**1. Introduction**

Hypertension, a chronic condition of elevation in blood pressure (BP), is a well-known risk factor for the development of cardiovascular diseases (CVDs), including heart failure, coronary heart diseases, myocardial fibrosis, and infarction [1], atherosclerosis [1,2], stroke [1,3], and kidney failure [4]. The pathogenesis of hypertension is multifaceted,

**Citation:** Chao, Y.-M.; Rauchová, H.; Chan, J.Y.H. Disparate Roles of Oxidative Stress in Rostral Ventrolateral Medulla in Age-Dependent Susceptibility to Hypertension Induced by Systemic L-NAME Treatment in Rats. *Biomedicines* **2022**, *10*, 2232. https:// doi.org/10.3390/biomedicines 10092232

Academic Editor: Marie-Louise Bang

Received: 20 July 2022 Accepted: 6 September 2022 Published: 8 September 2022

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and one of the well-characterized causes is endothelial dysfunction resulting from nitric oxide (NO) deficiency [5–7]. Circulatory NO, derived primarily from endothelial NO synthase (eNOS) in vascular endothelial cells [8], participates in BP regulation via maintenance of vascular tone through the classical NO/soluble guanyl cyclase/cyclic guanosine monophosphate signaling to decrease intracellular calcium in vascular smooth muscle cells and evoke vasodilatation [9,10]. In addition, NO regulates cardiovascular functions by exhibiting anti-inflammatory [11], anti-platelet aggregatory [12], and anti-proliferative [13] actions. Accordingly, deficiency in NO production and/or availability leads to impaired vascular relaxation, increased vascular resistance, augmented platelet aggregation, and vascular inflammation or proliferation; all of which have been demonstrated to underpin the pathophysiology of hypertension [6,14,15].

A rat model of hypertension induced by the systemic administration of Nω-Nitro-Larginine methyl ester hydrochloride (L-NAME), a NOS inhibitor, is widely used to mimic hypertension in human [16]. This treatment downregulates NOS expression in blood vessels [17], and reduces plasma NO levels [18], resulting in systemic vasoconstriction, increased vascular resistance, and hypertension [17–19]. L-NAME further aggravates the development of hypertension via antagonization of the anti-inflammatory, anti-platelet aggregatory, and/or anti-proliferative actions of NO [20,21]. In addition to its peripheral actions, L-NAME reportedly crosses the blood–brain barrier (BBB) to decrease NOS expression and inhibit NOS activity in brain [22–24], leading to brain-initiated hypertension. The neural mechanism(s) that underlies the pathogenesis of this form of hypertension induced by systemic L-NAME treatment is not fully understood.

The rostral ventrolateral medulla (RVLM), where sympathetic premotor neurons reside [25], is one of the major sites of action in the brain stem where NO participates in the neural control of cardiovascular functions [6,26–28]. Emerging evidence from preclinical studies in animal models suggests that NO in RVLM exhibits a predominantly sympathoinhibitory action in the regulation of sympathetic nerve activity (SNA) under physiological conditions [28,29]. Deficiency in tissue NO availability under various cardiovascular conditions, including hypertension [29], chronic heart failure [30], and metabolic syndrome [31,32], may lead to an increase in SNA because of the blunted sympathoinhibitory action of NO in RVLM. Moreover, NO participates in the neural control of cardiovascular systems via interactions with reactive oxygen species (ROS), in particular superoxide anion (O2 •−) [6,26,32]. Under physiological conditions, NO is generated from its substrate L-arginine through coupling of NOS [33]. This coupling capacity is reduced under hypertensive conditions, wherein NOS removes an electron from its cofactor, NADPH, and donates it to an oxygen molecule to generate O2 •− rather than NO [32–34]. The causal role of ROS at RVLM in systemic L-NAME treatment-induced hypertension, nonetheless, has not been fully elucidated.

Epidemiological studies indicate aging is a predominant risk factor for CVDs. The prevalence of hypertension increases markedly with aging, attributed primarily to alterations in the structure, responsiveness, function, and rigidity of vessel walls [35], as well as dysregulation of the autonomic nervous system [36]. Several theories have so far been proposed to explain the etiology of biological aging [37]; among them, tissue oxidative stress was postulated to be a common denominator [38]. Indeed, a wide array of studies suggests the engagement of ROS in age-related CVDs, including hypertension, atherosclerosis, atrial fibrillation, and stroke [38]. Despite evidence suggesting causal links between oxidative stress, aging, and CVDs, there is a paucity in the literature on the molecular mechanisms that underlie age-related dysregulation of redox homeostasis and its contribution to the development of CVDs.

Redox signaling is critically involved in cardiovascular pathophysiology, and oxidative stress in RVLM has been demonstrated unambiguously to play a pivotal role in neural mechanisms of hypertension via the increase in neurogenic sympathetic outflow [6,14,26–28]. However, there is no current information on whether redox homeostasis in RVLM is affected by systemic L-NAME treatment. Furthermore, exactly how aging changes the redox

state in RVLM, leading to its engagement in hypertension induced by systemic L-NAME treatment is unknown. This study therefore aims to investigate whether tissue oxidative stress in RVLM plays an active role in age-dependent susceptibility to hypertension in response to systemic L-NAME treatment, and to decipher the underlying molecular mechanisms. Our findings suggest that redox homeostasis in RVLM is tolerated in young (8 weeks of age) normotensive rats subjected to systemic L-NAME treatment; and NO deficiency, both in the vascular smooth muscle cells and RVLM, may likely be the key underlying molecular mechanisms. When animals turn adult (20 weeks of age), tissue ROS level in RVLM is augmented, leading to an enhanced increase in central sympathetic outflow that exacerbates hypertension induced by L-NAME.

#### **2. Materials and Methods**

#### *2.1. Animals*

Experiments were carried out in young (4 weeks old, *n* = 51) and adult (12 weeks old, *n* = 63) male normotensive Wistar-Kyoto (WKY) rats purchased from BioLASCO, Taipei, Taiwan. They were housed in animal rooms under temperature control (24 ± 0.5 ◦C) and 12 h light/dark (5 am to 5 pm) cycle. Standard laboratory rat chow (PMI Nutrition International, Brentwood, MO, USA) and tap water were available ad libitum. All animals were allowed to acclimatize for 14 days (young group) or housed to the age of 20 weeks (adult group) in an AAALAC-International accredited animal holding facility in Kaohsiung Chang Gung Memorial Hospital, Taiwan, before experimental manipulations. All experiments were performed in accordance with the guidelines for animal experimentation approved by our institutional animal care and use committee (no. 2018051701) as adopted and promulgated by the U.S. National Institutes of Health.

#### *2.2. Implantation of Osmotic Minipump*

Implantation of osmotic minipump into the peritoneal cavity or the cisterna magna was carried out according to previously reported procedures [32]. Briefly, for intraperitoneal (i.p.) implantation, animals were anesthetized with pentobarbital sodium (50 mg/kg, i.p.) and a midline incision of the abdominal cavity was made, followed by implantation of an Alzet® osmotic minipump (model 2002, DURECT Co., Cupertino, CA, USA) into the peritoneal cavity. The incision was closed with layered sutures. Some animals also received implantation of an additional micro-osmotic minipump into the cistern magna. For intracisternal (i.c.) implantation, we performed a midline incision of the dorsal neck, and dissected the underneath muscle layers to expose the dura mater between the foramen magnum and C1 lamina. The dura mater was perforated with a 22-gauge steel needle, followed by insertion of a polyvinylchloride tubing into the cistern magna using an Alzet® brain infusion kit. Observation of presence of cerebrospinal fluid at the outer end of the catheter assured the patency of the implantation. Tissue glue was used to seal the catheter to the dura mater and the incision was closed with layered sutures. A microosmotic minipump (Alzet® 1007D) was positioned subcutaneously in the neck, and was connected to the outer end of the catheter. No obvious neurological signs were observed after minipump implantation. All animals received postoperative intramuscular injection of procaine penicillin (1000 IU). Only animals that showed recovery and progressive weight gain after the operation were used in subsequent experiments.

#### *2.3. Blood Pressure and Heart Rate Measurement*

Systolic blood pressure (SBP) and heart rate (HR) were determined in conscious animals with a tail-cuff sphygmomanometer (MK-2000; Momuroki Kikai Co., Tokyo, Japan). Baseline values were measured on days 3 and 1 prior to, on the day (day 0) before osmotic pump implantation, and then on days 1, 3, 5, 7, 9, 11, and 14 following i.p. L-NAME (10 mg/kg/day; Sigma-Aldrich, MA, USA) or 0.9% saline (vehicle control) treatment. During the recording sessions (at 14:00–16:00), rats were placed in restraint holders and the tail warmed on a warming pad for 10–15 min to increase blood flow and improve data

acquisition. A full recording session consisted of 5 acclimatization cycles to optimize data acquisition, followed by 5 data acquisition cycles during which SBP and HR were recorded. The mean SBP and HR calculated for each time point were used for statistical analyses. It should be noted that SBP obtained by tail-cuff plethysmography in conscious rats has been validated to be comparable to those measured by radiotelemetry [32].

#### *2.4. Evaluation of Sympathetic Vasomotor Activity*

Sympathetic vasomotor activity was measured at the end of i.p. L-NAME infusion in young and adult animals from blood pressure recorded from a cannulated femoral artery in animals anesthetized via an anesthesia mask with isoflurane (5% for induction and 2% for maintenance). Each recording session lasted 30 min, and took place between 14:00 and 16:00. Temporal fluctuations in the low-frequency (LF, 0.25–0.8 Hz) component of the SBP signals were detected by continuous online, real-time spectral analysis based on fast Fourier transform using an arterial blood pressure analyzer (APR31a, Notocord, Le Pecq, France). The power density of the LF band was used as our experimental index to reflect sympathetic vasomotor tone [32].

#### *2.5. Measurement of Plasma Norepinephrine*

We measured plasma norepinephrine (NE) level by the o-phthalaldehyde (OPA) method using high-performance liquid chromatography (HPLC) with fluorescence detection. Plasma sample was mixed with ice-cold trichloroacetic acid, and centrifuged at room temperature. The supernatant was mixed with 4-fold methanol after being filtered through a syringe filter (0.22 μm; Chroma Technology Corp., Bellows Falls, VT, USA), centrifuged again at room temperature, and kept at −80 ◦C until analyses. NE was measured by HPLC as previously described [39], and comparing the area under the curve of each sample against standard NE solutions of known concentrations was used to compute the concentration (μg/μL). Each sample was analyzed in triplicates and the mean was used for statistical analyses.

## *2.6. Measurement of Serum Nitric Oxide*

Serum NO concentration was assessed indirectly by measuring the levels of nitrate and nitrite, the stable end-product of NO, with a NO colorimetric assay kit (Arbor assay kit, Ann Arbor, MI, USA) according to manufacturer's instructions. The absorbance of the solution was read on a microplate reader at 540 nm (ThermoFisher Scientific Inc., Waltham, MA, USA). Each sample was analyzed in triplicates and the mean was used for statistical analyses.

#### *2.7. Measurement of Plasma Malondialdehyde*

Assays for lipid peroxidation are commonly used for estimation of oxidative status. Levels of lipid peroxidation in plasma were measured by a malondialdehyde (MDA; a primary indicator of lipid peroxidation) assay kit (Biovision, Milpitas, CA, USA), following the protocol provided by the manufacturer. Briefly, plasma samples were reacted with thiobarbituric acid (TBA) at 95 ◦C for 60 min. A microplate spectrophotometer (ThermoFisher Scientific) was used to determine the level of MDA-TBA adduct with colorimetric absorbance read at 532 nm. Each sample was analyzed in triplicates and the mean was used for statistical analyses.

#### *2.8. Measurement of Plasma Proinflammatory Cytokines*

The levels of proinflammatory cytokines, including interleukin-1 β (IL-1β), IL-6, and tumor necrosis factor-alpha (TNF-α), in plasma, were measured using anti-rat ELISA Kits (ThermoFisher Scientific) according to the manufacturer's specification. Plasma was collected and centrifuged for 10 min at 4 ◦C. The supernatants were used immediately to measure the concentrations of proinflammatory cytokines. Positive and negative controls were included on each plate. The final concentration of the cytokines was calculated by converting the optical density readings against a standard curve. Each sample was analyzed in triplicate and the mean was used for statistical analyses.

#### *2.9. Tissue Collection and Protein Extraction from Rostral Ventrolateral Medulla*

Animals were deeply anesthetized at the end of the experiments with an overdose of pentobarbital sodium (100 mg/kg, i.p.), followed by intracardial infusion with 500 mL of warm (37 ◦C) normal saline. The brain stem was rapidly removed and immediately frozen on ice. Using a rodent brain matrix (World Precision Instruments, Sarasota, FL, USA) and based on the atlas of Watson and Paxinos [40], the medulla oblongata covering RVLM was blocked between 0.5 and 1.5 mm rostral to the obex. Tissue from bilateral RVLM was collected by micropunches, and was stored at −80 ◦C until use.

For total protein extraction from RVLM, tissue micropunches were homogenized using a Dounce grinder with a tight pestle in ice-cold lysis buffer mixed with a cocktail of protease inhibitors (Sigma-Aldrich, St. Louis, MO, USA) to prevent protein degradation. Solubilized proteins were centrifuged at 20,000× *g* at 4 ◦C for 15 min, and the total protein in supernatant was quantified by the Bradford assay with a protein assay kit (Bio-Rad, Hercules, CA, USA).

#### *2.10. Measurement of Reactive Oxygen Species in RVLM*

RVLM tissues were homogenized in sodium phosphate buffer (20 mM), centrifuged and the supernatant was collected for ROS measurement by electron paramagnetic resonance (EPR) spin trapping technique, as described previously [41]. EPR spectra were captured using a Brucker EMXplus spectrometer (Bruker, Ettlingen, Germany). Typical parameters were set at microwave power: 20 mW, modulation frequency: 100 kHz; modulation amplitude: 2 G; time constant: 655.36 ms; conversion time: 656 ms; sweep time: 335.87 s. We added a membrane-permeable superoxide dismutase (SOD; 350 U/mL) into the incubation medium to determine ROS specificity. Spectra represented the average of 6 scans. Each sample was analyzed in triplicate and the mean was used for statistical analyses.

#### *2.11. Measurement of Nitric Oxide in RVLM*

For measurement of NO in RVLM tissue, tissue micropunches were homogenized in lysis buffer, centrifuged, and the supernatant was stored at −80 ◦C until use after being deproteinized using a Centricon-30 filtrator (Microcon YM-30, Bedford, MA, USA). Tissue NO levels were determined based on chemiluminescence reaction with the purge system of a NO analyzer (Sievers NOA 280TM, Boulder, CO, USA) [32]. Each sample was analyzed in triplicate and the mean was used for statistical analyses.

#### *2.12. Measurement of Nitric Oxide Synthase Activity in RVLM*

Tissue NOS activity in RVLM was measured according to previously reported procedures [32]. RVLM tissues were lysed and centrifuged to obtain the supernatants, which were then used for detection of the enzyme activity following the manufacturer's instructions of a NOS activity assay kit (Merck KGaA, Darmstadt, Germany). After colorimetric reaction, the optical density was read using a microplate spectrophotometer (ThermoFisher Scientific) at an absorbance wavelength of 540 nm. Each sample was analyzed in triplicate and the mean was used for statistical analyses.

#### *2.13. Measurement of NADPH Oxidase Activity in RVLM*

NADPH oxidase activity of protein samples from RVLM was measured using the lucigenin-derived chemiluminescence method [32]. Briefly, the luminescence assay was performed in phosphate buffer with NADPH as the substrate. After dark adaptation, a tissue homogenate (100 μg protein) was added, and the chemiluminescence value was recorded. O2 •− production was measured with the addition of NADPH, in the presence or absence of an NADPH oxidase, diphenyleneiodonium. All measurements were conducted in the dark room with temperature maintained at 22–24 ◦C. Light emission was recorded by a Sirius Luminometer (Berthold, Germany). Protein concentrations were determined using a Bio-Rad protein assay kit (Bio-Rad Laboratories). Each sample was analyzed in triplicate and the mean was used for statistical analyses.

## *2.14. Measurement of Total Antioxidant Activity in RVLM*

Tissue antioxidant activity in RVLM was measured by a total antioxidant capacity assay kit (Sigma-Aldrich), following the protocol provided by the manufacturer. RVLM tissues were homogenized in lysis buffer, centrifuged, and the supernatant was used for analysis. The reaction was based on Cu2+ reduction by the small molecule antioxidants and the reduced Cu+ ion chelates with a colorimetric probe that was read with a standard 96 well spectrophotometric microplate reader at 570 nm. Antioxidant capacity was determined by comparison with Trolox, a water-soluble vitamin E analog that serves as an antioxidant standard. Each sample was analyzed in triplicate and the mean was used for statistical analyses.

#### *2.15. Measurement of Mitochondrial Respiratory Enzyme Activity in RVLM*

The mitochondrial fraction from RVLM tissue was isolated following procedures reported previously [42]. Purity of the mitochondrial-rich fraction was verified by the expression of the mitochondrial cytochrome *c* oxidase. Activities of mitochondrial respiratory chain enzymes were measured immediately after mitochondrial isolation, according to procedures reported previously [42] using a thermostatically regulated spectrophotometer (ThermoFisher Scientific). Enzyme activity was expressed in nmol/mg protein/min.

For the measurement of nicotinamide adenine dinucleotide (NADH) cytochrome *c* reductase (NCCR; enzyme for electron transport between ETC Complex I and Complex III) activity, mitochondrial fraction was incubated in a mixture containing K2HPO4 buffer, KCN, β-NADH, and rotenone at 37 ◦C for 2 min. After the addition of cytochrome *c* (50 μM), the reduction of oxidized cytochrome *c* was measured as the difference in the presence or absence of rotenone at 550 nm for 3 min at 37 ◦C.

For the determination of succinate cytochrome *c* reductase (SCCR; enzyme for electron transport between ETC Complex II and Complex III) activity, mitochondrial fraction was performed in the same buffer solution supplemented with succinate. After a 5 min equilibration at 37 ◦C, cytochrome *c* (50 μM) was added and the reaction was monitored at 550 nm for 3 min at 37 ◦C.

For the determination of cytochrome *c* oxidase (CCO, marker enzyme for ETC Complex IV) activity, mitochondria fraction was pre-incubated at 30 ◦C for 5 min in K2HPO4 buffer, then 45 μM ferrocytochrome *c* was added to start the reaction, which was monitored at 550 nm for 3 min at 30 ◦C. In all measurements, experiments were performed in triplicate, and the mean was used for statistical analyses.

#### *2.16. Measurement of ATP Levels in RVLM*

RVLM tissues were centrifuged at 10,000× *g* for 10 min after homogenization in a protein extraction solution (Pierce, Rockford, IL, USA), ATP concentration in the supernatant was determined by an ATP colorimetric assay kit (AbCam, Waltham, MA, USA) using a microplate reader (ThermoFisher Scientific). The ATP level was normalized to the protein concentration of the sample. Each measurement was performed in triplicate and the mean was used for statistical analyses.

#### *2.17. Western Blot Analysis*

We determined the expression levels of gp91phox, p22phox, p67phox, and p47phox subunits of NADPH oxidase, manganese dismutase (SOD2), nuclear factor erythroid 2-related factor 2 (Nrf2), nNOS, iNOS, eNOS, NADPH oxidase activator 1 (Noxa1), uncoupling protein 3 (UCP3), and GAPDH in total protein extracted from RVLM by Western blot analysis [43]. In brief, 8–12% SDS-polyacrylamide gel electrophoresis was used for protein separation, and a Bio-Rad miniprotein-III wet transfer unit (Bio-Rad) was employed to transfer samples onto polyvinylidene difluoride transfer membranes (Immobilon-P membrane; Millipore, Bedford, MA) for 1.5 h at 4 ◦C. The transfer membranes were then incubated with a blocking solution (5% nonfat dried milk dissolved in Tris-buffered saline-Tween buffer) for 1 h at room temperature. Primary antisera used in this study included goat polyclonal, rabbit polyclonal or monoclonal, or mouse monoclonal antiserum against gp91phox (1:5000; BD Biosciences, Sparks, MD, USA), p22phox, p67phox, and p47phox (1:5000; Santa Cruz Biotechnology, Santa Cruz, CA, USA), SOD2 (1:3000; Stressgen, San Deigo, CA, USA), Nrf2 (1:1000; Santa Cruz), nNOS, iNOS, and eNOS (1:1000; BD Biosciences), Noxa1 (1:1000, Santa Cruze), UCP3 (1:1000, Santa Cruz), and GAPDH (1:10,000; Merck). Membranes were subsequently washed three times with TBS-t buffer, followed sequentially by incubation with the secondary antibodies (1:10,000; Jackson ImmunoResearch, West Grove, PA, USA) for 1 h and horseradish peroxidase-conjugated goat anti-rabbit IgG or goat anti-mouse IgG (Jackson ImmunoResearch). An enhanced chemiluminescence Western blot detection system (GE Healthcare Bio-Sciences Corp., Piscataway, NJ, USA) was used to detect specific antibody–antigen complex. For the detection of eNOS or nNOS dimerization, nondenaturing, low-temperature sodium dodecyl sulfate polyacrylamide gel electrophoresis was used [32,34]. During the electrophoresis process and transfer of proteins to nitrocellulose membrane, buffers were placed in an ice-water bath and the whole apparatus was kept at 4 ◦C. ImageJ software (NIH, Bethesda, MD, USA) was used to quantify the number of detected proteins, which was expressed as the ratio to loading control (GAPDH).
