Metabolic adaptation of endothelial cells to substrate deprivation

Ognjen Culic, Ulrich K. M. Decking, and Jürgen Schrader

Institut für Herz- und Kreislaufphysiologie, Heinrich-Heine-Universität Düsseldorf, 40225 Düsseldorf, Germany


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Endothelial cells are known to be metabolically rather robust. To study the mechanisms involved, porcine aortic endothelial cells (PAEC), cultured on microcarrier beads, were perfused with glucose (10 mM) or with substrate-free medium. Substrate-free perfusion for 2 h induced an almost complete loss of nucleoside triphosphates (31P-NMR) and decreased heat flux, a measure of total energy turnover, by >90% in parallel microcalorimetric measurements. Heat flux and nucleoside triphosphates recovered after addition of glucose. Because protein synthesis is a major energy consumer in PAEC, the rate of protein synthesis was measured ([14C]leucine incorporation). Reduction or blockade of energy supply resulted in a pronounced reduction in the rate of protein synthesis (up to 80% reduction). Intracellular triglyceride stores were decreased by ~60% after 2 h of substrate-free perfusion. Under basal perfusion conditions, PAEC released ~30 pmol purine · mg protein-1 · min-1, i.e., 16% of the cellular ATP per hour, while ATP remained constant. Substrate deprivation increased the release of various purines and pyrimidines about threefold and also induced a twofold rise in purine de novo synthesis ([14C]formate). These results demonstrate that PAEC are capable of recovering from extended periods of substrate deprivation. They can do so by a massive downregulation of their energy expenditure, particularly protein synthesis, while at the same time using endogenous triglycerides as substrates and upregulating purine de novo synthesis to compensate for the loss of purines.

nucleotide; purine de novo synthesis; protein synthesis; energy deprivation


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

VASCULAR ENDOTHELIUM forms the biological interface between circulating blood elements and the interstitial space and is capable of synthesizing various autocrine and exocrine factors, which play an important role in the regulation of such processes as inflammation, coagulation, and vascular tone (3). For its performance, a sufficient supply of energy is therefore required. In endothelial cells, ATP is predominantly generated by glycolysis and O2 consumption is comparatively low (22).

Endothelial cells are known to be extremely tolerant to hypoxia (22, 24). Adaptation to hypoxia is likely to be crucial for the survival of vascular endothelial cells, since they are the first to be exposed to decreases in PO2. Low energy demand and high glycolytic activity might explain why the coronary endothelium is less severely injured than cardiomyocytes in ischemic and anoxic hearts (22). It was also suggested that a tight regulation of ATP and GTP turnover exists, which enables endothelial cells to maintain their high-energy phosphates during hypoxia (28). In view of the stable energy status, it is therefore not surprising that hypoxia was reported not to influence the endothelial production of such exocrine factors as NO, prostacyclin, and adenosine (23, 24).

In a clinical setting of ischemia, there is a lack of O2 and a lack of substrates as a consequence of cessation of tissue perfusion. Although cultured endothelial cells can very well tolerate an ambient PO2 of 0.1 Torr, they become extremely sensitive to lack of O2 when glucose becomes the limiting substrate (22). How well the energy metabolism of endothelial cells recovers from substrate depletion and which mechanisms are responsible have not been explored.

Protein synthesis was identified to be the main endothelial cell energy consumer, followed by actomyosin-ATPase, actin polymerization, and endoplasmic reticulum Ca2+-ATPase, Na+-K+-ATPase, and H+-ATPase (7). With use of control analysis, a hierarchy of ATP-consuming processes in mammalian cells was recently proposed (6). According to this concept, each ATP consumer has strong control over its own rate but very little control over the rates of the other ATP consumers. It has been shown that, in turtle hepatocytes, when ATP turnover rate was decreased by ~90% by anoxia, major energy-consuming processes such as protein and urea synthesis were downregulated by ~92% and 72%, respectively (18). It thus appears that processes not essential for the immediate needs of the cell are given up before those more critical for cellular integrity of ATP supply are compromised.

Adaptive mechanisms responsible for cell survival under conditions of substrate (energy) lack are only poorly understood. A close matching of ATPase flux with ATP synthase flux has been termed the "energy-coupling constraint" and is believed to represent a major principle of metabolic regulation (13). For pathways of ATP utilization and ATP synthesis, the dominant regulation is achieved through regulation of the active enzyme concentration, whereas fine tuning of flux is achieved by substrate-, product-, or modulator-mediated kinetic adjustments (13). In anoxic hepatocytes it has been proposed that maintenance of plasma membrane ion gradient, which is essential for cell survival, was maintained through parallel downregulation of Na+-K+-ATPase activity and ion influx through ion channel arrest (4). The role of actomyosin-ATPase in the regulation of ATP turnover rates was investigated in a comparative study in fast- and slow-twitch oxidative muscle fibers (12). It was shown that fluxes through the ATP left-right-arrow  ADP + Pi cycle were extremely well regulated in rest and with exercise, implying extraordinary precision of energy coupling in both conditions.

The aim of the present study was to define the kinetics and extent of metabolic downregulation as a consequence of lack of glucose by using 31P-NMR combined with microcalorimetry. Furthermore, the mechanisms responsible for the endothelial robustness were investigated by measuring purine and pyrimidine release, endogenous triglycerides as alternative substrate, and the rate of protein and de novo purine synthesis. The results demonstrate that endothelial cells have adopted several strategies to cope with substrate lack: they massively downregulate their energy expenditure (hibernation) and use triglycerides as alternative fuel while simultaneously upregulating purine de novo synthesis.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Isolation, Characterization, and Culturing of the Endothelial Cells

Porcine aortic endothelial cells (PAEC) were isolated from slaughterhouse material by scraping of the intimal layer. Cells were plated and cultured as previously described (25). Typically, cells were grown for 7 days on cell culture plates, then trypsinized and seeded on microcarrier beads and grown for another 5 days before they were used for the experiments. The purity of the culture was checked for smooth muscle cells with use of smooth muscle alpha -actin antibody staining. Endothelial cells were positively identified with indocarbocyanin-coupled, acetylated low-density lipoproteins, as described previously (25). Confluence of PAEC grown on beads was routinely assessed by phase-contrast microscopy.

Perfusion Experiments

Microcarrier beads (2 g) with endothelial cells were washed three times with PBS supplemented with glucose (10 mM). One-tenth of this amount was used for protein and nucleotide determination (see below). Cells were mixed with an equal volume of ice-cold perchloric acid (1 M) and centrifuged for 10 min (10,000 g). The pellet was used for protein determination according to Lowry et al. (19). The supernatant was subsequently neutralized with ice-cold KH2PO4 (2 M) and centrifuged again for 3 min (5,000 g), and the acid extracts were stored at -20°C.

The remaining microcarriers were carefully packed into a glass column (10 mm ID) and perfused with PBS containing glucose (10 mM). The perfusion rate was 1 ml/min. Superfusate was collected into the ice-cooled vials, which were immediately frozen until HPLC analysis. At the end of the perfusion experiment, carriers from the column were transferred into a tube, and acid extract was prepared according to the procedure described above.

HPLC Measurements

Nucleoside and nucleobase analysis. Nucleoside and nucleobase release of perfused endothelial cells was measured by reverse-phase HPLC (17). Typically, 200 µl of superfusate were injected on a 150-mm C18 Bondapak column (Waters) and eluted with ammonium acetate (26 mM, pH 5) and a solution consisting of 67% methanol-33% water. The gradient was changed from 5% methanol solution-95% ammonium acetate to 100% methanol solution. Ultraviolet light absorbance was simultaneously monitored at 254 and 290 nm. Methods of peak identification were coelution with standards, analysis of the absorption spectrum (photodiode array detection), and enzyme shift. Enzymes used for the shift assays were nucleoside phosphorylase, guanase, adenosine deaminase, hypoxanthine-guanine phosphoribosyl transferase, and xanthine oxidase.

Nucleotide determination. To determine nucleoside tri- (NTP), di- (NDP), and monophosphate (NMP) content of endothelial cells by HPLC, the cells were washed with PBS supplemented with glucose (10 mM), extracted with 1 M perchloric acid, and subsequently centrifuged. The supernatant was neutralized (K3PO4, pH 7.5) and centrifuged to remove precipitated KClO4, while the pellet consisting of protein and microcarrier beads was dissolved in 1 M NaOH. The supernatant was injected on a reverse-phase C18 column (4 µm Bondapak, Waters), and a linear elution gradient was used to separate AMP, IMP, ADP, and ATP, with tetrabutylammonium sulfate-KH2PO4 pH 3.0 changed to pH 5.4 and finally to 70% MeOH. HPLC peaks (254 nm) were identified by comparing the retention times with those of external standards and quantified by comparison of the integrated peak areas with those of the standards after interactive baseline correction. In addition, enzyme shift assays were performed. As a control, standard nucleotides were also treated with the relevant enzyme under conditions identical to those used for the probes. 5'-Nucleotidase was used to identify NMPs. After NMPs were derivatized into their respective nucleoside forms, subsequent (nucleoside) HPLC analysis was performed. Alkaline phosphatase shift was used to dephosphorylate NTPs or NDPs, and after derivatization their identity was confirmed in a separate HPLC analysis. Nucleotide content was related to the protein content (19) of the samples.

31P-NMR Spectroscopy

Nucleotide triphosphate content of endothelial cells was studied as previously described (7). Briefly, cells on microcarrier beads were transferred into a 10-mm NMR tube containing a bottom filter through which a central capillary was passed and used as the outflow line. The NMR tube was placed inside an NMR magnet, and temperature was maintained at 37°C. Perfusion conditions were identical to the experiments performed outside the magnet.

All NMR spectra were obtained on an AMX 400 WB NMR spectrometer (Bruker, Karlsruhe, Germany) connected to an Oxford/Spectrospin 9.4-T wide-bore magnet. A 10-mm, dedicated 31P-NMR probe was employed. The homogeneity of the magnetic field was adjusted by optimizing the free induction decay of the water proton signal. Each free induction decay consisted of 2,000 data points. Data were subsequently zero filled to 4,000, then subjected to exponential multiplication (line broadening 10 Hz), Fourier transformation, and manual phasing.

Spectral acquisition time was 1 h (2,048 scans) or 15 min (512 scans); pulse width was 21 µs with a pulse repetition time of 2 s. Peak areas were determined after manual zero- and first-order baseline correction for each individual peak. Quantification was achieved by relating the initial spectrum to the nucleotide content of an aliquot of each cell batch determined by HPLC (see above).

Microcalorimetry

The details about microcalorimetry measurements have been described previously (6). Briefly, PAEC were grown on microcarrier beads [medium 199 and HEPES-buffered 10% newborn calf serum (NCS)]. The cells were packed into a steel perfusion chamber (cell volume ~1 cm3) under sterile conditions and perfused at 37°C. The cells were permitted to equilibrate in the microcalorimeter for ~16 h to achieve a stable heat flux. A thermal activity monitor (model 2277 TAM, Thermometric) fitted with a flow microcalorimeter was used for the continuous microcalorimetry measurements. The flow rate was chosen to be 12 ml/h, which makes flow conditions comparable to NMR perfusion conditions. Experiments were usually performed in the 300- or 1,000-µW range, for which the instrument was calibrated before each experiment. Before each experiment, the calorimeter system (including tubing) was sterilized with 80% ethanol.

Protein Synthesis Determination

Cells grown to confluency in six-well plates were incubated with 1 ml of PBS (with or without 10 mM glucose). Whenever inhibitors were included, they were added separately as a 1:1,000 dilution from a stock solution. Radioactive [14C]leucine was added to a final concentration of 10 µCi/ml with no additional "cold" leucine. Cells were incubated for 1-4 h at 37°C, and then medium was removed and cells were washed twice with 2 ml of ice-cold PBS. Protein was precipitated by addition of 1 ml of 10% ice-cold TCA, and then the cells were washed twice with 1 ml of 5% TCA. Protein was subsequently solubilized with 2 ml of 1 M NaOH (56°C for 30 min). Solubilized material was mixed with scintillation fluid and counted.

Triglyceride Determination

Cells were grown in medium 199 + 10% NCS before the experiment. The experiment was initiated by replacing the cell culture medium with PBS + glucose (10 mM) or PBS (no glucose). After 2 h of incubation at 37°C, the cells were harvested with a rubber policeman in 2 ml of PBS. Lipid was extracted with 6 ml of chloroform-ethanol (2:1). The organic phase was dried and dissolved in 50 µl of ethanol (8). Triglycerides were measured enzymatically (kit 334-UV, Sigma Chemical) according to Bucolo and David (5). Glycerol (25 mg/ml) was used as standard. 1,2,3-Tripalmitoyl glycerol (881 mol wt) was considered to be the representative triglyceride form. About 1.5 × 106 cells were used per assay.

Determination of the Rate of Purine De Novo Synthesis

Purine de novo synthesis was assayed by measuring the incorporation of [14C]formate into total cellular purines and released purines, according to the method of Martin and Owen (20). PAEC grown on 100-mm cell culture dishes (~1 × 107 cells) were washed twice with PBS supplemented with 10 mM glucose. Five milliliters of incubation medium containing 1 mM [14C]formate (specific radioactivity 1 mCi/mmol) were then added to the cell monolayer. After 2 h of incubation at 37°C, the medium was separated from the cells. Cells were washed twice with 1 ml of cold PBS, and washings were combined with the medium fraction. Cells were lysed by addition of perchloric acid and further processed according to Allsop and Watts (1). Radioactivity of the precipitated purine complexes from the cell or the medium fraction was counted in a liquid scintillation counter, and the activity was expressed as nanomoles per hour per 106 cells. Cell number was determined indirectly by multiplying the measured protein concentration by an experimentally determined conversion factor (1 mg PAEC protein = 5.4 × 106 cells, n = 10).

Statistics

Values are means ± SD; n represents the number of data averaged. To compare data obtained under different conditions, a paired one- or two-sided Student's t-test was used. Results were considered significantly different when P < 0.05.

Materials

Microcarrier beads were from Nunc (Roskilde, Denmark). Cell culture media and media supplements were obtained from GIBCO BRL (Paisley, Scotland). Nucleosides and nucleobases (adenosine, inosine, hypoxanthine, xanthine, xanthosine, uric acid, guanine, guanosine, cytidine, cytosine, thymidine, uracil, uridine, dehydrouracil) and nucleoside phosphates (ATP, GTP, CTP, UTP, ITP, ADP, GDP, CDP, UDP, IDP, AMP, GMP, CMP, UMP, IMP), guanase, and 5'-nucleotidase were from Sigma Chemical (Deisenhofen, Germany). Adenosine deaminase, xanthine oxidase, and alkaline phosphatase were purchased from Boehringer Mannheim. [14C]leucine (specific radioactivity 300 mCi/mmol) and [14C]formate (specific radioactivity >50 mCi/mmol) were purchased from Amersham. All other chemicals were obtained from Merck (Darmstadt, Germany) and were of analytic grade.


    RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

To determine the influence of lack of exogenous substrate on intracellular NTP content, PAEC were grown on microcarrier beads, superfused in a column at 37°C, and analyzed by 31P-NMR spectroscopy. As shown in Fig. 1, NMR-visible NTP signals were stable during the initial 2 h of glucose perfusion. On switching to a glucose-free perfusion medium, the amount of NTP declined within 45 min, while the amounts of intracellular Pi and phosphomonoesters increased in parallel. No NTP could be detected by NMR in the 2nd h of glucose-free perfusion. On reperfusion with glucose-containing medium, the intracellular NTP reappeared and the gamma -NTP peak reached 65 ± 14% (n = 3) of the initial value after 2 h.


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Fig. 1.   Representative 31P-NMR spectra of porcine aortic endothelial cells (PAEC) perfused with medium containing substrate (+glucose) or medium without substrate (-glucose). About 108 cells grown on microcarrier beads were continuously perfused (1 ml/min, 37°C) with PBS supplemented with glucose (10 mM) or PBS without glucose. Each perfusion condition lasted for 2 h. Spectral acquisition time was 1 h (2,048 scans) or 15 min (512 scans), resulting in a lower signal-to-noise ratio in 15-min spectra. Peaks are as follows: Pii and Pie, intra- and extracellular inorganic phosphate, respectively; alpha -, beta -, and gamma -NTP, alpha -, beta -, and gamma -phosphate of nucleoside triphosphates.

To verify NMR data and because 31P-NMR does not permit differentiation of adenine from other cellular NTPs, HPLC analysis was undertaken using the protocol outlined in Fig. 1. From the data compiled in Fig. 2, in endothelial cells the GTP pool is one-half the size of the ATP pool. Uridine and cytidine nucleotides were below the limit of detection by HPLC. The total NAD+ + NADH pool was 2.88 ± 0.74 nmol/mg protein, which is ~40% of the ATP content and reflects the known glycolytic character of these cells (7). Two hours of glucose-free perfusion reduced tissue levels of ATP and GTP to 21.8 and 19.7%, respectively (Fig. 2); no NTP (NTP = ATP + GTP) was detectable by NMR under identical conditions (Fig. 1). Despite extensive purine release (see below), the amount of AMP remained virtually unchanged.


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Fig. 2.   Nucleotide content of PAEC before and after 2 h of substrate-free perfusion. Nucleotide content of cells grown in complete cell culture medium was compared with nucleotide content of cells that were perfused for 2 h with PBS + glucose (10 mM), then with PBS without glucose. Nucleotide content was analyzed in neutralized acid extract by HPLC. In parallel, protein content of extracted cells was determined. Values are means ± SD from at least 6 experiments.

In a separate experimental series the release of nucleosides and nucleobases was quantified under the conditions described in Fig. 1. As shown in Fig. 3, four different nucleosides and four nucleobases could be identified in the superfusate outflow. Under basal conditions, the release typically reached steady-state values 60 min after initiation of perfusion. Thereafter, release of all nucleosides and nucleobases was stable (Fig. 3). In the steady state the total release of purines and pyrimidines amounted to 45.5 ± 4 pmol · min-1 · mg protein-1, the ratio of purines to pyrimidines being 2.7:1.0 (n = 5). The ratio of adenosine-inosine-hypoxanthine to guanosine-guanine to uridine-uracil to cytidine was 5.6:5.7:3.3:1.0. Switching to glucose-free medium resulted in a transient increase of the concentrations of most of the metabolites, typically reaching a maximum 30 min after initiation of glucose-free perfusion. Only the concentrations of adenosine and uracil showed a more stable increase by factors of ~2.5 and 8.5, respectively. The ratio of adenosine-inosine-hypoxanthine to guanosine-guanine to uridine-uracil to cytidine was 6.1:6.5:6.4:1.0, whereas the ratio of total purines to pyrimidines released during 2 h of glucose-free perfusion was 1.4:1.0 (n = 5).


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Fig. 3.   Time course of nucleoside and nucleobase release from PAEC under control (PBS + 10 mM glucose) and energy-deprived conditions (PBS without glucose). Concentrations of nucleosides and nucleobases were determined by HPLC as shown in Fig. 4 and described in METHODS. Glucose-free perfusion was initiated 120 min after perfusion of PAEC was initiated. Open symbols, purines; filled symbols, pyrimidines. Values are means ± SD of 5 experiments. * Statistically significant increase in concentration of nucleosides in perfusate (paired Student's t-test, P < 0.05).

To compare the previous results with total energy turnover, parallel experiments using microcalorimetry were undertaken. Cells perfused with PBS + glucose showed stable heat flux of 223.2 ± 75.2 µW/cm3 microcarrier beads (n = 10). As shown in a representative recording in Fig. 4, perfusion with medium containing no exogenous substrate (glucose) for 2 h decreased heat flux to 8.5 ± 0.4% (n = 6) of its initial value. On reperfusion with glucose, heat flux fully recovered.


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Fig. 4.   Microcalorimetry recording of PAEC perfused with PBS with or without substrate. Concentration of glucose, when included, was 10 mM. Cells were perfused at 37°C. Perfusion protocol is described in Fig. 1 and 2 legends.

Because in a previous study protein synthesis was identified as the main energy-consuming process in PAEC (7), we investigated whether protein synthesis is inhibited when substrate-induced downregulation of energy turnover occurs. As shown in Fig. 5, [14C]leucine incorporation progressively decreased as the glucose concentration was lowered. Cycloheximide (20 µM) in the presence of glucose (10 mM) inhibited leucine incorporation by ~92%. When glucose was absent, [14C]leucine incorporation was reduced to ~23% of the initial value. Inhibition of oxidative phosphorylation by antimycin also significantly reduced protein synthesis (Fig. 5).


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Fig. 5.   Influence of energy deprivation on rate of protein synthesis in PAEC. Confluent cells grown on cell culture dishes (~106 cells) were washed and incubated for 2 h in presence of [14C]leucine. Incubation medium was PBS. Medium was removed, and cellular protein was precipitated with ice-cold 10% TCA. Protein was subsequently solubilized with NaOH-SDS solution and counted in a scintillation counter. Values are means ± SD from 4 different experiments. * Statistically significant difference compared with control (Student's t-test, P < 0.05).

When PAEC were perfused with PBS (without glucose), heat flux was massively downregulated, as shown in Fig. 4, and cycloheximide was unable to further decrease the heat flux (results not shown). This provides evidence in addition to [14C]leucine incorporation experiments that protein synthesis in PAEC was downregulated in the absence of the exogenous substrate.

To address the question of endogenous substrates used by PAEC during substrate-free perfusion, we measured the triglyceride content after cells were incubated for 2 h with or without glucose. As shown in Fig. 6, the triglyceride content of PAEC was significantly decreased after incubation without glucose.


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Fig. 6.   Triglyceride depletion due to lack of exogenous substrate. PAEC were incubated for 2 h in PBS with or without glucose. Cells were then harvested, and lipids were extracted with chloroform-ethanol. Triglycerides were determined enzymatically by measurement of glycerol content after triglyceride hydrolysis. Glycerol solution of 250 mg/dl was used as standard. Values are means ± SD from 7 different experiments. * Statistically significant decrease (P < 0.05) of triglyceride content in cells incubated without glucose compared with cells incubated with glucose.

Aside from protein synthesis, purine de novo synthesis is known to be a highly energy-requiring process. In a final experimental series the influence of substrate lack on purine de novo synthesis was studied using [14C]formate as a precursor. Purine de novo synthesis in PAEC was determined to be 0.99 nmol · 106 cells-1 · h-1, which is the sum of newly synthesized purines present intracellularly and those released into the extracellular medium. This value is similar to that reported by Hirai et al. (11). Surprisingly, purine de novo synthesis in glucose-deprived PAEC was stimulated by ~184% (Fig. 7). In the absence and presence of glucose, newly synthesized purines were identified not only in the cellular fraction but also in the supernatant (filled vs. open bars in Fig. 7).


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Fig. 7.   Purine de novo synthesis of PAEC. Confluent cells grown on 100-mm cell culture dishes were incubated for 2 h with 5 ml of medium (with or without 10 mM glucose) supplemented with 1 mM [14C]formate. Cells were separated from incubation medium, and cells and medium were further processed as described in METHODS. Filled bars, part of newly synthesized purines detected in intracellular fraction; open bars, fraction detected in extracellular (incubation) medium. Values are means ± SD of 4 experiments. * Statistically (2-sided t-test, P < 0.05) significant increase of de novo purine synthesis rate compared with cells incubated with PBS + glucose (10 mM).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The main finding of this study is that endothelial cells are capable of reversibly downregulating their energy expenditure when no exogenous substrate (glucose) was provided. They do so by reducing important energy-requiring reactions, such as protein synthesis, and using endogenous triglycerides as an alternative energy fuel. At the same time, however, they upregulate purine de novo synthesis to cope with the loss of purines into the effluent. This suggests that a hierarchy of metabolic reactions exists by which endothelial cells can compensate for and survive metabolic stress.

Previous studies have indicated that endothelial cells are very robust and tolerant to lack of O2 and substrate (7, 22, 24, 28). Here we demonstrate that, on lack of exogenous substrate, endothelial cells lose their cellular ATP within 2 h of perfusion but recover most of their adenine nucleotides when glucose is readministered (Fig. 1). With use of 31P-NMR, nucleotide triphosphates could not be detected after 2 h of substrate-free perfusion (Fig. 1), whereas under the same conditions ~20% of the initial ATP and GTP could still be measured using HPLC. The reason for this discrepancy is not fully clear. It could involve compartmentation of ATP and GTP into an NMR-visible (free) and an NMR-invisible (bound) pool (9).

An important strategy for survival of extended periods of substrate deprivation is downregulation of energy expenditure. Microcalorimetric measurements (Fig. 4) revealed that, in the absence of glucose, endothelial cells are able to drastically downregulate total energy turnover. This finding is similar to the previous report (7), where we showed that blocking the glycolysis by substrate reduction (PBS + 10% NCS) and 2-deoxyglucose (10 mM) caused ATP signals to disappear in the NMR spectrum. On the basis of the ATP content and measurements of basal energy turnover, the half-life of endothelial cell ATP was calculated to be ~8 s (7). Without a compensatory reduction of ATP expenditure, the decline in ATP induced by substrate deprivation should be complete within ~1 min. Endothelial cells do not contain measurable amounts of glycogen (unpublished results), but they could rely on endogenous fatty acids as an energy source, as indicated in Fig. 6. Basal triglyceride content of 17 µg/mg protein is very similar to a published value for human umbilical vein endothelial cells (8). The extent to which the triglycerides as alternative endogenous substrates can sustain basal metabolic rate in the absence of exogenous glucose is illustrated in the following calculation. Taking into account that the remaining heat flux on withdrawal of exogenous substrate is ~10 µW/mg protein, one can calculate that over 2 h cells liberate ~7.2 × 10-2 J. With the assumption that the average molecular weight of triglycerides is 900, complete oxidation of 10 µg of triglycerides (~1.1 × 10-8 mol) releases energy sufficient for the synthesis of ~3.0 × 10-6 mol ATP (~300 mol ATP can be synthesized from 1 mol tripalmitoyl glycerate) (26). Hydrolysis of 1 mol ATP was reported to release ~60.5 kJ/mol (15). Therefore, the expected overall released energy would amount to 18 × 10-2 J. This value is very close to the measured 7.2 × 10-2 J released as heat over 2 h of substrate-free perfusion. Therefore, it is reasonable to assume that triglycerides are most likely the major endogenous substrate responsible for the survival of PAEC without exogenous substrate.

In zoology the term hibernation signifies a form of dormancy by which an organism ceases its consumption of external nutrients concomitant with a metabolic depression of variable but controlled degree (13). Hibernation was later used to describe at the organ level a chronic adaptive reduction of myocardial contractile function in response to a reduction of coronary blood flow (29). More recently, hibernation was demonstrated at the level of isolated cardiomyocytes when ambient PO2 was systematically lowered (27). In most general terms, hibernation, therefore, describes the coordinated downregulation of metabolism when the supply of O2 or nutrients becomes limiting. This study demonstrates that endothelial cells have the ability to hibernate, which reversibly downregulates their energy turnover by ~90% when glucose as sole exogenous substrate is omitted.

Protein synthesis is the main energy-consuming reaction in endothelial cells, comprising at least 23% of total energy turnover (7). Protein synthesis in the hibernating endothelial cells was found to be profoundly downregulated, being almost fully inhibited when no external substrate was provided. Rat thymocytes resemble PAEC, since deprivation of energy-providing substrates for 2 h caused a 75-80% drop in the rate of protein synthesis (21). In fibroblasts a severe reduction in cellular ATP was associated with an ~50% inhibition of proteolysis (10). These observations are in line with the hypothesis that a hierarchy of ATP-consuming processes exists in mammalian cells, protein synthesis being the most sensitive to energy supply followed by RNA/DNA synthesis and substrate oxidation, Na+ cycling, and Ca2+ cycling (6).

A surprising finding of the present study is that, in view of the general reduction of energy expenditure, purine de novo synthesis was upregulated under conditions of substrate deprivation, suggesting that the functioning of this pathway may be essential for the survival of endothelial cells under conditions of metabolic stress. As to the regulation of de novo purine synthesis, it should be recalled that synthesis begins with phosphoribosyl pyrophosphate (PRPP) and requires 4 mol of ATP to synthesize 1 mol of IMP, which is then converted to AMP or GMP. The overall rate of purine nucleotide biosynthesis and the relative rates of formation of the two end products, adenylate and guanylate, are regulated by feedback control (16). Measurement of AMP, GMP, and IMP revealed that glucose-free perfusion of endothelial cells did not alter tissue levels of AMP but caused a significant reduction in the levels of GMP and IMP (Fig. 2). GMP and IMP are involved in three major feedback mechanisms, which cooperate in regulating de novo purine synthesis: PRPP synthase, glutamine-PRPP amidotransferase, and IMP dehydrogenase. The reduced levels of GMP and IMP can therefore biochemically explain the measured upregulation of de novo purine synthesis in vivo.

There is additional evidence for increased de novo purine synthesis in endothelial cells. Under steady-state conditions with constant levels of NTPs (Fig. 1), endothelial cells perfused on microcarrier beads release ~30 pmol purine · mg protein-1 · min-1, which is ~16% of the cellular NTP per hour under basal conditions. De novo purine synthesis measured in confluent monolayers is of similar magnitude: if it is assumed that 1 mg of total endothelial protein corresponds to ~5.4 × 106 cells (unpublished results), de novo purine synthesis can be calculated to be 15 pmol · mg protein-1 · min-1. During substrate-free perfusion, this value increases to ~27 pmol · mg protein-1 · min-1, which at least could partially compensate for the augmented loss of purines (Fig. 3). The rapid replenishment of nucleotide triphosphates, particularly ATP, on readmission of substrates (Fig. 1) can only be explained by the highly active de novo purine pathway, since the salvage pathway is likely to be only of minor importance in a flow-through system. The recovery of ATP most likely contributes to the rapid increase in energy turnover as reflected by heat flux measurements (Fig. 4).

The high rate of de novo purine synthesis in endothelial cells may shed some new light on previously published data. Zimmer et al. (30) reported de novo purine synthesis in the rat heart to be 8.4 ± 1.42 nmol · g-1 · h-1, which is equivalent to 60 pmol · mg protein-1 · min-1 when it is assumed that 1 g of myocardial tissue corresponds to ~140 mg of protein. The value 60 pmol · mg protein-1 · min-1 in the heart compares with ~900 pmol · mg protein-1 · min-1 measured in endothelial cells in the present study. If it is assumed that cardiac de novo synthesis resides exclusively in the endothelium, a compartment size of ~7% of the total heart can be calculated. Interestingly, morphometric measurements revealed that endothelial cells comprise ~5% of the heart on a volume basis (2). This suggests that endothelial cells may be the predominant site of adenine de novo synthesis in the heart.

In summary, the present study has demonstrated that endothelial cells can survive extended periods of substrate deprivation and can resume a normal metabolic rate thereafter. Three mechanisms appear to be responsible for this high tolerance to lack of substrates: 1) a reduction in energy expenditure by inhibition of protein synthesis and most likely other energy-requiring reactions to limit the energetic consequences of the reduced ATP synthesis, 2) mobilization of the endogenous triglycerides as an energy source, and 3) stimulation of de novo synthesis of adenine nucleotides to compensate for the loss of purines and to help replenish ATP in the phase following substrate readmission.


    ACKNOWLEDGEMENTS

We thank Eva Bergschneider for excellent work and valuable suggestions concerning various aspects of HPLC analysis and Adele Brand for technical help in preparation of the manuscript.


    FOOTNOTES

This research was funded in part by Deutsche Forschungsgemeinschaft Grant SFB 242.E1 and by the Center for Biological and Medical Research (Biomedizinisches Forschungszentrum) of the Heinrich-Heine-University Düsseldorf.

Address for reprint requests and other correspondence: J. Schrader, Dept. of Physiology, Heinrich-Heine-University Düsseldorf, PO Box 10 10 07, D-40001 Düsseldorf, Germany (E-mail: schrader{at}uni-duesseldorf.de).

Received 14 July 1998; accepted in final form 5 January 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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