Department of Molecular and Cellular Physiology, Louisiana Health Sciences Center, Shreveport, Louisiana
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ABSTRACT |
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Human umbilical vein endothelial cells (HUVECs) are an endothelial model of replicative senescence. Oxidative stress, possibly due to dysfunctional mitochondria, is believed to play a key role in replicative senescence and atherosclerosis, an age-related vascular disease. In this study, we determined the effect of cell division on genomic instability, mitochondrial function, and redox status in HUVECs that were able to replicate for ~60 cumulative population doublings (CPD). After 20 CPD, the nuclear genome deteriorated and the protein content of the cell population increased. This indicated an increase in cell size, which was accompanied by an increase in oxygen consumption, ATP production, and mitochondrial genome copy number and ~10% increase in mitochondrial mass. The antioxidant capacity increased, as seen by an increase in reduced glutathione, glutathione peroxidase, GSSG reductase, and glucose-6-phosphate dehydrogenase. However, by CPD 52, the latter two enzymes decreased, as well as the ratio of mitochondrial-to-nuclear genome copies, the mitochondrial mass, and the oxygen consumption per milligram of protein. Our results signify that HUVECs maintain a highly reducing (GSH) environment as they replicate despite genomic instability and loss of mitochondrial function.
replicative senescence; glutathione; cell size changes; genomic instability; human umbilical vein endothelial cells
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INTRODUCTION |
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HUMAN UMBILICAL VEIN ENDOTHELIAL CELLS (HUVECs) replicate a finite number of times in culture and are an endothelial model of replicative senescence. Reactive oxygen species (ROS) and oxidative damage to DNA (24), protein (7), and lipid (45) increase with age and have been implicated in aging and replicative senescence (19, 48). Oxidative stress has also been implicated in atherosclerosis, an age-related vascular disease, and homocysteine, a contributing factor to atherosclerosis that induces oxidative stress, accelerates endothelial senescence in culture (52). Atherosclerosis is hypothesized to be the result of a "response to injury" of the endothelium. The endothelium must constantly respond to changes in the vascular environment and experiences frequent insults from mechanical, hemodynamic, and inflammatory stresses, all of which enhance oxygen free radical production. In response to injury, endothelial cells must replicate to repair blood vessels. Telomere length is shorter in endothelial cells derived from vessels where there is greater hemodynamic stress, and presumably more endothelial cell replication to repair injury, than in cells isolated from areas of lower stress (10). Injury induced by balloon catheter denudation of rabbit carotid arteries has been shown to result in the accumulation of senescent endothelial cells (18). Studies of the endothelium at atherosclerotic lesions (9, 50) and in vessels from elderly people (30, 50) have detected multinucleated cells and changes in cell size, alterations characteristic of replicative senescence. Similar changes in gene expression have been demonstrated in endothelial cells aging in vivo and HUVEC models of senescence (23, 30). For example, replicative senescent HUVECs in culture overexpress intercellular adhesion molecule-1 (ICAM-1) (35) and endothelin (30), whereas atherosclerotic lesions have also been shown to overexpress ICAM-1 (42), and plasma endothelin levels from patients with advanced atherosclerosis are elevated (33). HUVEC "aging" in culture therefore has characteristics of endothelial cellular aging in vivo.
ROS are constantly produced in metabolizing cells in vivo. One source of ROS is the mitochondria that utilize ~85% of the cell's oxygen, of which 5-10% is reduced to ROS that can damage mitochondrial macromolecules (24, 47). Mitochondrial DNA deletions (13) and mutations (36) accumulate with age (14), which correlates with a decline in mitochondrial function (11, 17). Treatment of mice with the antioxidant N-acetylcysteine (NAC) protected against an age-related decline in activity of respiratory complex I in hepatic mitochondria (37). This is consistent with the idea that the mitochondria are damaged by ROS. After deterioration of the mitochondria, it has been proposed that ROS production from the mitochondria increases and leads to further oxidation of lipids, proteins, and nuclear DNA (4). Cells have established antioxidant defense mechanisms to protect against ROS, and these consist of detoxification enzymes and glutathione, which reacts with ROS and protects thiol groups from oxidation. Redox cycling of glutathione is a way of protecting macromolecules against cellular oxidative stress by maintaining levels of reduced glutathione (GSH). Alterations in antioxidant defense mechanisms or generation of an oxidative stress have been shown to alter the lifespan of organisms and the replicative life of cells. For example, overexpression of superoxide dismutase (SOD) and catalase extended the lifespan of Drosophila melanogaster (40). However, treatment of primary cells with ionizing radiation increased the rate of replicative senescence in culture (34, 39), and fibroblasts deficient in glucose-6-phosphate dehydrogenase (G6PD), an enzyme required to generate NADPH for the conversion of oxidized glutathione (GSSG) to GSH, also senesce at a faster rate (26).
We have used HUVECs replicating in culture to try to understand the
deterioration in the mitochondria and redox antioxidant systems of
replicating endothelial cells. A decline in the redox system in the
"larger" endothelial cells at sites of injury could make them more
susceptible to damage. Depletion of GSH in HUVECs in culture alters
their response to stimuli such as TNF- (12) and phorbol
ester (27), and glutathione peroxidase was shown to be
essential for endothelial survival during culture with peroxides (22). Many studies examine only young and senescent cells;
we have studied HUVECs during many rounds of replication by examining the population at different stages of its replicative life. In this
way, we have determined a time course of changes in genomic stability,
mitochondrial function, and redox status (GSH/GSSG) prior to senescence
to aid in the understanding of changes in endothelial cells in vivo
that replicate after injury and during the progression of vascular disease.
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MATERIALS AND METHODS |
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Isolation and Growth of HUVECs
HUVECs were harvested from umbilical cords by 0.25% collagenase treatment for 20 min at 37°C, as previously described (53). HUVECs were grown on plastic coated with 1 µg/cm2 human fibronectin (BTI, Stoughton, MA) in endothelial growth medium (EGM) (BioWhittaker, Walkersville, MD) containing 2% fetal bovine serum in a humidified atmosphere at 37°C and 5% CO2. Freshly isolated cells were seeded and grown to confluency in 25-cm2 flasks during a period of 3-5 days. At this point, cells were designated cumulative population doubling (CPD) zero. Cells were frozen in 90% fetal bovine serum and 10% DMSO. During growth, cell growth medium was changed every 2 days and cells removed from the plastic just reached confluency (~80-90% confluent) using 0.05% trypsin and 0.53 mM EDTA. Cells were centrifuged, and a portion was resuspended in growth medium and trypan blue (final concentration of 0.2%; Sigma, St. Louis, MO) before being counted with a hemocytometer. At each passage, the total number of cells and the viable number of cells in the culture were counted and the change in population doubling was calculated by the following formula: change in population doubling = ln (total cells harvested/viable cells seeded)Two separate sets of data were collected for each of the oxygen consumption, ATP production, and glutathione analyses, but the results presented in this paper are one complete set of data generated from samples taken at exactly the same time.
To confirm that the population consisted of endothelial cells, cells of CPD 7, 31, and 40 were incubated with acetylated low-density lipoprotein labeled with 1,1'-dioctadecyl-3,3,3'-tetramethylindo-carbocyanine perchlorate (BTI) at 10 µg/ml in growth medium for 4 h at 37°C, 5% CO2 and examined for fluorescence by microscopy (Nikon Optiphot-2 microscope; Nikon, Garden City, NY). Cells were also stained with an antibody against von Willebrand factor (Sigma) or VE-cadherin (Immunotech, Cedex, France). The cultures were positive for all of these endothelial cell markers. Normal human dermal fibroblasts from neonatal skin (BioWhittaker) were also stained for each of these markers as a negative control.
Senescence-Associated -Galactosidase
Nuclear Genome Content
Fluorescence in situ hybridization.
HUVECs were grown on chamber slides, washed in PBS, fixed in
methanol-acetic acid (3:1, vol/vol), and stored at 80°C before the
fluorescence in situ hybridization (FISH) assay was performed. The
samples were denatured at 73°C for 5 min in 70% formamide (Life
Technologies, Rockville, MD) in 2× saline-sodium citrate buffer (SSC;
1 × SSC: 0.15 M NaCl and 0.015 M sodium citrate, pH 7) and
sequentially dehydrated using an ethanol series (70, 85, and 100%).
The hybridization mixture, containing the chromosome enumeration probe
(SpectrumOrange CEP 4[
satellite]; Vysis, Downers Grove, IL)
and CEP hybridization buffer (Vysis), was denatured at 73°C for 5 min. Hybridization was carried out at 37°C overnight in a HYBrite
hybridization system (Vysis). Posthybridization washes consisted of
0.3% NP-40, 0.4× SSC at 73°C for 2 min, followed by a wash with
0.1% NP-40, 2× SSC at room temperature for 30 s. Nuclei were
counterstained with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI
II; Sigma). Samples were visualized by using an Olympus BX60
microscope, and 200 nuclei were examined for each CPD.
Chromosome number. To determine changes in ploidy, chromosomes were counted from chromosome spreads. Cells were arrested in metaphase by addition of colcemid (KaryoMax; GIBCO, Rockville, MD) at 0.5 µg/ml final concentration to the medium for 1 h at 37°C. Harvested cells were swollen in hypotonic KCl (0.075 M), fixed in 3:1 (vol/vol) methanol:acetic acid, and dropped onto cold, wet microscope slides. The chromosomes were stained with a Giemsa solution (Sigma), and 30 nuclei were examined for each CPD.
Quantitative PCR to Determine the Ratio of Mitochondrial-to-Nuclear Genome Copy Number
HUVECs of different CPDs were harvested, and the DNA was isolated by using the QIAamp DNA blood mini kit according to the manufacturer's recommendations (Qiagen, Valencia, CA). Quantitative PCR using the "Taqman" approach (41) was used to compare the amounts of PCR product for a primer set located in the D-loop of the human mitochondrial genome (GenBank accession nos. J01415, M12548, M58503, M63932, and M63933) and a single-copy gene encoding the RNA moiety of RNase P in the nuclear genome (predeveloped assay; Applied Biosystems, Foster City, CA). The primers and probe for the mitochondrial sequence were designed using the Primer Express 1.5 computer program (Applied Biosystems). The forward primer 5'-GTGAAATCAATATCCCGCACAAG-3', the reverse primer 5'-TCACTTTAGCTACCCCCAAGTGTT-3', and the TaqMan probe 5'-CTACTCTCCTCGCTCCGGGCCC-3' modified at the 5' terminus with a reporter dye (FAM, 6-carboxyfluorescein) and at the 3' terminus with a quencher dye (TAMRA, 6-carboxytetramethylrhodamine) were obtained from Applied Biosystems. Each primer/probe set was used to set up PCR reactions using 10 ng of HUVEC DNA: the reaction for the RNA moiety of RNase P was set up according to the manufacturer's protocol (Applied Biosystems), while the reaction for the mitochondrial D-loop contained 500 nM of forward primer, 300 nM of reverse primer, and 200 nM of probe in TaqMan Universal PCR master mix (Applied Biosystems) in a total volume of 50 µl. Reactions were placed at 50°C for 2 min, 95°C for 10 min, and then 95°C for 15 s (denaturation step) and 60°C for 1 min (annealing and extension steps) for 40 cycles in an ABI Prism 7700 sequence detection system (Applied Biosystems; Ref. 41). Reactions were performed in triplicate, and an average was determined for the detection-threshold cycle for the mitochondrial (CTm) and the nuclear probe (CTn). For each sample, the mitochondrial PCR was normalized to the nuclear PCR by subtracting the threshold cycles (CTmMitochondrial Mass Measurement
MitoTraker Green FM (Molecular Probes, Eugene, OR) was dissolved in DMSO at a concentration of 1 mM and diluted to 40 µM in DMSO. HUVECs were harvested by trypsinization, counted, centrifuged, and resuspended in EGM containing either 0.1% DMSO or 40 nM MitoTraker Green FM (Molecular Probes) at 106 cells/ml. Cells were incubated in the dark for 45 min at 37°C, followed by centrifugation and resuspension at 106 cells/ml in EGM containing 0.1 µg/ml propidium iodide. Samples were placed on ice and analyzed within 30 min using a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA). Green (530/30 nm) and red (>650 nm) fluorescence emissions were collected using logarithmic amplification. Vehicle (DMSO)-treated cells were used to determine the background cellular fluorescence. Data were collected from 10,000 cells that were negative for propidium iodide staining.Total Oxygen Consumption
HUVECs were medium changed 4-6 h before trypsinization. A known number of cells (2-6 × 106 cells) were centrifuged and resuspended in 3.3 ml of PBS (37°C). The cell suspension was placed in a closed vessel, and total oxygen consumption was measured per minute with a Yellow Springs Instrument model 5300 biological oxygen monitor, as previously described (29). After the rate of oxygen consumption had been determined, a 1-ml sample of the cell suspension was centrifuged to obtain a cell pellet. Cell pellets were dissolved in 0.1 M NaOH, and the protein concentration was determined according to Bradford (8), utilizing the Bio-Rad protein assay reagent (Bio-Rad, Hercules, CA). Rates of total oxygen consumption are expressed as nanomoles of oxygen consumed per minute per 106 cells or as nanomoles of oxygen consumed per minute per milligram of protein.Quantification of Cellular ATP
The HUVEC growth medium was changed 4-6 h before trypsinization. Cells were counted, and 0.3 to 0.5 × 106 cells were harvested by centrifugation, washed with PBS, and resuspended in cold PBS and cold 3 M perchloric acid to obtain a final concentration of 1 M perchloric acid. Samples were stored atQuantification of GSH and GSSG
Cellular GSH and GSSG were determined by the HPLC method of Reed et al. (44). HUVECs were medium changed 4-6 h before trypsinization. Cells were counted, and 0.3 to 0.5 × 106 cells were harvested by centrifugation, washed with PBS, and resuspended in 5% cold trichloroacetic acid (TCA). Samples were stored atDetection of Reactive Oxygen Species Using Dihydrorhodamine 123
HUVECs were grown on chamber slides and incubated with 5 µM dihydrorhodamine 123 (DHR) (Molecular Probes) in growth medium at 37°C, 5% CO2 for 30 min. Cells fluorescing due to the oxidation of DHR to rhodamine 123 were counted, and fluorescent images were obtained with the use of a Nikon Optiphot-2 microscope (Nikon) and a digital camera (SenSys Color 1400, Photometrics, Tucson, AZ). At least 300 cells were examined at each CPD, except for CPD 56 where 136 cells were used for the analysis.Redox Cycle Enzyme Assays
HUVECs were trypsinized and counted, and a known number of cells were harvested by centrifugation. Cell pellets were washed in PBS and stored atStatistical Analyses
Data are expressed as average ± SD unless stated otherwise. Analyses were performed by Dunnett post hoc test, using the SAS software, to a confidence level of 0.05. ![]() |
RESULTS |
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Protein Content of the Cell Population
During growth of the HUVECs, we observed an increase in cell size. To confirm this morphological observation, the protein content per cell was determined from cell pellets obtained after treatment of cells with 5% TCA (Fig. 1A). The average protein content per cell did not change until CPD 21.9, at which point the protein per cell doubled. From CPD 21.9 to 44.4, the protein content per cell was stable; however, further increases were seen between CPD 48 and 57.9, at which point the protein content per cell was approximately four to five times that of CPD 2.8. The increase in cell size was also confirmed by examining the light-scattering properties of the cells by flow cytometry. As can be seen in Fig. 1B, CPD 7.2 cells scatter the light to a lower extent than cells of CPD 33.2 and 49.8. The contour lines situated close together represent a high density of cells. It is evident that the majority of cells at these three stages of replicative life are of different sizes: small, medium, and large.
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To ensure cells were progressing towards senescence, we stained for
senescence-associated -galactosidase activity. At CPD 10, only 4%
of the population was positive for this stain. This increased steadily
with increasing CPD (21% at CPD 23.4, 38% at CPD 34, 42% at CPD
38.3, and 55% at CPD 44.8), which is consistent with previous findings
for cultures of fibroblasts and endothelial cells (30).
Genomic Instability
Endothelial cells have previously been found to exhibit nuclear genomic instability (50). Because an alteration in nuclear content could result in a change in cellular protein content, we determined, first, the ploidy within the cell population of a single chromosome (chromosome 4, Fig. 2) by FISH and, second, the average chromosome number per cell at different CPDs (Table 1). It is evident from the results of both of these experiments that the cell population before CPD 21 is predominantly diploid (70-80%). However, at CPDs >21, the cell population decreases in diploid cells and increases in triploid and tetraploid cells. From Table 1 (see legend), it appears that the cells at CPD 34 undergo a phase during which the population is predominantly triploid or tetraploid and emerge through this instability period (CPD 39) with a higher percentage of tetraploid than diploid cells, but a lower percentage of cells appearing triploid. This instability period, however, was not seen for chromosome 4 alone (Fig. 2), although the percentage of tetraploid and triploid cells did increase with increasing CPD. To further determine whether there was a tendency for certain chromosomes to selectively increase in number, we examined 10-15 metaphases of CPD 12, 22, and 39 for chromosomes 1, 2, and 4 using the chromosome paint WCP 1, 2, and 4 multicolor DNA probe (Vysis). The results indicated that the percentage of diploid decreased and the percentage of tetraploid increased with increasing CPD for each of the three chromosomes (data not shown) as found for chromosome 4 using FISH (Fig. 2). In both the FISH and chromosome painting experiments, we did detect a very low frequency of cells within the population that were >4 n. From the FISH study in which 200 nuclei were examined, ~1 to 2% of all the different populations were >4 n (data not shown). When examining the individual metaphases for chromosomes 1, 2, and 4, we did find instances in which one chromosome was tetraploid and the other two were diploid and at CPD 22 and 39 in which two chromosomes were tetraploid and one chromosome was diploid. However, this appeared to be random. There was no evidence from this study, which uses a small sample size, that a particular chromosome selectively increased in ploidy.
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Because we observed changes in mitochondrial function during the
replicative life of the HUVECs (see below), we postulated that the
mitochondrial genome copy number may also change during the alteration
in cell size. The literature indicates that mitochondrial function
declines while mitochondrial DNA mutations increase with age, but the
copy number has not been examined in a replicative senescent model. We
utilized quantitative PCR to compare the ratio of the copy number of
the mitochondrial genome to the nuclear genome. The primers were
designed to the D-loop of the mitochondria, which contains the heavy
strand origin of replication and appeared from the literature to be the
only region not deleted during aging. The nuclear region examined was
the gene encoding RNase P, which is a single-copy gene. As can be seen
from Fig. 3A, the ratio compared with CPD 6 was stable until CPD 28, where there is a 3.5- to
4-fold increase in the mitochondrial copies compared with the nuclear
copies. This ratio was still evident at CPD 44 but declined to ~2 at
CPD 48.
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Mitochondrial Mass
The increase in the number of copies of the mitochondrial genome at CPD 28 could have been due to an increase in the number of mitochondria. The only way to determine the number of mitochondria/cell is to perform electron microscopy studies. However, MitoTraker Green FM is a fluorescent probe that accumulates in mitochondria independent of mitochondrial membrane potential and can therefore be used to compare the mitochondrial mass of cell populations at different replicative age. Initial experiments determined that staining of cells was linear within a range of 20-200 nM MitoTraker Green FM, and we chose 40 nM because this dose provided a peak of stained cells that was clearly distinguishable from the background fluorescent signal. As shown in Fig. 3B, a significant although very small (~10%) increase in mass was detected at CPD 33.2, and a decrease (~32%) was detected at CPD 49.8 compared with cells at CPD 7.2.Mitochondrial Function
Approximately 85% of a cell's total oxygen consumption is used by the mitochondria to generate ATP. Given the changes in mitochondrial genome and mass (see Fig. 3), we wanted to examine how the mitochondrial function altered during the replicative life of endothelial cells and whether mitochondrial dysfunction occurs in large endothelial cells. To determine the mitochondrial oxygen consumption, we tried to measure the oxygen consumption after the cells had been treated with antimycin A. However, the solvents required to solubilize antimycin A (dimethylformamide, ethanol, or dimethyl sulfoxide) destroyed the HUVECs' ability to consume oxygen, probably due to membrane damage. We were therefore only able to quantify the rate of total cellular oxygen consumption per 106 cells (Fig. 4A) and per milligram of protein (Fig. 4, B and C). Duplicate measurements were performed at each CPD. Unexpectedly, we found that the oxygen consumption per 106 cells increased from 2.85 nmol · min
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The majority of ATP is produced via oxidative phosphorylation in the
mitochondria: 22 ATP molecules are produced for each glucose molecule
as opposed to 2 from glycolysis and 2 from the citric acid cycle. We
therefore examined cellular ATP and ADP content. Accurate measurements
of ADP were hampered by the small amounts of ADP in the HUVEC samples,
and significant changes were not seen in ADP levels (data not shown). A
significant increase in ATP per 106 cells was detected at
CPD 26 and higher (Fig. 5A)
when values were compared with CPD 2.8. Comparison of CPD 2.8 and CPD
57 showed a 4.8-fold increase of ATP per 106 cells. Upon
examining the ATP per milligram of protein (Fig. 5B), only
the value at CPD 53.7 was significantly different from CPD 2.8. The
HUVECs were therefore able to maintain ATP production even as the cells
grew larger, despite a decrease in oxygen consumption per milligram of
protein.
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Changes in Cellular Redox Status with Respect to GSH and GSSG
Cellular redox status has been found to be important in the decision of whether a cell proliferates or dies (3), and an oxidative stress in the cell can be detected by a change in the levels of GSH and GSSG. Because oxidative damage has been implicated in replicative senescence and aging, and mitochondrial dysfunction can generate ROS, we decided to define the redox status in the HUVECs as they progressed towards senescence. GSH significantly increased per 106 cells between CPD 26 and 57.9 compared with CPD 2.8 (Fig. 6A). This correlates with the increase in protein content of the cells (Fig. 1A). However, in examining the results of the GSH per milligram of protein (Fig. 6B), there is a significant increase in production of GSH between CPD 37.1 to 57 that cannot be explained simply by an increase in cell size. Rather, a compensatory stimulation in GSH production could occur during stress situations. Although a significant increase in GSSG per 106 cells was detected between CPD 33.7 and CPD 42.1 to 51.2 (Fig. 6C), significant differences were not detected in the measurements of GSSG per milligram of protein (Fig. 6D). The "stress" was therefore not reflected in the total cellular GSSG content. It is possible that a subcompartment of GSH in an organelle such as the mitochondria, which is only a small percentage of the total cellular glutathione, may have been affected. Studies of mitochondrial glutathione are prohibited by the large number of cells required to isolate and quantify GSH in the mitochondrial compartment. To obtain insight into the redox environment within the cells, the GSH:GSSG ratio was calculated by using the data obtained for the GSH and GSSG per 106 cells (Fig. 6E). The GSH:GSSG ratio increased as the cells increased in CPDs, indicating the HUVECs were maintaining a highly reduced environment. The increase in the ratio largely reflects the huge increase in cellular GSH (Fig. 6A).
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An increase (~4-fold) in GSSG per milligram of protein (Fig. 6D) and GSSG per 106 cells (Fig. 6C) was detected at the earliest time point in the culture (CPD 2.8), indicating an oxidative stress. This study used cells that had previously been frozen in liquid nitrogen. We have examined freshly isolated cells at similar CPDs for GSH and GSSG content and saw a similar transient increase (~2-fold) in the GSSG level (data not shown). This initial oxidative stress is likely explained by the primary cells adapting to life in cell culture.
Detection of ROS Generation
Because the average changes we observed in GSSG levels were small, we postulated that oxidant generation was occurring in either a subpopulation of the cells, resulting in GSH oxidation only in the subpopulation, or that there was selective oxidation of mitochondrial GSH. To visualize cellular ROS generation, we treated the population with the oxidant-sensitive nonfluorescent probe DHR. DHR oxidation results in rhodamine 123, which is fluorescent and accumulates in actively respiring mitochondria. We were unable to detect any fluorescent cells at CPD 6 and 17, and only 0.3% of the population fluoresced at CPD 22 (Fig. 7A). At CPD 29, however, 5% were fluorescing and this increased to 15% by CPD 44. A similar percentage of fluorescent cells was detected at CPD 51.2, 51.9, and 56.1. The rhodamine 123 signal was punctate and perinuclear (Fig. 7B), similar to that seen for mitochondrial probes. However, we were unable to colocalize this signal with that of a mitochondrial probe (MitoTraker Green, Molecular Probes) because the signal for rhodamine 123 was detectable with the filter used to image the MitoTraker Green. It is not possible from this study to determine whether the oxidation of DHR was mediated by mitochondria-derived ROS.
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Enzymes Involved in Maintaining Cellular Redox
Because of the observed changes in cellular redox (Fig. 6), we sought to examine the three enzymes of the glutathione redox cycle that are important in maintaining cellular redox status: glutathione peroxidase catalyzes the conversion of GSH to GSSG in the presence of hydroperoxides, GSSG reductase converts GSSG to GSH and requires NADPH, and G6PD converts NADP+ to NADPH in the pentose phosphate pathway. A deficiency of one of these enzymes would decrease function of the glutathione redox cycle, disrupt the reduced environment, and result in oxidation of macromolecules, which has been implicated in replicative senescence and aging. The levels of all three enzymes increased as the HUVECs increased in protein content until CPD 44.8, as shown by a maintenance of enzyme activity per milligram of protein (Fig. 8). Activities were also seen to increase per cell during this time (data not shown). At CPD 51.9, glutathione peroxidase increased per milligram of protein (Fig. 8A), whereas GSSG reductase (Fig. 8B) and G6PD (Fig. 8C) decreased per milligram of protein and per cell (data not shown). G6PD actually started to decline at CPD 44.8 (Fig. 8C). This latter enzyme decreased from 0.033 (CPD 6.3) to 0.017 units/mg protein at CPD 51.9. This result indicates that redox cycling is compromised in cells before replicative senescence.
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DISCUSSION |
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Although HUVECs are a culture model of cellular aging, they
demonstrate characteristics of cells aging in vivo. One obvious example
that we and others have found is the change in the size of the cells.
Endothelial cultures established from people of different ages have
shown a wide variation in the cell size (50-250 µm in diameter)
within a single population (50), and examination of
vessels from adults revealed larger endothelial cells interspersed among small endothelial cells (50). We observed a very
small percentage of large cells in our young cultures (<20 CPD), and these cells tended to stain positive for senescence-associated -galactosidase and were often multinucleated (data not shown). Similarly, multinucleated cells (2-10) were observed
in cultures established from vessels (50). As our cultures
increased in CPD, all the cells were seen to increase in size. To
ensure this observation was truly a change in cell size and not just an
artifact of the confluency of the culture, we determined the protein
content per cell. It was evident that the protein content of the
population doubled by CPD 21.9 and remained stable until CPD 44.4. Further increases were detected between CPD 48 and 57.9. Examination of the light scattering properties of three cell populations at three different replicative stages (Fig. 1B) demonstrated that the
cells did slowly increase in size and that the majority of the cells at
a particular age were a similar size. Cells at CPD 33.2 were intermediate in size compared with the small cells at CPD 7.2 and large
cells at CPD 49.8. General increases in cell size have been previously
documented for fibroblasts grown in culture (15). Kumazaki
(30) also correlated an increase in cell area with an
increase in expression of fibronectin mRNA per cell, using in situ
hybridization, for aortic endothelial cells aged in vivo as well as for
HUVECs replicating in culture. The twofold increase in protein content
per cell we detected (Fig. 1A) at CPD 21.9 did correlate
with an increase in the proportion of cells found to be tetraploid
(Table 1 and Fig. 2). Genomic instability does appear to be related to
an increase in cellular protein content. It will be interesting to
determine what the signal(s) is (are) for the two events and whether
genomic instability occurs before the cell size change or whether the
two occur concurrently. Our model suggests it is necessary to study the
population at ~CPD 20 to try to determine the changes in signal
transduction that lead to the genomic instability and protein increases
per cell. Ongoing efforts in our laboratory are devoted to this area of investigation.
This increase in cell size and production of protein would be expected to require an increase in energy and metabolism; however, many studies examining different tissues from aged animals have demonstrated a decrease in mitochondrial function (reviewed in Ref. 47), an increase in mitochondrial proton leak, and a decreased oxygen consumption (25). To address this question, we examined the mitochondrial genome copies relative to nuclear genome, the total cellular oxygen consumption, and cellular ATP content on a per cell basis. All three parameters increased at CPD 26-28, consistent with enhanced oxidative metabolism: the mitochondrial genome-to-nuclear genome ratio increased 3.5- to 4-fold by CPD 28 (Fig. 3A), the total oxygen consumption increased 1.5- to 2-fold at CPD 26-28 (Fig. 4A), and ATP increased ~2-fold at CPD 26 (Fig. 5A). Although we were unable to determine the mitochondrial oxygen consumption, our data suggest that the endothelial cells are able to adapt at this midreplicative life stage to a demand for energy. The existence of this transition point where the cell population increases in size and can maintain oxidative phosphorylation has never been shown before, probably because most studies examine only young and senescent cells, the beginning and end stages of the cell's replicative life. The increase in oxygen consumption, mitochondrial genome copy number, and ATP production could be explained by replication of the mitochondria, decreased mitochondrial turnover, or an increase in biochemical function of the mitochondria. It is possible that the increase in mitochondrial copy number could be due to an increase in ploidy of the mitochondria and/or the formation of "gigantic" mitochondria. Enlarged mitochondria have been isolated from old animals (51). To understand this further, we compared the mitochondrial mass of 10,000 viable cells at CPD 7.2, 33.2, and 49.8 by using MitoTraker Green FM (Fig. 3B). The amount of mitochondrial replication or decreased mitochondrial turnover required to account for a twofold change in function would be expected to substantially increase the mitochondrial mass, whereas the conversion of small mitochondria to large mitochondria or an increase in biochemical function could occur without a large change in the mass. From this study, it appears that although the biochemical function of the mitochondria and the ploidy increased, there was not a substantial change in mitochondrial mass at CPD 33.2, indicating that the mitochondria did not increase in number. It is possible that the mitochondria may have fused to form enlarged mitochondria.
The ratio of the mitochondrial and nuclear genome was stable until CPD 44 (Fig. 3A), as was also found for the oxygen consumption (Fig. 4A). However, at CPD 48 the mitochondrial-to-nuclear ratio declined as did the mitochondrial mass (Fig. 3B). It is likely that the decrease in the mitochondrial-to-nuclear copy number ratio at CPD 48 was due to a decrease in the number of mitochondria, consistent with a decreased mitochondrial mass. Even at CPD 48 the mitochondrial-to-nuclear genome ratio did not decrease below that of CPD 6. Barazzoni et al. (5) detected a decrease in mitochondrial genome copy number in the liver and skeletal muscle, but not in the heart, of old rats. Hypertrophy has also been found to occur with age in the left ventricular myocardium of humans and rodents (31). Rat models of hypertrophy have shown that mitochondrial DNA content increases with an increase in cell size (43). It appears that age-related alterations in the mitochondrial genome number differ depending on the tissue, and variations with age may be related to cell size. Interestingly, the mitochondrial copy number increased to a greater extent than the oxygen consumption and ATP production per 106 cells. Our PCR reaction for the mitochondrial genome is performed over a small region (75 bp) of the D-loop, which contains the origin of replication of the heavy strand. Our comparison of mitochondrial-to-nuclear genome would not have taken into account the extent to which the mitochondrial genome was deleted or mutated, and this may explain the discrepancy.
At CPD 26, the oxygen consumption (Fig. 4A) and the ATP (Fig. 5A) increased by the same amount (~2-fold), which suggests the increase in ATP production was linked to oxidative phosphorylation. However, further increases in cell size at CPD >44 did not result in a further increase in the rate of oxygen consumption, as seen by the decline in oxygen consumption per minute per milligram of protein (Fig. 4C) and a plateau of the oxygen consumption per minute per 106 cells (Fig. 4A). Examination of the ATP production per milligram of protein did not decline with increasing CPD, as would be expected from a decline in mitochondrial function. This suggests that the endothelial cells in mid-late replicative life support their energy needs by glycolysis. In agreement with our work, studies using cultured fibroblasts have also demonstrated that "old" cells were not deficient in energy production, but did utilize more glucose and produce more lactate than "young" cells (21), consistent with an increase in glycolytic activity.
Redox cycling of glutathione is a way that macromolecules can be protected against cellular oxidative stress. Studies have examined the GSH or GSH/GSSG in old and young animals and have found an increase in GSSG (38, 46). We examined the level of GSH and GSSG with respect to the increase in endothelial cell size and protein content and found that endothelial cells maintained a reducing environment, even at the end of their replicative life (Fig. 6). This is reflected in significant increases in cellular GSH as determined per cell or per milligram of protein (Fig. 6, A and B). Because GSH production is stimulated by stress within the cell, this increase infers that the cells were under a form of stress. Curiously, the GSSG per milligram of protein did not significantly increase, even though a significant increase was seen between CPD 42-48 for GSSG per 106 cells. Small increases in GSSG or small quantities of ROS not detectable by probes such as DHR may act as a signal of stress in the cell. It is possible that examination of the mitochondrial glutathione pool may reveal that an oxidative stress was predominantly manifested in the mitochondria. Unfortunately, direct quantification of mitochondrial GSH was prohibited by the large number of cells required to isolate mitochondria for this assay. Notwithstanding, our studies indicate that the overall cellular reducing environment is not compromised in large endothelial cells, consistent with a lack of a major oxidative stress. In fact, the GSH:GSSG ratio increased approximately fourfold (Fig. 6E). A previous study using fibroblasts showed that the GSH per milligram of protein was the same in young and senescent cells, but the senescent cells were not able to increase the GSH content to the same extent as young cells when grown in oxygen tensions >35% (1). On the basis of this finding and our results of decreased GSSG reductase and G6PD (see below), we predict that large endothelial cells of greater replicative age should be more susceptible to oxidative stress. This aspect warrants further investigation. However, in the absence of an imposed oxidant, we find the percentage of cells that exhibit oxidative stress (as measured by DHR oxidation) in the population was zero or very low until CPD >44. At CPD >44, ~15% of the population showed an oxidative stress. From our experience, the cells at this late stage were very vulnerable to physical damage. It is possible that these cells were undergoing apoptosis or necrosis and were unable to make the transition to senescent cells.
The redox cycling of glutathione is dependent upon glutathione peroxidase, GSSG reductase, and G6PD. The activity of these enzymes was increased as HUVEC protein content increased. However, toward the end of the HUVECs' replicative life, G6PD and GSSG reductase activity decreased both per milligram of protein and per 106 cells, whereas glutathione peroxidase increased (Fig. 8). Similar changes were seen in hepatocytes from old rats (46), although other studies using various types of tissues have shown enzyme activities to increase (32), decrease (2), and stay the same (6). Glutathione peroxidase has been shown to be essential for the survival of HUVECs after treatment with peroxides (22). Even though the cells have an increased ability to remove damaging peroxides, a decline in G6PD and GSSG reductase limits the cells' ability to convert GSSG to GSH, which may decrease the overall detoxification capacity and eventually result in GSH depletion. The importance of G6PD was demonstrated by the finding that fibroblasts deficient in this activity senesced at a faster rate in culture (26).
Endothelial cells similar in morphology to our large HUVECs are situated in atherosclerotic lesions and at sites of increased vascular stress and injury. Previous studies have only examined the beginning and the end of replicative life, namely young and senescent cells. In contrast, we examined a number of endpoints related to oxidative stress at multiple CPDs as the cells increased in replicative age to try to understand more about the endothelial cells in vessels that have to replicate to repair injury. Endothelial cells at sites of injury during the progression of vascular disease will likely be at different stages of replicative life. Our results are consistent with the existence of two transition points during the replicative life of endothelial cells: the first occurs where the cells change in size (CPD 22 in this population) but are able to adapt to their new morphology in terms of redox status and oxidative phosphorylation capacity, and the second transition was seen ~10 population doublings before replicative senescence and after the initiation of mitochondrial dysfunction, where the cells show a decline in their ability to recycle GSSG to produce GSH. This work suggests that although endothelial cells adapt and replicate to repair injury, at the end of their replicative life they are extremely vulnerable to damage, and loss of these endothelial cells could result in a perpetuation of vessel injury and endothelial dysfunction.
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ACKNOWLEDGEMENTS |
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We thank L. Coe for isolation of the HUVECs, Dr. B. J. Williams for help with the FISH analysis, D. Chervenack for operating the flow cytometer, and Drs. Grisham and Granger for critical reading of the manuscript.
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FOOTNOTES |
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This work was supported by National Institutes of Health Grants AG-16410, DK-44510, and DK-43785.
Address for reprint requests and other correspondence: L. Harrison, Dept. of Molecular and Cellular Physiology, Louisiana Health Sciences Center, 1501 Kings Highway, Shreveport, LA 71130 (E-mail: lclary{at}lsuhsc.edu).
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.
August 22, 2002;10.1152/ajpcell.00092.2002
Received 28 February 2002; accepted in final form 19 August 2002.
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