© 2000 by European Society of Cardiology
Copyright © 2000, European Society of Cardiology
Temporal regulation of endothelial ET-1 and eNOS expression in intact human conduit vessels exposed to different intraluminal pressure levels at physiological shear stress
Clinical Experimental Research Laboratory, Heart and Lung Institute, Sahlgrenska University Hospital/Östra, Göteborg University, SE 416 85 Gothenburg, Sweden
* Corresponding author. Tel.: +46-31-343-5921; fax: +46-31-191-416 sverker.jern{at}hjl.gu.se
Received 23 March 2000; accepted 24 May 2000
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Abstract |
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 Top Abstract 1 Introduction2 Material and methods3 Results4 DiscussionReferences |
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Objective: By using a computerized vascular perfusion model, we investigated temporal effects of sub-acute pressure elevation on vasomotor behavior and expression of endothelin-1 (ET-1) and endothelial nitric oxide synthase (eNOS) in intact human conduit vessels. Methods: Paired umbilical veins were perfused during 1.5, 3 and 6 h under high/low intraluminal pressure (40/20 mmHg) and at identical shear stress level of 10 dyn/cm2. ET-1 and eNOS gene and protein expression was quantified with real-time reverse-transcribed polymerase chain reaction and quantitative immunohistochemistry, respectively. Results: Pressure induced differential temporal regulation patterns of ET-1 and eNOS gene expression. During the high pressure condition, eNOS mRNA was upregulated after 3 h and leveled off after 6 h of perfusion, while ET-1 mRNA was elevated after 6 h perfusion. Immunohistochemistry verified synchronal changes at the protein level. Significant vasodilation was observed after 3 h in the high-pressure system. Conclusion: Thus, subacute pressure elevation exerts differential effects on the endothelial eNOS/ET-1 expression, which dynamically regulate the vasomotor tone.
KEYWORDS Endothelial function; Mechanotransduction; Endothelins; Nitric oxide; Gene expression; Veins
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1 Introduction |
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TopAbstract1 Introduction2 Material and methods3 Results4 DiscussionReferences |
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The vascular endothelium is continuously exposed to three major types of fluid dynamic forces: shear stress, compressive and circumferential stretch force. It is now commonly accepted that flow-mediated ‘conducted vasodilatation’ in the vascular network is due to the shear-sensing property of the endothelial cells [1,2]. Acute elevation of intraluminal pressure is known to induce myogenic response in the vascular wall [3]. In addition to acute flow-induced vasodilation, which is crucial for short-term modulation of hemodynamic homeostasis, shear stress exerts also subacute and chronic effects on vascular mediators released from endothelial cells [4]. Unfortunately, data from chronic perfusion experiments have been obtained in cell cultures and therefore do not provide any direct evidence on the possible physiological relevance of the findings. It has become increasingly clear, however, that these molecular observations must be related to the overall function of the intact organ. In the present study, we report subacute effects of elevated intraluminal pressure on intact conduit vessels at levels of gene and protein expression, as well as physiological function.
Endothelin-1 (ET-1) is a potent endothelium-derived vasoconstrictor, which causes prolonged increases in vascular tone [5]. ET-1 has been identified as one of the important vasoactive substances associated with pathophysiological conditions such as hypertension, ischemic heart disease, and congestive heart failure [5,6]. In vitro studies of effects of mechanical stress on ET-1 production have provided somewhat conflicting results. While some investigators have shown a transient shear-induced upregulation of ET-1 mRNA expression followed by a chronic downregulation, others have reported a monophasic upregulation of ET-1 [7,8]. Pressure, on the other hand, has been found to enhance the production of ET-1 [9]. Recently, Ziegler et al. demonstrated an increase of ET-1 mRNA expression in cultured cells exposed to a combination of pressure and low shear stress in an ingenious experimental flow device [10]. This finding underscores the importance studying the effects of mechanical stress ET-1 changes in a physiologically relevant setting, in which the complex interplay between mechanical forces can be simulated.
Nitric oxide (NO) as a pivotal vasodilator, antiaggregatory, and anti-proliferative substance, has been shown to be regulated by shear stress both in vitro and in vivo at levels of enzyme activity, protein synthesis, and gene expression [7,11,12]. Pressure in the absence of flow has been shown to depress the production of NO in cultured cells [9].
At the level of endothelial cells, ET-1 has been shown to release NO through ETB receptors, while NO inhibits the production of ET-1 [13]. At the level of smooth muscle cells, NO and ET-1 causes opposite responses, and NO may also increase the expression of ET receptors [14]. Studies using ET receptor antagonists and NO inhibitor indicate that there is a short-term dynamic relationship between ET-1 and NO pathways [15,16]. In cultured endothelial cells, fluid mechanical forces have been shown to influence both mediators separately [8,10]. This negative feedback mechanism is probably of substantial importance for an intact organ.
Due to the opposing actions of these two mediators and the complex interactions between them, it is of great interest to investigate how NO and ET-1 combine to maintain and regulate vasomotor behavior in an intact vessels. To enable an integrative physiological and molecular biological study, we have developed a computerized whole-vessel vascular perfusion model in which conduit arteries or veins can be exposed to well-defined combinations of fluid mechanical forces. In the present study, we investigated the effects of subacute (1.5–6 h) pressure elevation combined with physiological shear stress on vasomotor behavior and vascular NO/ET-1 pathways in intact human conduit vessels.
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2 Material and methods |
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TopAbstract1 Introduction2 Material and methods3 Results4 DiscussionReferences |
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2.1 Vascular perfusion model
The perfusion system has recently been described in detail elsewhere [17]. Shear stress is calculated through the formula
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where 
is wall shear stress, 
P is the pressure drop over the vessel, L is the vessel length, 
viscosity of the fluid, and Q is the flow through the vessel, i.e. shear stress was calculated on-line by continuous measurement of pressure drop and flow-rate. Reynold's number was kept below 400 during the present work to ensure a laminar flow profile.
A schematic of the perfusion system is depicted in Fig. 1. Perfusion circuits are driven by hydrostatic pressure created by the vertical distance between the up- and down-stream reservoirs (1,2). The vascular segment (3) is mounted in a perfusion chamber (4), which is placed in a 37°C water-bath (Tempette Junior TE-8J, Technie, Cambridge, UK). A peristaltic pump (Minipuls 3, Gilson, Villiers le Bel, France; 5) is used to transport the perfusion medium from the lower to the upper reservoir. Constant temperature of 37±0.1°C in the perfusion medium is ensured by a heat exchanger (D720 Pediatric size, Dideco, Mirandola, Italy; 6).
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The perfusion model is operated by a computerized control and feedback system. Data from up- and down-stream pressure transducers (DPT-600, pvb Medizintechnik, Kirchseeon, Germany; 7, 8) and electromagnetic flow meter detector (Toshiba ultrasmall electromagnetic flow meter detector model 334, Toshiba, Tokyo, Japan; 9) are recorded and digitized through a data acquisition PCI-MIO-16XE-50 board (National Instruments, Austin, USA). Digital signals are fed into a Macintosh Power PC Computer 7600/120 MHz (Apple Computer, Santa Clara, CA, USA; 10), equipped with a custom-assembled program developed by us in LABVIEW 4.0. Through computerized control of the height regulator [11] and the proportionating solenoid valve (type 2821 with control electronics type 1094, Bürkert, Ingelfingen, Germany; 12), various combinations of hydrodynamic perfusion parameters can be generated. The software permits continuous real-time monitoring of perfusion pressure (P1, P2), mean intraluminal pressure [(P1+P2)/2], flow-rate, pH values, shear stress, vascular resistance (defined as pressure drop/flow-rate), and Reynold's number. pH of the perfusion medium is continuously measured by digital pH meters (Microprocessor pH meter RE 357, EDT Instruments, Dover Kent, UK; 13). The superfusion and perfusion medium is bubbled by a gas mixture of 90% N2–5% CO2–5% O2 gas (AGA Gas, Stockholm, Sweden; 14). Gas delivery to each circuit is separately controlled by the computer sub-routines through magnetic valves (Type 200, Bürkert, Ingelfingen, Germany; 15) to keep pH at predefined target levels.
2.2 Experimental protocol
The studies were approved by the local Ethics Committee of the Göteborg University, and were conducted according to the principles of the Declaration of Helsinki.
2.2.1 Preparation procedure
Human umbilical cords were collected from the maternity ward of the hospital immediately after vaginal delivery. The umbilical cord was divided into two segments for perfusion in parallel. Each perfused segment had a length of at least 20 cm. Placental and fetal segments were randomized to high or low-pressure circuits to eliminate any systematic variation due to differences between the two vascular segments. Both vessel segments were carefully cannulated and rinsed with PBS solution to remove any remaining blood. Prepared vessels were kept in situ in the organ perfusion chamber and connected to the perfusion circuit. After a 10-min non-recirculating wash-out period, vessels were equilibrated for another 20 min under constant mean intraluminal pressure and flow-rate of 20 mmHg and 10 ml/min, respectively.
2.2.2 Perfusion under high versus low intraluminal pressure
Paired umbilical veins were perfused under high (target 40 mmHg) or low (target 20 mmHg) intraluminal pressure at constant shear stress of 10 dyn/cm2. Three series of perfusion experiments were performed for 1.5, 3 and 6 h.
2.3 Immunohistochemistry
Paired umbilical veins were perfused according to each of the perfusion protocols. At the end of each experiment, tissue samples were collected for immunohistochemical study. Briefly, a vascular segment of 
1 cm in the middle of each umbilical cord was cut out and fixed in 4% formalin at room temperature for at least 24 h. The specimens were embedded in paraffin, sectioned into 5-µm slices. Paired slices from same experiment were mounted on Superfrost Plusglasses (Menzel, Merck, Poole, UK). Sections were then deparaffinized in xylene, rehydrated in graded alcohol. A 0.01% solution of Protease type XXIV in 0.05 M Tris buffer, pH 7.6, containing 0.01% calcium chloride was used for antigen retrieving. Together with coverplates (Shandon Life Sciences, Pittsburg, PA, USA), the slides were placed in cassettes (Labex Instrument, Helsingborg, Sweden) with Cadenza buffer (Shandon).
For immunohistochemical staining, universal streptavidin/biotin immunoperoxidase detection system with diaminobenzidine (DAB) chromogen (OmniTags Plus, Immunon, Lipshaw, USA) was used. To quench endogenous peroxidase activity and reduce background staining, sections were treated for 5 min at room temperature with 3% hydrogen peroxidase. Non-specific binding of antibodies was reduced by treatment with a protein blocking agent. The slides were incubated with a 1:20 dilution of mouse monoclonal antibody against eNOS (606-259-1550, Affinity Research Products, Mamhead Castle, UK) for 30 min. Monoclonal antibody against ET-1 (Sigma) was used to detect ET-1 antigen with 1:15 dilution. Non-immune mouse serum incubated preparations were used as negative control (Immunon). After wash in buffer, the sections were incubated with a biotinylated secondary antibody for 30 min and streptavidin–peroxidase reagent for another 30 min. In the end, chromogen solution was added and the slides were counterstained with hematoxylin. After washing in water, the sections were dehydrated and mounted with resin-based mounting medium.
Matched high/low pressurized vascular preparations were examined pairwise by light microscopy (Olympus BX-60, Olympus, Tokyo, Japan). Hematoxylin counterstaining revealed intact structure of the vessel wall in all stimulated umbilical veins in both high and low pressurized veins. Lack of immunostaining in negative controls verified the specificity of the eNOS and ET-1 antibody. Three randomly selected area of each vessel preparation were digitized with 400 times enlargement with an utility software C-2.1 through a digital camera (Olympus DP10). Staining intensity of the obtained digital images were thereafter measured with an automatic digital image analysis routine (Fig. 2). Briefly, positive staining area was selected automatically by using a positive staining standard color. The selected area was spectrally separated in red (R), blue (B), green (G) and grey (Gr) and total optical density was calculated through the following formula: OD=area for positive stainingx(1/red intensity+1/blue intensity+1/green intensity+1/grey intensity). Average optical density for each vessel preparation was compared pairwise.
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2.4 Quantification of eNOS mRNA by reverse transcriptase real-time PCR
2.4.1 Isolation of total RNA
Following perfusion, endothelial cells were eluted by incubating each vessel with 0.1% collagenase for 12 min at 37°C. The incubation protocol was optimized to ensure a yield of >90% endothelial cells as validated by immunocytochemistry. The treatment resulted in an almost complete removal of endothelial cells as verified by scanning electron microscopy. The incubation medium was rinsed out by PBS and the cell suspension was then centrifuged for 10 min at 260 g, and the cell pellet was resuspended in denaturing solution (solution D; 4 M guanidinium thiocyanate, 25 mM sodium citrate, pH 7, 0.5% sarcosyl and 0.1 M 2-mercaotoethanol) according to Chomczynski. Total RNA concentrations were determined by A260 and A280 spectrophotometric measurements (A260/A280=1.7–1.9) using a Gene Quant II RNA/DNA calculator (Amersham Pharmacia Biotech, Uppsala, Sweden). Integrity of precipitated total RNA was determined by 1% agarose gel electrophoresis during denatured conditions.
2.4.2 Reverse transcription
Reverse transcription of 1 µg total RNA was carried out in a total volume of 20 µl reaction mixture consisting of 5 mM MgCl2, 1 mM dNTP mix, 50 mM KCl, 10 mM Tris–HCl (pH 8.3), 2.5 µM random hexamer, 1 U/µl RNA-guard, 2.5 U MuLV reverse transcriptase. Samples were incubated at 20°C for 10 min, at 42°C for 15 min, at 99°C for 5 min, and finally at 5°C for 5 min.
2.4.3 Quantitative real-time RT-PCR assay
Relative quantification of mRNA was performed on a ABI Prism® 7700 Sequence Detector (Perkin-Elmer Applied Biosystems, Foster City, CA, USA). Briefly, the assay uses the 5'nuclease activity of Taq polymerase to cleave a reporter dye from a non-extendable hybridization probe during the extension phase of the PCR reaction. The fluorogenic probe is labeled with a reporter dye (FAM; 6-carboxy-fluorescein; emission maximum 518 mm) at the 5' end and a quencher dye (TAMRA; 6-carboxy-tetramethyl-rhodamine; emission maximum 582 nm) at its 3' end via a linker arm nucleotide (LAN). When the probe is intact, reporter dye emission is quenched due to the physical proximity of the reporter and quencher dyes. During the extension phase, the reporter dye is released and the increase in dye emission is monitored in real-time. The threshold cycle (CT) is defined as the fractional cycle number at which the reporter fluorescence reaches a certain level (i.e. usually 10 times the standard deviation of the baseline). As shown by Higuchi et al. [18], there is a linear relationship between CT and the log of initial target copy number. For amplicons designed and optimized according to the manufacturer's guidelines, amplification efficiency is typically close to one, i.e. product accumulation increases two-fold until the plateau phase is reached.
2.4.4 Oligonucleotides for TaqMan® PCR assay
Oligonucleotide primers and TaqMan probes were designed by using PRIMER EXPRESS version 1.0 (Perkin-Elmer Applied Biosystems) based on sequences from the GenBank database as follows: eNOS (accession: D26607
[GenBank]
) [19] and β-actin (accession: E00829
[GenBank]
) [20] (Table 1). Constitutively expressed β-actin was selected as endogenous control to correct for potential variation in RNA loading or efficiency of the amplification reaction. To verify that the amplification product was a target gene, the amplified RT-PCR products of one sample were sequenced.
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2.4.5 PCR amplification
All reactions were performed using the ABI PRISM® 7700 sequence detector (Perkin-Elmer Applied Biosystems) equipped with a Gene-amp PCR System 9600 and a CCD (charge-coupled device) camera for measuring the fluorescent emission spectra from 500 to 650 nm. Frosted microAmp optical tubes (part no. N801-0933, Perkin-Elmer) were used as reaction tubes and tube caps were specially designed to prevent light scattering. Reaction conditions were programmed on a Power Macintosh 7100 (Apple Computer) linked directly to the Model 7700 Sequence Detector.
For amplification of the 258-bp eNOS product, one µl of cDNA diluted 1:4 was added to the PCR reaction mixture consisting of Taqman buffer A, 5 mM MgCl2, 0.2 mM dNTP mix (20 mM dUTP and 10 mM of dATP, dCTP, and dGTP), 1.25 U Taq gold polymerase, 0.5 U AmpErase® UNG, 15 pmol of the forward primer, 2.5 pmol of the reverse primer, and 5 pmol probe in a final volume of 50 µl.
For amplification of the 71-bp ET-1 product, 1 µl of cDNA diluted 1:4 was added to the PCR reaction mixture consisting of Taqman buffer A, 5 mM MgCl2, 0.2 mM dNTP mix (20 mM dUTP and 10 mM of dATP, dCTP, and dGTP), 1.25 U Taq gold polymerase, 0.5 U AmpErase UNG, 0.3 pmol of the forward primer, 0.3 pmol of the reverse primer, and 0.1 pmol probe in a final volume of 50 µl.
For amplification of the 76-bp β-actin product, 1 µl of cDNA diluted 1:4 was added to the PCR reaction mixture consisting of Taqman buffer A, 5 mM MgCl2, 0.2 mM dNTP mix (20 mM dUTP and 10 mM of dATP, dCTP, and dGTP), 1.25 U Taq gold polymerase, 0.5 U AmpErase UNG, and 15 pmol of each primer, and 5 pmol probe in a final volume of 50 µl.
Thermal cycling conditions included the following steps: 2 min at 50°C, then the reaction mixture was preheated for 10 min at 95°C before the PCR cycles started. A 50-cycle two-step PCR was performed consisting of 15 s at 95°C and 1 min at 60°C (Abi Prism 7700). All samples were amplified simultaneously in triplicate in one assay run.
2.4.6 Methodological validation
The average amount of extracted RNA from the endothelial cells was approximately 5 µg per 20 cm umbilical vessel. The quantity of total cellular RNA extracted from each vessel segment was similar in vessels exposed to high versus low pressure. Transcript levels of the endogenous control β-actin were independent of pressure stimulation. Also, no effect of either stimulation conditions was observed when β-actin mRNA levels were expressed relative to GAPDH (data not shown). A standard curve was obtained by performing amplifications of the three target gene cDNAs in a series of two-fold serial template dilutions of total endothelial cell RNA from 1:1 through 1:32. For both mRNAs, linear inverse correlation were observed between CT values (cycles at threshold lines) and the amount of applied cellular RNA (R2=0.999, 0.995 and 0.999 for eNOS, ET-1 and β-actin, respectively) (Fig. 3). The input amount (TGia) was determined by the following formula: TGia=[(cell containing CT value)–b]/m, where b is the y-intercept and m is the slope of the standard curve (Perkin-Elmer Applied Biosystems; User Bulletin no. 2, December, 1997). Normalized amount of target gene is expressed as ratio between the target and control gene cDNA.
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2.5 Drugs
Unless otherwise state, all reagents and drugs were purchased from Sigma (St. Louis, MO, USA). PCR consumables were purchased from Perkin-Elmer Applied Biosystems.
2.6 Statistical analysis
Data are expressed as mean and standard error of the mean (SEM) unless otherwise stated. Parametric methods (ANOVA and t-test) were used for evaluation of changes in response to the different perfusion conditions. For comparison of gene expression during the different stimulation conditions, the ratios of eNOS and ET-1 over β-actin cDNA were evaluated after logarithmic transformation of data. Contrast analysis was applied when the overall ANOVA indicated a significant main effect of treatment or a significant interaction effect. Significance tests were considered significant at P<0.05 (two-tailed test).
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3 Results |
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TopAbstract1 Introduction2 Material and methods3 Results4 DiscussionReferences |
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3.1 Hemodynamic data during 6 h perfusion
Hemodynamic data from all experiments included in the studies are shown in Fig. 4. During high and low pressure conditions, average pressure levels were 39.9±0.0 and 20.0±0.0 mmHg, respectively (t-test, P<0.0001). Mean shear stress was maintained at identical levels in both systems (10.0±0.0 dyn/cm2; NS). The average flow-rate during the whole experiment was 79.0±2.2 and 52.5±1.6 ml/min in the high and low-pressure systems, respectively (t-test, P = 0.0001).
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3.2 Quantification of eNOS and ET-1 mRNA
High compared to low intraluminal pressure caused significant different temporal regulation patterns of the two target genes (ANOVA target genextime, P = 0.01). At 1.5 and 3 h of pressure stimulation, the ET-1/β-actin ratio reached a level of 175±44% (log t-test, P = 0.16) and 145±33% (P = 0.60) of the baseline level. After 6 h of perfusion, the ET-1/β-actin ratio increased to 211±54% (log t-test, P = 0.023) in the high compared to the low pressure condition.
The eNOS/β-actin ratio showed an initial transient decrease to 75±24% (log t-test, P = 0.14, NS), but then increased significantly to 305±90% of baseline after 3 h of perfusion (log t-test, P = 0.049). After 6 h of perfusion, eNOS/β-actin ratio levelled off at 167±34% (log t-test, P = 0.09, NS) (Fig. 5).
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3.3 Immunohistochemistry
In the perfused veins, immunoreactivity for eNOS was detected mainly in the endothelium, but also and to a substantial extent in the smooth muscle cells layer. Digital spectral analysis of the paired vessel preparations revealed approximately 120% higher eNOS immunoreactivity in all high-pressure stimulated veins compared to those perfused under low pressure conditions after 3 h perfusion (Fig. 6). No systematic differences in the intensity of immunostaining were detected in the 1.5- and 6-h experimental series (data not shown).
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Immunostaining optical density of ET-1 was increased by approximately 130% after 6 h of perfusion in the high compared to low pressure system in all sections, as revealed by digital spectral analysis (Fig. 6). No systematic differences were observed between the high and low-pressure systems after 1.5 or 3 h of perfusion.
3.4 Change of vascular resistance during 6 h perfusion
Percentage changes in vascular resistance in high and low-pressure systems during 6 h of perfusion are shown in Fig. 7. In vessels perfused under low-pressure condition, a transient vasoconstriction was observed during the 3rd and 4th hour of perfusion and the vascular resistance returned to baseline level after 6 h of perfusion. By contrast, in high pressure perfused vessels a significant vasodilatation occurred after 3 h of perfusion, which lasted throughout the 4th hour and returned to baseline level at the end of the perfusion period. Percentage changes of vascular resistance were 84.9±6.5% and 120.1±15.8% (contrast analysis, P = 0.0032) after 3 h and 84.1±7.7% and 121.8±17.0% (P = 0.0017) after 4 h of perfusion in the high and low-pressure systems, respectively.
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4 Discussion |
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TopAbstract1 Introduction2 Material and methods3 Results4 DiscussionReferences |
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In the present study, we demonstrate that subacute elevation of intraluminal pressure in the presence of physiological shear stress increases the expression of ET-1 in the vascular endothelium and induces a transient upregulation of eNOS expression. Gene expression of the two mediators was differentially regulated by pressure. The dynamic regulation patterns were accompanied by significant changes in intracellular protein levels and were shown to be of functional importance for the vasomotor behavior of the vessel. Our findings indicate that increased intraluminal pressure may have a transient effect on the vascular wall in terms of vasodilatation, which may be due to an elevated eNOS/ET-1 ratio. Sustained elevation of intraluminal pressure tends to diminish the initial vasodilatation by increased ET-1 and an unchanged eNOS expression.
Regulation of ET-1 mRNA expression by shear appears to be cell-type specific and dependent on the magnitude of shear stress. Recent findings suggest that in venous endothelial cells, there appears to be a threshold level of shear below which ET-1 is down-regulated and above which an up-regulation is observed [8]. Shear-induced ET-1 expression has also been shown to depend on cell shape change in the vascular endothelium. Isolated pressure seems to enhance ET-1 release partially through activation of phospholipase C and protein kinase C in human umbilical vein endothelial cells (HUVECs) [9]. However, effects of combined intraluminal pressure and shear on vascular ET-1 production are poorly understood. Our present findings provide evidence that intraluminal pressure exerts significant modulating effects on ET-1 gene expression independently of the shearing force.
We have recently shown that shear stress rather than intraluminal pressure appears to be the major modulating factor for NO metabolism. A 6-h period of high-shear perfusion under physiological pressure significantly increased both eNOS gene and protein expression, and endowed the endothelial cells with a markedly enhanced capacity for stimulated NO production [21]. By contrast, prolonged (6 h) perfusion at elevated intraluminal pressure and normal shear stress had no effect on NO metabolism. Whereas a similar stimulatory effects of shear stress on NO production has been demonstrated previously in vitro, exposure of endothelial cells to increased strain (suggested to simulate an isolated increase in intraluminal pressure) has been shown to down-regulate NO production [9]. However, when the combined effects of both the flow and pressure induced forces were investigated in the present study, the integrated effects were found to be different from those predicted from the in vitro observations. Indeed, short-term elevation of intraluminal pressure caused a transient upregulation of eNOS expression between 3 and 4 h after onset of high-pressure stimulation. Interestingly, this stimulatory effect on the NO pathway is apparently of functional significance since it was found to be associated with a significant transient vasodilation with a similar temporal pattern. The importance of evaluating these effects in living whole-vessel preparation is further underscored by the complex interactions between the ET-1 and eNOS pathways at various cellular and inter-cellular levels.
Somewhat surprisingly, intraluminal pressure at physiological shear stress increased immunoreactivity for eNOS not only in the endothelial cells but also in the vascular smooth muscle cell layers. The mechanism by which eNOS is upregulated in the smooth muscle cells is unclear. However, we have recently observed a similar induction in vessels exposed to elevated shear stress [21]. The validity of this observation was verified by similar immunostaining patterns with a second highly specific antibody, and Western blotting demonstrated distinct bands of the expected molecular size of 140 kDa in both cell types. The specificity of the antibodies was also verified by peptide competition experiments by using eNOS specific blocking peptides. These findings clearly indicate that eNOS is present not only in the endothelium but also in the smooth muscle cells of the umbilical vein. Bloch et al. have previously shown that eNOS was expressed in smooth muscle cells in human corpus cavernosum [22], while Papadaki et al. found iNOS in cultured smooth muscle cells [23]. Thus, presence of various NOS isoforms may be tissue-dependent.
Chronic exposition to elevated shear stress combined under conditions of physiological pressure induced amplitude-dependent vasodilation in human umbilical veins (Gan, 2000, in preparation). This vascular negative feedback mechanism is important for the hemodynamic homeostasis and accounts for the subacute adaptation of the vascular wall to increased hemodynamic load [24]. Acute and subacute elevation of intraluminal pressure may occur in conduit vessels during impaired vasodilation in distal resistance arteries. Pressure induced transient vasodilation we reported in the present study, may serve as a secondary feedback mechanism through enhanced NO pathway, which is capable of dilating adjacent down-stream vessels. However, this vasodilation was short-lasting and was counterbalanced by increased ET-1 expression during sustained pressure elevation.
A potential limitation of the present study is that it was performed in umbilical veins and caution must therefore be exercised when attempting to extrapolate our findings to other conduit vessels. However, many previous in vitro studies in this field have been performed in HUVEC cultures [9,25]. The current experiments can therefore be regarded as extensions of these observations to a whole-vessel level. Furthermore, use of vessels of non-human origin involves potential pitfalls, since substantial differences in vessel phenotype expression may exist between human and non-human vascular tissues due to variations in genetic set-up. For instance, the bovine eNOS promoter region contains nine shear-stress responsive elements whereas the corresponding region in man contains two, of which only one has been shown to confer shear responsiveness [26].
In summary, the intact vascular wall responds to acute elevation of intraluminal pressure by a subacute compensatory vasodilatation, which may be a consequence of/is associated with an up-regulated NO and down-regulated ET-1 expression. Sustained elevation of pressure alters the balance between the NO and ET-1 systems by inducing an elevated ET-1 expression, and the initial vasodilating effect of pressure elevation is diminished. By using the computerized vascular perfusion model, we show that the dynamic molecular and physiological responses of the vascular wall to complex hemodynamic forces can be studied in an intact living vessel.
Time for primary review 21 days.
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Acknowledgements |
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The authors are indebted to laboratory technicians Hannele Korhonen and Camilla Ejdestig for excellent technical assistance throughout the study. This work was supported by grants from the Swedish Medical Research Council (project no. 09046), the Bank of Sweden Tercentenary Foundation, the Swedish Heart–Lung Foundation, the Swedish Hypertension Society, the Åke Wiberg Foundation, and the Gothenburg Medical Society.
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