Capillary hemodynamics and contracting skeletal muscle oxygen pressures in male rats with heart failure: Impact of soluble guanylyl cyclase activator

Abstract

In heart failure with reduced ejection fraction (HFrEF), nitric oxide-soluble guanylyl cyclase (sGC) pathway dysfunction impairs skeletal muscle arteriolar vasodilation and thus capillary hemodynamics, contributing to impaired oxygen uptake $\left({\stackrel{.}{\text{VO}}}_2\right)$ kinetics. Targeting this pathway with sGC activators offers a new treatment approach to HFrEF. We tested the hypotheses that sGC activator administration would increase the O₂ delivery $\left({\stackrel{.}{\text{QO}}}_2\right)\text{-to-}{\stackrel{.}{\text{VO}}}_2$ ratio in the skeletal muscle interstitial space (PO₂is) of HFrEF rats during twitch contractions due, in part, to increases in red blood cell (RBC) flux (f_RBC), velocity (V_RBC), and capillary hematocrit (Hct_cap). HFrEF was induced in male Sprague-Dawley rats via myocardial infarction. After 3 weeks, rats were treated with 0.3 mg/kg of the sGC activator BAY 60–2770 (HFrEF + BAY; n = 11) or solvent (HFrEF; n = 9) via gavage b.i.d for 5 days prior to phosphorescence quenching (PO₂is, in contracting muscle) and intravital microscopy (resting) measurements in the spinotrapezius muscle. Intravital microscopy revealed higher f_RBC (70 ± 9 vs 25 ± 8 RBC/s), V_RBC (490 ± 43 vs 226 ± 35 μm/s), Hct_cap (16 ± 1 vs 10 ± 1%) and a greater number of capillaries supporting flow (91 ± 3 vs 82 ± 3%) in HFrEF + BAY vs HFrEF (all P < 0.05). Additionally, PO₂is was especially higher during 12–34s of contractions in HFrEF + BAY vs HFrEF (P < 0.05). Our findings suggest that sGC activators improved resting ${\stackrel{.}{\text{QO}}}_2$ via increased f_RBC, V_RBC, and Hct_cap allowing for better ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ matching during the rest-contraction transient, supporting sGC activators as a potential therapeutic to target skeletal muscle vasomotor dysfunction in HFrEF.

1. Introduction

Heart failure (HF) is a prevalent chronic disease with high morbidity and mortality [1]. Following myocardial infarction (MI), multiple organ system dysfunction coalesces to constrain skeletal muscle oxygen delivery $\left({\stackrel{.}{\text{QO}}}_2\right)$ [2]. Thus, HF severity is often classified by the impaired O₂ transport measured via peak oxygen uptake tests (${\stackrel{.}{\text{VO}}}_2$ peak) [3,4]. Within skeletal muscle in HF with reduced ejection fraction (HFrEF) there is decreased nitric oxide (NO) bioavailability, impaired endothelium-dependent arteriolar dilation, capillary involution, and compromised capillary hemodynamics at rest and during muscle contractions, resulting in decreased skeletal muscle perfusive and diffusive O₂ transport [5-7]. Accordingly, skeletal muscle in HFrEF has a limited ability to match ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ spatially and temporally resulting in greater deoxygenation (i.e., decreased interstitial O₂ partial pressures, PO₂is) following the onset of contractions [8].

Ferreira and colleagues [9] found that the NO-donor sodium nitroprusside could acutely improve the microvascular O₂ driving pressure (PO₂mv) in contracting skeletal muscle of HFrEF rats. However, there has been substantial interest in stimulating the soluble guanylyl cyclase (sGC) vasodilation cascade (i.e., downstream of NO). sGC activators have shown efficacy for increasing enzymatic activity of sGC to generate cyclic guanosine monophosphate (cGMP) independently of NO [10]. sGC activators preferentially target the NO-insensitive, oxidized, pathological, and heme-deficient form of sGC present in HFrEF [10]. In fact, long term treatment with the sGC activator, Ataciguat, in HFrEF-induced rats, reduced mitochondrial superoxide production, enhanced cardiac capillary density, reduced left ventricular filling pressures, and overall showed more effective cardioprotection in comparison to angiotensin-converting enzyme therapy [11]. sGC activators have greater pharmacological activity in diseased (heme-deficient) versus healthy isolated vascular smooth muscle cells by inducing endothelium-independent smooth muscle vasodilation, thereby demonstrating the potential for improving skeletal muscle oxygenation and improved capillary hemodynamics in HFrEF [10]. Therefore, we hypothesized that treatment with a sGC activator would increase the ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ ratio in the skeletal muscle interstitial space (PO₂is) of HFrEF rats at rest and during twitch contractions due, in part, to increases in red blood cell (RBC) flux (f_RBC), velocity (V_RBC), and capillary tube hematocrit (Hct_cap).

2. Methods

2.1. Animals

All procedures were approved by the Institutional Animal Care and Use Committee of Kansas State University and conducted according to the National Research Council Guide for the Care and Use of Laboratory Animals. Experiments were performed on male Sprague-Dawley rats (n = 20; 3–4 mo old, Charles River Laboratories, Wilmington, MA). Upon arrival animals were maintained in accredited (Association for the Assessment and Accreditation of Laboratory and Animal Care) animal facilities under a 12:12 h light:dark cycle with food and water provided ad libitum.

2.2. Myocardial infarction (MI) procedure

MI was induced via surgical ligation of the left main coronary artery [12]. Rats were anesthetized with a 2.5% isoflurane-O₂ mixture and intubated for mechanical ventilation with a rodent respirator (model 680, Harvard Apparatus, Holliston, MA) for the duration of the surgical procedure. The heart was exposed through a left thoracotomy in the fifth intercostal space and the left main coronary artery was ligated with 6–0 silk suture ~1–2 mm distal to the edge of the left atrium. Ampicillin (50 mg kg ⁻¹ i.m.) was injected into the closed incision to reduce susceptibility to infection. Thereafter, analgesic agents bupivacaine (1.5 mg kg ⁻¹ s.c.) and buprenorphine (0.01–0.05 mg/kg i.m.) were administered. All rats were then monitored closely for ~6 h post-MI for the development of arrhythmias and signs of stress. Rats were assessed daily for signs of distress (appetite, weight loss/gain, gait/posture, etc.) and care was administered as appropriate according to an intensive 14-day post-operative plan conducted in conjunction with the university veterinary staff.

2.3. Echocardiography

21 days following MI echocardiography was performed to assess left ventricular (LV) dysfunction. Transthoracic echocardiography was performed using a commercially available system (Logiq S8; GE Health Care, Milwaukee, WI) with an 18 MHz linear transducer (L8-18i) as previously described [8]. Rats were initially anesthetized with a 2.5% isoflurane-O₂ mixture and placed on a heating pad to maintain core temperature. Standard two-dimensional and M-mode images from the midpapillary level were obtained with frame rates of >50 frames/s. Ventricular dimensions and wall thicknesses were obtained from M-mode measurements over four consecutive cardiac cycles. LV internal dimensions and posterior wall (PW) thicknesses were measured at end systole (LVIDs; PWs) and end diastole (LVIDd; PWd). Fractional shortening (FS) was calculated from the measurements of LV chamber diameters: FS = [(LVIDd − LVIDs)/LVIDd] × 100. LV end-systolic (LVESv) and end-diastolic (LVEDv) volumes were calculated using the Teichholz formula: LV volume = (7.0/2.4 + LV dimension) × LV dimension³. Stroke volume was calculated as: SV = LVEDv- LVESv. Ejection fraction (EF) was calculated as: EF = [LVEDv- LVESv/LVEDv] × 100.

2.4. Drug dosing

Following echocardiography rats were randomly assigned to activator (HFrEF + BAY; n = 11) or control HFrEF (HFrEF; n = 9) groups. The sGC activator BAY 60–2770 (Bayer AG, Pharmaceuticals, Wuppertal, Germany) was weighed and mixed with 10% Transcutol, 20% Cremophor, (Sigma Aldrich, St. Louis, MO), and 70% water to obtain a dose with the final concentration of 0.3 mg/kg BAY 60–2770 in a 1 ml volume. The HFrEF group was given solvent only. Either BAY 60–2770 or solvent was administered via oral gavage. To mimic clinical protocols, the drug/solvent was administered b.i.d. for 5 days prior to the following post-MI experiments.

2.5. Surgical preparations

After completion of the dosing regimen rats were anesthetized initially with a 5% isoflurane-O₂ mixture and subsequently maintained on 2.5% isoflurane-O₂ while positioned on a heating pad to maintain core temperature at ~37 °C as measured via rectal thermometer. The carotid artery was isolated and cannulated and a 2-French catheter-tip pressure transducer (Millar Instruments, Houston, TX, USA) was advanced into the LV for measurements of systolic and diastolic pressures and changes in LV pressure over time (LV dp/dt). Upon completion of the LV measurements the transducer was removed and the carotid artery was cannulated with a catheter (PE-10 connected to PE-50, Intra-Medic polyethylene tubing, Clay Adams Brand, Benton, Dickson and Company, Sparks, MD, USA) for measurement of mean arterial pressure (MAP) and heart rate (HR) (Digi-Med BPA; Model 400). The caudal artery was catheterized for administration of pentobarbital sodium anesthesia (50 mg/ml) and arterial blood sampling. Rats were then transitioned to pentobarbital sodium anesthesia (20 mg/kg body wt.) given intra-arterially while concentrations of isoflurane were decreased and subsequently discontinued. The depth of anesthesia was regularly monitored via toe pinch and palpebral reflex, with pentobarbital anesthesia supplemented (3.5–7.0 mg/kg) as necessary.

The overlying skin and fascia were carefully removed from the mid-dorsal-caudal region of the rat and the left spinotrapezius muscle was exposed in a manner that did not disrupt vascular supply to the muscle [13]. This muscle was chosen for its similar fiber-type composition and oxidative capacity to the untrained human quadriceps muscle [14]. Silver wire electrodes were then sutured (6-0 silk) to the rostral (cathode) and caudal (anode) regions of the muscle to induce twitch contractions. The exposed muscle was continuously superfused with Krebs-Henseleit bicarbonate-buffered solution (4.7 mM KCl, 2.0 mM CaCl₂, 2.4 mM MgSO₄, 131 mM NaCl, and 22 mM NaHCO₃; pH = 7.4; equilibrated with 5% CO₂–95% N₂ at 38 °C) and the surrounding tissue was covered with Saran Wrap (Dow Industries, Indianapolis, IN) to minimize dehydration.

2.6. Experimental protocol

Following surgical preparations phosphorescence quenching was used to determine PO₂is at rest and during contractions using a frequency domain phosphorimeter (PMOD 500; Oxygen Enterprises, Philadelphia, PA) as previously described [15,16]. Oxyphore G4 (Pd-meso-tetra-(3,5-dicarboxyphenyl)-tetrabenzo-porphyrin) was injected locally into the muscle (3–4 injections at 10-μM concentration) using a 29-G needle. Care was taken to avoid damaging any visible vasculature. This probe allows dynamic visualization of the tissue PO₂is levels of the spinotrapezius muscle [17]. Following injections, the spinotrapezius muscle was covered with Saran wrap to protect the muscle from dehydration during a 15-min period which allows for diffusion of Oxyphor G4 throughout the muscle into the interstitial space. A bifurcated light guide was positioned ~3–4 mm above the surface of the exposed muscle in a field absent of large vessels. To minimize any phosphorescent interference all measurements were performed in a dark room with minimal light exposure.

2.7. Contractions protocol

Twitch contractions were electrically evoked (1Hz, 6–7 V, 2-ms pulse duration) with a Grass S88 Stimulator (Quincy, MA) for 180 s PO₂is was measured and recorded at 2 s intervals throughout the duration of the protocol. This stimulation protocol elicits a 4-5-fold increase in blood flow and a 7-fold increase in metabolic rate consistent with moderate intensity exercise [18,19]. Immediately following PO₂is measurements approximately 0.3 ml of blood was sampled from the caudal artery catheter for the determination of arterial blood lactate ([La⁻]), pH, P_CO2, %O₂ saturation, and systemic hematocrit (Nova Stat Profile M; Nova Biomedical, Waltham, MA).

2.8. Analysis of interstitial PO₂ kinetics

The kinetics analyses of the PO₂is response was measured utilizing the Stern-Volmer relationship. Direct measurement of phosphorescence lifetime yielded PO₂is via the following equation:

$$P{O}_2=[{\tau }_0-1]∕{\kappa }_{𝒬}x{\tau }_0$$

where ${\kappa }_{𝒬}$ is the quenching constant and τ and τ₀ are the phosphorescence lifetimes at the ambient O₂ concentration and in the absence of O₂, respectively. In tissues at 32.3 °C (mean spinotrapezius muscle surface temperature was ~32 °C) the parameters for G4 are as follows: k_Q of 258 mmHg⁻¹ s⁻¹ and τ₀ of 226 μs [17]. As these parameters are dependent on pH and temperature of the muscle, neither of which change sufficiently to impact the PO₂is measurement during the contraction protocol, the resolved phosphorescence lifetime is determined by the PO₂is. The kinetics analyses of the PO₂is responses were conducted using 10 s of resting data and the 180-s contraction bout using a monoexponential plus time delay model:

$$P{O}_2(t)=P{O}_{2(BL)}-{\Delta }_{1}P{O}_2\left[1-e^{-\frac{t-TD}{\tau }}\right]$$

or a monoexponential plus time delay using a secondary component when required:

$$P{O}_2(t)=P{O}_{2(BL)}-{\Delta }_{1}P{O}_2\left[1-e^{-\frac{t-TD}{\tau }}\right]+{\Delta }_{2}P{O}_2\left[1-e^{-\frac{t-T{D}_2}{{\tau }_2}}\right]$$

where PO₂ (t) represents the PO₂is at any point in time, PO_{2 (BL)} is the baseline before the onset of contractions, Δ₁ and Δ₂ are the primary and secondary amplitudes, TD and TD₂ are the time delays for each component, and τ and τ₂ are the time constants (i.e., the time required to reach 63% of the amplitude) for the primary and secondary amplitudes. The mean response time was calculated as the sum of the model-derived TD and τ. When the secondary component model was necessary, the primary amplitude was constrained to not exceed the nadir value to maximize accuracy of the primary response kinetics. The goodness of model fit was determined using the following criteria: 1) the coefficient of determination; 2) sum of squared residuals; 3) visual inspection and analysis of the model fits to the data and the residuals. Since the second amplitude (Δ₂PO₂, undershoot of the PO₂) was often monoexponential in nature, it was manually calculated by subtracting the difference between the PO₂is at the end of contractions minus the nadir value of PO₂is during contractions.

2.9. Intravital microscopy

Following PO₂is measurements, electrode sutures were gently removed. The caudal end of the spinotrapezius was carefully exteriorized and sutured (6-0 silk) to a thin wire horseshoe-shaped manifold at five or six equidistant points around the perimeter. The rat was placed on a water-heated Lucite platform with the spinotrapezius positioned such that a microvascular field within the midcaudal and dorsal surface was observed using an intravital microscope (Nikon, Eclipse E600-FN; × 40 objective; 0.8 numerical aperture) equipped with a noncontact illuminated lens and a high-resolution color monitor (total viewing area = 270 × 210 μm; Sony Trinitron PVM-1954Q, Ichinonya, Japan). The muscle was transilluminated to ensure clear resolution of the sarcomere A-bands. The final magnification ( × 1184) was confirmed by initial calibration of the system using a stage micrometer (MA285, Meiji Techno). This magnification is adequate for measuring all essential structural and hemodynamic variables [6,20]. The spinotrapezius muscle was maintained at physiological sarcomere length (~2.7 μm) throughout the subsequent observation period, and any exposed tissue was continuously superfused with the Krebs-Henseleit solution. Once the spinotrapezius muscle was positioned on the platform, 5–10 microvascular viewing fields were each recorded for ~1–1.5 min. Recordings were time referenced and stored for subsequent offline analysis.

2.10. Capillary data analysis

5-7 fields were chosen randomly from each rat for analysis on the basis of clear visualization of sarcomeres, fibers, and capillaries. Capillaries supporting RBC flow were assessed in real time, and each capillary was placed into one of two categories: 1) normal flow = 30 s of continuous, or 2) impeded flow or stopped flow for >10 of 30 s. These criteria were used for determination of percentage of flowing capillaries [i.e. (number of capillaries supporting RBC flow/total number of visible capillaries per area), × 100]. For all capillaries in which hemodynamics were assessed and where the capillary endothelium was clearly visible on both sides of the lumen, capillary luminal diameter was measured (2–4 measurements/capillary) with calipers accurate to ± 0.25 mm (±0.17 μm at × 1184 magnification). Capillary lineal density was determined as the number of capillaries crossing a line drawn perpendicular to the fiber axis.

Examination of the microvascular fields was conducted by frame-by-frame analysis techniques (Dartfish Video Software, Fribourg, Switzerland). Sarcomere length was determined from sets of 10 consecutive in-register sarcomeres (i.e., distance between 11 consecutive A-bands) measured parallel to the muscle fiber longitudinal axis. This measurement was repeated 3–4 times where sarcomeres were visible to obtain a mean sarcomere length for each viewing field. V_RBC was determined in all capillaries that were continuously perfused by following the RBC path length. f_RBC was measured by counting the number of RBCs in a capillary passing an arbitrary point for several frames. For each capillary in which hemodynamic data were measured, Hct_cap was calculated by the following equation:

$${\text{Hct}}_{\text{cap}}=({\text{volume}}_{\text{RBC}}\mathrm{x}{f}_{\text{RBC}})∕[\pi \mathrm{x}({\mathrm{d}}_{\mathrm{c}}∕2{)}^2\mathrm{x}{V}_{\text{RBC}}]$$

where volume_RBC is red blood cell volume, which was taken to be 61 μm³ [21] and capillaries were approximated as circular in cross section [22]. Following intravital microscopy the animals were euthanized with an overdose of pentobarbital sodium (>100 mg/kg).

2.11. Heart failure classification

Following experimentation, the heart was excised and the chambers were separated. The LV was opened by making an incision through the interventricular septum, from the base to the apex and was spread out onto a wax sheet. A digital photograph of the endocardium was taken to demarcate between the remodeled (luminous) from healthy (dark red) myocardium. The image was printed and infarct size determined by planimetry as previously described [8,23].

2.12. Western immunoblotting

sGC is a heterodimer consisting of two subunits, α and β [24]. In this study we only investigated the concentration of sGCβ₁ within the spinotrapezius muscle, because the literature suggests the β₁ subunit is the common dimerizing subunit for all isoforms of sGC and its deletion results in loss of both isoforms [25] making it key to the NO-sGC cascade. To quantify the sGCβ₁ subunit expression, samples were homogenized in RIPA buffer (Sigma-Aldrich, St. Louis, MO) containing a protease inhibitor cocktail (1:100 vol:vol; Sigma-Aldrich). A BeadBug 6 homogenizer (Benchmark Scientific, Edison, NJ) with 3.0 mm zirconium beads was used to homogenize the tissues. Although sGCβ₁ expression was measured in the entire spinotrapezius muscle, which may be influenced by varying cell types, we expect this to be largely representative of sGCβ₁ expression in the myocytes. Following 3–30 s cycles of 4350 rpm with 30 rests, homogenates were centrifuged at 1500 g for 10 min. The supernatant was collected and protein concentration determined following Qubit protocol. Jess Simple Western System (Protein Simple, San Jose, CA, USA) was performed following the instruments protocol. Proteins were loaded at 2 mg/ml with primary antibody (sGCβ₁, Novus, NBP1-89784) concentration of 1:50, which is the appropriate antibody concentration with the Jess system. GAPDH (Novus, NB300-221) served as a housekeeping protein. Compass simple western software (V 4.1.0) was used to calculate heights, area, and signal/noise ratio. All samples were run in duplicate and repeated at least twice.

2.13. Statistical analysis

All PO₂is curve fitting and statistical analyses were performed using a commercially available software package (SigmaPlot 12.5, Systat Software, San Jose, CA). Student’s paired and unpaired t-tests were performed, when appropriate, to determine differences in morphometric, pressure, and microscopy measurements followed by Pearson correlation tests. A student’s unpaired t-test was used to detect differences in PO₂is kinetic parameters between HFrEF + BAY and HFrEF. A two-way repeated measures ANOVA was used to assess temporal interactions (Group x Time). Holm-sidak post hoc tests were used for multiple comparisons when significant differences were detected. Data are presented as means ± SE. Significance was accepted at P < 0.05.

3. Results

3.1. Hemodynamic and morphometric data

20 rats (HFrEF + BAY, n = 11; HFrEF, n = 9) were analyzed for hemodynamic and morphometric data. Prior to dosing the rats, measurements of LV function were assessed via transthoracic echocardiography. No differences were found between groups in any echocardiographic measurements (fractional shortening 20.5 ± 3.0 vs. 21.0 ± 2.9%; ejection fraction 44.7 ± 5.1 vs. 46.3 ± 4.8%; stroke volume 0.8 ± 0.1 vs. 0.9 ± 0.1 ml/beat, HFrEF + BAY vs HFrEF; all p > 0.05). Importantly, LV end-diastolic pressure (LVEDP) (taken immediately prior to terminal experiments) and the index of MI (% of endocardial surface area infarcted; assessed via planimetry) are indicative of moderate HFrEF [23,26] (Table 1) and did not differ between groups.

Table 1.

Hemodynamic and morphometric data.

	HFrEF	HFrEF + BAY	P-value
Infarct Size (%)	28 ± 3	29 ± 2	0.83
LVEDP (mmHg)	14 ± 1	16 ± 1	0.46
LV + dP/dt (mmHg/s)	8082 ± 264	7709 ± 397	0.60
LV/BW (mg/g)	2.00 ± 0.04	1.96 ± 0.07	0.90
RV/BW (mg/g)	0.62 ± 0.04	0.62 ± 0.03	0.91
Lung/BW (mg/g)	3.17 ± 0.10	3.75 ± 0.12*	0.01
Lung Weight (g)	1.58 ± 0.05	1.70 ± 0.08	0.16
Body Weight (g)	500 ± 12	454 ± 10*	0.02
Initial Body Weight (g)	370 ± 11	356 ± 11	0.41

Data are means ± SE. Analyzed via student’s unpaired t-test. LVEDP, left ventricular end-diastolic blood pressure; LV, left ventricle; +dP/dt, rise in blood pressure over time; RV, right ventricle; BW, body weight.

^*P < 0.05 vs HFrEF; HFrEF, n = 9; HFrEF + BAY, n = 11.

3.2. Phosphorescence quenching

19 rats were assessed for the PO₂is measurements due to one not completing the protocol (HFrEF + BAY n = 10; HFrEF n = 9). Mean PO₂is profiles at rest and during contractions are presented in Fig. 1. There were no significant differences in MAP or HR during contractions (Table 2). Following contractions there was a lower lactate concentration in HFrEF + BAY (1.06 ± 0.07 vs 1.34 ± 0.12 mmol/L), but no differences in blood gases (PaO₂: 86.9 ± 2.5 vs 87.2 ± 1.8 mmHg; SaO₂: 95.8 ± 0.4 vs 95.6 ± 0.4%) or pH (7.4 ± 0.01, both groups) were observed, HFrEF + BAY vs HFrEF, respectively. No significant differences were identified for the kinetics parameters between HFrEF + BAY and HFrEF (P > 0.05; Table 3). However, PO₂is values were particularly higher in HFrEF + BAY rats (P < 0.05; Fig. 1) during 12–34 s of contractions.

Fig. 1. Interstitial PO₂ from rest to contractions in spinotrapezius muscle.

Data are means ± SE. Analyzed via two-way ANOVA (Group x Time). Group average spinotrapezius interstitial PO₂ (PO₂is) at rest and during electrically induced contractions for HFrEF + BAY (open circles; n = 10) and HFrEF (filled circles; n = 9); contractions beginning at 0s. *P < 0.05 (shaded area; 12–34 s).

Table 2.

Central hemodynamics during phosphorescence quenching.

	HFrEF	HFrEF + BAY	P-value
Heart Rate (bpm)	359 ± 6	357 ± 5	0.80
MAP (mmHg)	107 ± 5	105 ± 2	0.53

Data are means ± SE. Analyzed via student’s unpaired t-test. MAP, mean arterial pressure. HFrEF, n = 9; HFrEF + BAY, n = 11.

Table 3.

Interstitial PO₂ kinetics parameters of spinotrapezius at rest and during 180 s of contractions.

	HFrEF	HFrEF + BAY	P-value
PO2(BL), mmHg	23.0 ± 1.9	19.2 ± 1.2	0.08
Δ1PO2, mmHg	11.9 ± 1.2	13.0 ± 2.1	0.61
T, S	12 ± 1	15 ± 1	0.09
TD, S	9 ± 1	6 ± 1	0.12
MRT, S	26 ± 4	24 ± 4	0.73
PO 2(Nadir)	7.4 ± 1.3	9.9 ± 1.3	0.12
Δ2PO2, mmHg	5.1 ± 1.2	3.4 ± 0.6	0.46
PO2(End), mmHg	12.9 ± 1.7	13.3 ± 1.6	0.57
AUC, mmHg	1100 ± 123	1321 ± 114	0.26
ΔPO2/ T, mmHg/S	1.0 ± 0.1	0.9 ± 0.1	0.31

Data are means ± SE. PO_2(BL), baseline interstitial PO₂; Δ1PO₂, primary amplitude; τ, timepoint of τ; TD, time delay, MRT, mean response time; PO_2(nadir), lowest response of PO₂ profile; Δ2PO₂, secondary amplitude, PO_2(End), interstitial PO₂ at end of contractions; AUC, area under the curve, total PO₂ throughout experimentation; ΔPO₂/τ, change in PO₂ for a given τ. Compared via two-tailed unpaired t tests.

3.3. RBC hemodynamics

17 rats (HFrEF + BAY; n = 9, HFrEF; n = 8) were utilized for intravital capillary hemodynamic measurements due to two animals losing cardiovascular stability during measurements. Sarcomere length was not different across groups and conditions, ranging from 2.4 to 2.7 μm. No differences in capillary lineal density for all capillaries were seen in HFrEF + BAY vs HFrEF (P = 0.33). Increased f_RBC, V_RBC. Hct_cap, and % of capillaries with RBC flow were observed in HFrEF + BAY vs HFrEF (P < 0.05; Fig. 2). V_RBC was positively and linearly correlated with f_RBC (R = 0.958; Fig. 3).

Fig. 2. Capillary hemodynamics in spinotrapezius muscle at rest.

Data are means ± SE. Analyzed via student’s unpaired t-test. A. RBC flux in skeletal muscle capillaries supporting flow. B. RBC velocity in skeletal muscle capillaries supporting flow. C. Capillary hematocrit in skeletal muscle capillaries. D. Percentage of capillaries supporting RBC flow. HFrEF + BAY n = 9; HFrEF n = 8; all * P < 0.05.

Fig. 3. Relationship between average RBC velocity and flux in capillaries supporting continuous flow in HFrEF + BAY and HFrEF.

(P < 0.05).

3.4. Western immunoblotting

18 rats (HFrEF + BAY; n = 10, HFrEF; n = 8) were used to assess sGCβ₁ expression within the spinotrapezius muscle. Group means were not statistically significant (P = 0.17); however, group averages show BAY 60–2770 allowing for an increase in sGCβ₁ expression compared to the HFrEF group (Fig. 4).

Fig. 4. Western immunoblotting.

Data are means ± SE. A. Normalized protein expression in HFrEF (n = 8) vs HFrEF + BAY (n = 10) (P > 0.05). B. Representative blotting of sGCβ1 expression in HFrEF vs HFrEF + BAY.

4. Discussion

This is the first investigation, to our knowledge, to study the effects of sGC activation (via BAY 60–2770) on capillary hemodynamics and PO₂is in the skeletal muscle of HFrEF rats. sGC activator increased PO₂is at the onset of contractions (12–34 s; Fig. 1) and improved capillary hemodynamics (increased f_RBC, V_RBC, and Hct_cap; Fig. 2A-D). This supports the notion that sGC activators may target dysfunctional sGC to allow for improved ${\stackrel{.}{\text{QO}}}_2-{\stackrel{.}{\text{VO}}}_2$ matching in the skeletal muscle of HFrEF rats at rest and following the onset of contractions. As previously demonstrated in HFrEF rats, there is a significantly lower PO₂is [8], capillary f_RBC, V_RBC and percentage of capillaries flowing [6] which impairs ${\stackrel{.}{\text{QO}}}_2$ and likely contributes to exercise intolerance. Novel therapeutic agents, such as BAY 60–2770, have been developed to increase sGC bioactivity >200-fold, regardless of redox state. However, western immunoblotting data herein show no significant differences in sGCβ₁ expression in the spinotrapezius in HFrEF + BAY rats (P = 0.17).

4.1. Resting hemodynamic measurements

Endothelial dysfunction and diminished skeletal muscle blood flow in HFrEF are due, in part, to decreased NO bioavailability via endothelial NO synthase (eNOS) uncoupling, increased reactive O₂ species, and ultimately sGC oxidation [27,28]. Determining the temporal and spatial distribution of RBCs, which is impaired in HFrEF [6,7], is imperative to understanding the effect of sGC activation on PO₂is and therefore blood-myocyte O₂ exchange. Importantly, capillary hemodynamics at rest in HFrEF rats exhibit a decreased f_RBC and V_RBC, which has been shown to improve with dietary nitrate supplementation [29,30]. Therefore, we hypothesized that sGC activator administration would increase the proportion of capillaries supporting flow and RBC dynamics (f_RBC, V_RBC, and Hct_cap). Indeed, in the skeletal muscle of HFrEF rats, following sGC activator administration, more capillaries supported RBC flow and there was an augmented capillary f_RBC (Fig. 2). The particulate nature of blood dictates that the skeletal muscle surface area per capillary available for O₂ exchange is a function of Hct_cap and capillary length in RBC-flowing capillaries [31,32]. In HFrEF + BAY there were ~10% more capillaries supporting RBC flow, which could contribute to an increased ${\stackrel{.}{\text{VO}}}_2$ with contractions. Crucially, in HFrEF those capillaries not supporting RBC flow at rest do not start flowing with contractions [7]. More capillaries supporting RBC flow in HFrEF with a greater Hct_cap allows for greater perfusive and diffusive O₂ delivery (DO₂) with higher V_RBC, also promoting longitudinal recruitment of capillary surface area, thereby allowing better matching of ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$.

Interestingly, the 3–4 mmHg higher PO₂is in HFrEF + BAY rats at rest prior to contractions was not statistically different. It is possible that this reflects, in part, an increased ${\stackrel{.}{\text{VO}}}_2$ as permitted by the greater ${\stackrel{.}{\text{QO}}}_2$ . Another potential explanation for this finding resides in the spatial heterogeneity of ${\stackrel{.}{\text{QO}}}_2$ among individual fiber fields [6,20,31], where it is possible to have an increased f_RBC, V_RBC, and Hct_cap in a smaller region of skeletal muscle, whereas changes in PO₂is reflect a larger sample composed as the sum of smaller regions with variable ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ ratios. Notably, the elevated f_RBC, V_RBC, and Hct_cap in HFrEF + BAY at rest were associated with increased dynamic ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ matching as metabolic demands dramatically increase (i.e. 12-34 s during contractions) (Fig. 1).

In accordance with our findings, other studies have substantiated the pharmaceutical potency of sGC activators and stimulators in improving endothelial function following MI [11,33]. Importantly, in vivo, it was shown that the use of sGC activator BAY 58–2667, an analogue of BAY 60–2770, increased whole protein expression of the heme-binding sGCβ₁ subunit in pathological vessels [34]. However, we measured sGCβ₁ subunit expression in the spinotrapezius and documented no significant increase in sGCβ₁ among groups.

4.2. Contracting skeletal muscle measurements

sGC activators, which have been previously under-studied in skeletal muscle, can decrease progressive cardiac remodeling following MI [11, 34] and improve vascular smooth muscle function throughout the pulmonary vasculature [35,36]. Much of the literature supporting sGC activators as a therapeutic in HFrEF has focused on improving cardiac and pulmonary function [11,35], however, improving the peripheral vascular function in HFrEF via sGC activators offers a potential therapeutic avenue to improve exercise tolerance and promote effective exercise rehabilitation.

Skeletal muscle PO₂is kinetics reflect the dynamic matching between ${\stackrel{.}{\text{QO}}}_2$ and ${\stackrel{.}{\text{VO}}}_2$ within the interstitial space during transitions in metabolic demand [8,15,16]. In health, with the onset of muscular contractions there is a rapid increase in ${\stackrel{.}{\text{QO}}}_2$ (initial phase ~20 s) to meet mitochondrial ${\stackrel{.}{\text{VO}}}_2$ elevated demands [32,37-39]. This elicits shear-stress mediated vasodilation in locomotory skeletal muscle [40], followed by the secondary phase which reflects, in part, feedback control to maintain ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ matching [41]. However, peripheral derangements in rats with moderate HFrEF reduce capillary f_RBC and V_RBC [6]. This decrease in shear-stress on the luminal side of the endothelium hampers eNOS expression [9] and thereby impairs perfusive and diffusive O₂ delivery across the rest-to-contraction transition [6]. Notably, a more precipitous fall in PO₂is following the onset of contractions in HFrEF, compared to health, is indicative of a slower increase in ${\stackrel{.}{\text{QO}}}_2$ relative to ${\stackrel{.}{\text{VO}}}_2$ [8]. Prior studies from our laboratory demonstrated that 5 days of dietary nitrate supplementation in HFrEF rats enhances ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ matching (i.e., PO₂is, 29), potentially via increased percentage of capillaries supporting blood flow and greater f_RBC at rest and during contractions [30]. Within the capillary bed, there is a vast surface area for O₂ exchange, but diminished bulk blood flow and altered RBC distribution among capillaries limits ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ matching in skeletal muscles at rest [6] and during contractions in HFrEF [7,8].

Increased PO₂is in HFrEF + BAY compared to HFrEF demonstrated herein is expected to speed ${\stackrel{.}{\text{VO}}}_2$ kinetics, thereby decreasing the O₂ deficit and reliance upon anaerobic pathways for ATP regeneration. This will help support sustained exercise bouts by reducing intracellular metabolic perturbations that accelerate glycogenolysis and contribute to exhaustion [42,43]. Evidence supporting this notion, including the lower blood lactate concentration following twitch contractions in HFrEF + BAY compared to HFrEF herein indicate either a reduced glycolysis during contractions, improved lactate removal, or both [2,42,44]. Improving metabolism is imperative in HFrEF, because previous studies have shown accelerated glycolysis and glycogenolysis in response to an increased proportion and recruitment of type II fibers in HFrEF skeletal muscle [44-46] therefore decreasing their exercise capacity. Moreover, as DO₂ is determined by the number of RBCs adjacent to the contracting skeletal muscle fibers [31], the ~60% Hct_cap increase with sGC activation will elevate DO₂. This, combined with the enhanced perfusive O₂ delivery, may allow the HFrEF + BAY rats to speed their ${\stackrel{.}{\text{VO}}}_2$ kinetics further increasing exercise tolerance.

4.3. Clinical implications

Improving exercise tolerance is imperative to decreasing the morbidity and mortality in HFrEF patients. This study indicates that sGC activators may be beneficial to combating the exercise intolerance characteristic of HFrEF patients by increasing skeletal muscle ${\stackrel{.}{\text{QO}}}_2$ via activation of oxidized sGC. Although this study utilizes single muscle twitch contractions, evidence suggests that targeting individual muscles or muscle groups in patient populations can improve peripheral perfusive and diffusive O₂ transport and exercise capacity [47].

HFrEF patients function largely in the O₂-delivery-dependent region with respect to ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ kinetics (to the left of the “tipping point”, 2). Thus, any elevated muscle O₂ delivery has the potential to speed ${\stackrel{.}{\text{VO}}}_2$ kinetics and enhance exercise tolerance [2]. With increased sodium, and thus fluid retention, the progression and severity of HFrEF worsens. Interestingly, in HFrEF, sGC activators have recently been shown to decrease sympathetic outflow and increase renal perfusion (via increases in cGMP) [48]. There was a decreased body weight with HFrEF + BAY which may be due to decreased fluid retention or appetite in the BAY group, or potentially increased production of brown adipose tissue in place of the less metabolically active white adipose tissue [49].

4.4. Experimental considerations

The present investigation assessed the use of sGC activators in male rats. Women have a lower risk of developing heart disease until post-menopausel, when the incidence increases dramatically. Recent data suggests that women rely on greater basal NO bioavailability in HFrEF [15], therefore, future studies warrant investigating PO₂is and capillary hemodynamics in female rats and ovariectomized female rats receiving the sGC activator. In addition, assessing capillary hemodynamics microscopically presents challenges regarding small sampling area/volume and variability among sampled fields. These were overcome partially by using rigorous techniques on several different locations per muscle to limit errors in assessing capillary hemodynamics. Furthermore, participation of multiple independent blinded investigators provided confidence in quality control. Lastly, sGC activators are capable of activating the oxidized heme on sGC, therefore, in order to assess the entirety of its effects in the microcirculation a larger MI%, (i.e., >40% of LV damage) resulting in greater increases in ROS (i.e., more pathological sGC), is necessary in order to assess the utility of sGC activators across the broader range of HFrEF severity possible in the rat post-MI model [50].

5. Conclusions

This study is the first to demonstrate efficacy of administration of a sGC activator in a HFrEF model in the skeletal muscle microcirculation and its ability to increase ${\stackrel{.}{\text{QO}}}_2$ relative to ${\stackrel{.}{\text{VO}}}_2$ at rest and during the exercise (muscle contraction) transient. Specifically, at rest the sGC activator increased the proportion of capillaries supporting flow, and elevated f_RBC, V_RBC, and Hct_cap. PO₂is was also higher during the rest-contraction transient suggesting improved ${\stackrel{.}{\text{QO}}}_2\text{-to-}{\stackrel{.}{\text{VO}}}_2$ matching during submaximal exercise. The increased components of perfusive and diffusive O₂ transport in HFrEF + BAY demonstrated herein have significant implications for sGC activators as a potential therapeutic avenue in HFrEF to improve exercise tolerance and quality of life.