1 Institut für Herz- und Kreislaufphysiologie, Heinrich-Heine-Universität Düsseldorf, 40225 Düsseldorf, Germany; and 2 Institute for Cell Biology, ETH-Hönggerberg, CH-8093 Zürich, Switzerland
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ABSTRACT |
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To characterize the isoenzyme distribution of creatine kinase
(CK) in endothelial cells (ECs) and its functional role during substrate depletion, ECs from aorta (AECs) and microvasculature (MVECs)
of pig and rat were studied. In addition, high- energy phosphates were
continuously monitored by 31P NMR spectroscopy in pig AECs
attached to microcarrier beads. CK activity per milligram of protein in
rat AECs and MVECs (0.08 ± 0.01 and 0.15 ± 0.08 U/mg,
respectively) was <3% of that of cardiomyocytes (6.46 ± 1.02 U/mg). Rat and pig AECs and MVECs displayed cytosolic BB-CK, but no
MM-CK. Gel electrophoresis of mitochondrial fractions of rat and pig
ECs indicated the presence of mitochondrial Mi-CK, mostly in dimeric
form. The presence of Mia-CK was demonstrated by indirect
immunofluorescence staining using Mia-CK antibodies. When
perifused with creatine-supplemented medium, phosphocreatine (PCr)
continuously increased with time (1.2 ± 0.6 nmol · h1 · mg protein
1),
indicating creatine uptake and CK activity. Glucose withdrawal from the
medium induced a rapid decrease in PCr, which was fully reversible on
glucose addition, demonstrating temporal buffering of an energy
deficit. Because both cytosolic and mitochondrial CK isoforms are
present in ECs, the CK system may also contribute to energy
transduction ("shuttle hypothesis").
endothelium; transport; energy metabolism; phosphorus-31 nuclear magnetic resonance spectroscopy
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INTRODUCTION |
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CELLS AND TISSUES WITH INTERMITTENTLY high and fluctuating energy requirements, such as skeletal and cardiac muscle, brain, retina, and spermatozoa, depend on the immediate availability of large amounts of energy. In these cells and tissues, the enzyme creatine kinase (CK; ATP:creatine N-phosphoryl transferase, EC 2.7.3.2.) has been proposed to play a key role in energy metabolism (38). This role may involve 1) the replenishment of ATP through transfer of the phosphoryl group from phosphocreatine (PCr) to ADP ("buffer function") but also 2) a facilitated intracellular energy transduction ("creatine kinase shuttle hypothesis"), linking mitochondrial ATP generation to cytosolic sites of ATP consumption via mitochondrial and cytosolic CK isoenzymes (2, 25, 38).
Two cytosolic subunit isoforms, "ubiquitous" brain-type B-CK and "sarcomeric" muscle type M-CK as well as two mitochondrial subunit isoforms, ubiquitous Mia-CK and sarcomeric Mib-CK, are synthesized in a tissue-specific manner. The cytosolic subunits combine to form enzymatically functional homodimers and heterodimers, MM-CK, MB-CK, and BB-CK isoenzymes, whereas the mitochondrial isoforms form either homodimers or homooctamers (26).
The vascular endothelium is generally regarded as a nonexcitable tissue with a rather constant energy demand imposed by processes as diverse as cytoskeletal contraction (myosin ATPase), protein synthesis, and ion transport (Na+-K+-ATPase) (5). The endothelium is characterized by a high glycolytic capacity and can survive extended periods of hypoxia (18). These properties suggest that cytosolic isoforms of CK are not essential in endothelial cells (ECs) and may not be present at all, as is the case, e.g., in hepatocytes (12). It was therefore rather surprising that almost equal amounts of BB-CK and MM-CK were suggested to be present in human umbilical vein ECs (15). BB-CK activity, but no MM-CK, was also observed in rat liver ECs (35), while no evidence of mitochondrial CK was obtained. In fact, the release of BB-CK from liver has even been used as an index of endothelial cell injury (24, 35).
While there appears to be agreement that ECs contain CK, several important questions remain. 1) What is the functional role of CK in ECs? 2) Do vascular ECs contain both MM-CK and BB-CK or do they display BB-CK activity only? In normal adult hearts, in the absence of hypertrophy, cardiac myocytes predominantly express MM-CK and MB-CK. Thus, in the case where ECs would exclusively express BB-CK, one might be able to use a release of this latter CK isoform to discriminate between cardiac myocyte and endothelial cell injury. 3) Are there differences between macrovascular and microvascular ECs, e.g., between ECs from aorta and coronary capillaries? 4) Are both cytosolic and mitochondrial isoforms of CK expressed? In this case, a prerequisite for the CK shuttle hypothesis would be fulfilled. If only the cytosolic isoform were present, the CK system might simply operate as a temporary high-energy phosphate buffer.
To address these questions, microvascular ECs from the coronary system of pig and rat as well as macrovascular aortic ECs from these two species were analyzed and compared with rat cardiomyocytes. Isoenzyme distribution and total CK activity were assessed. Moreover, the role of CK in the intact cell was investigated by 31P-NMR spectroscopy of perifused porcine aortic ECs grown on microcarrier beads; in this system the total creatine pool was either increased by creatine supplementation or the cells were subjected to a reduced energy status by withdrawing glucose supply.
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MATERIALS AND METHODS |
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Preparation of macrovascular ECs from porcine aortas.
Macrovascular ECs were isolated from porcine thoracic aortas obtained
from a local slaughterhouse as described in detail before (30). In brief, isolated cells were suspended in culture
medium [M199 with Earle's salts (GIBCO, Karlsruhe, Germany)
containing 20% newborn calf serum (NCS), 250 IU/ml penicillin, 250 µg/ml streptomycin, 12.5 µg/ml amphotericin B, and 50 µg/ml
gentamicin] and plated onto 100-mm plastic culture dishes (Falcon,
type 3003). Cultures were incubated at 37°C with 5% CO2
in room air. Four hours after plating, the culture medium was renewed.
After 2 days of incubation the medium was renewed again, reducing the
concentration of antibiotics to 100 IU/ml penicillin, 100 µg/ml
streptomycin, and 5 µg/ml amphotericin B. Gentamicin was omitted at
this time. Five to seven days after plating, cells reached confluency
and were harvested at this point. Cultures were washed twice with PBS,
pH 7.4, scraped off the dishes with a rubber policeman, immediately frozen in MSH buffer (in mM: 220 mannitol, 70 sucrose, 10 HEPES, 0.2 EDTA, 1 2-mercaptoethanol, and 1 sodium azide, pH 7.4) and stored at
70°C. This procedure was repeated for all cell types.
Preparation of microvascular ECs from porcine hearts. Microvascular ECs (MVECs) were isolated from porcine hearts obtained from a local slaughterhouse. The procedure used was similar to the isolation of MVECs from rat hearts described by Piper et al. (22). In brief, an artery of the right ventricle was cannulated and rinsed with ice-cold Krebs-Henseleit solution [containing (in mM) 116 NaCl, 4.7 KCl, 1.1 MgSO4, 1.17 KH2PO4, 25 NaHCO3, 2.5 CaCl2, 8.3 glucose, and 2 pyruvate, pH 7.4, when equilibrated with carbogen (95% O2-5% CO2)] to remove the remaining blood and to define the area of tissue supplied by this vessel. This piece of the right ventricle was cut out and mounted onto a perfusion device. The flow was adjusted to ~10 ml/min. After an initial perfusion period with Krebs-Henseleit buffer at 37°C, the perfusate was switched to a buffer, containing (in mM) 110 NaCl, 2.6 KCl, 1.2 MgSO4, 25 NaHCO3, and 11 glucose and continuously gassed with carbogen through the tip of a Pasteur pipette to maintain a pH of 7.4. The effluent was collected in a beaker placed underneath the tissue and was discarded. After 20 min the perfusion was switched to a recirculating mode and continued for another 45 min with 80 ml of fresh buffer containing 20 mg of collagenase and 15 µl of a 100 mM CaCl2 stock solution. The ventricular tissue was taken off the cannula, treated with a tissue chopper, and transferred into a glass beaker containing the recirculated medium and 800 mg of BSA, fraction V. During an incubation period of 30-40 min at 37°C the mixture was gassed with carbogen and frequently resuspended with a 10-ml disposable pipette. The dissolved tissue was filtered through a nylon mesh, and the filtrate was centrifuged at 25 g for 5 min. The supernatant was transferred to a Teflon beaker, and 300 mg of BSA, 15 mg of trypsin (1:250), and 45 µl of CaCl2 stock solution (100 mM) were added. During an incubation period of 30 min at 37°C, the mixture was gassed again with carbogen and stirred in a shaking water bath. The cell suspension was centrifuged at 250 g for 10 min and the pellet resuspended in a culture medium, similar to that of porcine aortic ECs but containing 10% NCS, 10% FCS, and no amphotericin B. After another centrifugation step and resuspension, the cells were plated onto 100-mm plastic culture dishes and handled as indicated above.
Preparation of ECs from rat aortas. Thoracic aortas were isolated from 200- to 300-g male Wistar rats following removal of the hearts (see above). The vessels were immersed in ice-cold PBS, and adjacent connective and adipose tissue were removed. The aortas were opened in a longitudinal direction, and segments of ~2-3 mm length were cut. They were placed with the intima facing downward into a 100-mm plastic culture dish and were allowed to adhere to the bottom of the dish by incubation at 37°C (with 5% CO2) for about 1 h. Culture dishes were cautiously filled with culture medium (see Preparation of macrovascular ECs from porcine aortas) and incubated again at 37°C (with 5% CO2). Every other day, culture medium containing reduced concentrations of antibiotics was renewed. When ECs started to grow from underneath the aortic segments, the tissue was removed and cells were cultured until they reached confluency.
Preparation of microvascular ECs and cardiomyocytes from rat
hearts.
Microvascular ECs and adult cardiac myocytes were isolated from 200- to
300-g male Wistar rats as described by Piper et al. (21-23) and kindly provided by T. Stumpe from our
laboratory. In brief, following perfusion with a Ca2+-free
Krebs-Henseleit buffer, hearts were treated with collagenase (0.1%) in
a recirculating mode, and then were minced and filtered. Centrifugation
enabled the separation of ECs (supernatant) and cardiomyocytes
(pellet). Cardiomyocytes were washed with increasing Ca2+
concentrations and purified by centrifugation (4% BSA). The rat heart
ECs were incubated with trypsin for 30 min, centrifuged, and
transferred to culture plates. Culture medium and incubation conditions
were similar to those for pig heart MVECs. Aliquots of cell suspensions
obtained from different experiments were immediately frozen and stored
at 70°C.
Purity of cell culture.
Purity of EC cultures was determined by phase-contrast microscopy and
immunofluorescence. The uptake of low-density lipoproteins (LDLs) in
live ECs was performed with indocarbocyanin-coupled acetylated LDLs (10 µg/ml, for 4 h at 37°C; Paesel+Lorei, Frankfurt, Germany)
(36). Contamination with fibroblasts or smooth muscle cells was studied by immunofluorescent staining with a monoclonal mouse
antibody to -smooth muscle actin (1:50, 30 min; Sigma, Deisenhofen,
Germany) (1). Endothelial cultures used in this study were
>95% pure.
Preparation of mitochondrial and cytosolic extracts.
Mitochondrial and cytosolic extracts from rat or pig heart muscle and
brain were prepared as described previously (41). In
brief, following tissue homogenization and one centrifugation step (750 g), the supernatant was centrifuged at 6,000 g, separating mitochondria (pellet) and cytosolic extract (supernatant). The pellet
was washed and centrifuged (6,000 g) two times before the mitochondria were resuspended in 5-30 ml of buffer and
subsequently extracted. Fractions were stored in MSH buffer at 70°C
until used as reference samples for cellulose polyacetate electrophoresis.
NMR spectroscopy of ECs. ECs grown on microcarrier beads were studied as previously described (6). They were transferred to a 10-mm NMR tube that contained a bottom filter (pore diameter 35 µm); a central capillary passing through the filter was used as outflow line. Within the NMR-sensitive volume, a standard capillary containing a defined amount of methylenediphosphonic acid (MDP) was fixed close to the center of the tube. Cells on microcarriers sedimented rapidly and formed a homogeneous column that was perfused from top to bottom at 1 ml/min with a HEPES buffer, containing (in mM) 150 NaCl, 2.7 KCl, 1.2 KH2PO4, 1.2 MgSO4, 1.0 CaCl2, 5.6 glucose, 0.5 creatine, and 10 HEPES, as well as 10% NCS, adjusted to pH 7.4 and equilibrated with room air. The NMR tube was placed inside the magnet, and temperature was maintained at 37°C. Protein content per gram of carrier was determined in every experiment (16). For this purpose an aliquot of the cells was washed in serum-free HEPES buffer.
All NMR spectra were obtained on an AMX 400 WB NMR spectrometer (Bruker, Karlsruhe, Germany) that was connected to an Oxford/Spectrospin 9.4 Tesla wide-bore magnet. A 10-mm broadband probe head (Bruker) was employed. The homogeneity of the magnetic field was adjusted by optimizing the free induction decay (FID) of the water proton signal. A near-Lorentzian line shape with a line width at half height of 12-20 Hz was achieved in every experiment. Each FID consisted of 2,048 data points. Data were subsequently zero-filled to 4,096, followed by exponential multiplication (line broadening 25 Hz), Fourier transformation and manual phasing. The NMR-sensitive volume (1.75 ml) contained ~5.4 mg protein of ECs on 1.04-g microcarriers. Fully relaxed spectra were the result of 2,048 90° scans (pulse width 27 µs) with a pulse repetition time of 15 s. Partially saturated spectra were obtained using 1,800 70° scans and a pulse interval of 3 s. Peak areas were determined after manual zero and first-order baseline correction for each individual peak and referenced to the external standard (MDP) that contained a known amount of phosphates, using the integration software of the NMR spectrometer (UXNMR; Bruker). Because the longitudinal relaxation time T1 of MDP was measured to be 5.68 s, an appropriate saturation factor (0.933) was taken into account in spectra obtained at a pulse interval of 15 s.HPLC. To compare the NMR measurements with conventional biochemical techniques, aliquots of ECs were extracted with 1 M perchloric acid. The extracts were neutralized and centrifuged, and the supernatant was injected and separated on a µBondapak C18 4-µm column (Waters, Eschborn, Germany) while ultraviolet absorption at 254 nm was detected. A linear elution gradient was employed changing from 36.8 mM KH2PO4/2.95 mM tetrabutylammoniumsulfate (TBAS), pH 3.0, to 14.7 mM KH2PO4/2.95 mM TBAS, pH 5.4, and finally to 70% methanol. Chromatogram peaks were identified by comparing the retention times of the samples with those of external standards containing AMP, ADP, and ATP, and peaks were quantified by comparing the integrated peak areas with those of the external standards.
CK activity. CK activity was determined according to the method of Szasz et al. (32). A commercially available test kit (Boehringer Mannheim, Germany) was used for these assays. Thawed samples were sonicated and, following centrifugation, the supernatants were used for measurements of enzyme activity. Protein content of all cell suspensions was determined according to the Lowry method.
Cellulose polyacetate electrophoresis. Cellulose polyacetate electrophoresis was performed on Cellogel strips (Bio-tec-Fischer, Reiskirchen, Germany) in veronal buffer (pH 8.6) at room temperature as described earlier (40). CK isoenzymes were separated at a constant voltage of 150 V for 45 min (see Figs. 4 and 5) or at 100 V for 90 min (see Figs. 6 and 7). Separated isoenzymes were visualized by an overlay gel technique (39) in the presence and absence of 15 µM P1,P5-di(adenosine-5') pentaphosphate (Ap5A), which inhibits adenylate kinase activity (14).
Immunofluorescence labeling. ECs were grown in monolayers on glass coverslips placed in plastic culture dishes (35 mm, Falcon). Cells were fixed overnight with 3% paraformaldehyde and 0.2% glutaraldehyde in ice-cold PBS, pH 7.4. The fixative was exchanged with a mixture of 0.3% paraformaldehyde and 0.02% glutaraldehyde in ice-cold PBS, and cells were stored at 4°C until use. All immunolabeling steps were carried out at room temperature. Fixed cells were washed three times with Tris-buffered saline (TBS) containing 50 mM Tris · HCl and 150 mM NaCl, pH 7.4, and the fixative was quenched by incubation for 10 min with TBS supplemented with 0.1 M glycine. After permeabilization with 0.2% Triton X-100 in TBS for 20 min, cells were incubated for 20 min with TBS containing 0.5% BSA, 0.2% gelatin, and 2% horse serum (referred to as TBG) to saturate nonspecific protein-binding sites. After a 90-min incubation period in a moist chamber with either rabbit anti-chicken B-CK or anti-chicken ubiquitous mitochondrial CK (Mia-CK) antibodies and, in parallel, also with the corresponding preimmune IgG [previously prepared and characterized in the lab of T. Wallimann (10)], each diluted 1:500 with TBG, cells were washed three times for 10 min each with 0.2% Triton X-100 in TBS and three times with TBS to remove unbound antibodies. The cells were then incubated for 75 min with rhodamine-conjugated goat anti-rabbit IgG (Pierce) diluted 1:500 with TBG. After additional washing steps (as the ones described above), the cells were mounted in buffered polyvinyl alcohol Lennette medium (13) containing p-phenylenediamine (1 mg/ml; Sigma) as anti-fading agent. Photographs were taken on a Zeiss universal fluorescence microscope using Kodak T-Max 400 DIN black/white film.
Statistics. Data are given as means ± SD. Differences in CK activities between different types of ECs were evaluated using Student's t-test. P < 0.05 was considered to be statistically significant.
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RESULTS |
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31P NMR of porcine aortic ECs.
Energy metabolism of perifused porcine macrovascular ECs from the aorta
(AECs) was investigated by 31P NMR spectroscopy. In fully
relaxed spectra, peak areas represent the concentration of phosphorus
nuclei of the respective compounds. A representative example is
shown in Fig. 1. Resonances of inorganic phosphate, PCr, NAD, and nucleotide triphosphates (-,
- and
-phosphate resonances) were identified, as well as peaks in the phosphomonoester and phosphodiester region. The
-phosphate of the
nucleotide triphosphates was quantified to be 22.8 ± 3.8 nmol/mg protein (n = 3), similar to ATP measurements obtained
in acid extracts of the same cell batches by HPLC. Thus ATP is the
dominant nucleotide in AECs. While ADP was below the NMR detection
threshold in perifused AECs, the ADP/ATP ratio in extracts was ~1:7.
This suggested that a considerable portion of ADP is protein bound and
NMR invisible. A further peak at
12.3 parts per million (ppm) could not be identified and may represent uridine diphosphoglucose. The
inorganic phosphate peak represents almost exclusively the inorganic
phosphate of the perifusion medium (1.2 mM
KH2PO4), since the intracellular volume of the
ECs (~30 µl) was small compared with the surrounding medium. The
amount of phosphomonoesters, i.e., metabolites like glucose 6-phosphate
and fructose 6-phosphate, was about twice the size of the endothelial
ATP pool, consistent with the high glycolytic capacity of EC.
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CK activity in ECs of rats and pigs.
CK activities were measured in cell preparations from rat and
pig (Table 1). In the rat, ECs isolated
from hearts (MVECs) or thoracic aortas (AECs) showed a significantly
lower CK activity compared with cardiomyocytes. Rat MVEC CK activity
was ~1/40 and AEC CK activity only ~1/80 of that of cardiomyocytes.
Enzyme activities of macrovascular ECs from pig and rat were in a
similar range (0.06 and 0.08 U/mg protein). Both in pig and rat,
microvascular ECs displayed a higher CK activity than the corresponding
AECs.
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Cytosolic CK isoenzymes in ECs.
Extracts of macro- and microvascular ECs from rat and pig were analyzed
by cellulose polyacetate electrophoresis to separate cytosolic and
mitochondrial CK isoenzymes. Tissue extracts of heart and brain from
both species were included as standards. As can be seen in Fig.
4A, macro- and microvascular
ECs from rat contain mainly cytosolic BB-CK, thereby corresponding in
enzyme pattern to rat brain (lane 4). MM-CK, typical for
heart (lane 3) and skeletal muscle, as well as the
heterodimer MB-CK could not be detected. Similar results were obtained
with AECs and MVECs from pig (Fig. 4B). The fainter bands
seen especially in MVECs from rat (Fig. 4A, lane 2) and pig
(Fig. 4B, lane 2) represent adenylate kinase (see below;
Fig. 5).
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Mitochondrial CK isoenzymes in ECs.
Analysis of enriched mitochondrial fractions from rat and pig micro-
and macrovascular ECs revealed some enzymatic activity at cathodic
positions indicative for mitochondrial Mi-CK isoenzymes, which have a
higher isoelectric point than the cytosolic isoforms (26)
(Fig. 6). Although these Mi-CK activity
bands are rather weak, no obvious differences between micro- and
macrovascular ECs of the same species were noticeable (lanes
1 and 4 and 2 and 3). However,
the bands of mitochondrial CK isoenzymes obtained from rat displayed a
more cathodic position than those of pig.
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Immunofluorescence staining of porcine aortic ECs.
Indirect immunofluorescence staining of porcine aortic ECs with
anti-B-CK antibodies revealed a relatively homogeneous, rather strong
staining of all ECs sparing the nuclei (Fig.
8A). In contrast, the same
staining procedure for ubiquitous Mia-CK (Fig.
8B) revealed a rather weak, mostly perinuclear and spotted
staining, typical for mitochondria (7), while no staining
was seen in the control (Fig. 8C). This indicated (in line
with the biochemical data) that B-CK is the prominent cytosolic CK
isoform in ECs, with only minor amounts of Mia-CK present
as the ubiquitous mitochondrial CK isoform in ECs. The presence of both
B-CK and Mia-CK would indicate a prerequisite fulfilled for
a PCr circuit working in these ECs.
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DISCUSSION |
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By monitoring the cellular energy status noninvasively by 31P-NMR spectroscopy, the present study demonstrates reversible alterations in PCr concentration and thus functional CK activity in ECs. The only cytosolic isoform detected in cultured macro- and microvascular ECs of pig and rat was BB-CK. In addition, the presence of a mitochondrial isoform of CK, most likely Mia-CK, was observed in ECs of both rat and pig. The finding of both mitochondrial (albeit at low levels) and cytosolic BB-CK (at significantly higher levels) isoforms in conjunction with functional data strongly suggests that the PCr circuit is operative also in ECs.
CK isoforms in ECs. In the past, conflicting evidence had been provided regarding the cytosolic CK isoforms expressed in ECs. While some groups observed exclusively BB-CK in the endothelial compartment (35), others reported the presence of both BB-CK and CK-MM in human ECs derived from the umbilical vein (15). This was surprising, considering the fact that MM-CK is predominantly expressed in skeletal and cardiac muscle. However, since ECs are also characterized by a rather large contribution of myosin ATPase to total energy turnover (5), a functional similarity of ECs to muscular cells may be assumed. In the light of the present data, it seems likely that the previous identification of MM-CK in ECs was confounded by a similar migration pattern of CK and adenylate kinase: in our hands, the putative MM-CK band disappeared when gels were treated with Ap5A to inhibit adenylate kinase activity. Moreover, the band developed also in the absence of creatine phosphate. Thus the only cytosolic isoform present in pig and rat ECs was BB-CK.
The expression and activity of CK varies between different vascular beds. Both in pig and rat, the BB-CK expression in microvascular ECs by far exceeded the expression in macrovascular ECs derived from the aorta. In fact, the presence of BB-CK in pig aortic ECs was hardly discernible (see Figs. 4 and 5). The different expression correlated with CK activity, which in the pig was sixfold greater in microvascular than in macrovascular ECs (Table 1). The major quantitative differences in CK activity imply differences in energy turnover and cellular function. It is well conceivable that ECs from the coronary circulation, playing a substantial role in the regulation of coronary flow by the formation of various mediators, are characterized by a higher energy turnover or a more fluctuating energy demand than cells from the aorta. Previous studies did not report the presence of the mitochondrial isoform of CK in ECs of hepatic and myocardial origin (3, 28, 35). Lack of mitochondrial CK would imply that the cytosolic CK could only operate as a temporal energy buffer. In the present work, however, weak bands were detected in both rat and pig ECs, clearly representing mitochondrial CK. Although no definite assignment of the mitochondrial isoform was feasible on the basis of gel electrophoresis, indirect immunofluorescence staining for ubiquitous Mia-CK revealed the presence of this mitochondrial isoform in porcine ECs (Fig. 7). In the liver, the release of BB-CK in reperfusion following ischemia has been taken as a marker of endothelial cell injury (24, 35). Similarly, in the heart, following cardioplegia during valve replacement surgery, substantial differences in the time course of serum BB-CK and MM-CK activity were reported (33), suggesting these isoforms to be released from different compartments. Consistent with the smaller size of the endothelial compartment and the lower CK activity, the arterio-venous difference for BB-CK was much smaller than for MM-CK. A distinct release of BB-CK has also been observed following myocardial infarction (17, 34). However, in contrast to the liver, BB-CK release from the heart is unlikely to be an exclusive marker of endothelial injury. BB-CK is also present in smooth muscle cells (11) and cardiomyocytes, as indicated, e.g., by the faint BB-CK band in Fig. 5B. Moreover, it is well known that cardiac hypertrophy is often associated with a fetal shift in CK isoenzyme distribution, i.e., an increased myocardial expression of BB-CK (29). Finally, when comparing BB-CK and MM-CK release following cardioplegia (33), a ratio of 1:10 was observed. Because the endothelial compartment comprises only 3% of the total cardiac volume and endothelial CK activity per mg protein is rather low, it is unlikely that the endothelium contributed up to 10% to total cardiac CK release. This further underlines the notion that BB-CK release into the coronary sinus is not a selective marker of endothelial injury.CK activity and functional role in ECs.
Compared with cardiomyocytes, ECs are characterized by a rather low
activity of CK (2.5% of cardiomyocytes). Considering the different
cellular functions of ECs and cardiomyocytes, the lower CK activity in
ECs may be considered to be paralleled by a lower energy turnover.
However, several lines of evidence suggest that the energy turnover of
ECs is in the same range as that of cardiomyocytes. In the presence of
physiological substrate concentrations, the endothelial oxygen
consumption was reported to be 8 nmol · min1 · mg protein
1
(18), which is about one-third of that of stimulated
cardiomyocytes (31). Moreover, when total energy turnover
of perifused ECs on microcarrier beads was determined by
microcalorimetry, a total heat flux of 230 µW/mg protein was found
(5). This is comparable to the total energy turnover of
cardiomyocytes, as revealed by the following calculation: cardiac
oxygen consumption being ~5 µmol · min
1 · g
1 (110 µl · min
1 · g
1) and the
average caloric equivalent being 20 kJ/l O2 gives an approximate energy turnover of 2.2 J · min
1 · g
1 (37 mJ · s
1 · g
1). Assuming a
protein content of 160 mg/g wet weight, an energy turnover of 231 mJ
· s
1 · mg protein
1 can be
estimated, virtually identical to the total heat flux of ECs.
Therefore, it is not only the total CK activity but also the ratio of
CK activity to energy turnover that is rather low in ECs. This,
together with the low CK turnover, made it impossible to use
31P NMR magnetization transfer to measure the CK flux in
perifused endothelial cell (unpublished results).
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ACKNOWLEDGEMENTS |
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We thank Dr. Thomas Stumpe for isolating rat cardiac myocytes and endothelial cells, Dr. Ognjen Culic for providing pig aorta endothelial cells on microcarrier beads, and Claudia Kirberich, Eva Bergschneider, and Else Zanolla for skillful technical assistance.
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FOOTNOTES |
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Financial support by the Deutsche Forschungsgemeinschaft (Grant SFB 242 E1 to J. Schrader), the Biomedical Research Center of the Heinrich-Heine-University (BMFZ), the Swiss Foundation for Muscle Diseases, and the Swiss National Science Foundation (Grant 31-33907.92 to T. Wallimann) are gratefully acknowledged.
Present address of M. Wyss: F. Hoffmann-La Roche AG, Building 241/865, CH-4070 Basel, Switzerland.
Address for reprint requests and other correspondence: U. Decking, Dept. of Cardiovascular Physiology, Heinrich-Heine-Univ. Düsseldorf, PO Box 10 10 07, 40001 Düsseldorf, Germany (E-mail: decking{at}uni-duesseldorf.de).
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. Section 1734 solely to indicate this fact.
Received 18 July 2000; accepted in final form 22 January 2001.
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