AJP - Heart Fuel your research with LabChart
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 280: H1222-H1231, 2001;
0363-6135/01 $5.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (9)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Figueroa, X. F.
Right arrow Articles by Boric, M. P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Figueroa, X. F.
Right arrow Articles by Boric, M. P.
Vol. 280, Issue 3, H1222-H1231, March 2001

In vivo assessment of microvascular nitric oxide production and its relation with blood flow

X. F. Figueroa, A. D. Martínez, D. R. González, P. I. Jara, S. Ayala, and M. P. Boric

Unidad de Regulación Neurohumoral, Departamento de Ciencias Fisiológicas, Facultad de Ciencias Biológicas, Pontificia Universidad Católica de Chile, Santiago, Chile 6513492


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

To assess the hypothesis that microvascular nitric oxide (NO) is critical to maintain blood flow and solute exchange, we quantified NO production in the hamster cheek pouch in vivo, correlating it with vascular dynamics. Hamsters (100-120 g) were anesthetized and prepared for measurement of microvessel diameters by intravital microscopy, of plasma flow by isotopic sodium clearance, and of NO production by chemiluminescence. Analysis of endothelial NO synthase (eNOS) location by immunocytochemistry and subcellular fractionation revealed that eNOS was present in arterioles and venules and was 67 ± 7% membrane bound. Basal NO release was 60.1 ± 5.1 pM/min (n = 35), and plasma flow was 2.95 ± 0.27 µl/min (n = 29). Local NO synthase inhibition with 30 µM Nomega -nitro-L-arginine reduced NO production to 8.6 ± 2.6 pmol/min (-83 ± 5%, n = 9) and plasma flow to 1.95 ± 0.15 µl/min (-28 ± 12%, n = 17) within 30-45 min, in parallel with constriction of arterioles (9-14%) and venules (19-25%). The effects of Nomega -nitro-L-arginine (10-30 µM) were proportional to basal microvascular conductance (r = 0.7, P < 0.05) and fully prevented by 1 mM L-arginine. We conclude that in this tissue, NO production contributes to 35-50% of resting microvascular conductance and plasma-tissue exchange.

nitric oxide chemiluminescence; subcellular endothelial nitric oxide synthase distribution; Nomega -nitro-L-arginine; vascular conductance; hamster cheek pouch


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

BLOOD FLOW CONTROL is a dynamic process that depends on a balance of vasoconstrictor and vasodilator signals. Nitric oxide (NO) is an important vasodilator produced by endothelial cells in response to biochemical or mechanical stimuli (18). NO is generated from L-arginine (L-Arg) by the enzyme NO synthase (NOS), which is constitutive in endothelial cells (eNOS) (20). Many vasoactive substances depend on endothelial NO release to exert their vasodilatory action (18, 28). In addition, blood flow shear stress on endothelial cells stimulates tonic NO production participating in the control of vascular tone (7). Systemic NOS blockade with L-arginine analogs results in substantial blood pressure elevation, indicating that basal NO production contributes to reduce total vascular resistance (7, 26, 28). Furthermore, increased blood pressure is observed after targeted disruption of the eNOS gene (12).

Most evidence supporting eNOS involvement in vascular function was obtained in isolated vessels, or in whole animals, using pharmacological methods. However, the actual importance of microvascular NO production in the control of flow and solute exchange in vivo is unclear. The hamster cheek pouch is an experimental model allowing direct in vivo visualization of a microcirculatory bed with minimal surgical trauma. Several studies have used short-term local NOS inhibition to assess the participation of NO on vascular homeostasis in this tissue. For instance, there are conflicting reports on the role of NO in the regulation of venular permeability that NO mediates (15, 22) or does not mediate (9) the increase in macromolecular efflux induced by inflammatory agents. However, microvascular NO production has seldom been measured, and its importance in the maintenance of microvascular conductance and small solute exchange has not been characterized. In addition, the regional distribution of the eNOS source for NO release is not well established.

In cultured endothelial cells derived from large vessels, eNOS is largely found in the microsomal fraction, in an inhibitory association with caveolin (17), and it was proposed that cytosolic eNOS may represent the active enzyme pool (21). Therefore, it is important to explore the subcellular distribution of microvascular eNOS in an intact tissue, preserving the dynamic environment and the variety of stimuli acting on endothelial cells in vivo.

The aims of this study were 1) to characterize vascular and subcellular location of eNOS in the hamster cheek pouch microcirculation, 2) to directly quantify microvascular NO release in vivo, and 3) to characterize its importance on microvessel diameters and exchangeable flow. In a parallel study, we report the changes produced in these variables upon stimulation with acetylcholine, an endothelium-dependent vasodilator (Figueroa XF, González DR, Martinez AD, Ayala S, Duran WN, and Boric MP, unpublished observations).


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Animal and drug sources. Adult, 100- to 120-g male golden Syrian hamsters (Mesocricetus aureatus) were obtained from our university animal facilities. All studies were conducted following institutional and international guidelines for the welfare of animals in compliance with the Helsinki Declaration and the "Guiding Principles in the Care and Use of Laboratory Animals" endorsed by the American Physiological Society. Unless specified, all biochemical reagents and inhibitors were purchased from Sigma Chemical (St. Louis, MO) and chemicals of analytic grade were purchased from E. Merck, (Darmstaad, Germany). Monoclonal and polyclonal primary anti-human eNOS antibodies, and human endothelial cell standard lysate were purchased from Transduction Labs (Lexington, KY). Horseradish peroxidase-conjugated goat anti-rabbit and anti-mouse secondary antibodies were obtained from Pierce (Rockford, IL).

Preparation of the Cheek Pouch for Intravital Microscopy

The hamsters were anesthetized with pentobarbital sodium (60 mg/kg ip). The trachea, left carotid artery, and left jugular vein were cannulated, and the right cheek pouch was prepared for intravital microscopy as previously described (4, 6). A lucite plate and a fiber optic bundle were introduced through the mouth into the pouch to immobilize and to transilluminate the tissue. A skin incision was performed to expose the pouch, the avascular layer of connective tissue was cleared, and the observation chamber was placed on top of the pouch and secured to the skin. The hamster was placed on the stage of a Nikon Optiphot microscope and the cheek pouch was superfused with bicarbonate buffer of (in mM) 125 NaCl, 1.17 MgSO4, 2 CaCl2, and 20 NaHCO3 (pH 7.4, 37°C) equilibrated with 95% N2-5% CO2 at 1 ml/min. A peristaltic pump was used to move the superfusion solution into the observation chamber and out to a fraction collector. A glass cover slide was used to isolate the observation chamber from room air and to prevent optical disturbances. Drugs were applied topically without interrupting the superfusate flow, using a sideline near the observation chamber input, or dissolved in the superfusion medium when long application periods (>60 min) were needed.

Microvascular flow and conductance determinations. Once surgery was complete, a 45-min period was allowed for stabilization followed by an intravenous injection of a 0.2 ml saline bolus containing 2 × 106 counts/min of sodium-22 radioisotope (22NaCl, NEZ-081, New England Nuclear; Boston, MA), used as a highly diffusible tracer. Another 20-min period was allowed for equilibration of the radioactive tracer between the plasma and the extracellular compartment. The experiment was started by collecting the cheek pouch superfusate output every 2.5 min. Arterial carotid pressure was registered continually on a Grass polygraph. Duplicate 20-µl arterial blood samples were taken approximately once every hour. Radioactivity of superfusate and plasma samples was determined in a Wallac-Turku gamma counter, and clearance of 22Na was calculated by the [superfusate]-to-[plasma] radioactivity content ratio (4, 6). In addition, the ratio of sodium clearance divided by mean arterial pressure was calculated at every sampling period as an index of relative vascular conductance (RVC). We have reported that changes in RVC correlated closely with changes in microvessel diameters induced by vasodilator (3) and vasoconstrictor agents (4, 6).

Vessel diameters. The microcirculatory network was examined with a ×10-LWD Leitz objective. At given intervals before, during, and after the exposure to the different drugs, a few selected fields were recorded on videotape by using a projection lens, a TV camera, and a VHS recorder. Vessel diameters were measured with a video caliper (Texas A&M) during playback at a magnification of ×900, with an accuracy of ±0.5 µm. Arterioles and venules were classified according to their branching order, assuming the largest arteriole or venule to be of first order and increasing one order each time a vessel divided into two of similar size. In the present study, A4 refers to arterioles 5-15 µm in diameter; A3 refers to arterioles 16-30 µm in diameter; A2 refers to arterioles 31-45 µm in diameter; and A1 refers to arterioles 46-70 µm in diameter, respectively. In the case of the venules, V4 corresponds to vessels 12-25 µm in diameter, V3 refers to vessels 26-45 µm in diameter, and V2 refers to venules 46-65 µm diameter.

Determination of NO Production

In separate experiments, NO released to the superfusate was quantified with a NO analyzer (NOA, Sievers 280) that detects the specific chemiluminescence generated by the NO-ozone reaction with a threshold of 0.5-1.0 pmol. This method is the most sensitive and accurate for NO measurements available (1). However, for a more efficient determination of NO production in vivo, we also measured nitrite, its first oxidized product (5).

Use of a radioactive tracer was precluded in these experiments to avoid contaminating the equipment. Great care was taken in the preparation of buffers and sample handling. Only freshly obtained tridistilled water was used. No bubbles were allowed to enter the observation chamber during application of test substances, because they produced significant artifact readings. Samples were immediately sealed with parafilm to minimize exposure to room air and were measured within 6 h of collection (5). A T connector was placed in the inlet line, immediately before the observation chamber to obtain background buffer samples every 10 min. The reduction chamber of the NO analyzer was filled with 8 ml of glacial acetic acid containing 100 mg of potassium iodide at room temperature. Fifty microliters of each sample was injected to this chamber to rapidly reduce nitrites to NO. The equipment detected the chemiluminescence of the newly formed NO gas. Calibration of the equipment was performed daily using standards of 10-1,000 nM sodium nitrite. Background readings were subtracted. Results are expressed as the net concentration of NO plus nitrite (NO-nitrite) found in the superfusate output (pmol/ml), which equals NO production in the exposed tissue (pmol/min).

Identification of eNOS

Immunocytochemistry. The exposed cheek pouch was excised, submerged in embedding media (Histo-Prep), and immediately frozen in liquid nitrogen. Five-micrometer-thick tissue sections were obtained in a cryostat at -25°C and mounted on microscope slides using 1% silane A-174 as glue. Cuts were fixed with 70% ethanol at -20°C for 20 min and washed with Tris-saline 0.05 M, pH 7.3 (3× for 5 min), followed by 20 min incubation in 3% H2O2 in methanol to block endogenous peroxidases. Slices were washed; incubated in 5 mM EDTA, 1% fish gelatin, 0.05% Nonidet P-40, 1% IgG free-BSA, and 1% goat serum for 30 min at 4°C to saturate unspecific binding sites; and then incubated with rabbit polyclonal primary anti-eNOS antibody 1:200 overnight at 4°C. After being washed with Tris-saline buffer, the slices were incubated 1 h at room temperature with a anti-rabbit secondary peroxidase-conjugated antibody 1:100, washed 3× for 5 min, and developed by incubating 20 min at room temperature with 0.1% 3,3'-diaminobenzidine and 0.1% H2O2 in the dark. The slides were washed again and dehydrated in graded ethanol concentrations (50-100%) followed by xylol and were covered with a cover slide. Slices subjected to a similar procedure, but without primary antibody, were used as controls for the specificity of staining.

Western blot. The exposed cheek pouch area (80-100 mg) was excised and transferred to 500 µl of cold antiprotease-lysis buffer (1 µg/ml aprotinin, 1 mM benzamidine, 10 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, 200 µg/ml soybean trypsin inhibitor, 5 mM EGTA, 100 mM Tris, pH 7.4). The tissue was homogenized with an Ultraturrax for 15 s on ice. The homogenate was centrifuged at 10,000 g during 30 min at 4°C. The pellet was discarded, and the supernatant was ultracentrifuged at 100,000 g for 90 min (4°C) to separate the microsomal (pellet) and cytosolic (supernatant) fractions. The pellet was resuspended in 100 µl of 100 mM of Tris, pH 7.4 containing 1% SDS. After protein content was determined by the Bradford method, both fractions were separated by SDS-PAGE (7.5% gel, 70 µg per lane). Prestained molecular weight markers (Bio-Rad) and human endothelial cell lysate were used as standards and positive control, respectively. Proteins were blotted onto a nitrocellulose membrane (Life Technologies), which was blocked overnight with 5% nonfat milk in Tris pH 7.4 at 4°C. This was followed by incubation with a monoclonal primary anti-eNOS antibody 1:2,500 for 3 h at room temperature and 1 h of incubation with anti-mouse secondary antibody, and developed by a 15-min incubation with 0.01% 3,3'-diaminobenzidine, 0.5% H2O2 in the dark. Western blots were scanned and submitted to densitometric analysis using NIH image software. To estimate the proportion of eNOS present in the microsomal and cytosolic fractions, for each animal, the apparent concentration of the enzyme as determined by Western blot (densitometric units/µg protein) was multiplied by the corresponding total protein content of the fraction.

Experimental Protocols

Microvascular dynamics. A first set of experiments was performed to characterize the effect of inhibiting local endogenous NO synthesis on microvascular blood flow and vessel diameters. All experiments started with a 30-min baseline collection period.

To assess the effects of a short-term NOS inhibition, 10 µM Nomega -nitro-L-arginine (L-NNA) was applied for 30 min, followed by 60 min of drug washout. This L-NNA concentration was chosen based on the inhibitory efficacy of this analog as reported in other preparations (13, 19). The period of application was chosen based on our previous experience with vasoactive drugs and the response delays in our experimental system (3, 4, 6).

In a second series, we evaluated the effects of long-term eNOS inhibition, by applying 1, 10, or 30 µM L-NNA for 120 min. This protocol was used to explore the most effective L-NNA concentration.

The effects of L-arginine (L-Arg), the endogenous substrate for eNOS (20), were studied in a third set of experiments. First, we tested for the possible direct action 1 mM L-Arg, applied for 60 min. Second, we assessed whether L-Arg counteracted the effects of L-NNA. To do this, the cheek pouch was superfused with 1 mM L-Arg throughout the experiment, and either 10 µM (n = 7 cheek pouches) or 30 µM L-NNA (n = 5 cheek pouches) was applied for 30 min.

Direct determination of microvascular NO release. From the results of the previous experiments, three groups of hamsters were used to study NO release under control conditions and during eNOS inhibition. Ten hamsters were superfused for 30 min with normal buffer to measure baseline NO release. Thereafter, five hamsters were kept as time controls (90 min), whereas the other five hamsters were superfused with 30 µM L-NNA over 90 min. A third group of five hamsters was superfused with 1 mM L-Arg during the whole experiment (120 min), and 30 µM L-NNA was applied during 90 min after a 30-min baseline, as in the second group.

Analysis

Results are shown as means ± SE. Paired Student's t-test and two-way ANOVA were used to assess significance of variations along time within groups. Comparisons between groups were made using unpaired Student's t-test. Significance of differences was set at 0.05. To perform a regression analysis on the net effect of L-NNA inhibition on RVC, and to calculate absolute baseline NO release, we included data from similarly treated hamsters of a parallel study aimed to elucidate the effects of acetylcholine on NO production and eNOS distribution (Figueroa et al., unpublished observation). The added data correspond to basal and L-NNA application periods, before any stimulation with other drugs.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Identification and Distribution of eNOS

Immunocytochemical analysis revealed positive staining for eNOS in the endothelial layer of the hamster cheek pouch microvessels (Fig. 1). A clear and homogeneous staining was visible in arterioles and venules of all different branching orders but was less prominent in venular endothelium. Whereas postcapillary venules show a clear eNOS reaction, only occasionally was a faint labeling observed in capillary-sized vessels. No stain was detected in tissues not incubated with anti-eNOS antibody, confirming the specificity of labeling.


View larger version (100K):
[in this window]
[in a new window]
  		Fig. 1.   Immunocytochemical localization of endothelial nitric oxide synthase (eNOS) in hamster cheek pouch microvessels. A and D: consecutive serial frozen sections stained with hematoxiline and anti-eNOS antibody, respectively. Specific eNOS staining is observed in the endothelial lining of arterioles (A3, 16-30 µm in diameter and A4, 5-15 µm in diameter, solid arrows) and venules (V3, 26-45 µm in diameter and V2, 46-65 µm in diameter, open arrows), but it is absent in interstitial tissue despite the presence of cell nuclei. Calibration bar = 50 µm. B and C: higher magnification of the same partially constricted A3 (B, cross section) and A4 (C, oblique section) arterioles shown in D. Note the strong labeling of ridged arteriolar endothelium (thick arrow); in comparison, a faint positive reaction is detected in small vessels, likely to be capillaries (thin arrows). Bar = 20 µm. E: positive reaction in V3 and V2 venules (thin and thick open arrows, respectively). Bar = 20 µm. F: eNOS reaction in an A4 arteriole (oblique section), showing a prominent staining in the perinuclear region of an endothelial cell (thick arrow) Bar = 10 µm. G: continuous eNOS staining in the endothelium of a postcapillary venule (thin open arrow) merging a V3 venule (thick open arrow). Bar = 20 µm.

The presence of eNOS was further confirmed by Western blot analysis of cheek pouch homogenates (Fig. 2). Subcellular fraction and densitometric analysis of seven paired microsomal-cytosolic samples demonstrated that eNOS was about eight times more concentrated in the microsomal pool. After correcting by the total protein content of each homogenate (1.29 ± 0.17 mg of cytosolic fraction and 0.35 ± 0.04 mg of microsomal fraction), we calculated that 67 ± 7% of total detectable cheek pouch eNOS was membrane bound (Fig. 2).


View larger version (30K):
[in this window]
[in a new window]
  		Fig. 2.   Subcellular distribution of eNOS. Top: representative Western blot of cytosolic (Cyt) and microsomal (Mic) eNOS fractions of hamster cheek pouch homogenates [70 µg protein (prot.) per lane]. Human aortic endothelial cell (HAEC) lysate was used as a positive control (10 µg). Middle: relative eNOS concentration determined by densitometric analysis of Western blot. Bottom: total eNOS content, as determined from the protein content and band intensity of each sample. Error bars represent means ± SE.

Functional Relevance of Endogenous NO Production on Microvascular Blood Flow

Total exchangeable plasma flow of the superfused cheek pouch, as assessed by sodium clearance show small fluctuations in control conditions (Fig. 3). These fluctuations were likely due to changes in systemic perfusion pressure, because the relative vascular conductance (RVC) index was stable in the control period (Figs. 3 and 4), confirming previous reports (3, 4, 6). Topical application of 10 µM L-NNA over 30 min produced a slow and persistent decay in sodium clearance and RVC, without affecting systemic arterial pressure (Fig. 3). Whereas RVC reduction reached significance at 20 min of L-NNA application, sodium clearance was significantly reduced only after the drug superfusion period ended. Sodium clearance and RVC continued to decrease during the first 15 min of drug washout, reaching a 25-30% reduction compared with baseline, and slowly returned toward baseline in the next 60-75 min. However, even after 90 min of drug washout, RVC was not completely recovered (not shown). Two-way ANOVA confirmed significant variations in sodium clearance [F(41, 164) = 4.72, P < 1 × 10-10] and RVC [F(41, 164) = 4.62, P < 1 × 10-12], and no variations in mean arterial pressure [F(41, 164) = 0.44, not significant (NS)] during the experimental period.


View larger version (19K):
[in this window]
[in a new window]
  		Fig. 3.   Effect of local NOS inhibition on microvascular flow and vessel diameters shown by simultaneous determination of plasma flow (Na Clearance), relative vascular conductance (RVC), mean arterial pressure (MAP), and arteriolar and venular diameters in the hamster cheek pouch microcirculation (means ± SE). The number of vessels in each branching order is shown in parenthesis. Nomega -nitro-L-arginine (L-NNA) was applied in the superfusion medium during a 30-min period (horizontal bars). Shaded symbols denote significant differences relative to the average basal value (P < 0.05, paired t-test). Open symbols denote no significant difference from the average baseline value. A1, arterioles 46-70 µm in diameter; A2, arterioles 31-45 µm in diameter; V3, venules 12-25 µm in diameter.



View larger version (22K):
[in this window]
[in a new window]
  		Fig. 4.   Time and dose dependence of L-NNA on RVC. Top: RVC changes induced by sustained application of 10 µM or 30 µM L-NNA (horizontal arrows). Values are means ± SE. Shaded symbols show significant differences in respect to baseline (P < 0.05, paired t-test). Open symbols denote no significant difference from the average baseline value. Middle: average RVC from several experimental groups determined at baseline and after 45 min of superfusion with 10 or 30 µM L-NNA. (*P < 0.05 vs. basal, paired t-test). Bottom: correlation analysis between the change in RVC after 45 min of superfusion with 10 µM (n = 14) or 30 µM L-NNA (n = 17) and basal RVC, using data from the middle. A significant linear regression (P < 0.05) was found with either inhibitor concentration.

The reductions in sodium clearance and RVC were paralleled by constriction of arterioles and venules of every branching order (Fig. 3). In this particular experimental series, there was a transient constriction of A2 (-12.4 ± 4.4%) and A4 arterioles (-5.7 ± 3.6%) 2-5 min after the onset of L-NNA application, which may explain the tendency for a decrease in sodium clearance and RVC observed at that moment. Small arterioles (A4, A3) showed a faster constriction compared with A1 and A2, which presented a longer delay. All arteriolar orders were consistently constricted at the end of L-NNA application. The maximal constriction, expressed as percentage of control diameter was 14.4 ± 5.4% in A1, 8.5 ± 4.4% in A2, 10.8 ± 3.8% in A3, and 13.1 ± 3.2% in A4. Compared with arterioles, venules of all branching orders showed a proportionally stronger constriction. The maximal diameter reduction in venules was 20.0 ± 4.5% in V4, 18.6 ± 3.6% in V3, and 25.3 ± 6.7% in V2. The time course of venular constriction was similar to the changes in sodium clearance and RVC.

The same parallel pattern among sodium clearance, RVC, and vessel diameters was observed for all drug applications, and therefore, only RVC changes are depicted and used for analysis.

Sustained superfusion with 1 µM L-NNA produced no effect on RVC (n = 3, data not shown). In contrast, superfusion with 10 and 30 µM L-NNA produced a steady reduction of microvascular flow (Fig. 4) [10 µM, F(23, 138) = 5.29, P < 5 × 10-10; 30 µM, F(23, 138) = 9.50, P < 1 × 10-18]. The slope of RVC reduction was proportional to the concentration of inhibitor; however, in both cases, RVC stabilized at ~60% of baseline (after 60-70 min with 10 µM and 45-60 min with 30 µM).

The net reduction in RVC attained after 45 min of superfusion with 10 or 30 µM L-NNA was directly proportional to the initial RVC value. As expected, the slope for the regression curve was steeper with 30 µM L-NNA than with 10 µM L-NNA (Fig. 4). This result is consistent with the idea that tissues with higher NO production are more vasodilated, and therefore, they show a larger change after NOS blockade. However, it may be possible that variations in basal flow correspond to differences in tissue thickness and/or vessel density among hamsters. In that case, thicker or more vascularized tissues may present larger actual flow reductions at equal degrees of vasoconstriction. To correct for this possibility, an additional correlation analysis was performed between the fractional reduction in RVC induced by L-NNA (expressed as percentage of basal RVC) versus the absolute baseline RVC (in nl · min-1 · mmHg-1). This correlation was also significant (r = 0.385, n = 31 hamsters, P < 0.05), confirming that tissues with higher blood flow responded to L-NNA with a higher degree of vasoconstriction.

The vasocontractile effects of L-NNA were completely prevented when this inhibitor was applied in the presence of 1 mM L-Arg (Fig. 5). The sole superfusion with this concentration of eNOS substrate tended to increase RVC; however, the changes were significant only in one of three groups of hamsters. Topical L-Arg did not modify systemic arterial pressure in any group. A control 60-min application of 1 mM L-Arg did not produce significant changes in RVC [Fig. 5, top, F(47, 188) = 0.82, NS]. Whereas, a 120-min application not only prevented the effects of a 30-min application of 10 µM L-NNA but also resulted in a significant increment of RVC to 20 ± 6% above baseline [Fig. 5, bottom, F(45, 270) = 1.55, P < 0.039]. In a third group, also superfused with 1 mM L-Arg for 90 min, but challenged for 60 min with 30 µM L-NNA, RVC remained constant. RVC values (in nl · min-1 · mmHg-1) were 27.1 ± 6.7 before, and 27.2 ± 5.4, 24.2 ± 4.4, and 25.3 ± 5.0 after 30, 45, and 60 min of NOS blockade, respectively [n = 4 hamsters, F(35, 105) = 0.798, NS].


View larger version (25K):
[in this window]
[in a new window]
  		Fig. 5.   Prevention of L-NNA effects with arginine. Top: RVC (means ± SE) before, during, and after a 60-min superfusion with 1 mM of L-arginine (L-Arg) (thin bar). Bottom: a 30-min application of 10 µM L-NNA, similar to Fig. 3 (thick horizontal bar) was given during continuous superfusion with 1 mM L-Arg (thin bar). Shaded symbols indicate significant differences with respect to baseline (P < 0.05, paired t-test). Open symbols denote no significant difference from the average baseline value.

Direct measurement of microvascular NO production. The chemiluminescence technique allowed us to obtain reproducible and consistent readings of NO-nitrite content in the hamster cheek pouch superfusate at every sampling interval (Fig. 6). The net baseline microvascular NO production ranged between 30 and 100 pmol/min. In control preparations, average NO output remained around 50-60 pmol/min during the 100-min observation period but showed significant fluctuations [F(18, 72)= 2.34, P < 0.006]. Application of 30 µM L-NNA caused a strong reduction in NO superfusate content, which became significantly lower than baseline or time-matched controls after 25-30 min [Fig. 6, F(20, 80)= 6.11, P < 2 × 10-8]. By pooling different series, we reduced NO release to 8.6 ± 2.6 pmol/min after 45-70 min L-NNA application (n = 9 hamster cheek pouches). Similar to RVC determinations, the effect of 30 µM L-NNA on NO release was fully prevented by superfusion with 1 mM L-Arg [Fig. 6 bottom, F(18, 90)= 1.08 NS]. There was no statistical difference in basal NO release between control and L-Arg-treated tissues.


View larger version (21K):
[in this window]
[in a new window]
  		Fig. 6.   Nitric oxide (NO) production in the hamster microcirculation in vivo. Top: NO-nitrite released to cheek pouch superfusate under control conditions or during local inhibition of NOS with 30 µM L-NNA (horizontal bar). Bottom: 30 µM L-NNA was applied as above (thick horizontal bar), during continuous superfusion with 1 mM L-Arg (thin bar). Values are means ± SE. Shaded symbols indicate significant differences relative to baseline (P < 0.05, paired t-test). Open symbols denote significant difference from the average baseline value. *P < 0.05, significant difference compared with time-matched controls by unpaired t-test.

We calculated that locally produced NO largely accounts for the NO-nitrite recovered in the superfusate, whereas the contribution of circulating NO-nitrites is only ~7%. The basal NO release detected in all hamsters studied was 60.1 ± 5.1 pmol/min (n = 35 hamsters). Circulating plasma nitrite concentration was 1.4 ± 0.37 µM (n = 9). Assuming that sodium and nitrite have similar vascular distribution and permeability, we used the average baseline sodium clearance (2.95 ± 0.27 µl/min, n = 29 hamsters) to estimate that the maximal nitrite delivery from plasma to the interstitium is 4.13 pmol/min (2.95 µl/min × 1.4 µM, Fig. 7).


View larger version (30K):
[in this window]
[in a new window]
  		Fig. 7.   Schematic representation of NO production and release in the hamster cheek pouch microcirculation. Total plasma flow through the exposed cheek pouch area is ~3 µl/min, as estimated from plasma clearance of radioactive sodium. According to the circulating nitrite concentration, this plasma flow brings in ~4 pmol/min of nitrites. The total NO-nitrite content in the superfusate output, measured by chemiluminescence using mild reducing conditions, is ~60 pmol/min. Therefore, the local measurable production of NO amounts to ~ 56 pmol/min. The fraction of NO that is further oxidized into nitrates, for instance in the presence of hemoglobin in the blood compartment, is undetected.

To confirm the above assumption, cheek pouch nitrite clearance was actually determined in a few hamsters that infused intravenously with sodium nitrite to step-increase their plasma nitrite concentration 10-50 times above baseline. This maneuver did not affect systemic arterial pressure or microvessel diameters, but caused a rapid rise in superfusate nitrite content proportional to the change in plasma concentration. After baseline NO-nitrite release was subtracted, a nitrite clearance of ~3 pmol/min was determined (n = 4).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In this study, by the combined use of intravital microscopy with NO chemiluminescence, clearance of isotopic tracer, and NOS blockade, we quantified local microvascular NO production and characterized its relative importance on the control of vessel diameters and exchangeable flow in the hamster cheek pouch. We found that this tissue produces ~1 pmol NO · min-1 · mg wet wt-1. Microvascular NO production is directly correlated with vascular conductance and accounts for about one-third to one-half of resting blood flow. In addition, by using Western blot we found that under resting conditions in vivo microvascular eNOS is mainly membrane bound, although a relevant fraction (30-35%) is also found in the cytosolic compartment.

Microvascular Exchangeable Flow

We used sodium clearance as a measure of actual plasma flow through exchange vessels in the superfused tissue. This method allowed us to continuously assess microvascular flow with a good time resolution, based on the following considerations. First, because sodium permeability-surface product across the microvessel wall is much larger than blood flow, sodium extraction approaches 100% during a single passage through the exposed-superfused tissue. Second, back diffusion of the tracer into the blood compartment is negligible because the interstitium is constantly washed at a rate of 1 ml/min, which is two orders of magnitude larger than blood flow. Therefore, sodium clearance is mainly a flow-limited process equivalent to actual plasma flow. The 2.5-min sampling interval reasonably accounts for tracer turnover from plasma, through the interstitial and superfusion chamber compartments, as demonstrated by the fact that if microcirculation is suddenly arrested by killing the animal with intravenous KCl, sodium clearance decays exponentially with a half-time of 2.0-2.5 min. The half-time for washout of the superfusion chamber is 1.5 min; thus sodium washout from microvessels and the interstitial space takes ~1 min.

A more stable parameter, compared with sodium clearance, is obtained when fluctuations in arterial pressure are corrected by calculating RVC. We reported (4, 6) that microvessel diameters and RVC remain stable for up to 4 h in control buffer-superfused tissues.

Detection of NO

This study is the first to characterize NO production in the hamster microcirculation in vivo. We successfully adapted the chemiluminescence method to continuously detect NO release into the superfusate buffer. Because of the short half-life of authentic NO in biological media, an efficient assessment of NO production is met by measuring its oxidized products nitrites and/or nitrates (1). Chemiluminescence detection requires the reduction of these species back to NO. In this study, we used mild reducing conditions to reduce only nitrites to NO, because we have established that this procedure minimizes background noise and prevents interference from chemicals containing amino groups (5). Therefore, our readings represent the sum of NO-nitrite released directly toward the adventitia of microvessels, plus diffusion of NO and nitrite from the lumen to the interstitial space (Fig. 7). In our experimental conditions, the absence of oxygen in the superfusion buffer minimized NO oxidation to nitrate. Nevertheless, our measurement underestimates the total amount of NO produced because we discard the NO fraction further oxidized to nitrate in the tissue. After this work was submitted, similar levels of basal NO production in this tissue (~50 pmol/min) have been reported measuring superfusate chemiluminescence during continuous superfusion (8) or stop-flow conditions (16).

We discarded the idea that NO-nitrite produced in other tissues of the hamster significantly contribute to the amount detected in the cheek pouch superfusate because plasma NO-nitrite content was rather low. The assumption that total plasma-borne nitrite recovered in the superfusate is equal to the product of sodium clearance and plasma nitrite concentration is validated by the fact that nitrite and sodium clearances were similar (~3 µl/min). This analysis rules out the possibility that the variations in our NO measurements actually were the consequence of changes in microvascular flow.

The local enzymatic origin of the NO-nitrite found in the superfusate is further confirmed by the observation that topical L-NNA almost abrogated the measured NO signal (Fig. 6) whereas it reduced blood flow by only one-third to one-half of control (Figs. 3 and 4). It is reasonable to assume that most microvascular NO comes from endothelial cells as the product of eNOS activity. However, we cannot discard a possible contribution from other sources, like neuronal NOS reported in hamster microvascular smooth muscle cells (24). The use of specific antagonists of neuronal NOS may clarify this point.

Distribution of eNOS in the Hamster Cheek Pouch

By using cryostat and peroxidase-antiperoxidase immunocytochemistry techniques, we detected the presence of eNOS in the endothelium of arterioles and also in venules of all branching orders. These results agree with a recent report demonstrating the presence of eNOS in endothelium throughout the hamster vascular tree using similar methods (24) and with a study done in parallel using fixed hamster cheek pouch tissue (8). With the methods used, we did not clearly distinguish cheek pouch capillaries, precluding to definitively resolve the presence of eNOS in these vessels. Likewise, Segal et al. (24) did not find eNOS in cheek pouch capillaries, although they reported detection of eNOS in hamster skeletal muscle capillaries.

Our subcellular distribution analysis revealed that in vivo two-thirds of hamster cheek pouch eNOS is found in the microsomal fraction. This value is similar to that reported in freshly isolated endothelial cells (11) and is somehow lower than the estimated membrane-associated enzyme fraction reported in perfused rat lungs (23). Interestingly, the in vivo membrane-bound fraction of eNOS reported here is considerably lower than values reported for cultured endothelial cells (~85-90%), either naive (27) or transfected (25). As studied in cultured cells derived from large vessels, translocation of eNOS to the cytosol may represent activation of eNOS (10, 21). In this regard, the sizable fraction of cytosolic eNOS found in intact microvessels in vivo may reflect the effect of constant physiological stimuli favoring eNOS release from the membrane in the microcirculation. In support of this notion, we found that microsomal eNOS content is significantly reduced after stimulation with acetylcholine in the hamster cheek pouch in vivo (Figueroa et al., unpublished data).

Functional Relevance of Microvascular NO

Topical inhibition of NOS slowly reduced NO release (Fig. 6) in parallel with significant reductions in microvascular flow and arteriolar and venular diameter (Figs. 3 and 4). This was expected because we confirmed the presence of eNOS in venules and arterioles, and Bohlen (2) reported that both types of vessels produce NO in response to physiological stimulation. Compared with arterioles, venules showed a more intense constriction (Fig. 3) despite a less marked eNOS expression (Fig. 1). This enhanced venular constriction may involve active contraction and also passive effects due to a reduced blood volume entering the venules after an increase in upstream resistance. A similar finding was observed after application of vasoconstrictors such as endothelin (4), neuropeptide Y (NPY), and norepinephrine (6). In all these studies, the time course and magnitude of changes in RVC or sodium clearance correlated best with changes in diameter of large venules (V3-V2), likely reflecting total blood flow converge into these vessels.

We found a significant correlation between basal RVC and the net effect of NOS inhibition on either the absolute or fractional RVC values. A first interpretation of this correlation is that those tissues with higher blood flow present higher NO production and respond to L-NNA with a higher degree of vasoconstriction. Although we did not directly assess this assumption because we could not measure NO release and RVC simultaneously, the correlation on the results with L-NNA emphasizes the importance of local NO production in the maintenance of microvascular blood flow. Tissues with low RVC (<15 nl · min-1 · mmHg-1) did not constrict in response to L-NNA application, and low RVC levels were observed in most pouches after prolonged L-NNA application, when NO release was negligible. Furthermore, we found a very good match among the time course of reductions in NO production (Fig. 6), microvessel diameters, and RVC (Figs. 3 and 4) induced by NOS blockade with L-NNA.

Our results with L-Arg clearly show that the vasoconstrictor effect of L-NNA in the cheek pouch is due to inhibition of NO production. An important pharmacokinetic barrier for L-NNA action was evidenced because local application of NOS antagonists slowly (30-45 min) reduced NO production, and the recovery process of NOS blockade was even slower. Therefore, care has to be taken when interpreting results obtained during short exposures to this inhibitory agent.

The observation that L-Arg in some groups caused a significant rise in RVC may be related to the length of application, or to a lower basal RVC value (Fig. 5). In this way, L-Arg may induce dilation in the more contracted tissues, in a reciprocal relationship as established for L-NNA effects (Fig. 4). Nevertheless, because superfusion with 1 mM L-Arg did not cause a consistent increase in RVC (Fig. 5) and did not increase NO release (Fig. 6), it seems that substrate availability is not a step-limiting process for NO production in this tissue.

Considering all other flow-control mechanisms present in the living tissue, it is noteworthy that the sole inhibition of microvascular NO production elicits this major reduction in exchangeable flow. Because drugs were applied topically, the participation of systemic regulatory mechanisms is unlikely; however, we can assume that local myogenic and metabolic compensatory mechanisms were fully acting in our experimental preparation (14). In our experience, the hamster cheek pouch microvessels respond to vasoconstrictors, such as norepinephrine and NPY, with distal vasodilation and reactive hyperemia (6). In contrast, L-NNA-induced vasoconstriction was long lasting, affecting all branching orders without signs of downstream vasodilation, resembling the slowly developing and long-lasting vasoconstriction induced by endothelin (4). In addition, norepinephrine and NPY-induced vasoconstriction are stronger and more prolonged after NOS inhibition (5a, 6), further confirming the importance of microvascular NO production as the major factor contributing to blood flow control in this tissue.


    ACKNOWLEDGEMENTS

This study was funded in part by the Chilean National Fund for Science and Technology Fondo Nacional de Ciencia y Tecnología Grants 1971222 and 2990079 and Grant Líneas Complementarias 8990008.


    FOOTNOTES

Address for reprint requests and other correspondence: M. P. Boric, Depto. Ciencias Fisiológicas, P. Universidad Católica de Chile, Casilla 114-D, Santiago, Chile 6513492 (E-mail: mboric{at}bio.puc.cl).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 8 November 1999; accepted in final form 17 October 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Archer, S. Measurement of nitric oxide in biological models. FASEB J 7: 349-60, 1993[Abstract].

2.   Bohlen, HG. Mechanism of increased vessel wall nitric oxide concentrations during intestinal absorption. Am J Physiol Heart Circ Physiol 275: H542-H550, 1998[Abstract/Free Full Text].

3.   Boric, MP, and Albertini R. Atrial natriuretic peptide increases microvascular blood flow and macromolecular escape during renin infusion in the hamster. Microcirc Endothelium Lymphatics 6: 67-88, 1990[ISI][Medline].

4.   Boric, MP, Donoso V, Fournier A, St Pierre S, and Huidobro-Toro JP. Endothelin reduces microvascular blood flow by acting on arterioles and venules of the hamster cheek pouch. Eur J Pharmacol 190: 123-133, 1990[ISI][Medline].

5.   Boric, MP, Figueroa XF, Donoso MV, Paredes A, Poblete I, and Huidobro-Toro JP. Rise in endothelium-derived NO after stimulation of rat perivascular sympathetic mesenteric nerves. Am J Physiol Heart Circ Physiol 277: H1027-H1035, 1999[Abstract/Free Full Text].

5a.   Boric, MP, Martinez A, Ayala S, and Huiobro-Toro JP. Neuropeptide trysoine (NPY) potentiates microvascular hyperemia post nondrenaline (NA) in the hamster: involvement of NPY receptor subtypes and EDRF-NO (Abstract). Microcirculation 2: 84, 1995.

6.   Boric, MP, Martinez A, Donoso MV, and Huidobro-Toro JP. Neuropeptide Y is a vasoconstrictor and adrenergic modulator in the hamster microcirculation by acting on neuropeptide Y1 and Y2 receptors. Eur J Pharmacol 294: 391-401, 1995[ISI][Medline].

7.   Chu, A, Chambers DE, Lin CC, Kuehl WD, and Cobb FR. Nitric oxide modulates epicardial coronary basal vasomotor tone in awake dogs. Am J Physiol Heart Circ Physiol 258: H1250-H1254, 1990[Abstract/Free Full Text].

8.   Duran, WN, Seyama A, Yoshimura K, Gonzalez DR, Jara PI, Figueroa XF, and Boric MP. Stimulation of NO production and of eNOS phosphorylation in the microcirculation in vivo. Microvasc Res 60: 104-111, 2000[ISI][Medline].

9.   Feletou, M, Bonnardel E, and Canet E. Bradykinin and changes in microvascular permeability in the hamster cheek pouch: role of nitric oxide. Br J Pharmacol 118: 1371-1376, 1996[ISI][Medline].

10.   Goetz, RM, Thatte HS, Prabhakar P, Cho MR, Michel T, and Golan DE. Estradiol induces the calcium-dependent translocation of endothelial nitric oxide synthase. Proc Natl Acad Sci USA 96: 2788-2793, 1999[Abstract/Free Full Text].

11.   Hecker, M, Mulsch A, Bassenge E, Forstermann U, and Busse R. Subcellular localization and characterization of nitric oxide synthase(s) in endothelial cells: physiological implications. Biochem J 299: 247-252, 1994.

12.   Huang, PL, Huang Z, Mashimo H, Bloch KD, Moskowitz MA, Bevan JA, and Fishman MC. Hypertension in mice lacking the gene for endothelial nitric oxide synthase. Nature 377: 239-242, 1995[Medline].

13.   Ishii, K, Chang B, Kerwin JF, Jr, Huang ZJ, and Murad F. Nomega -nitro-L-arginine: a potent inhibitor of endothelium-derived relaxing factor formation. Eur J Pharmacol 176: 219-223, 1990[ISI][Medline].

14.   Johnson, PC, and Smiesko V. Overview of regulatory mechanisms in the microcirculation. In: Flow-Dependent Regulation of Vascular Function, edited by Bevan JA, Kaley G, and Rubanyi GM.. New York: Oxford University Press, 1995, p. 261-273.

15.   Mayhan, WG. Nitric oxide accounts for histamine-induced increases in macromolecular extravasation. Am J Physiol Heart Circ Physiol 266: H2369-H2373, 1994[Abstract/Free Full Text].

16.   Mayhan, WG, Sharpe GM, and Anding P. Agonist-induced release of nitric oxide during acute exposure to nicotine. Life Sci 65: 1829-1837, 1999[ISI][Medline].

17.   Michel, JB, Feron O, Sacks D, and Michel T. Reciprocal regulation of endothelial nitric-oxide synthase by Ca2+- calmodulin and caveolin. J Biol Chem 272: 15583-15586, 1997[Abstract/Free Full Text].

18.   Moncada, S, Palmer RM, and Higgs EA. Nitric oxide: physiology, pathophysiology, and pharmacology. Pharmacol Rev 43: 109-142, 1991[ISI][Medline].

19.   Moore, PK, al-Swayeh OA, Chong NW, Evans RA, and Gibson A. L-NG-nitro arginine (L-NOARG), a novel, L-arginine-reversible inhibitor of endothelium-dependent vasodilatation in vitro. Br J Pharmacol 99: 408-412, 1990[ISI][Medline].

20.   Palmer, RM, Ashton DS, and Moncada S. Vascular endothelial cells synthesize nitric oxide from L-arginine. Nature 333: 664-666, 1988[Medline].

21.   Prabhakar, P, Thatte HS, Goetz RM, Cho MR, Golan DE, and Michel T. Receptor-regulated translocation of endothelial nitric-oxide synthase. J Biol Chem 273: 27383-27388, 1998[Abstract/Free Full Text].

22.   Ramirez, MM, Quardt SM, Kim D, Oshiro H, Minnicozzi M, and Duran WN. Platelet activating factor modulates microvascular permeability through nitric oxide synthesis. Microvasc Res 50: 223-234, 1995[ISI][Medline].

23.   Rizzo, V, McIntosh DP, Oh P, and Schnitzer JE. In situ flow activates endothelial nitric oxide synthase in luminal caveolae of endothelium with rapid caveolin dissociation and calmodulin association. J Biol Chem 273: 34724-34729, 1998[Abstract/Free Full Text].

24.   Segal, SS, Brett SE, and Sessa WC. Codistribution of NOS and caveolin throughout peripheral vasculature and skeletal muscle of hamsters. Am J Physiol Heart Circ Physiol 277: H1167-H1177, 1999[Abstract/Free Full Text].

25.   Sowa, G, Liu J, Papapetropoulos A, Rex-Haffner M, Hughes TE, and Sessa WC. Trafficking of endothelial nitric-oxide synthase in living cells. Quantitative evidence supporting the role of palmitoylation as a kinetic trapping mechanism limiting membrane diffusion. J Biol Chem 274: 22524-22531, 1999[Abstract/Free Full Text].

26.   Tolins, JP, Palmer RM, Moncada S, and Raij L. Role of endothelium-derived relaxing factor in regulation of renal hemodynamic responses. Am J Physiol Heart Circ Physiol 258: H655-H662, 1990[Abstract/Free Full Text].

27.   Venema, VJ, Zou R, Ju H, Marrero MB, and Venema RC. Caveolin-1 detergent solubility and association with endothelial nitric oxide synthase is modulated by tyrosine phosphorylation. Biochem Biophys Res Commun 236: 155-161, 1997[ISI][Medline].

28.   Whittle, BJ, Lopez-Belmonte J, and Rees DD. Modulation of the vasodepressor actions of acetylcholine, bradykinin, substance P and endothelin in the rat by a specific inhibitor of nitric oxide formation. Br J Pharmacol 98: 646-652, 1989[ISI][Medline].

Am J Physiol Heart Circ Physiol 280(3):H1222-H1231
0363-6135/01 $5.00 Copyright © 2001 the American Physiological Society


This article has been cited by other articles:


Home page
 J. Appl. Physiol.Home page
M. Kavdia and A. S. Popel
Contribution of nNOS- and eNOS-derived NO to microvascular smooth muscle NO exposure
J Appl Physiol, July 1, 2004; 97(1): 293 - 301.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
M. Kavdia, N. M. Tsoukias, and A. S. Popel
Model of nitric oxide diffusion in an arteriole: impact of hemoglobin-based blood substitutes
Am J Physiol Heart Circ Physiol, June 1, 2002; 282(6): H2245 - H2253.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
M. Kavdia, N. M. Tsoukias, and A. S. Popel
Model of nitric oxide diffusion in an arteriole: impact of hemoglobin-based blood substitutes
Am J Physiol Heart Circ Physiol, June 1, 2002; 282(6): H2245 - H2253.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (9)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Figueroa, X. F.
Right arrow Articles by Boric, M. P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Figueroa, X. F.
Right arrow Articles by Boric, M. P.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online