Temperature effects on morphological integrity and Ca²⁺ signaling in freshly isolated murine feed artery endothelial cell tubes

Abstract

To study Ca²⁺ signaling in the endothelium of murine feed arteries, we determined the in vitro stability of endothelial cell (EC) tubes freshly isolated from abdominal muscle feed arteries of male and female C57BL/6 mice (5–9 mo, 25–35 g). We tested the hypothesis that intracellular Ca²⁺ concentration ([Ca²⁺]_i) responses to muscarinic receptor activation would increase with temperature. Intact EC tubes (length: 1–2 mm, width: 65–80 μm) were isolated using gentle enzymatic digestion with trituration to remove smooth muscle cells. A freshly isolated EC tube was secured in a chamber and superfused at 24 (room temperature), 32, or 37°C. Using fura-2 dye, [Ca²⁺]_i was monitored (ratio of fluorescence at 340- to 380-nm wavelength) at rest and in response to bolus doses of ACh (20 nmol to 200 μmol). The morphological integrity of EC tubes was preserved at 24 and 32°C. Based on the Ca²⁺ K_d values we determined for fura-2 (174 nM at 24°C and 146 nM at 32°C), resting [Ca²⁺]_i remained stable for 180 min at both 24 and 32°C (27 ± 4 and 34 ± 2 nM, respectively), with peak responses to ACh (20 μmol) increasing from ∼220 nM at 24°C to ∼500 nM at 32°C (P < 0.05). There was no difference in responses to ACh between EC tubes from male versus female mice. When EC tubes were maintained at 37°C (typical in vivo temperature), resting [Ca²⁺]_i increased by ∼30% within 15 min, and gaps formed between individual ECs as they retracted and extruded dye, precluding further study. We conclude that EC tubes enable Ca²⁺ signaling to be evaluated in the freshly isolated endothelium of murine feed arteries. While Ca²⁺ responses are enhanced by approximately twofold at 32 versus 24°C, the instability of EC tubes at 37°C precludes their study at typical body temperature.

intracellular [Ca²⁺] ([Ca²⁺]_i) is a key controller of endothelial cell (EC) function. Endothelium-dependent regulation of vasomotor tone is effected through Ca²⁺-dependent production of diffusible autacoids (e.g., nitric oxide and metabolites of arachidonic acid) (7, 13, 31, 51) and the opening of intermediate-conductance (K_Ca3.1) and small-conductance (K_Ca2.3) Ca²⁺-activated K⁺ channels (6, 32), which collectively provide negative feedback to smooth muscle cell (SMC) contraction. Changes in EC [Ca²⁺]_i also regulate vascular permeability (1, 47, 50) and infection (28) as well as proliferation and angiogenesis (24, 38). Physiological agonists, including ACh, ANG II, bradykinin, and serotonin, act via G protein-coupled receptors to stimulate the release of Ca²⁺ from the endoplasmic reticulum (ER) and the influx of Ca²⁺ through the plasma membrane (39). The rapid release of Ca²⁺ from internal stores followed by the activation of store-operated Ca²⁺ entry (SOCE) together with receptor-activated Ca²⁺ entry (RACE) is manifest as the classic biphasic Ca²⁺ response with an initial “peak” followed by a sustained “plateau” (10, 11, 49).

The nature of Ca²⁺ signaling in preparations of isolated ECs has been determined in large part using cell culture (12, 28, 35, 36). However, ECs in cell culture have altered phenotypes compared with native ECs, often including the loss of muscarinic receptors (5, 12, 48). While new insight into Ca²⁺ signaling of intact ECs in resistance arteries (32) and arterioles (46) has been afforded through transgenic mice (30), much less is known about Ca²⁺ signaling in the native endothelium, particularly in the absence of blood flow or surrounding SMCs, both of which can affect vasoactive EC signaling pathways (16, 26, 52). In light of the increasing application of murine models to study the vasculature, our goal was to isolate intact EC “tubes” from mice to provide new insight into the nature of Ca²⁺ signaling in the murine circulation. The first study (11) of Ca²⁺ signaling in EC tubes used relatively short (length: ∼800 μm) segments of hamster cremaster arterioles. A limitation of these earlier experiments was the potential loss (by washout) of the EC tube during superfusion. Therefore, a key technical goal for the present experiments was to obtain intact EC tubes of sufficient length to be secured at each end during continuous superfusion.

To study Ca²⁺ signaling properties of the intact murine endothelium from resistance vessels, we determined the in vitro stability of EC tubes freshly isolated from abdominal muscle feed arteries (AFAs), which enabled us to obtain long (1–2 mm) intact segments devoid of branching. The previous study (11) of [Ca²⁺]_i responses in hamster EC tubes was performed at room temperature. Whereas the Q₁₀ effect on biological systems is well documented (2, 3, 44), little is known of how temperature affects Ca²⁺ signaling in ECs of the microcirculation or whether such effects differ between sexes. Thus, a second goal was to obtain measurements at temperatures that more closely approximate normal physiological conditions. Using EC tubes prepared from AFAs of male and female C57BL/6 mice, we tested the hypothesis that Ca²⁺ responses to muscarinic receptor activation would increase with temperature. Despite reported differences between sexes in EC [Ca²⁺]_i (21, 29, 41), we further hypothesized that the effect of temperature on [Ca²⁺]_i of EC tubes would be similar for males and females. In addition, we determined the Ca²⁺ K_d values of fura-2 in our system, which enabled estimates of [Ca²⁺]_i within freshly isolated EC tubes of murine resistance vessels for the first time.

MATERIALS AND METHODS

Animal Care and Tissue Sampling

All procedures were approved by the Institutional Animal Care and Use Committee of the University of Missouri (Columbia, MO) and were performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Mice were maintained on a 12:12-h light-dark cycle at ∼23°C with standard rodent diet and water ad libitum. Male and female C57BL/6 mice bred at the University of Missouri (5–9 mo, 25–35 g) were anesthetized with an intraperitoneal injection of pentobarbital sodium (60 mg/kg). The abdomen was shaved, and a midline incision was made through the skin. Temperature recorded with a thermocouple placed under the abdominal skin was ∼37°C, indicating that AFAs (superior epigastric artery) typically reside at core temperature. The origin of each AFA was exposed near the sternum and ligated (6-0 silk suture) to maintain a column of blood within the vessel lumen to facilitate the resolution of the vessel during microdissection. Abdominal muscles were rapidly excised and immediately placed into ice-cold (4°C) dissection buffer (pH 7.4, 285–290 mosM) composed of (in mM) 137 NaCl, 5.6 KCl, 1 MgCl₂, 10 HEPES, 10 glucose, 0.01 sodium nitroprusside, and 0.1% BSA (no. 10856, United States Biochemical, Cleveland, OH). Chemicals and reagents were obtained from Sigma-Aldrich (St. Louis, MO) unless otherwise stated. The anesthetized mouse was euthanized by an overdose of pentobarbital followed by cervical dislocation.

Microdissection of AFAs and Isolation of EC Tubes

After being equilibrated for 10 min to relax SMCs, the abdominal muscles were transferred to a dissection chamber containing dissection buffer (at 4°C) and pinned onto a transparent silicone rubber block (Sylgard 184, Dow-Corning, Midland, MI). Bilaterally, each AFA courses underneath its respective rectus abdominus muscle in parallel to the linea alba. While being viewed through a stereomicroscope (MZ8, Leica Microsystems, Bannockburn, IL), an AFA was dissected from the point of ligation to the first major branch point (isolated segment length: 3–3.5 mm). Cannulation pipettes were pulled (P-97, Sutter Instruments, Novato, CA) from borosilicate glass capillaries (G150T-4, Warner Instruments, Hamden, CT). Using a microforge, the tip outer diameters were broken to 50–80 μm and heat polished. The AFA segment was cannulated at one end, and the lumen was flushed with chilled dissection buffer to remove residual blood. Each AFA segment was then cut into two to three smaller segments (each ∼1–2 mm long) and transferred into 12 × 75-mm culture tubes on ice containing 4 ml of dissection buffer.

Intact EC tubes were isolated according to procedures developed for hamster arterioles (11, 23) and adapted here for the mouse. Briefly, AFA segments were transferred to a 12 × 75-mm culture tube containing 1 ml of preheated (37°C) dissociation buffer (pH 7.4, 290–295 mosM) composed of 137 mM NaCl, 5.6 mM KCl, 1 mM MgCl₂, 10 mM HEPES, 10 mM glucose, 2 mM CaCl₂, 0.1% BSA, 0.62 mg/ml papain (P-4762), 1.5 mg/ml collagenase (C-8051), and 1.0 mg/ml DTT (D-8255). Segments were incubated for 30 min at 37°C and then transferred to enzyme-free dissociation buffer at room temperature. An individual AFA segment was transferred to a recording chamber (RC-27N, Warner) on the stage of an inverted microscope (Nikon Diaphot, Garden City, NY). During visual inspection at ×40 magnification, the surrounding SMCs were dissociated from the EC tube using gentle trituration through the heat-polished tip (internal diameter: ∼120 μm) of a micropipette pulled (P-97) from borosilicate glass capillary tubes (1B100-4, World Precision Instruments, Sarasota, FL).

The freshly isolated EC tube was positioned at the center of the recording chamber at room temperature (24°C). Each end was pressed gently against the chamber bottom (a 24 × 50-mm coverslip) using a blunt-tip holding micropipette (heat-polished borosilicate glass, external diameter: ∼100 μm) secured in three-axis micromanipulators (DT3–100, Siskiyou Design, Grants Pass, OR). The EC tube was extended to approximately in situ length by retracting each holding micropipette while gentle pressure was maintained on each end of the EC tube. Thus secured, the EC tube was continuously superfused (2.5 ml/min) with physiological salt solution (PSS; 290–295 mosM, pH 7.38–7.40 from 24–37°C) composed of (in mM) 137 NaCl, 5.6 KCl, 1 MgCl₂, 10 HEPES, 10 glucose, and 2 CaCl₂.

Morphological Integrity of EC Tubes

We assessed the morphological integrity of AFA EC tubes at three temperatures, which spanned room temperature to typical body core temperature. In the recording chamber, the EC tube was superfused at 24°C for ∼15 min to equilibrate. The superfusion solution was then either maintained at 24°C or raised to 32 or 37°C in three equal 10-min increments (each 1/3 of the total difference between 24°C and the new target temperature) during a 30-min period using a platform heater (PH-6, Warner) and an in-line heater (SH-27B, Warner) controlled by a dual temperature controller (TC-344B, Warner). The EC tube was superfused at the respective temperatures for 1 h, after which it was examined under a Nikon E800 microscope using differential interference contrast imaging. Images were acquired using ×40 Plan Fluor objective (numerical aperture: 0.75) coupled to a cooled charge-coupled device (CCD) megapixel camera (SPOT, Diagnostic Instruments, Sterling Heights, MI) interfaced to a personal computer.

Ca²⁺ Photometry

Ca²⁺ photometry was performed (4, 14) with an IonOptix system (Milford, MA). Using a ×20 objective (Nikon Fluor20, numerical aperture: 0.75), fura-2 dye was excited alternately (250 Hz) at 340 and 380 nm while fluorescence emission was collected at 510 nm and expressed as the ratio of fluorescence at 340- to 380-nm wavelength (F₃₄₀/F₃₈₀).

Determination of K_d values for fura-2 dye.

To estimate values for EC tube [Ca²⁺]_i, we determined the K_d values of fura-2 dye for Ca²⁺ using our photometry system. The pentapotassium salt form of fura-2 dye (F1200, Invitrogen, Carlsbad, CA) was added to 11 solutions of known free [Ca²⁺] (Calcium Calibration Kit 1, C3008MP, Invitrogen) prepared as previously described (4). Briefly, a glass coverslip was partitioned into compartments using thin strips of vacuum grease. The respective calibration solutions were prepared by serial dilution, 1 μl was added to each compartment, and a coverslip was placed on top to prevent evaporation. Polystyrene beads (diameter: 15 μm, Polysciences, Warrington, PA) within each solution maintained a constant optical path length between the two coverslips. Autofluorescence was recorded (calibration solution without dye), and fura-2 fluorescence in each [Ca²⁺] solution was recorded three times and averaged. After measurements at 24°C, coverslips were heated to 32°C, and the fluorescence was measured again. Minimum and maximum fluorescence ratios were determined in 0 μM [Ca²⁺] (R_min) and 39 μM [Ca²⁺] (R_max), respectively. The constant β was calculated from the minimum and maximum fluorescence during excitation at 380 nm with 0 μM [Ca²⁺] divided by that with 39.8 μM [Ca²⁺]. By plotting these data as log{β[(R − R_min)/(R_max − R)]} versus log[Ca²⁺], the log K_d value was determined from the x-axis intercept (see Fig. 2).

Fig. 2.

Double logarithmic plots for determining the Ca²⁺ K_d value of fura-2 at 24 and 32°C. The plot of log{β[(R − R_min)/(R_max − R)]} versus log Ca²⁺ concentration ([Ca²⁺]) with values obtained from known free [Ca²⁺] standards provides log K_d as the x-axis intercept from y = 0 of the linear regression equation. R_min and R_max are the minimum and maximum fluorescence ratios, respectively, R is the observed ratio of fluorescence at 340- to 380-nm wavelength (F₃₄₀/F₃₈₀), and β was calculated from the minimum and maximum fluorescence during excitation at 380 nm with 0 μM [Ca²⁺] divided by that with 39.8 μM [Ca²⁺]. A: K_d at 24°C = 174 nM. B: K_d at 32°C = 146 nM. Summary values are means ± SE; n = 6 each.

Fura-2 dye loading and Ca²⁺ measurements in EC tubes.

A freshly isolated EC tube was secured in the recording chamber and equilibrated for ∼15 min during superfusion (2.5 ml/min) with PSS at 24°C. During this period, fura-2 AM dye (F14185, Invitrogen) was dissolved in DMSO and diluted to 5 μM in PSS (final DMSO concentration: <0.5%). The fura-2 AM dye solution was added to the chamber, and the EC tube was incubated for 30 min without flow. Dye loading was followed by superfusion with PSS for 30 min to wash out excess dye and allow fura-2 within cells to deesterify. During this equilibration period, temperature was maintained at 24°C or raised to either 32 or 37°C over 30 min in 10- min increments as described above. Autofluorescence values at 510 nm during excitation at 340 and 380 nm were recorded before dye loading and subtracted from the respective recordings. The imaging window for collecting fluorescence emission was 320 μm long and adjusted to the width of each EC tube (65–80 μm) while it was observed on a digital video monitor (LCD1550V, NEC, Rancho Dominguez, CA) using transmitted light from a halogen lamp (600-nm long-pass filter) directed to a CCD camera (LCL-902C, Watec, Orangeburg, NY).

Determination of constants for evaluating EC [Ca²⁺]_i.

[Ca²⁺]_i for EC tubes was calculated using the following equation: [Ca²⁺]_i (in nM) = K_d × β_{EC tube} × (R − R_{min,EC tube})/(R_{max,EC tube} − R) (22), where R is the observed F₃₄₀/F₃₈₀. During our attempts to titrate [Ca²⁺]_i in the presence of ionomycin, we found that EC tubes became leaky and lost fura-2 dye, as evidenced by the loss of fluorescence. However, we were able to determine R_min, R_max, and β values in EC tubes through appropriate timing. After being loaded with fura-2 AM dye, EC tubes were superfused with PSS containing 0 mM Ca²⁺ and 5 mM EGTA (Sigma) for 1 h to ascertain R_{min,EC tube}. The superfusion solution was then changed to one containing 3 μM ionomycin and 10 mM Ca²⁺, and R_{max,EC tube} was measured within the first min of the peak increase in fluorescence. β_{EC tube} was calculated from the ratio of minimum and maximum fluorescence values during excitation at 380 nm under the respective conditions of superfusion. This protocol was performed at both 24 and 32°C (n = 4 each) to obtain the respective values. Using the K_d values of fura-2 dye for Ca²⁺ determined in solution (above), R_{min,EC tube}, R_{max,EC tube}, and β_{EC tube} were used to estimate [Ca²⁺]_i in EC tubes.

Stability of resting [Ca²⁺]_i in EC tubes over time.

After the 30-min period of washout and temperature equilibration, resting baseline F₃₄₀/F₃₈₀ at time 0 was recorded. F₃₄₀/F₃₈₀ was then recorded every 15 min for 10 s at 250 Hz over a period of up to 3 h. For each 10-s recording, F₃₄₀/F₃₈₀ was averaged after subtracting the respective values of autofluorescence. Due to the duration of these measurements, a given EC tube was studied at one temperature only, at either 24, 32, or 37°C for this series of experiments.

Transient activation of muscarinic receptors.

A key goal of this study was to determine how the dynamic range of transient [Ca²⁺]_i responses in EC tubes to the activation of muscarinic receptors was influenced by temperature. Preliminary experiments established that a 200-μl bolus of ACh delivered from a pipette into the recording chamber at the site of superfusion inflow (i.e., just upstream from the EC tube) evoked the classic biphasic peak and plateau in [Ca²⁺]_i (10, 11, 49). Since the actual ACh concentration contacting the EC tube could not be determined, stimuli are presented as the molar amount (in μmol) of ACh delivered in the 200-μl bolus. An identical bolus of blue dye indicated that flow through the chamber was laminar, and time controls confirmed that [Ca²⁺]_i responses of EC tubes to a 200-μmol bolus of ACh were reproducible for up to 3 h at 24°C (data not shown).

For paired comparisons within each EC tube, [Ca²⁺]_i responses to incremental boluses of ACh were first recorded at 24°C and (after 30 min of reequilibration using 10-min increments as described above) again at 32°C. After each bolus of ACh, F₃₄₀/F₃₈₀ was allowed to return to baseline before the next bolus was delivered. As the fluorescence response to each bolus recovered completely within 1–3 min, <30 min was required to record responses to the full range of ACh stimuli at each temperature. Thus, including the 30-min period for temperature reequilibration, each EC tube was studied for a total duration of ≤ 1.5 h during this series of experiments. Ca²⁺ responses to ACh were not studied at 37°C due to the loss of EC tube integrity within 1 h (see results).

Recorded F₃₄₀/F₃₈₀ values were converted to [Ca²⁺]_i as described above. For each ACh stimulus, the total response was calculated as the area under the curve (AUC) of F₃₄₀/F₃₈₀ and [Ca²⁺]_i, with values for AUCs expressed in arbitrary units. The resting baseline value was taken as the average F₃₄₀/F₃₈₀ or [Ca²⁺]_i for 5 s before each ACh stimulus. To provide further insight with respect to how temperature affected [Ca²⁺]_i responses, the kinetics of transient responses to 20 μmol ACh were evaluated. These measurements (shown in Fig. 6A) included the following: time to peak, which was taken from when [Ca²⁺]_i deviated from resting baseline until it reached its highest value, and rate of rise to the peak response, which was the slope during this period. Peak duration was defined as the interval that the slope of the trace at its highest point was zero; peak [Ca²⁺]_i was the average [Ca²⁺]_i during this time. Plateau duration began when the recording stabilized after the initial peak and ended when F₃₄₀/F₃₈₀ declined to 50% of the peak ratio. The average [Ca²⁺]_i during this time was the plateau average [Ca²⁺]_i. Values for AUCs and response amplitudes are presented after the resting baseline F₃₄₀/F₃₈₀ or [Ca²⁺]_i was subtracted.

Fig. 6.

Kinetics of the [Ca²⁺]_i transient induced by ACh. A: representative [Ca²⁺]_i transient resulting from a 20-μmol bolus of ACh. Brackets indicate the initial peak (1) and secondary plateau (2) phases. B: summary data of the time to peak at both 24 and 32°C for males and females. *P < 0.05 vs. 24°C. C: summary data of the peak rate of rise at both 24 and 32°C for males and females. *P < 0.05 vs. 24°C. D: summary data of peak [Ca²⁺]_i at both 24 and 32°C for males and females. *P < 0.05 vs. 24°C. E: summary data of the peak duration at both 24 and 32°C for males and females. *P < 0.05 vs. 24°C. F: summary data of the plateau duration at 24 and 32°C for males and females. G: summary data for plateau average [Ca²⁺]_i at 24 and 32°C for males and females. *P < 0.05 vs. 24°C. There were no significant differences between males and females at either 24 or 32°C for any of the analyses performed (n = 6 mice/group). Actual values are shown in Table 1.

Statistical Analysis

One EC tube was studied per mouse. Data were analyzed using GraphPad Prism software (version 5.0, La Jolla, CA). Summary data are reported as means ± S.E. Differences in the AUCs between bolus-response curves were determined using two-way ANOVA. Peak and plateau Ca²⁺ dynamics were analyzed using one-way ANOVA. Post hoc comparisons were performed using Bonferroni tests. Differences were accepted as statistically significant with P < 0.05.

RESULTS

Morphological Integrity of EC Tubes

The morphological integrity of EC tubes was evaluated after 1 h of incubation at each temperature. As shown in Fig. 1A, for EC tubes held at 24°C, individual ECs exhibited an elongated fusiform shape while being well connected to neighboring cells on all sides. At 32°C, individual ECs remained connected to each other in the presence of slight shortening and rounding of nuclei (Fig. 1B). Thus, during incubation at 24 and 32°C, >90% of EC tubes maintained morphological integrity, which enabled us to perform reliable experiments over several hours. In contrast, during incubation at 37°C, individual ECs retracted from each other and from their holding pipettes, creating large gaps throughout the tube (Fig. 1C). None of the EC tubes remained suitable for further study after 1 h at 37°C.

Fig. 1.

Morphology of endothelial cell (EC) tubes maintained at 24, 32, and 37°C. A: an EC tube after 1 h at 24°C. ECs remained elongated and connected to each other in all directions. B: an EC tube after 1 h at 32°C. ECs remained connected to each other with modest cell shortening. C: an EC tube after 1 h at 37°C. Individual cells have retracted away from each other, and multiple rounded cells are present. Areas that appear out of the focal plane illustrate gaps that have opened between cells in the upper layer of the tube. Scale bars = 50 μm. Images were obtained using differential interference contrast.

Ca²⁺ Photometry

Determination of K_d and EC [Ca²⁺]_i at 24 and 32°C.

As shown in Fig. 2, plotting our Ca²⁺ calibration data as log{β[(R − R_min)/(R_max − R)]} versus log [Ca²⁺] enabled log K_d values to be determined from the x-axis intercept. These experiments yielded K_d values of 174 nM at 24°C and 146 nM at 32°C (n = 6). After determining the Ca²⁺ K_d value of fura-2 in our system, we measured the minimum and maximum F₃₄₀/F₃₈₀ values as well as the ratio of the maximum to minimum fluorescence signal of the 380-nm wavelength (β) in EC tubes (n = 4). These experiments provided values for calculating [Ca²⁺]_i at 24 and 32°C, respectively: R_{min,EC tube} = 0.600 and 0.618, R_{max,EC tube} = 11.244 and 10.198, and β_{EC tube} = 9.951 and 10.220 (22).

Resting [Ca²⁺]_i in feed artery EC tubes.

Resting levels of [Ca²⁺]_i in mouse feed artery ECs have not previously been determined. To obtain these values, we measured the fura-2 ratio in freshly isolated, unstimulated EC tubes from AFAs and used these measurements to calculate [Ca²⁺]_i. There were no significant differences in resting F₃₄₀/F₃₈₀ values between EC tubes obtained from male (n = 6) versus female (n = 6) mice at either 24°C (0.751 ± 0.02 vs. 0.772 ± 0.04) or 32°C (0.855 ± 0.02 vs. 0.812 ± 0.02). Similarly, resting [Ca²⁺]_i in EC tubes was not significantly different between sexes at 24°C (25 ± 4 vs. 29 ± 7 nM) or 32°C (38 ± 3 vs. 31 ± 3 nM). When data from males and females were pooled (n = 12), the F₃₄₀/F₃₈₀ value at 24°C (0.762 ± 0.02) was significantly lower than that at 32°C (0.833 ± 0.01, P < 0.05). However, resting [Ca²⁺]_i was not significantly different between temperatures (27 ± 4 nM at 24°C vs. 34 ± 2 nM at 32°C, P = 0.11).

Stability of resting [Ca²⁺]_i over time in EC tubes.

As shown in Fig. 3, resting [Ca²⁺]_i of EC tubes remained stable for at least 3 h while maintained at 24 or 32°C (n = 5 each). There were no differences in resting [Ca²⁺]_i between the respective temperatures. In contrast, for EC tubes studied at 37°C (n = 5), resting [Ca²⁺]_i increased rapidly such that the initial measurement after the 30-min equilibration (i.e., time 0 in Fig. 3) was already above values recorded at 24 or 32°C and remained significantly elevated thereafter. Reliable data could no longer be obtained after 90 min at 37°C due to loss of the fluorescence signal, which was attributed to dye extrusion. Therefore, to evaluate the effects of temperature on Ca²⁺ responses of EC tubes to ACh, subsequent experiments were performed at 24 and 32°C.

Fig. 3.

Resting [Ca²⁺]_i in EC tubes over time at 24, 32, and 37°C. Resting [Ca²⁺]_i was measured every 15 min for 3 h. At 24 and 32°C, [Ca²⁺]_i remained stable for the duration of these experiments. At 37°C, resting [Ca²⁺]_i increased rapidly and was significantly higher than that at 24°C within 15 min. After 90 min at 37°C, loss of the fluorescence signal indicated that cells had extruded the fura-2 dye, precluding accurate measurements beyond this time. *P < 0.05 vs. 24°C (n = 5 per group). Data are from EC tubes obtained from male mice and are presented as F₃₄₀/F₃₈₀ values after autofluorescence was subtracted.

[Ca²⁺]_i responses to transient activation of muscarinic receptors.

Representative traces of fura-2 fluorescence signals in response to ACh are shown in Fig. 4. For boluses of >0.02 μmol, F₃₄₀/F₃₈₀ increased with the amount of ACh delivered; responses recorded at 24°C (Fig. 4A) were consistently less than those recorded at 32°C (Fig. 4B). This difference between temperatures was maintained when F₃₄₀/F₃₈₀ values were converted to [Ca²⁺]_i (Fig. 4, C and D). Integrating the AUC of F₃₄₀/F₃₈₀ (Fig. 5A) and [Ca²⁺]_i (Fig. 5B) confirmed the greater [Ca²⁺]_i responses to ACh at 32 versus 24°C, with values for 2, 20, and 200 μmol ACh at 32°C essentially double the respective values recorded at 24°C (P < 0.05). There were no significant differences in F₃₄₀/F₃₈₀ or [Ca²⁺]_i responses to ACh stimuli between EC tubes obtained from male versus female mice at either 24 or 32°C (Fig. 5, C–F), indicating that the effects of ACh and temperature on [Ca²⁺]_i of EC tubes were independent of sex.

Fig. 4.

[Ca²⁺]_i transients of EC tubes in response to ACh. Representative recordings are shown from an EC tube (male) in response to progressive increases in ACh (0.02, 0.2, 2.0, 20, and 200 μmol, each delivered in a bolus of 200 μl). Time scale = 100 s for A–D. A: changes in F₃₄₀/F₃₈₀ at 24°C. B: changes in F₃₄₀/F₃₈₀ at 32°C. C: changes in [Ca²⁺]_i at 24°C. D: changes in [Ca²⁺]_i at 32°C. There were no detectible responses below 0.02 μmol ACh.

Fig. 5.

Integrated [Ca²⁺]_i responses to ACh in EC tubes. Left: calculated from F₃₄₀/F₃₈₀; right: calculated from [Ca²⁺]_i. A: area under the curve [AUC; in arbitrary units (AU²)] analysis calculated from F₃₄₀/F₃₈₀ values to ACh responses curves at 24 and 32°C. Data for EC tubes from males (n = 6) and females (n = 6) were pooled (n = 12). B: AUC analysis calculated from [Ca²⁺]_i to ACh responses curves at 24 and 32°C. Data for EC tubes from males (n = 6) and females (n = 6) were pooled (n = 12). *Responses at 32°C were significantly different from responses at 24°C (P < 0.05). C: AUC analysis calculated from F₃₄₀/F₃₈₀ values to ACh responses at 24°C comparing males (n = 6) and females (n = 6). D: AUC analysis calculated from [Ca²⁺]_i to ACh responses at 24°C comparing males (n = 6) and females (n = 6). E: AUC analysis calculated from F₃₄₀/F₃₈₀ values to ACh responses at 32°C comparing males (n = 6) and females (n = 6). F: AUC analysis calculated from [Ca²⁺]_i to ACh responses at 32°C comparing males (n = 6) and females (n = 6). There were no significant differences at any dose of ACh between males and females at either 24 or 32°C.

The initial phase of the biphasic Ca²⁺ transient (Fig. 6A, phase 1) reflects the internal release of Ca²⁺ from the ER (10, 49). In response to a standardized (20-μmol) bolus of ACh, increasing temperature from 24 to 32°C enhanced the kinetics of the initial Ca²⁺ transient in EC tubes. The time to peak (Fig. 6B) decreased (P < 0.05), the peak rate of rise (Fig. 6C) increased (P < 0.05), the peak [Ca²⁺]_i (Fig. 6D) increased (P < 0.05), and the peak duration (Fig. 6E) decreased (P < 0.05) at 32 versus 24°C. The second phase of the biphasic [Ca²⁺]_i transient (Fig. 6A, phase 2) reflects the influx of Ca²⁺ across the plasma membrane (10, 49). Whereas increasing temperature from 24 to 32°C had no significant effect on plateau duration (Fig. 6F), it more than doubled the plateau average [Ca²⁺]_i (P < 0.05; Fig. 6G). As with AUC values, there were no significant differences in [Ca²⁺]_i dynamics between EC tubes isolated from male versus female mice (Fig. 6, B–G; the respective values are shown in Table 1).

Table 1.

Kinetics of [Ca²⁺]_i transients in EC tubes from male and female C57BL/6 mice at 24 and 32°C

	24°C		32°C
	Males	Females	Males	Females
Time to peak, s	2.45 ± 0.017	2.17 ± 0.33	1.43 ± 0.18	0.95 ± 0.11
Ratio of rise, dratio/dt	0.36 ± 0.03	0.66 ± 0.18	1.34 ± 0.22	2.54 ± 0.41
Peak Ca2+, nM	203.3 ± 11.7	240.5 ± 22.4	463.1 ± 54.7	542.8 ± 54.0
Peak Duration, s	0.73 ± 0.08	0.55 ± 0.02	0.41 ± 0.05	0.23 ± 0.04
Plateau duration, s	49.87 ± 6.26	57.19 ± 8.87	59.30 ± 13.16	47.57 ± 5.26
Plateau Ca2+, nM	155.8 ± 3.8	176.9 ± 9.7	345.4 ± 27.2	417 ± 47.7

Values are means ± SE obtained from intracellular Ca²⁺ concentration ([Ca²⁺]_i) transients resulting from a 20-μmol bolus of ACh in endothelial cell (EC) tubes from male (n = 6) and female (n = 6) C57BL/6 mice.

DISCUSSION

For vessels controlling peripheral resistance and tissue blood flow, key signaling events governing endothelium-dependent vasodilation are triggered by elevations in EC [Ca²⁺]_i. However, little is known of Ca²⁺ signaling in the freshly isolated intact endothelium of resistance vessels, particularly in the absence of luminal blood flow or surrounding SMCs. In the present study, intact EC tubes of 1–2 mm long were freshly isolated from feed arteries of mouse abdominal skeletal muscle and secured in a chamber during continuous superfusion for several hours. We showed that transient, biphasic [Ca²⁺]_i responses to muscarinic receptor activation increase with the mass of ACh delivered and that [Ca²⁺]_i responses at 32°C were double those at 24°C. These relationships hold for EC tubes obtained from both male and female mice. Furthermore, although the morphological integrity of EC tubes was preserved for at least 3 h during experiments at 24 and 32°C, EC retraction at 37°C disrupted tube morphology within the first hour of incubation. Whereas the retraction of ECs coincides with a significant elevation in resting [Ca²⁺]_i and suggests Ca²⁺-dependent activation of EC contractile proteins, we cannot exclude the possibility that cell retraction triggered the increase in [Ca²⁺]_i that we observed. Beyond 90 min, loss of the fluorescence signal attributable to dye extrusion further precluded study at 37°C. Thus, compared with room temperature (24°C), studying EC tubes at 32°C more closely approximates typical physiological temperature (37°C) of these feed arteries and enhances the dynamic range of [Ca²⁺]_i signaling without compromising EC tube integrity.

EC Tubes as an Experimental Model

EC tubes provide a unique model for investigating the biophysical properties of the freshly isolated feed artery endothelium. The primary strength of this preparation is the ability to study signaling events that are intrinsic to native ECs of the resistance vasculature independent from the influence of blood flow or from surrounding SMCs, tissue parenchyma, and associated nerves. A previous study (11) of [Ca²⁺]_i in EC tubes was performed using preparations from hamster cremaster muscle arterioles to investigate the role of membrane potential during agonist-stimulated increases in [Ca²⁺]_i. However, those experiments relied on a single concentration of agonist applied continuously for several minutes and were performed entirely at room temperature. In the present study, the preparation of longer segments allowed EC tubes to be secured at each end during continuous superfusion, enabling experiments of longer duration to be performed reliably. In turn, our data demonstrate that [Ca²⁺]_i in EC tubes increases with the amount of ACh delivered (Fig. 4). Furthermore, the adaption of the EC tube preparation to mice opens the door for studying how altering the expression of specific proteins (e.g., through genetic manipulation) influences the biophysical properties of the native endothelium from resistance vessels that govern tissue blood flow in the intact system. In addition to Ca²⁺ responses after G protein-coupled receptor activation, this model may also prove useful in studying signaling events elicited by other agonists, in understanding the effects of manipulating ion channel activity and expression on endothelial function, and determining how such relationships may be altered in diseased states. A particularly promising area for future studies using this model entails the regulation of cell-to-cell communication along the microvascular endothelium (14).

An important limitation to the EC tube model results from the loss of myoendothelial junctions, which have been recognized as important microdomains for EC Ca²⁺ signaling (33). Furthermore, because SMCs are removed during trituration, EC tubes cannot be used to study endothelium-dependent regulation of vasomotor tone (7, 17, 19) or the influence of SMCs on EC [Ca²⁺]_i (16, 26, 52). Remarkably, EC retraction occurred when temperature was elevated above ambient (24°C). Although modest cell shortening was apparent at 32°C, cell retraction leading to intercellular gaps was readily apparent for EC tubes studied at 37°C (Fig. 1C) and coincided with significant increases in EC [Ca²⁺]_i (Fig. 3). The retraction of ECs may reflect the activation of contractile proteins regulated by the phosphorylation of myosin II regulatory light chains by myosin light chain kinase, a Ca²⁺/calmodulin-dependent enzyme (8, 25, 34). EC retraction may also result from a Ca²⁺-independent mechanism whereby myosin II serves as a substrate for p21-activated kinase 2 (9, 42). In ECs, Cdc-42 can activate p21-activated kinase 2, leading to the phosphorylation of myosin II regulatory light chains and EC contraction (50, 53). It is also possible that EC retraction triggered the increase in [Ca²⁺]_i. Although contractile proteins of ECs are integral to the regulation of pore formation and permeability (50), further study is needed to elucidate the mechanism by which cell retraction is activated at increased temperatures in EC tubes and whether retraction is dependent on the increase in [Ca²⁺]_i.

Estimation of [Ca²⁺]_i in EC Tubes

This study is the first to estimate [Ca²⁺]_i of freshly isolated, intact EC tubes isolated from mice. Our resting values for [Ca²⁺]_i of ∼30 nM are close to the value of 52 nM calculated for ECs within arterioles of the rat cremaster muscle (17). They are also in agreement with values reported for ECs cultured from swine coronary arteries (15), bovine aortae (43), and pulmonary arteries (37). Each of these studies estimated resting EC [Ca²⁺]_i values of ∼50 nM. In contrast, a study (45) using primary cultures of explanted mouse aortic ECs reported a resting [Ca²⁺]_i of 103 nM, which is closer to resting values obtained for ECs of isolated coronary arteries (29) and middle cerebral arteries (20) from rats (range: 90–174 nM). It should also be recognized that earlier studies used a predetermined value for the Ca²⁺ K_d value of Fura-2 [e.g., 282 nM (20, 29)], in contrast to determining K_d values under the conditions of their experiments, as done here. Where the effects of temperature have been studied on bradykinin-induced Ca²⁺ signaling in cultured ECs, the effective Ca²⁺ K_d value of fura-2 in solution decreased by ∼20 nM upon warming from 24 to 32°C (40), which is consistent with our present findings (Fig. 2). While our values for the Ca²⁺ K_d value of fura-2 were also determined using calibrated solutions of free [Ca²⁺], our estimates of [Ca²⁺]_i in EC tubes are based on actual measurements of R_{min,EC tube}, R_{max,EC tube}, and β_{EC tube} in our experimental preparation. Applying higher values for the Ca²⁺ K_d value of fura-2 would increase our estimates of EC [Ca²⁺]_i accordingly.

Brief (bolus) delivery of ACh was used to induce reproducible Ca²⁺ transients in EC tubes. ACh was selected as an agonist that acts via G protein-coupled receptors and that is commonly used to investigate endothelium-dependent relaxation of vascular SMCs in vivo and in vitro (4, 14, 17, 19, 33, 45, 46). The present data illustrate that both the initial peak and sustained plateau were evoked reproducibly with ACh boluses of >0.02 μmol (Fig. 4) and that both components increased dramatically when temperature was increased from 24 to 32°C (Figs. 4 and 6). It should be recognized that ACh-mediated signaling events cannot be studied in cultured ECs when muscarinic receptors are no longer expressed (5, 12, 48). However, the EC tube model presented here enables the use of ACh as a well-characterized agonist for investigating the nature of Ca²⁺ signaling in the intact endothelium freshly isolated from resistance vessels. Based on the present findings, a question for future studies concerns how the functional expression of muscarinic (or other G protein-coupled) receptors may be influenced by the temperature at which experiments are performed.

Effects of Temperature on Components of EC Ca²⁺ Signaling

Our calibrations with fura-2 dye enabled estimates of actual [Ca²⁺]_i through a range of stimulus intensities while addressing the temperature-dependence of ACh-induced Ca²⁺ transients. The binding of ACh to muscarinic receptors activates phospholipase C to generate 1,4,5-inositol trisphosphate (IP₃), which induces the rapid release of Ca²⁺ from IP₃ receptors of the ER, resulting in the initial peak phase of the ACh-induced Ca²⁺ transient (10, 11, 49). Increasing temperature from 24 to 32°C more than doubled the initial peak [Ca²⁺]_i (Fig. 6D) while significantly shortening the duration of this component of the fluorescence response, as reflected by the reduced time to peak (Fig. 6B) and accelerated peak rate of rise (Fig. 6C). These effects of warming may result from greater generation of IP₃, enhanced sensitivity of IP₃ receptors, and/or increased Ca²⁺ content within the ER. Because IP₃ receptors are sensitive to [Ca²⁺]_i, enhanced initial release of Ca²⁺ from the ER at warmer temperature could initiate a feedforward mechanism, further promoting Ca²⁺ release through IP₃ receptors until [Ca²⁺]_i attained sufficient levels to inhibit further release (18). Because negative feedback upon IP₃ receptors occurs at higher [Ca²⁺]_i [e.g., >500 nM (18)], such regulation would be most likely to have occurred in response to the highest ACh stimuli studied here (20 and 200 μmol; Fig. 3). In turn, the shortened peak duration at 32 versus 24°C (Fig. 6E) suggests that the sequestration of [Ca²⁺]_i by sarco(endo)plasmic reticulum Ca²⁺-ATPase (SERCA) is enhanced by warming, that Ca²⁺ extrusion from the cell by plasma membrane Ca²⁺-ATPase is accelerated, and/or that IP₃ dissociates more rapidly from its receptors.

The release of Ca²⁺ from the ER activates SOCE, which, together with RACE, accounts for the plateau phase of the [Ca²⁺]_i response to ACh (10, 11, 49). Whereas temperature had little effect on plateau duration (Fig. 6F), the estimated plateau [Ca²⁺]_i at 32°C was approximately twofold greater than the respective values at 24°C (Fig. 6G). This lack of change in plateau duration suggests that there may be little effect of temperature on SOCE and RACE signaling pathways. Alternatively (and more likely), a new balance may be achieved between Ca²⁺ release from the ER and its influx and reuptake via SERCA, such that the plateau duration is maintained at a higher level of [Ca²⁺]_i. Thus, the increase in Ca²⁺ entering via SOCE and RACE at 32 versus 24°C may coincide with greater Ca²⁺ release from the ER. With these relationships identified in the present study, additional experiments are needed to determine how temperature affects the respective components of [Ca²⁺]_i signaling dynamics in EC tubes.

Sex and EC Ca²⁺

In ECs of isolated mesenteric arterioles (21), middle cerebral arteries (20), and coronary arteries (29) as well as in freshly isolated valvular ECs (41) obtained from rats, resting [Ca²⁺]_i in females was consistently greater than corresponding values in males. Across these studies, this difference between sexes was attributed to the actions of estrogen in female animals. In the present study, we found no difference between EC tubes isolated from male versus female mice in either resting [Ca²⁺]_i or elevations in [Ca²⁺]_i due to ACh-induced Ca²⁺ transients (Figs. 4–6). This lack of a sex effect on EC Ca²⁺ may reflect a species difference between the present murine model and the rat models used previously. Moreover, sex differences in vascular function may also depend on the vascular bed (27), and ours is the first study to investigate whether sex differences are apparent in either resting [Ca²⁺]_i or responses to ACh in the endothelium of resistance vessels of skeletal muscle. For C57BL/6 mice, the present findings illustrate that EC tubes obtained from AFAs of either sex manifest similar behavior in Ca²⁺ responses to ACh at both 24 and 32°C. At the same time, our data do not exclude the possibility of sex differences in the physiology and/or pharmacology of EC tubes in response to other vasoactive agents or experimental interventions.

Summary and Conclusions

This study demonstrates the viability of EC tubes from murine resistance vessels for in vitro study for the first time. In so doing, we present a model for investigating [Ca²⁺]_i in the freshly isolated intact feed artery endothelium independent from the influence of blood flow (i.e., luminal shear stress), surrounding SMCs (e.g., myoendothelial coupling) or tissue parenchyma, each of which are known to influence [Ca²⁺]_i in ECs. When maintained at 24°C, ECs remain elongated and fusiform in appearance. Despite modest cell shortening at 32°C, ECs remain well attached to each other, and the integrity of EC tube morphology is well maintained. In contrast, when studied at 37°C, cell retraction is pronounced, and the structural integrity of EC tubes is lost within 1 h, coincident with a significant rise in [Ca²⁺]_i and loss of fluorescence, attributable to dye extrusion. The dynamic range of [Ca²⁺]_i responses to ACh effectively doubles at 32 versus 24°C. With no difference in behavior between EC tubes obtained from male versus female mice, we conclude that experiments performed at 32°C provide an acceptable balance between maintaining the structural and functional integrity of EC tubes and understanding their behavior at physiological temperatures.

GRANTS

This work was supported by National Institutes of Health (NIH) Grants R37-HL-041026 and R01-HL-086483. M. J. Socha was supported by NIH Training Grant T32-AR-048523.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).