Institut für Herz- und Kreislaufphysiologie, Heinrich-Heine-Universität Düsseldorf, 40225 Düsseldorf, Germany
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ABSTRACT |
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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 protein1 · 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
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INTRODUCTION |
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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 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.
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METHODS |
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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 musclePerfusion 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 atThe 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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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
-NTP peak reached 65 ± 14% (n = 3) of the initial value after 2 h.
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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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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 · min1 · 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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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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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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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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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
cells1 · 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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DISCUSSION |
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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 × 102 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
protein1 · 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 · g1 · 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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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REFERENCES |
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![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Allsop, J.,
and
R. W. E. Watts.
Purine de novo synthesis in liver and developing rat brain and the effects of some inhibitors of purine nucleotide interconversions.
Enzyme
30:
172-180,
1983[Medline].
2.
Anversa, P.,
G. Olivetti,
and
M. Melissari.
Stereological measurements of cellular and subcellular hypertrophy and hyperplasia in the papillary muscle of adult rat.
J. Mol. Cell. Cardiol.
12:
781-795,
1980[Medline].
3.
Bassenge, E.,
and
G. Heusch.
Endothelial and neuro-humoral control of coronary blood flow in health and disease.
Rev. Physiol. Biochem. Pharmacol.
116:
77-165,
1990[Medline].
4.
Buck, L. T.,
and
P. W. Hochachka.
Anoxic suppression of Na+-K+-ATPase and constant membrane potential in hepatocytes: support for channel arrest.
Am. J. Physiol.
265 (Regulatory Integrative Comp. Physiol. 34):
R1020-R1025,
1993
5.
Bucolo, G.,
and
H. David.
Quantitative determination of serum triglycerides by the use of enzymes.
Clin. Chem.
19:
476-481,
1973
6.
Buttgereit, F.,
and
M. D. Brand.
A hierarchy of ATP-consuming processes in mammalian cells.
Biochem. J.
312:
163-167,
1995[Medline].
7.
Culic, O.,
M. L. H. Gruwel,
and
J. Schrader.
Energy turnover of vascular endothelial cells.
Am. J. Physiol.
273 (Cell Physiol. 42):
C205-C213,
1997
8.
Endersen, M. J.,
B. Lorentzen,
and
T. Henriksen.
Increased lipolytic activity and high ratio of free fatty acids to albumin in sera from woman with preeclampsia leads to triglyceride accumulation in cultured endothelial cells.
Am. J. Obstet. Gynecol.
167:
440-447,
1992[Medline].
9.
Geisbuhler, T.,
R. A. Altschuld,
R. W. Trewyn,
A. Z. Ansel,
K. Lamka,
and
G. P. Brierly.
Adenine nucleotide metabolism and compartmentalization in isolated adult rat heart cells.
Circ. Res.
54:
536-546,
1984[Abstract].
10.
Gronostajski, R. M.,
A. B. Pardee,
and
A. L. Goldberg.
The ATP dependence of the degradation of short- and long-lived proteins in growing fibroblasts.
J. Biol. Chem.
260:
3344-3349,
1985[Abstract].
11.
Hirai, H.,
Y. Hayashi,
T. Koizumi,
N. Nakanishi,
T. Fukui,
and
A. Ichikawa.
Fibroblast growth factor-dependent metabolism of hypoxanthine via the salvage pathway for purine synthesis in porcine aortic endothelial cells.
Biochem. Pharmacol.
45:
1695-1701,
1993[Medline].
12.
Hochachka, P. W.,
M. S. C. Bianconcini,
W. S. Parkhouse,
and
G. P. Dobson.
On the role of actomyosin ATPase in regulation of ATP turnover rates during intense exercise.
Proc. Natl. Acad. Sci. USA
88:
5764-5768,
1991[Abstract].
13.
Hochachka, P. W.,
and
G. O. Matheson.
Regulating ATP turnover rates over broad dynamic work ranges in skeletal muscles.
J. Appl. Physiol.
73:
1697-1703,
1992
14.
Hochachka, P. W.,
and
G. N. Somero.
Off-switches in metabolism: from anhydrobiosis to hibernation.
In: Biochemical Adaptation, edited by P. W. Hochachka,
and G. N. Somero. Princeton, NJ: Princeton University Press, 1984, p. 204-249.
15.
Kammermeier, H.,
P. Schmidt,
and
E. Jüngling.
Free energy change of ATP-hydrolysis: a causal factor of early hypoxic failure of the myocardium?
J. Mol. Cell. Cardiol.
14:
267-277,
1982[Medline].
16.
Kornberg, A.,
and
T. A. Baker.
Purine nucleotide synthesis de novo.
In: DNA Replication (2nd ed.). New York: Freeman, 1990, p. 55-62.
17.
Kroll, K.,
U. K. M. Decking,
K. Dreikorn,
and
J. Schrader.
Rapid turnover of the AMP-adenosine metabolic cycle in the guinea pig heart.
Circ. Res.
73:
846-856,
1993[Abstract].
18.
Land, S. C.,
L. T. Buck,
and
P. W. Hochachka.
Response of protein synthesis to anoxia and recovery in anoxia-tolerant hepatocytes.
Am. J. Physiol.
265 (Regulatory Integrative Comp. Physiol. 34):
R41-R48,
1993
19.
Lowry, O. H.,
N. J. Rosebrough,
A. L. Farr,
and
R. J. Randall.
Protein measurement with the Folin phenol reagent.
J. Biol. Chem.
193:
265-275,
1951
20.
Martin, D. W.,
and
N. T. Owen.
Repression and derepression of purine biosynthesis in mammalian hepatoma cells in culture.
J. Biol. Chem.
247:
5477-5485,
1972
21.
Mendelsohn, S. L.,
S. K. Nordeen,
and
D. A. Young.
Rapid changes in initiation-limited rates of protein synthesis in rat thymic lymphocytes correlate with energy change.
Biochem. Biophys. Res. Commun.
79:
53-60,
1977[Medline].
22.
Mertens, S.,
T. Noll,
R. Spahr,
A. Krützfeldt,
and
H. M. Piper.
Energetic response of coronary endothelial cells to hypoxia.
Am. J. Physiol.
258 (Heart Circ. Physiol. 27):
H689-H694,
1990
23.
Richards, J. M.,
I. F. Gibson,
and
M. Martin.
Effects of hypoxia and metabolic inhibitors on production of prostacyclin and endothelium-derived factor by pig aortic endothelial cells.
Br. J. Pharmacol.
102:
203-209,
1991[Abstract].
24.
Shryock, J. C.,
R. Rubio,
and
R. M. Berne.
Release of adenosine from pig aortic endothelial cells during hypoxia and metabolic inhibition.
Am. J. Physiol.
254 (Heart Circ. Physiol. 23):
H223-H229,
1988
25.
Spahr, R.,
and
H. M. Piper.
Microcarrier cultures of endothelial cells.
In: Cell Culture Techniques in Heart and Vessel Research. Berlin: Springer, 1990, p. 220-229.
26.
Stryer, L.
Biochemistry (4th ed.). New York: Freeman, 1996, p. 610-611.
27.
Stumpe, T.,
and
J. Schrader.
Phosphorylation potential, adenosine formation, and critical PO2 in stimulated rat cardiomyocytes.
Am. J. Physiol.
273 (Heart Circ. Physiol. 42):
H756-H766,
1997
28.
Tretyakov, A. V.,
and
H. W. Farber.
Endothelial cell tolerance to hypoxia.
J. Clin. Invest.
95:
738-744,
1995[Medline].
29.
Tubau, J. F.,
and
S. H. Rahimtoola.
Hibernating myocardium: a historical perspective.
Cardiovasc. Drugs. Ther.
6:
267-271,
1992[Medline].
30.
Zimmer, H.-G.,
C. Trendelenburg,
H. Kammermeier,
and
E. Gerlach.
De novo synthesis of myocardial adenine nucleotides in the rat.
Circ. Res.
32:
635-642,
1973[Medline].