating that taxol did induce microtubule assembly (data not shown). Vessels treated with both taxol and vinblastine resembled control vessels more than vessels treated with vinblastine alone and retained filamentous staining (Fig. 8). Thus taxol treatment appeared to effectively antagonize vinblastine-induced microtubule disruption.
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Taxol Treatment Alters Metabolism Similarly to Vinblastine Treatment and in an Additive Manner
Metabolic incubations were performed in vessels treated with either taxol alone (10 µM) or taxol followed by vinblastine to determine if stabilization of microtubules could block the metabolic effect of vinblastine (n = 5). Surprisingly, taxol treatment produced effects on carbohydrate metabolism that were very similar to those produced by vinblastine (Fig. 9). In taxol-treated PCMV, glycolysis ([2-13C]lactate peak intensity) was decreased to 90.0 ± 2.4% of control (mean ± SE, P = 0.03). Treatment of PCMV with both taxol and vinblastine produced an additional decrease in the production of [2-13C]lactate to 73.3 ± 4.0% of control (mean ± SE, P = 0.002). Neither taxol alone nor taxol plus vinblastine affected gluconeogenesis, as the [1-13C]glucose peak intensity in these groups was 97.2 ± 4.3 and 94.5 ± 8.4% (mean ± SE, not significant) of self-paired controls, respectively. Neither taxol nor taxol plus vinblastine produced increased mixing of glycolytic and gluconeogenic intermediates.
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Vinblastine alone produced an ~12% decrease in [2-13C]lactate production, whereas taxol alone produced a 10% decrease. The combination of the two agents resulted in a 27% decrease. Thus it appears that vinblastine and taxol have additive (or possibly synergistic) effects on glycolysis. Because these agents have opposite affects on microtubule polymerization, it appears that the overall microtubule assembly state of the cell has little importance for regulation of carbohydrate metabolism. Rather, the availability of binding sites for glycolytic enzymes (see DISCUSSION) on the tubulin molecule appears to be a more important determinant of glycolytic rate. Thus enzyme-tubulin associations may play a significant role in the regulation of glycolysis, but not gluconeogenesis.
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DISCUSSION |
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Compartmentation of Carbohydrate Metabolism in PCMV
We used 13C-NMR to examine the metabolic fates of [2-13C]glucose and [1-13C]FBP in VSM of freshly isolated PCMV. PCMV metabolized [1-13C]FBP primarily to [1-13C]glucose (gluconeogenesis) while simultaneously metabolizing [2-13C]glucose to [2-13C]lactate (glycolysis). Although FBP is a glycolytic intermediate, PCMV used [1-13C]FBP almost exclusively for gluconeogenesis with little [3-13C]lactate production (glycolysis). However, [2-13C]FBP from [2-13C]glucose breakdown was readily used for glycolysis. Therefore, cells had access to both [1-13C]FBP and [2-13C]FBP (from [2-13C]glucose), yet [1-13C]FBP was used primarily for gluconeogenesis while [2-13C]FBP was used for glycolysis. Thus [2-13C]FBP (derived from exogenous glucose) and exogenous [1-13C]FBP were in nonequilibrating pools within the cytoplasm, demonstrating that the intermediates of glycolysis and gluconeogenesis mix incompletely in PCMV. This result is inconsistent with classical models of glycolysis and gluconeogenesis, which assume free diffusion of enzymes and substrates within the cytoplasm. According to these models, exogenous [1-13C]FBP should be metabolized no differently than [2-13C]FBP derived from exogenous [2-13C]glucose. Thus [1-13C]FBP should be converted largely to [3-13C]lactate rather than [1-13C]glucose. These data suggest that glycolysis and gluconeogenesis occur in separate compartments of VSM of PCMV. This phenomenon has been demonstrated repeatedly (7, 13, 17, 18, 20, 21) and is referred to as "compartmentation of carbohydrate metabolism." Cerebral microvessels are only several layers of cells thick and thus are well suited to pharmacological manipulation of cellular structures such as microtubules. Therefore, cerebral microvessels are a useful system for studies of the structural basis of compartmentation of carbohydrate metabolism.Compartmentation of Metabolism in PCMV Is Likely to Be Intracellular
PCMV are a heterogeneous tissue containing VSM, endothelium, and some trapped red blood cells. Thus it is possible that one of these cell types utilized [2-13C]glucose for glycolysis while another utilized [1-13C]FBP for gluconeogenesis (intercellular compartmentation). However, we believe that this is unlikely. First, a similar compartmentation of glycolysis and gluconeogenesis has been demonstrated in hog carotid artery (18), a tissue composed almost entirely of VSM. Second, when metabolism is measured in PCMV of different sizes (separated by different mesh sizes), the percentage of endothelial cell volume to total cell volume changes by more than a factor of two (see below), yet the ratio of glycolysis to gluconeogenesis does not change significantly. Immunofluorescence microscopy demonstrated that PCMV are mostly terminal arterioles containing one or more layers of VSM and a single layer of endothelium (Fig. 2). VSM cells of terminal arterioles are 5-7 µm thick near the nucleus and ~1.0 µm thick outside the nucleus, whereas endothelial cells range from 2 µm thick at the nucleus to 0.15 µm thick outside the nucleus (40). Because the nucleus does not take part in energy metabolism, its contribution to cell volume can be ignored. In microvessels isolated using the 35-µm mesh (assuming a diameter of 10 µm, with a single 1-µm layer of VSM and a single 0.15-µm layer of endothelium), 88% of the cross-sectional area is composed of VSM, whereas only 12% is composed of endothelium. If the mesh size is increased to 105 µm (assuming a diameter of 20 µm and two 1-µm smooth muscle cell layers), 94% of the volume is smooth muscle and 6% is endothelium. Finally, microvessels isolated using the 210-µm mesh (assuming microvessels 30 µm in diameter, with an average of 2.5 smooth muscle cell layers) would be composed of ~95% VSM and 5% endothelium. Therefore, this preparation is largely composed of VSM, and the percentage of endothelial cell mass is more than twofold higher in 10-µm microvessels than in 30-µm microvessels. However, the data from the metabolic studies demonstrate that, as vessel size decreased, the ratio of glycolysis to gluconeogenesis did not change significantly (Fig. 3). Therefore, the measured metabolism appears to occur in the predominant cell type, VSM, and the observed compartmentation is likely to be an intracellular compartmentation.Finally, any contribution of trapped red cells to glycolysis can be ignored, as pig red cells do not possess glucose transporters (24) and appear to utilize inosine for energy (54). Thus, although we cannot completely discount the possibility of intercellular compartmentation, it appears unlikely to account for our results.
What Is the Basis for Compartmentation of Glycolysis and Gluconeogenesis in VSM of PCMV?
If the compartmentation of metabolism that we observed is intracellular, the apparent inability of exogenous [1-13C]FBP to mix with [2-13C]FBP derived from exogenous glucose must be due to spatial separation of these two intermediates within the cell. Such a separation could occur in several ways. First, [1-13C]FBP probably enters the cell at sites distinct from sites of glucose entry. FBP is negatively charged and is therefore unlikely to enter cells by diffusion across cellular membranes. It appears that FBP utilizes a transport system distinct from the glucose transport system in VSM, possibly entering cells via dicarboxylate transporters (10, 12). Thus localization of dicarboxylate transporters and glucose transporters to separate microdomains of the plasma membrane could result in FBP and glucose entering the cell at different points. Recent studies on caveolae support the existence of membrane microdomains containing specific protein components (1, 36).However, the existence of separate transport sites for glucose and FBP is not sufficient to explain our results. Once inside the cell, classical models of diffusion suggest that [2-13C]glucose and its metabolic products (including [2-13C]FBP) should rapidly attain equilibrium with [1-13C]FBP and its products. Our results demonstrate that this equilibrium is not achieved. Therefore, a structural basis for compartmentation must exist within the cytoplasm itself. Far from being a freely diffusing solution of proteins, substrates, and cofactors, the cytoplasm of intact cells may be highly organized. An interlacing structure composed of microtubules, F-actin, and intermediate filaments (the microtrabecular lattice) is visible in electron micrographs (53). Much of the cytoplasm may be organized around this structure (8). Thus association of glycolytic enzymes with cellular structures such as microtubules represents one possible basis for compartmentation.
Localized Enzyme Systems May Facilitate Channeling of Metabolic Intermediates
It has been suggested that metabolic intermediates may be channeled (directly transferred) from one enzyme to the next enzyme in the pathway, rather than released into solution to diffuse to the next enzyme (for a detailed treatment of channeling, see Ref. 42). Because glycolytic enzymes and intermediates are present at similar concentrations in the cell, exogenously supplied glycolytic intermediates will not be able to enter the pathway freely, since there will be few unoccupied active sites (42). The compartmentation of glycolysis and gluconeogenesis that we have demonstrated is consistent with the existence of metabolite channeling, since exogenously supplied FBP has very limited access to the glycolytic pathway in our system. In addition, we have previously provided evidence in support of channeling of glycolytic intermediates in VSM cells (11). Localization of glycolytic enzymes to microtubules may facilitate the channeling process by keeping interacting enzymes in close proximity to each other.Associations of Glycolytic Enzymes with F-actin
Glycolytic enzymes have been shown to interact with F-actin, and an F-actin binding domain has been identified in aldolase (37). Unfortunately, the physiological significance of these interactions is difficult to study in intact cells due to the lack of effective disrupters of F-actin. Cytochalasins have a variety of effects on F-actin but cannot be relied on to depolymerize F-actin (9); in fact, cytochalasins cause actin polymerization in some systems (39). In preliminary studies on our model, we were unable to detect any effect of either cytochalasin D or latrunculin B on actin organization. Although the actin cytoskeleton is almost certainly involved in the organization of metabolism, studies on this aspect of metabolism will have to wait for the development of more effective actin-disrupting drugs.Interactions Between Microtubules and Glycolytic Enzymes
Over the years, considerable evidence has been collected in vitro suggesting that some glycolytic enzymes also bind to microtubules in vivo. Phosphofructokinase (27), aldolase, pyruvate kinase, the muscle isoform of lactate dehydrogenase (LDHM), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) interact with tubulin and/or microtubules (23, 51). Under conditions that simulate molecular crowding, glucose-6-phosphate kinase and phosphoglycerate kinase interact with tubulin as well (51). Aldolase, GAPDH, LDHM, and pyruvate kinase bind microtubules with dissociation constants ranging from 1 to 4 µM, suggesting that these interactions occur at physiological concentrations of enzyme and tubulin (51). In contrast, yeast hexokinase (23), glucose phosphate isomerase (23), enolase, triosephosphate isomerase, phosphoglyceromutase, and the heart isoform of lactate dehydrogenase (LDHH; see Refs. 23 and 51) do not interact with microtubules. The disparity in the ability of LDHM and LDHH to bind tubulin suggests that the interactions of glycolytic enzymes with microtubules may be tissue-specific.The specific nature of the associations between glycolytic enzymes and
microtubules is supported by the existence of a glycolytic enzyme-binding domain on the tubulin molecule. GAPDH,
LDHM, aldolase, and pyruvate
kinase bind to a site on the COOH-terminus of -tubulin (48) located
between residues 408-451 (49). Phosphofructokinase does not bind
to the COOH-terminus of the
-subunit (47) but appears to bind to a
site near the vinblastine-binding site (46).
Physiological Relevance of Associations of Glycolytic Enzymes with Microtubules
The physiological relevance of glycolytic enzyme/microtubule interactions remains unknown. It may be that binding of glycolytic enzymes to tubulin regulates metabolism or allows for production of energy near energy-consuming processes associated with microtubules (50). In vitro, tubulin generally inhibits the catalytic activity of glycolytic enzymes. Tubulin inhibits GAPDH (33, 48), LDHM (31), aldolase (48), and phosphofructokinase (27). The inhibitory effect of tubulin appears to be specific, since it does not inhibit LDHH, which does not bind tubulin (31). However, there is also evidence that binding of glycolytic enzymes to microtubules in vivo increases rather than decreases catalytic activity (5).Glycolytic enzyme binding to microtubules is well documented, but its physiological significance remains obscure. Thus we disrupted microtubules of PCMV with vinblastine to determine whether association of glycolytic enzymes with these structures is responsible for compartmentation of [1-13C]FBP and [2-13C]glucose metabolism in VSM of PCMV. Although the microtubule network was substantially disrupted by vinblastine, the 13C-labeled intermediates of glycolysis and gluconeogenesis still failed to mix completely. Because dissolution of the microtubule network did not result in a breakdown of compartmentation, the binding of glycolytic enzymes to microtubules is not sufficient in itself to produce compartmentation. However, because compartmentation of two metabolic pathways based on separate spatial localization would require at least one unique location for each pathway, this result is not necessarily surprising.
Most interestingly, microtubule disruption affected glycolysis and gluconeogenesis differently. Glycolytic rate was significantly reduced by microtubule disruption, whereas gluconeogenic rate was unchanged. These results suggest that the glycolytic pathway is at least partially localized to microtubules and that associations between glycolytic enzymes and microtubules play a role in the regulation of glycolysis.
It is possible that vinblastine treatment may have altered vessel diameter and thus altered the size distribution of microvessels used for metabolic and imaging measurements in the vinblastine-treated group compared with controls. A decrease in the average diameter of vinblastine-treated microvessels could allow some of these microvessels to pass through the mesh and be lost. However, no difference in pellet size was noted between treated and control groups at the conclusion of the metabolic experiments, suggesting that such a change in vessel size did not occur. In any case, vessel size does not affect the ratio of glycolysis to gluconeogenesis (see Fig. 3). Finally, examination of control and treated vessels using transmitted light did not reveal any differences in appearance between groups. Therefore, it is unlikely that vinblastine treatment altered microvessel diameter and microvessel selection or that any alterations of this type could account for our results.
To determine whether the decrease in the glycolytic rate after
vinblastine treatment was due to loss of microtubule structures or to
competition for glycolytic enzyme binding sites, we treated PCMV with
the microtubule-stabilizing drug taxol. Because the effect of taxol on
the microtubule network opposes that of vinblastine, we expected that
the metabolic effects of taxol would also oppose those of vinblastine.
Surprisingly, we found that the effects of taxol on metabolism were
very similar to the effects of vinblastine. Taxol significantly
decreased glycolytic rate, with no detectable effect on gluconeogenic
rate. In addition, the taxol and vinblastine effects appeared to be
additive, since treatment of PCMV with both agents produced a greater
decrease than treatment with either alone. These data suggest that the
overall integrity of the microtubule network does not regulate
glycolytic rate but that the availability of binding sites for the
glycolytic enzymes on the tubulin molecule does play a role in
metabolic regulation. The glycolytic enzyme phosphofructokinase
competes with vinblastine for binding to tubulin (46). Thus
displacement of phosphofructokinase could account for the decrease in
glycolytic rate produced by vinblastine. Recently, pyruvate kinase has
been shown to inhibit taxol-induced microtubule polymerization (45),
suggesting that taxol and pyruvate kinase may compete for binding.
Pyruvate kinase binds to the COOH-terminus of -tubulin, as do most
of the glycolytic enzymes that are known to bind to microtubules (49).
The region of
-tubulin identified as the glycolytic enzyme binding
domain contains helix 12 and a disordered segment of 10 residues at the
extreme COOH-terminus that has not yet been resolved (35). Recent
models of the microtubule suggest that taxol binds to
-tubulin at a
site near the lateral contacts between protofilaments in the
microtubule (35). Although the taxol-binding region of the
-subunit
does not appear to be closely opposed to the glycolytic enzyme binding
domain, it is possible that binding of taxol to
-tubulin causes a
conformational change that interferes with the interaction of
glycolytic enzymes with the COOH-terminus of
-tubulin, or vice
versa. This idea is supported by the apparent competition between
pyruvate kinase and taxol (45). Thus the additional decrease in
glycolytic rate produced by taxol may be due to displacement of
pyruvate kinase or other glycolytic enzymes from the
-tubulin
COOH-terminus, an effect that would be additive to the effect of
vinblastine. Vinblastine treatment does not completely dissociate
microtubules into tubulin monomers; rather, it is likely that small
microtubule segments continue to exist after vinblastine treatment.
Because taxol further decreases glycolytic rate in vinblastine-treated vessels, the association of glycolytic enzymes with these small segments can continue to organize metabolism, even in the absence of an
intact microtubule network.
A decrease in glucose transport activity might directly inhibit glycolysis by decreasing the presentation of substrate. However, our results cannot be explained by alterations in glucose transport after drug treatment. Vinblastine has no effect on glucose transport in cultured VSM cells (6). Although vinblastine and colchicine have been shown to decrease glucose transport in C6 glioma cells (41), stabilization of microtubules by taxol increased glucose transport in the same study. Our results with taxol are not consistent with glucose transport alterations of this type, since we found that both taxol and vinblastine decreased glycolytic flux (Figs. 7 and 9). Therefore, our data cannot be explained by alterations in glucose transport activity secondary to alterations in microtubule assembly.
In contrast to their effects on glycolytic rate, neither vinblastine nor taxol altered gluconeogenic rate. Because gluconeogenesis was unaffected by either treatment and mixing of the intermediates of gluconeogenesis with those of glycolysis did not increase, the enzymes specific to gluconeogenesis are probably not localized to microtubules in vivo. Thus the gluconeogenic pathway may be localized elsewhere within the cytoplasm of VSM cells.
The results of the present study suggest that glycolytic enzymes interact with microtubules and that this interaction alters pathway flux in vivo. However, the overall state of microtubule polymerization does not appear to be as important to metabolic regulation as does the availability of binding sites for the glycolytic enzymes on the tubulin molecule. These data suggest that glycolytic enzyme-microtubule associations mainly function to keep the enzymes of the pathway localized near each other, allowing for efficient channeling of intermediates. This novel aspect of the organization and regulation of carbohydrate metabolism in vivo should be explored further.
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ACKNOWLEDGEMENTS |
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The assistance of Kimberly McNutt, Lela Hall, and Brian Kleiber in scoring immunofluorescent images is appreciated. We thank Dr. Michael Rovetto for helpful discussions and Dr. Michael Sturek for helpful advice and for providing the microscopy equipment used for this project. The technical assistance of Tina Roberts is appreciated. Pig brains were provided by Excel (Marshall, MO).
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FOOTNOTES |
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This work was supported by an Established Investigator Grant from the American Heart Association (to C. D. Hardin), National Heart, Lung, and Blood Institute Training Grant T32 HL-07094 (to P. G. Lloyd), and National Science Foundation Instrumentation Grant CHE-89-08304.
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 19 January 1999; accepted in final form 11 August 1999.
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