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. 2018 Aug 18;596(18):4361–4374. doi: 10.1113/JP275611

Chronic exercise impairs nitric oxide pathway in rabbit carotid and femoral arteries

Patricia Marchio 1,†, Solanye Guerra‐Ojeda 1,†, José M Vila 1, Martín Aldasoro 1, Soraya L Valles 1, Carlos Soler 1, Maria D Mauricio 1,✉
PMCID: PMC6138298  PMID: 29968308

Abstract

Key points

  • Some of the beneficial effects of exercise in preventing vascular related diseases are mediated by the enhancement of endothelial function where the role of nitric oxide (NO) is well documented, although the relevance of calcium activated potassium channels is not fully understood.

  • The impact of oxidative stress induced by training on endothelial function remains to be clarified.

  • By evaluating different endothelial vasodilator pathways on two vascular beds in a rabbit model of chronic exercise, we found a decreased NO bioavailability and endothelial nitric oxide synthase expression in both carotid and femoral arteries.

  • Physical training induced carotid endothelial dysfunction as a result of an increase in oxidative stress and a reduction in superoxide dismutase expression.

  • In the femoral artery, the lower production of NO was counteracted by an increased participation of large conductance calcium activated potassium channels, preventing endothelial dysfunction.

Abstract

The present study aimed to evaluate the effects of chronic exercise on vasodilator response in two different arteries. Rings of carotid and femoral arteries from control and trained rabbits were suspended in organ baths for isometric recording of tension. Endothelial nitric oxide synthase (eNOS), Cu/Zn and Mn‐superoxide dismutase (SOD), and large conductance calcium activated potassium (BKCa) channel protein expression were measured by western blotting. In the carotid artery, training reduced the relaxation to ACh (10–9 to 3 × 10–6 m) that was reversed by N‐acetylcysteine (10–3 m). l‐NAME (10–4 m) reduced the relaxation to ACh in both groups, although the effect was lower in the trained group (in mean ± SEM, 39 ± 2% vs. 28 ± 3%). Physical training did not modify the relaxation to ACh in femoral arteries, although the response to l‐NAME was lower in the trained group (in mean ± SEM, 41 ± 5% vs. 17 ± 2%). Charybdotoxin (10–7 m) plus apamin (10–6 m) further reduced the maximal relaxation to ACh only in the trained group. The remaining relaxation in both carotid and femoral arteries was abolished by KCl (2 × 10–2 m) and BaCl2 (3 × 10–6 m) plus ouabain (10–4 m) in both groups. Physical training decreased eNOS expression in both carotid and femoral arteries and Cu/Zn and Mn‐SOD expression only in the carotid artery. BKCa channels were overexpressed in the trained group in the femoral artery. In conclusion, chronic exercise induces endothelial dysfunction in the carotid artery as a result of oxidative stress. In the femoral artery, it modifies the vasodilator pathways, enhancing the participation of BKCa channels, thus compensating for the impairment of NO‐mediated vasodilatation.

Keywords: Nitric oxide, endothelial dysfunction, calcium activated potassium channels, oxidative stress, chronic exercise

Key points

  • Some of the beneficial effects of exercise in preventing vascular related diseases are mediated by the enhancement of endothelial function where the role of nitric oxide (NO) is well documented, although the relevance of calcium activated potassium channels is not fully understood.

  • The impact of oxidative stress induced by training on endothelial function remains to be clarified.

  • By evaluating different endothelial vasodilator pathways on two vascular beds in a rabbit model of chronic exercise, we found a decreased NO bioavailability and endothelial nitric oxide synthase expression in both carotid and femoral arteries.

  • Physical training induced carotid endothelial dysfunction as a result of an increase in oxidative stress and a reduction in superoxide dismutase expression.

  • In the femoral artery, the lower production of NO was counteracted by an increased participation of large conductance calcium activated potassium channels, preventing endothelial dysfunction.

Introduction

Cardiovascular diseases are one of the leading causes of mortality in the western population and almost half of them could be prevented by lifestyle modification (Nocon et al. 2008; Thijssen et al. 2010; Piepoli et al. 2016). Control of cardiovascular risk factors such as hypercholesterolaemia, hyperglycaemia, obesity or hypertension, coupled with physical exercise, are effective strategies for reducing primary and secondary vascular events and risk of death from cardiovascular diseases by 20–40% (Nocon et al. 2008; Vanhoutte et al. 2009; Thijssen et al. 2010; Piepoli et al. 2016).

During exercise, blood flow is redistributed, increasing towards the active skeletal muscles at the expense of a reduction to inactive or less active organs such as renal and splanchnic circulation (McAllister, 1998; Jendzjowsky and DeLorey, 2012). The increase of shear stress during exercise is a powerful stimulus to training‐induced vascular adaptations, including enhancement of endothelial function and arterial remodelling (Green et al. 2011). The degree of participation of endothelial relaxing factors such as nitric oxide (NO), prostacyclin (PGI2) and endothelium‐derived hyperpolarizing factor (EDHF) in the regulation of vascular tone differs along the arterial tree. NO prevails in large conduit arteries, whereas EDHF is more relevant in smaller vascular beds, such as coronary arteries or resistance vessels (Chen et al. 2001; Villar et al. 2006). Most studies in health and disease demonstrate that physical training enhances endothelial‐dependent vasodilatation mainly by releasing NO and, to a lesser extent, EDHF and PGI2 (Chen et al. 2001; Heylen et al. 2008). This response is dependent on the vascular bed and duration of training (Chen and Li, 1993; McAllister et al. 1996, 2005). In addition, modality and intensity of training also influence vascular adaptations. The increased frequency and intensity of physiological physical forces in the vascular system, including shear stress, is a powerful inductor of intermittent vascular generation of reactive oxygen and nitrogen species (RONS) (Kojda and Hambrecht, 2005). Depending on concentration, and as a result of a phenomenon known as hormesis, the increase of RONS production could be beneficial or detrimental for the biological functions. In this sense, low to moderate concentrations of RONS such as superoxide anion (O2 −) or its dismutated product hydrogen peroxide produced during moderate physical training induce the expression of endogenous anti‐oxidant enzymes such as mitochondrial superoxide dismutase (MnSOD) or glutathione peroxidase. By contrast, excessive production of RONS induced by high intensity exercise leads to cell damage by oxidizing molecules and impairing their biological function (Gomez‐Cabrera et al. 2008; Powers et al. 2011). Therefore, the present study aimed to evaluate the effects of chronic exercise on vascular function in conduit arteries exposed to normal or elevated shear stress. Accordingly, we have chosen (i) the carotid artery as a representative vessel for the cerebral circulation and for its clinical relevance as being a predictor of vascular diseases (Chen and Li, 1993; Kemi et al. 2005) and (ii) the femoral artery as a conduit vessel supplying an active skeletal muscle area (Harris et al. 2010).

Methods

Ethical approval

This study was approved by the Bioethics Committee of the School of Medicine of the University of Valencia, Spain. Animals were handled according to the European Directive 2010/63/EU. Animals were acquired from Granja San Bernardo, Navarra, Spain, and transported under conditions specified in the Real Decreto 53/2013, which establishes the basic rules applicable for the protection of animals used in experimentation and other scientific purposes (Spain). Rabbits were housed according to our institutional guidelines (12:12 h light/dark cycle at a constant room temperature of 22°C and 60% humidity, with standard rabbit chow and water available ad libitum). The animals were killed with pentobarbital 100 mg kg−1 IV.

The authors understand the ethical principles under which The Journal of Physiology operates and confirm that the methodology utilized in this study complies with the animal ethics checklist outlined in Grundy (2015).

Animals and study design

Thirty‐two male New Zealand white rabbits (Oryctolagus cuniculus) were used in the present study. After 4 days of familiarization with the running on a motor‐driven treadmill, the animals were divided into two experimental groups: trained (n = 16) and control (n = 16). Animals that did not run adequately during familiarization because they stopped frequently or ran irregularly were excluded from the study. Animals in the control group were housed in their cages, whereas rabbits in the trained group were submitted to a protocol of chronic exercise. Briefly, rabbits ran 5 days a week for 6 weeks at 0.33 m s–1. Each training session was divided into six periods consisting of 4 min of running separated by 1 min of rest (Such et al. 2008). Once the training protocol was completed, blood samples were taken from the marginal vein of the ear in both groups of study. Then, animals were killed and arteries and soleus muscles were removed. Samples were stored in liquid nitrogen at −80°C or in 4% paraformaldehyde at 4°C until analysis, except for the artery rings used for the study of vascular reactivity, which were placed in chilled Krebs–Henseleit solution.

Drugs for organ bath were prepared daily in physiological saline solution. All substances and reagents used in this study were purchased from Sigma Chemical Co. (St Louis, MO, USA) unless otherwise specified.

Citrate synthase activity assay

To confirm the effectiveness of the training protocol, the citrate synthase activity was measured in the soleus muscle by the spectrophotometric method of Srere (1969). Activity is expressed as product formed in relation to protein in tissue (μmol min–1 g–1).

Organ bath experiments

Arteries were cleaned and cut into rings (3 mm in length) for isometric recording of tension. Two stainless steel L‐shaped pins were introduced through the lumen of the vessel. One pin was fixed to the wall of the organ bath and the other was connected to a force‐displacement transducer (FT03; Grass Instruments, West Warwick, RI, USA). Changes in isometric force were recorded on a Macintosh computer (Apple Corp., Cupertino, CA, USA) using Chart, version 7 and a MacLab/8e data acquisition system (AD Instruments). Each ring was suspended in a 4 mL bath containing modified Krebs–Henseleit solution composed of (mm) 115 NaCl, 4.6 KCl, 1.2 MgCl2·6H2O, 2.5 CaCl2, 25 NaHCO3, 11.1 glucose and disodium 0.01 EDTA. The solution was equilibrated with 95% O2 and 5% CO2 to obtain a pH in the range 7.30–7.40. Temperature was held at 37°C. The optimal resting tension for carotid and femoral arteries was 1 g. The arterial rings were allowed to attain a steady level of tension during a 2 h accommodation period. The contractile capacity of vascular smooth muscle was then evaluated by the maximal response to KCl (60 mm). The endothelium was considered functional if immediate relaxation to a single concentration of ACh (10–6 m) in rings precontracted with noradrenaline was ≥70%. Arterial rings without functional endothelium in control conditions were excluded from the study. To evaluate endothelial‐dependent vasodilatation, concentration–response curves to ACh (10–9 to 3 × 10–6 m) were performed in arterial rings precontracted with noradrenaline (10–7 to 10–6 m, to obtain a contraction of ∼75% of the maximal contraction to KCl of each vascular segment) under the conditions: (i) in the absence of inhibitors (control response); (ii) in the presence of indomethacin (10–5 m) to inhibit the production of PGI2; (iii) in the presence of l‐NAME (10–4 m) to inhibit the eNOS; (iv) in the presence of indomethacin plus l‐NAME to inhibit the production of both PGI2 and NO; and (v) in the presence of indomethacin plus l‐NAME combined with KCl (2 × 10–2 m) to block K+ channel activity. To investigate the nature of K+ channel activation, the concentration–response curves to ACh were performed in the presence of indomethacin plus l‐NAME combined with the inhibitors: (i) charybdotoxin (10–7 m), a blocker of both intermediate and large conductance Ca2+‐activated K+ channels (IKCa and BKCa, respectively), plus apamin (10–6 m), a blocker of small conductance Ca2+‐activated K+ channels (SKCa); (ii) glibenclamide (10–6 m), a selective blocker of ATP‐sensitive K+ channels; (iii) barium (BaCl2) (3 × 10–6 m), a blocker of inwardly rectifying K+ (KIR) channels; and (iv) in the presence of BaCl2 (3 × 10–6 m) plus ouabain (10‐4 m), an inhibitor of Na+, K+‐ATPase. To evaluate which type of Ca2+‐activated K+ channels was implicated in the vasodilator response to ACh, we separately incubated femoral segments with apamin (10–6 m), TRAM‐34 (10−6 m), an IKCa blocker, charybdotoxin (10–7  m) and iberiotoxin (10–7 m), a BKCa blocker. To evaluate the presence of oxidative stress, carotid rings were incubated with the RONS scavenger N‐acetylcysteine (NAC) (10–3 m) and relaxation to ACh (10–9 to 3 × 10–6 m) was evaluated. Endothelial independent relaxation was assessed by concentration–response curves to sodium nitroprusside (10–9 to 3 × 10–6 m). Control and experimental (after 20 min of incubation for inhibitors and NAC) concentration–response curves were obtained from separate preparations in both study groups.

Western blot analysis

Vascular tissues were homogenized in lysis buffer (0.125 m Tris‐HCl, pH 6.8, 2% SDS, 19% glycerol and 1% v/v protease inhibitors) and centrifuged at 12,000 g for 15 min at 4°C. Protein concentration was determined using a modified Lowry method (Peterson, 1977). Then, 0.5% (v/v) 2‐mercaptoethanol and 1% bromophenol blue were added and the samples were heated for 5 min at 90°C. Proteins (40 μg) were separated on SDS‐PAGE gels and transferred to nitrocellulose membranes in a humid environment using a transfer buffer (25 mm Tris, 190 mm glycine and 20% methanol). The following primary antibodies were incubated overnight at 4°C: polyclonal eNOS antibody (Ab‐1177; dilution 1:1000; catalogue no. 21170‐2; Signalway‐Antibody, College Park, MD, USA); polyclonal Cu/Zn‐SOD antibody (SOD1; dilution 1:500; catalogue no. ADI‐SOD‐101; Enzo Life Scences, Farmingdale, NY, USA); monoclonal Mn‐SOD antibody (SOD2; dilution 1:500; catalogue no. sc‐137254; Santa Cruz Biotechnology, Santa Cruz, CA, USA); and polyclonal Maxi Potassium channel alpha antibody (BKCa; dilution 1:500; catalogue no. ab3586; Abcam). After incubation, membranes were washed three times with wash buffer TBS + Tween 20 and incubated for 1 h at room temperature with the secondary antibodies anti‐rabbit IgG, HRP‐linked antibody (dilution 1:1000; catalogue no. 7074; Cell Signaling Technology, ) or goat anti‐mouse IgG (H+L) secondary antibody, HRP (dilution 1:10,000; catalogue no. 31430; from Thermo Fisher Scientific, Waltham, MA, USA). Membranes were washed three times and the enhanced chemiluminescence method (Amersham Biosciences, Barcelona, Spain) was used for antibody detection. Autoradiography signals were assessed using the digital image system ImageQuant LAS 4000 (GE Healthcare, Little Chalfont, UK). The primary antibody monoclonal α‐tubulin (B‐7) (dilution 1:1000; catalogue no. sc 5286; Santa Cruz Biotechnology) was used as a control for the amount of protein.

Plasma determination of malondialdehyde

Malondialdehyde (MDA) was measured by high‐performance liquid chromatography (HPLC) as described previously (Wong et al. 1987). The hydrolysis of lipoperoxides in plasma and the subsequent formation of an adduct between TBA and MDA is the basis of this methodology. This adduct can be detected by HPLC in reverse phase and quantified at 532 nm. The technique was performed under isocratic conditions where the mobile phase is a mixture of monopotassium phosphate 50 mm (pH 6.8) and acetonitrile (70:30). Concentrations of MDA in the samples were determined by comparison with standards.

Morphometric analysis

Carotid and femoral arteries were collected and cut into rings (1 cm in length) and fixed in with 4% paraformaldehyde dissolved in PBS. The arteries were further dehydrated in alcohol and embedded with paraffin blocks. Four independent serial cross‐sections (5 μm) were obtained, fixed overnight on slides, and subsequently stained with haematoxylin and eosin. For morphometric analysis, digital images were acquired (TIFF format, 36‐bit colour, 2560 × 1920 pixels) with a DFC450C digital camera and a DMI 3000 microscope (Leica Microsystems, Wetzlar, Germany) and analysed with Image J (NIH, Bethesda, MD, USA). We estimated the intima and media thickness calculating the mean of four measures per image at 0, 90, 180 and 270 by drawing a line across the intima‐media layer. The lumen diameter (d) was calculated as d = perimeter/π (Fernandes‐Santos et al. 2009).

Statistical analysis

Values are expressed as the mean ± SEM or mean ± SD. Relaxation is expressed as a percentage of inhibition of noradrenaline‐induced contraction. EC50 values (concentration of agonist producing half‐maximum effect) were determined from individual concentration–response curves by non‐linear regression analysis and are expressed as pD 2 (–log EC50). Values were compared by an unpaired t test and one‐ or two‐way ANOVA with Scheffe's test as a post hoc test. The number of animals is indicated by n. P < 0.05 was considered statistically significant. Statistical analysis was performed using SPSS, version 24.0 (IBM Corp., Armonk, NY, USA).

Results

Efficacy of training

Soleus muscle citrate synthase activity was 5.30 ± 0.16 μmol and 7.33 ± 0.17 μmol (mean ± SD) product formed min–1 g–1 protein in the control and trained groups, respectively (P = 0.001, n = 4 in each group). This 38% increase in citrate synthase activity indicates the effectiveness of the training protocol. There were no differences in the weight of soleus muscle between groups (1.12 ± 0.13 g vs. 1.28 ± 0.24 g (mean ± SD) for the control and trained groups, respectively, P = 0.42).

Relaxation responses to ACh in carotid artery

Figure 1 shows the concentration–response curves to ACh (10–9 to 3 × 10–6 m) in carotid artery rings precontracted with noradrenaline from control and trained rabbits. Physical training reduced the maximal relaxation to ACh. Incubation with NAC (10–3 m) did not modify ACh‐induced relaxation in the control group, although it did reverse the impaired relaxation to ACh in the trained group, suggesting that the training protocol induced vascular oxidative stress. Figure 2 shows the effects of the different pharmacological inhibitors on the response to ACh in the control and trained groups. Indomethacin (10–5 m) did not change the relaxation to ACh. However, the addition of l‐NAME (10–4 m) significantly reduced maximal relaxation to ACh in both groups. The incubation of indomethacin plus l‐NAME had no greater effect than l‐NAME alone, indicating that there was no interaction between indomethacin and l‐NAME (Table 1). The effect of l‐NAME was lower in the trained group (in mean ± SEM, 39 ± 2% vs. 28 ± 3.0% for the control and trained groups, respectively, P = 0.035), suggesting a lower bioavailability of NO in trained rabbits. Moreover, the endothelium‐dependent relaxation after exposure to indomethacin and l‐NAME was further reduced by KCl (2 × 10−2 m) in both groups, indicating that the relaxant response to ACh also involved the activation of K+ channels. In both groups, the combination of charybdotoxin (10–7 m) and apamin (10–6 m) had no additional effects on the ACh‐induced relaxation in the presence of indomethacin (10–5 m) plus l‐NAME (10–4 m). BaCl2 (3 × 10–6 m) plus ouabain (10–4 m) abolished the remaining relaxation to ACh. Neither glibenclamide (10–6 m), nor barium (3 × 10–6 m) modified the response to ACh (data not shown). The pD 2 values and maximal relaxations in arteries from control and trained groups are presented in Table 1.

Figure 1. Curves to ACh of carotid rings.

Figure 1

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Relaxation–response curves to ACh of carotid rings from control and trained rabbits and in the presence of NAC (10–3 m) in the trained group. Data are the mean ± SEM.

Figure 2. Acetylcholine curves for carotid rings in the presence of indomethacin, indomethacin plus L‐NAME and indomethacin plus L‐NAME combined with charybdotoxin plus apamin, or KCl or BaCl2 plus ouabain.

Figure 2

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Relaxation–response curves to ACh of carotid rings from control and trained rabbits in the absence of inhibitors and in the presence of indomethacin (10–5 m), indomethacin plus l‐NAME (10–4 m) and indomethacin plus l‐NAME combined with charybdotoxin (10–7 m) plus apamin (10–6 m), or KCl (2 × 10–2 m) or BaCl2 (3 × 10–6 m) plus ouabain (10–4 m). Data are the mean ± SEM.

Table 1.

pD 2 values and E max to ACh in carotid artery from control and trained rabbits in the different experimental conditions

Control group Trained group
ACh n pD 2 E max n pD 2 E max
Control 16 7.26 ± 0.04 90 ± 2 16 7.19 ± 0.05 81 ± 2**
NAC 10−3 m 5 7.36 ± 0.12 87 ± 2 5 7.84 ± 0.12* 91 ± 5*
Indomethacin 10−5  m 4 7.45 ± 0.11 92 ± 3 5 7.36 ± 0.06 82 ± 5
l‐NAME 10−4  m 4 6.87± 0.03* 58 ± 1* 5 7.03 ± 0.10 59 ± 5*
Indomethacin 10−5  m + l‐NAME 10−4  m 5 6.89 ± 0.09* 55 ± 8* 7 7.36 ± 0.02 58 ± 6*
Indomethacin 10−5  m + l‐NAME 10−4  m + KCl 20 mm 4 6.52 ± 0.06*# 30 ± 2*# 3 6.75 ± 0.12*# 30 ± 7*#
Indomethacin 10−5 m + l‐NAME 10−4 m + charybdotoxin 10−7 m + apamin 10−6  m 6 7.04 ± 0.11* 46 ± 4* 8 7.29 ± 0.08 47 ± 4*
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Data are the mean ± SEM. n, number of rabbits; pD 2, –log m of substance causing 50% of the maximal contraction; E max, maximal response; NAC, N‐acetylcysteine. * P < 0.05 vs. control within the same group. ** P < 0.05 vs. control group. # P < 0.05 vs. indomethacin 10−5 m + l‐NAME 10−4 m within the same group.

Relaxation responses to ACh in femoral artery

ACh (10–9 to 3 × 10–6 m) caused a concentration‐dependent relaxation in femoral artery without differences between groups (Fig. 3). Figure 4 shows how the different pharmacological inhibitors modify the vasodilator response in the control and trained groups. Indomethacin (10–5 m) did not change the relaxation to ACh. The effect of l‐NAME was lower in the trained group (in mean ± SEM, 41 ± 5% vs. 17 ± 2% for the control and trained groups, respectively, P = 0.025), suggesting a lower participation of NO in the trained rabbits. The combination of indomethacin plus l‐NAME had the same effect as the administration of l‐NAME alone (Table 2). Moreover, the endothelium‐dependent relaxation after exposure to indomethacin and l‐NAME was further reduced by KCl (2 × 10–2 m) in both groups, indicating that relaxant response to ACh also involved the activation of K+ channels. However, charybdotoxin plus apamin further reduced the maximal relaxation to ACh only in the trained group, indicating that physical training enhanced the participation of Ca2+‐activated K+ channels in the endothelium‐dependent vasodilatation. The blockade obtained with charybdotoxin plus apamin was similar to the blockade obtained with charybdotoxin alone or iberiotoxin. However, the incubation with apamin or TRAM‐34 had no further effect on ACh than the exposure to indomethacin and l‐NAME. Consequently, the channels implicated in this response were BKCa (Fig. 5 A).

Figure 3. Curves to ACh of femoral rings.

Figure 3

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Relaxation–response curves to ACh of femoral rings from control and trained rabbits. Data are the mean ± SEM.

Figure 4. Acetylcholine curves for femoral rings in the presence of indomethacin, indomethacin plus L‐NAME and indomethacin plus L‐NAME combined with charybdotoxin plus apamin, or KCl or BaCl2 plus ouabain.

Figure 4

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Relaxation–response curves to ACh of femoral rings from control and trained rabbits in the absence of inhibitors and in the presence of indomethacin (10–5 m), indomethacin plus l‐NAME (10–4 m) and indomethacin plus l‐NAME combined with charybdotoxin (10–7 m) plus apamin (10–6 m), or KCl (2 × 10–2 m) or BaCl2 (3 × 10–6 m) plus ouabain (10–4 m). Data are the mean ± SEM.

Table 2.

pD 2 values and E max to ACh in femoral artery from control and trained rabbits in the different experimental conditions

Control group Trained group
Ach n pD 2 E max n pD 2 E max
Control 10 7.37 ± 0.09 80 ± 3 10 7.16 ± 0.07 83 ± 3
Indomethacin 10−5 m 4 7.31 ± 0.13 81 ± 8 6 7.03 ± 0.09 84 ± 6
l‐NAME 10−4  m 4 7.18 ± 0.33 48 ± 1* 3 6.61 ± 0.21* 67 ± 1*
Indomethacin 10−5  m + l‐NAME 10−4  m 6 6.58 ± 0.12* 47 ± 6* 8 6.80 ± 0.20* 69 ± 8*
Indomethacin 10−5  m + l‐NAME 10−4  m + KCl 20 mm 4 7.15 ± 0.22# 21 ± 8*# 3 6.55 ± 0.21 19 ± 4*#
Indomethacin 10−5  m + l‐NAME 10−4  m + charybdotoxin 10−7  m + apamin 10−6  m 6 7.28 ± 0.13# 30 ± 6* 6 7.35 ± 0.30 38 ± 6*#
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Data are the mean ± SEM. n, number of rabbits; pD 2, –log m of substance causing 50% of the maximal contraction; E max, maximal response; NAC, N‐acetylcysteine. * P < 0.05 vs. control within the same group. # P < 0.05 vs. indomethacin 10−5 m + l‐NAME 10−4 m within the same group.

Figure 5. Blockade of remaining relaxation to Ach and protein expression levels of BKCa channels in femoral artery.

Figure 5

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A, control represents the remaining relaxation to ACh after blockade with indomethacin (10−5 m) plus l‐NAME (10−4 m) in femoral artery from the trained group. Effects of apamin (10−6 m) (APA), TRAM‐34 (10−6 m) (TRAM), charybdotoxin (10−7 m) (CHTX), iberiotoxin (10−7 m) (IBTX), and the combination of charybdotoxin (10−7 m) + apamin (10−6 m) (CHTX + APA) on the remaining relaxation to ACh after blockade with indomethacin (10−5 m) + l‐NAME (10−4 m). Data are the mean ± SEM. B, BKCa channels protein expression by WB analysis in femoral artery from control (C) and trained (T) group. A representative immunoblot is shown and α‐tubulin was used as the control amount of protein. Data are the mean ± SD of four independent experiments. * P < 0.05 vs. control.

The remaining relaxation was abolished by BaCl2 (3 × 10–6 m) plus ouabain (10–4 m) in both groups. Neither glibenclamide (10–6 m), nor BaCl2 alone (3 × 10–6 m) modified the response to ACh (data not shown). Table 2 shows the pD 2 values and maximal relaxations in arteries from control and trained groups.

Relaxation responses to sodium nitroprusside

Physical training did not modify the response to the endothelium independent vasodilator sodium nitroprusside in any of the vessels studied (Fig. 6).

Figure 6. Curves to sodium nitroprusside of carotid and femoral rings.

Figure 6

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Relaxation–response curves to sodium nitroprusside of carotid and femoral rings from control and trained rabbits. Data are the mean ± SEM.

Effects of physical training on eNOS, Cu/Zn‐SOD, Mn‐SOD and BKCa channel protein expression

Physical training significantly decreased eNOS expression in both vessels studied (P = 0.019 and P = 0.040 for carotid and femoral arteries, respectively, n = 4 in each group), whereas Cu/Zn and Mn‐SOD expression were only decreased in the carotid artery (P = 0.002 and P = 0.008 for Cu/Zn and Mn SOD, respectively, n = 4 in each group) (Fig. 7). BKCa expression was significantly increased in the trained group in the femoral artery (P < 0.001, n = 4 in each group) (Fig. 5 B).

Figure 7. Protein expression levels of eNOS, Mn‐SOD and Cu/Zn‐SOD.

Figure 7

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Protein expression levels of eNOS, Mn‐SOD and Cu/Zn‐SOD in carotid and femoral rings from control (C) and trained (T) rabbits. A representative immunoblot of each protein is shown and α‐tubulin was used as the control amount of protein. Data are the mean ± SD of four independent experiments. * P < 0.05 vs. control.

Plasma determination of malondialdehyde

MDA levels were similar in both groups (1.71 ± 0.38 μm vs. 1.79 ± 0.49 μm (mean ± SD) for the control and trained groups, respectively, P = 0.87, n = 8 in each group), indicating that exercise training did not induce systemic oxidative stress.

Morphometric analysis

There were no differences in the lumen diameter of carotid artery between groups (1.36 ± 0.19 mm vs. 1.46 ± 0.12 mm (mean ± SEM) for the control and trained groups, respectively, P = 0.21, n = 8 in each group). We did not find differences in the femoral lumen diameter with training either (1.14 ± 0.18 mm to 1.26 ± 0.19 mm (mean ± SEM) for the control and trained groups, respectively P = 0.18, n = 8 in each group). The intima‐media thickness was 130 ± 6 μm vs. 127 ± 5 μm (mean ± SEM) for the control and trained groups (P = 0.54, n = 8 in each group) in carotid the artery and 102 ± 4 μm and 108 ± 7 μm (mean ± SEM) for the control and trained groups (P = 0.25, n = 8 in each group) in the femoral artery, respectively, indicating that training did not modify the thickness of vascular wall of either the carotid or femoral artery (Fig. 8).

Figure 8. Representative morphological analysis.

Figure 8

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Representative morphological analysis showing haematoxylin and eosin staining in carotid and femoral artery sections from the control and trained groups. L, lumen; TI, tunica intima; TM, tunica media; TA, tunica adventitia. Scale bars = 400 or 100 μm.

Discussion

The main finding of the present study is that 6 weeks of physical training induces non‐uniform vascular responses in carotid and femoral rings. Arteries from control and trained rabbits have different pathways for endothelium‐dependent relaxation. This involves both nitric oxide‐dependent and independent pathways, where NO independent relaxation is mediated by the activation of K+ channels.

The efficacy of a training protocol is frequently assessed by the increase in the activity of the enzyme citrate synthase (Heylen et al. 2008). This increment can be observed from the first or second week of training and usually precedes vascular tone adaptations (Delp and Laughlin, 1997). In the present study, we observed a 38% increase in citrate synthase activity in the soleus muscle of trained rabbits compared to controls, indicating the effectiveness of the training protocol (McAllister et al. 2005; Jendzjowsky and DeLorey, 2012).

Vasodilator response

Previous studies in both animals and humans evidence that endothelial‐dependent relaxation is either unaltered (McAllister et al. 1996, 2005), enhanced (Delp and Laughlin, 1997; McAllister et al. 2005; Spence et al. 2013) or decreased (Bergholm et al. 1999) by physical training as a result of changes in NO bioavailability. Our findings obtained in the carotid artery from trained rabbits show a decreased relaxant response to ACh but not to sodium nitroprusside, consistent with a reduction in the synthesis, release or bioavailability of endothelial NO. Regarding bioavailability, two main mechanisms affect NO inactivation in the vascular wall. The first one is that NO dioxygenases, such as myoglobin, haemoglobin‐α and cytoglobin, oxidize NO to nitrate (Liu et al. 2017). The other mechanism is the interaction of NO with O2 − to form RONS (Wink and Mitchell, 1998; Förstermann, 2010), which most probably occurs in the carotid artery from the trained group because NAC reversed the endothelial dysfunction. However, in the femoral artery, training did not modify the relaxation induced by ACh and sodium nitroprusside. This response cannot be generalized to other vessels and species. For example, physical training enhanced relaxation to ACh in the rat aorta but not in the carotid artery (Chen Hi et al. 1996). Nevertheless, an increase (Kemi et al. 2004, 2005) in ACh‐dependent relaxation has been reported in the rat carotid artery in response to exercise. Discrepancies in these results could be related to exercise modality and the duration of training. In this sense, studies in humans indicate that regional differences in vascular remodelling are related to structural properties, shear stress pattern during exercise and training modality (Spence et al. 2013; Black et al. 2016). By contrast to the femoral artery, the carotid artery is not directly exposed to acute bouts of blood flow during exercise; hence, training‐induced adaptations would become evident over the long term.

Participation of prostanoids and NO in the response to ACh

Relaxation of vascular smooth muscle by ACh may be caused by the release of prostaglandins from endothelial cells (Villar et al. 2006). Indomethacin, an inhibitor of cyclooxygenase, did not affect the response to ACh in either control or trained rabbits, suggesting that prostaglandins did not play a role in endothelium‐dependent vasodilatation. Moreover, our experiments show that the relaxation to ACh was partially reduced by the NOS inhibitor l‐NAME in both carotid and femoral arteries, indicating that other factors than NO or prostanoids contribute to ACh‐induced relaxation. In both carotid and femoral arteries, the effect of l‐NAME on ACh‐induced relaxation was lower in trained rabbits compared to controls. In addition, our training modality decreased eNOS protein expression in both vascular beds. Thus, together, the lower effect of l‐NAME and the decrease in eNOS expression provide a unifying explanation for the impairment of ACh‐stimulated endothelium‐dependent NO vasodilatation observed in carotid and femoral arteries from trained rabbits.

Vascular oxidative stress

Oxidative stress is characterized by an increase of RONS. The main sources of them in the vascular wall are NADPH oxidase, xanthine oxidase, uncoupled eNOS and mitochondrial respiratory chain enzymes (Förstermann, 2008, 2010; Li et al. 2014). NADPH oxidase is the predominant under physiological conditions. The vascular wall also contains anti‐oxidant systems such as SOD, catalase, glutathione peroxidases, thioredoxin system, peroxideroxin and haeme oxygenase‐1 (Förstermann, 2008; Hsieh et al. 2014).

Impaired relaxation to ACh in the carotid artery was reversed by the RONS scavenger NAC, a synthetic N‐acetyl derivative of the amino acid l‐cysteine. Cysteine is crucial for the biosynthesis of glutathione, the most important intracellular defence against RONS. The thiol groups of glutathione reduce RONS by accepting their unpaired electrons. It has been demonstrated that NAC infusion increases cysteine blood levels and inhibits glutathione oxidation (Medved et al. 2004; Matuszczak et al. 2005). Consequently, by increasing the glutathione levels, NAC could reduce O2 − bioavailability, preventing the formation of other RONS, such as hydroxyl radical or peroxynitrite. O2 − reacts with NO to generate peroxynitrite, which interacts with DNA, proteins and lipids, modifying cell signalling. Peroxynitrite depletes thiols groups and reduces NO and superoxide bioavailability (Förstermann et al. 2010; Powers and Jackson, 2008). Therefore, oxidative stress reduces the amount of bioactive NO and increases the levels of peroxynitrite, which in turn suppresses eNOS expression and forms nitrotyrosine in the artery walls, with both processes being related to endothelial dysfunction (El‐Remessy et al. 2010). It has been demonstrated that chronic physical training induces an increase in anti‐oxidant enzymes in response to oxidative stress (Gomez‐Cabrera et al. 2008; Vina et al. 2012), shifting the balance between oxidant and anti‐oxidant systems in favour of the anti‐oxidant pathway. However, high intensity training impairs endothelium‐dependent relaxation that is associated with a proinflammatory state and oxidative stress (Bergholm et al. 1999; Sun et al. 2008). Accordingly, intense aerobic physical training in men decreased circulating levels of anti‐oxidants (Bergholm et al. 1999). Furthermore, a study comparing a protocol of moderate physical training in both previously trained and untrained rats found that the untrained group had lower levels of anti‐oxidant parameters in response to oxidative stress (Elikov, 2016). In this regard, we also found that Mn‐SOD and Cu/Zn‐SOD expression were significantly reduced in the carotid artery in the trained group. Superoxide dismutase is crucial for protecting against oxidative stress and hence reducing the formation of peroxynitrite. Moreover, increased levels of NO upregulate SOD in vascular smooth muscle cells, preventing superoxide production in a concentration‐dependent manner (Fukai et al. 2000). Consequently, a lower production or bioavailability of NO could reduce SOD expression. Therefore, a possible explanation for our findings is that the increased RONS production induced by training could decrease NO bioavailability and eNOS expression. Furthermore, the decreased expression of the anti‐oxidant enzymes could be explained by the decreased eNOS expression and NO production. Thus, the decrease in NO participation in response to ACh in the carotid artery would be related to the increase of oxidative stress induced by training. It is noteworthy that the oxidative stress only occurs in the carotid artery, and not in the femoral artery or at a systemic level, because plasma levels of MDA are similar in both groups. Hence, the remaining question is why the femoral artery is affected to a lesser extent by RONS damage compared to the carotid artery. Mechanical forces applied on endothelium, such as shear stress, an increase in circumferential stretch or high intraluminal pressure, can induce changes in biochemical pathways in a process known as mechanotransduction (Davies, 2009). Depending on blood flow pattern, shear stress induces both pro and anti‐oxidants enzymes, thus regulating RONS production (Laurindo et al. 1994; Abdulnour et al. 2006; Hsieh et al. 2014). On the basis of our results, we speculate that the training could induce oxidative stress in an unequal way, creating a favourable redox environment in arteries undergoing elevated shear stress, or bouts of high intraluminal pressure, such as the femoral artery, whereas, in other vascular beds, such as carotid artery, exercise can induce a damage pathway mediated by oxidative stress. In addition, as shown by the results of the present study, the femoral artery has a smaller radius than carotid artery. Because shear stress is inversely proportional to the internal radius of the vessel (Laughlin et al. 2008), we can assume that shear stress is greater in femoral artery. The results of some studies suggest that exposure to an increased shear stress leading to an increment of blow flow during exercise is the main signal with respect to triggering the endothelial adaptations (Miller and Burnett, 1992; Harrison et al. 1996; Nadaud et al. 1996). Indeed, our results indicate that the femoral artery demonstrates endothelial adaptations such as an increase in the EDHF pathway via BKCa that compensates for low NO bioavailability and maintains vasodilatation.

Role of Ca2+‐activated K+ channels on vascular function

Laughlin et al. (2008) concluded that increased shear stress during exercise contributes to an anti‐atherogenic phenotype of arterial endothelial cells. Hence, we can hypothesize that the range of shear stress that exercise produces in femoral artery leads to a ‘beneficial phenotype’, commonly characterized by an increase in the NO pathway (Spence et al. 2013; Black et al. 2016). However, we found that training decreases NO bioavailability in both arteries. The NO pathway is not the only one susceptible to changes induced by exercise. Both stretch (Fleming, 2004) and cyclic strain can increase endothelium‐dependent hyperpolarization (EDH) via Ca2+‐activated K+ channels, especially when NO is depressed (Quilley et al. 1997). The EDHF is a diffusible factor released by endothelium that hyperpolarizes vascular smooth muscle cells. An increase in endothelial calcium concentration stimulates Ca2+‐activated K+ channels and this process activates the EDH. It is known that, in small resistance arteries, both IKCa and SKCa participate in this pathway (Edwards et al. 1998), whereas, in other vascular beds, the EDH can act through the BKCa (Martinez‐León et al. 2003). Ca2+‐activated K+ channels are located in endothelial microdomains and let K+ efflux through them. This K+ efflux can be the diffusible EDHF, thus stimulating the Na+,K+‐ATPase and KIR channels in the vascular smooth muscle. At the same time, myoendothelial gap junctions allow spread of the hyperpolarizing current, which is responsible for the persistent relaxation of EDH (Mather et al. 2005).

Our results show that the remaining endothelial‐dependent relaxation after the inhibition of the NO/PGI2 pathway in both carotid and femoral arteries was further reduced by increasing the extracellular K+ concentration, by blocking Ca2+‐activated K+ channels or by blocking Na+, K+ ‐ATPase plus KIR channels. Incubation with charybdotoxin plus apamin or iberiotoxin reduced the maximal relaxation to ACh only in the femoral arteries from the trained group. In addition, the protein expression of BKCa was increased in the femoral artery from trained group, indicating that physical training enhances the participation of BKCa channels in the endothelium‐dependent vasodilatation in this vessel. There is little direct evidence for the involvement of EDHF in exercise (Clifford and Hellsten, 2004). Studies in rat thoracic aorta (Chen et al. 2001; Heylen et al. 2008), rat mesenteric artery (Chen et al. 2001) and pig coronary arteries (Xie et al. 2013) found an increased contribution of K+ channels in endothelium‐dependent vasodilatation with training. Our findings add evidence indicating that the involvement of BKCa channels in the relaxation induced by ACh in the femoral artery may be an adaptive mechanism to compensate for the decrease of the NO‐dependent relaxation induced by training, allowing the maintenance of vasodilatation. This adaptive response is absent in the carotid artery, reflecting the failure of counteractive mechanisms to prevent the impairment of endothelial vasodilatation evoked by training in this vessel. Based on the studies discussed previously, we can assume that, during exercise, both stretch and cyclic strain are higher in the femoral than in the carotid artery (Laughlin et al. 2008; Green et al. 2017) and could plausibly explain why we found a compensatory mechanism to maintain vasodilatation in response to ACh in the femoral artery.

Vascular remodelling

Exercise training can induce vascular remodelling affecting both diameter and arterial wall thickness. These structural adaptive mechanisms depend on the intensity, duration and type of exercise, and vascular bed. In this regard, many studies conclude that the arterial diameter increases in trained subjects (Green et al. 2017) with no changes (Rowley et al. 2011; Thijssen et al. 2012) or a decrease (Rowley et al. 2012; Spence et al. 2013) in wall thickness. One possible explanation is that vascular remodelling induced by training allows the vascular wall to support a higher tension in the presence of lower thickness (Green et al. 2017). However, in our model, functional adaptations occur without structural adaptations in the vessels.

Conclusions

The evidence is conclusive concerning the benefits of physical activity with respect to decreasing all‐cause and cardiovascular mortality (Thijssen et al. 2010; Piepoli et al. 2016). In this sense, it has been demonstrated that our training model increases myocardial electrical stability in an isolated heart preparation, suggesting that it could protect against re‐entrant ventricular arrhythmias (Such et al. 2008) and thus reduce the risk of sudden death. However, these beneficial effects do not necessarily occur in blood vessels. Indeed, our results reveal that 6 weeks of physical training induces endothelial dysfunction in the carotid artery as a result of oxidative stress and modifies the vasodilator pathways in response to ACh in the femoral artery. In this vessel, a lower production of NO is counteracted by an increase in the participation of BKCa channels, compensating for the impairment of NO‐mediated vasodilatation. Taken together, these results suggest that the changes in vascular reactivity induced by our training protocol are dependent on the vascular bed and the presence of compensatory mechanisms to prevent impaired vasodilatation.

Additional information

Competing interests

The authors declare that they have no competing interests.

Author contributions

All experiments were performed at the Vascular Laboratory at the Department of Physiology of University of Valencia. MDM, PM, JMV designed the study. CS trained the rabbits. PM, MDM, CS, SG‐O and SV performed the experiments and acquired the data. MDM, PM, JMV, SG‐O, MA and SV analysed the results. MDM, PM, JMV, MA and SV interpreted the data for the work. PM, JMV and MDM wrote the manuscript. All authors revised it critically for important intellectual content. All authors qualify for authorship, and all those who qualify for authorship are include as authors. All authors approved the final version of the manuscript submitted for publication and agree to be accountable for all aspects of the work.

Funding

No funding was received for the present study.

Acknowledgements

We wish to thank Professor Luis Such and GRELCA Research Group, Department of Physiology, School of Medicine, University of Valencia, Spain, who provided the animal model. We also wish to thank Professor Jose Viña, head of the Freshage Research Group, Department of Physiology, School of Medicine, University of Valencia, Spain, and Sonia Priego (Microscopía confocal, UCIM, University of Valencia, Spain) for their technical assistance.

Biographies

Patricia Marchio obtained her Master's Degree and PhD in Physiology under the direction of MD Mauricio and JM Vila from Department of Physiology at the University of Valencia, Spain.

Solanye Guerra‐Ojeda obtained her Master's Degree in Biomedical Engineering at the Polytechnic University of Valencia. Currently, she is a PhD candidate in Physiology at the University of Valencia. The laboratory's research is focused on vascular biology.

Edited by: Harold Schultz & Giovanni Mann

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