AJP - Heart BIOPAC complete lab solutions
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 278: H658-H665, 2000;
0363-6135/00 $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 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 Web of Science (7)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Fulton, D.
Right arrow Articles by Sessa, W. C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Fulton, D.
Right arrow Articles by Sessa, W. C.
Vol. 278, Issue 2, H658-H665, February 2000

SPECIAL COMMUNICATION
Quantification of eNOS mRNA in the canine cardiac vasculature by competitive PCR

David Fulton1, Andreas Papapetropoulos1, Xiaoping Zhang2, John D. Catravas3, Thomas H. Hintze2, and William C. Sessa1

1 Department of Pharmacology, Boyer Center for Molecular Medicine, Yale University School of Medicine, New Haven, Connecticut 06536-0812; 2 Department of Physiology, New York Medical College, Valhalla, New York 10595; and 3 Vascular Biology Center and Department of Pharmacology and Toxicology, Medical College of Georgia, Augusta, Georgia 30912-2500


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The goal of the present study was to develop a competitive PCR assay to measure changes in the expression of endothelial nitric oxide synthase (eNOS) mRNA levels throughout the canine vascular tree. A partial sequence of canine eNOS cDNA (1.86 kb), inducible NOS (1.95 kb), and neuronal NOS (1.16 kb) was cultured from canine aortic endothelial cells, LPS-treated canine splenic vein endothelial cells, and from canine left ventricle, respectively. Competitor eNOS cDNA (eNOS-C) was constructed via recombinant PCR. Thus, with the use of a standard curve competitive PCR with eNOS-C, the amount of eNOS mRNA in 500 ng of total RNA was greatest in the circumflex > right coronary artery > left anterior descending coronary artery > aorta. The isolation of coronary microvessels from the left ventricle was associated with an enrichment of endothelial cell markers such as eNOS, von Willebrand factor, and caveolin-1, an observation supported by the detection of up to 15-fold higher levels of eNOS mRNA in coronary microvessels relative to the larger arteries. The ability to quantify changes in eNOS mRNA levels throughout the canine vasculature should provide greater insight into the molecular mechanisms of how this gene is regulated in physiological and pathophysiological states.

nitric oxide; nitric oxide synthase; microvessels; complimentary deoxyribonucleic acid; endothelium


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

NITRIC OXIDE SYNTHASES (NOS) comprise a family of three distinct genes, neuronal (nNOS), inducible (iNOS), and endothelial (eNOS) isoforms that are expressed in various tissues in the nervous, immune, and cardiovascular systems (16). In the cardiovascular system, the abnormal production or bioactivity of endothelium-derived nitric oxide (NO) is recognized as a common feature of diseases associated with endothelial dysfunction. Attenuated endothelium-dependent relaxation has been reported in human and in animal models of heart failure, diabetes, and hypertension (2, 3, 13, 14, 21, 22, 30). Conversely, endothelial function is enhanced in conditions of high flow (18) and exercise (24). The enzymatic production of NO, which accounts at least in part for the modification of endothelial function, is a highly regulated event. The major trigger for NO production is the elevation of intracellular calcium and the binding of calcium-calmodulin to eNOS (12). In addition, subcellular localization, fatty acid acylation (15, 23, 25), serine (17) and tyrosine phosphorylation (6), and the binding of accessory proteins such as caveolin (4, 7) and heat shock protein 90 (5) can all influence NO release. However, NO production in vivo can also be chronically regulated through changes in the expression of eNOS. Although it has often been categorized as a constitutively expressed gene, eNOS mRNA and protein levels have been shown to be reduced in heart failure (27) and augmented in exercise (24) and conditions of high flow (19), suggesting transcriptional or posttranscriptional regulation of expression.

The use of canine models to understand cardiac physiology and pathophysiology is underscored by the similar physiology of the canine cardiovascular system with that of the human. Of the many genes involved in coordinating vascular function, eNOS-derived NO plays a central role in the control of vascular resistance and thus blood flow (8, 26). Previous studies in canine cardiovascular models have demonstrated that aortic expression of eNOS mRNA and protein can be up- or downregulated and that changes in expression correlate with changes in vascular function (24, 27). However, little is known about the molecular control of NO synthases in the cardiac vasculature. The similarity of canine NOS isoforms to those cloned for other species has yet to be thoroughly characterized, and the limited sensitivity of Northern blot analysis has thus far constrained the understanding of eNOS regulation to isolated aortic endothelial cells.

Therefore, the objective of the current study was to determine the cDNA sequences of the canine NOS isoforms and to develop a competitive PCR assay to specifically and accurately measure eNOS mRNA expression levels across the full spectrum of cardiac vessels. A major application of this technique was to quantitate eNOS gene expression in the canine coronary microvasculature, the site of coronary resistance and blood flow control.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cloning of NO Isoforms

eNOS. Canine eNOS cDNA clones were obtained from RT-PCR amplification of total RNA from cultured aortic endothelial cells (AEC). Canine AECs were cultured in medium 199 containing 10% (vol/vol) iron-supplemented FCS, 5% (vol/vol) FCS, 100 IU/ml penicillin, and 100 µg/ml streptomycin. Confluent AECs were washed with PBS, and total RNA was extracted with TRIZOL (Life Technologies, Rockville, MD). The RNA pellet was resuspended in diethyl pyrocarbonate-treated H2O and was quantitated spectrophotometrically by absorbance at lambda  = 260 nM. cDNA was synthesized from total RNA (1 µg). Total RNA was incubated with random hexamers (2.5 µM; Applied Biosystems) and RNase inhibitor (20 units; Boehringer Mannheim) for 10 min at 70°C and was quickly chilled on ice. Random hexamers were allowed to anneal at 25°C for 10 min before the addition of reverse transcriptase (200 units Superscript II; Life Technologies), first-strand buffer (50 mM Tris · HCl, 75 mM KCl, 5 mM MgCl2), dNTP (0.5 mM), and dithiothreitol (10 mM). RT was carried out at 42°C for 50 min, followed by inactivation of the RT enzyme at 70°C for 15 min. Ten percent of the RT reaction was used for PCR. Putative canine eNOS primers (set A: sense, 5'-ATGTTCAACTACATCTGCAAC-3'; antisense, 5'-TTCCACAGGGAC GAGGTGGTC-3') were designed from the conserved regions of human (M95296), bovine (M95674), and mouse eNOS (U53142) cDNAs (nucleotides 618-1184 of human eNOS cDNA). The PCR reaction mixture contained 2.5 units of Taq polymerase (Boerhringer Mannheim), 0.2 µM primers, 50 mM KCl, 10 mM Tris · HCl, and 1.5 mM MgCl2. cDNA was amplified for 35 cycles under the following conditions, denaturation at 94°C (1 min), annealing at 55°C (1 min), and extension at 72°C (1 min). PCR products were separated by electrophoresis on a 1% agarose gel (0.5 µg/ml ethidium bromide) in 1× 0.040 M Tris-acetate and 0.001 M EDTA and were visualized on an ultraviolet transilluminator. The resultant PCR product (566 bp) was cloned directly into PCR II (TA cloning vector; Invitrogen, Carlsbad, CA) and sequenced on both strands with an automated fluorescent sequencer. Based on the known partial sequence of canine eNOS, successive clones were obtained via primer walking in both 5' and 3' directions.

iNOS. Expression of iNOS mRNA was initiated by the administration of lipopolysaccharide (LPS; 1 µg/ml for 48 h) to cultured canine splenic vein endothelial cells. PCR primers for canine iNOS were designed based on regions of homology with human iNOS (accession number, L24553) and were positioned to span the corresponding region of the eNOS clone (equivalent, based on alignment to nucleotides 593-2538 and 443-2559 of human iNOS and eNOS, respectively; sense, 5'-CAGGGTGGAAGCGGT AACAAA-3' and antisense, 5'-GGTGAGGGCCTGGCTGAG TGA-3'). Total RNA from cultured splenic endothelial cells was isolated, and 1 µg was reverse transcribed as described above. Amplification of cDNA by PCR (94°C melting, 55°C annealing, and 72°C extension for 2 min) produced an approx 2-kb band that was cloned into PCR II. Canine iNOS was sequenced as described above.

nNOS. A partial nNOS clone was derived from the myocardium of a male mongrel dog. Approximately 100 mg of left ventricle were snap-frozen in liquid nitrogen and pulverized with a mortar and pestle, and fragments were homogenized in 1 ml of TRIZOL. Total RNA was extracted and reverse transcribed as described above. Canine nNOS primers were selected as described previously for iNOS [equivalent, based on alignment to nucleotides 1418-2574 and 1-1400 of human nNOS (U17327) and eNOS, respectively; sense, 5'-GGAATCC AGGTGGACAGAG-3' and antisense, 5'-TAGTTGAGCATCTCCTGGTGG-3']. PCR conditions were identical to those described for iNOS. A single amplification product of 1.4 kb was cloned into PCR II and sequenced as described above.

PCR Primers

On the basis of the known sequence of canine eNOS, primer design (Table 1) was optimized (DNASTAR) to enhance stability, reduce both primer-dimers and hairpins, and also to span introns. Extrapolating from the genomic sequence of human eNOS, the sense primer for eNOS has a complementary sequence on exon 6 and the antisense primer on exon 9. Primers for iNOS and nNOS were also designed based on the known sequence of canine iNOS and nNOS clones as described above. von Willebrand factor (vWF), caveolin-1 (Cav-1), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primers were designed based on published sequences (AF099154, U47060, and J04038, respectively).

                              
View this table:
[in this window]
[in a new window]
  		Table 1.   Primer sequences for PCR amplification of canine cDNAs

The specificity of primers for eNOS, iNOS, and nNOS was determined by PCR. eNOS primers (0.2 µM; Table 1) were incubated with 200 pg of canine eNOS cDNA template, 200 pg of canine iNOS, and 200 pg of canine nNOS and were amplified by PCR as described above under the following conditions for 35 cycles: 94°C denaturing (45 s), 61°C annealing (45 s), and 72°C extension (45 s). Specific primers for iNOS and nNOS (Table 1) were also tested against respective templates and eNOS cDNA.

Construction of Competitor from eNOS cDNA

To quantify eNOS mRNA levels, a competitor (eNOS-C) with an internal deletion and identical, competing flanking sequences was constructed. Two primers were designed (sense, 5'-CCAGAACTCTTTGCTTGAGCAACATGCTGC-3'; antisense, 5'-GCAGCA TGTTGCTCAAGCAAAGAGTTCTGG-3'), each composed of two 15 nucleotides domains. The first 15 nucleotides of each primer were the reverse complement of the last 15 nucleotides of the other, and the remaining 15 nucleotides of each primer correspond to regions of canine eNOS, separated by 94 bp (Fig. 1A). PCR was performed as described above for 30 cycles using 5 ng of canine eNOS cDNA as a template and the above primers (0.2 µM) with reciprocal primers for eNOS (set A). The resultant products were purified on a 1% agarose gel, extracted (Qiagen, Valencia, CA), and blunt ended to remove terminal adenines. Ten percent (10 µl) of the PCR reaction was combined with 2 µl dNTP, 5 mM MgCl2, and 1 unit of T4 polymerase (Boehringher Mannheim) and was incubated for 15 min at 12°C. The reaction mixture was heated to 95°C for 5 min to inactivate the polymerase and was purified through a spin column (Qiagen). Each fragment was diluted 1:100 and was combined with 0.2 µM of primer set A. PCR was performed as described above, and the resultant fragment (Fig. 2, B-C) was purified on an agarose gel, extracted, cloned into PCR II, and sequenced.


View larger version (18K):
[in this window]
[in a new window]
  		Fig. 1.   Generation of the endothelial nitric oxide synthase (eNOS) competitor. A: complementary primers were designed spanning a gap of 79 bp, and separate PCR reactions were conducted using both sense and antisense primers. Resultant PCR products are combined (B), and PCR amplification was conducted again using sense and antisense primers, resulting in the generation of an eNOS fragment containing a 79-bp deletion (C).



View larger version (39K):
[in this window]
[in a new window]
  		Fig. 2.   Isolation of nitric oxide synthase (NOS) isoform cDNAs and characterization of gene expression in canine heart and blood vessels. As seen in A, eNOS specific primers recognize only canine eNOS and not inducible NOS (iNOS) or neuronal NOS (nNOS) cDNAs. Primers for eNOS (0.2 µM) were incubated with cDNA (200 pg) encoding eNOS, iNOS, and nNOS and were amplified by PCR for 35 cycles. Primers for iNOS and nNOS (0.2 µM) were also tested against respective templates (n = 2). B: expression of endothelial cell markers, eNOS, von Willebrand factor (vWF), and caveolin-1 (Cav-1) were enriched relative to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and ribosomal RNAs during the isolation of coronary microvessels. Slight enrichment of iNOS and nNOS is found in microvessels. Total RNA (1 µg) from the left ventricle, a mixture of microvessels and myocytes, and purified microvessels representing the start, midpoint, and end of the isolation procedure was reverse transcribed and amplified by PCR for 25 (Cav-1, vWF, and GAPDH) and 35 (eNOS, iNOS, and nNOS) cycles. Results are representative of 2 separate experiments.

To control for reverse transcription, a cRNA competitor was synthesized. eNOS-C cRNA was prepared using a T7 in vitro transcription system (MEGAscript; Ambion, Austin, TX). eNOS-C cDNA (5 µg) was linearized overnight with Hind III (20 units) and was purified on an agarose gel as described above. Linearized plasmid (1 µg) was incubated with ribonucleotides and T7 polymerase for 6 h at 37°C, and cRNA was synthesized as described by the manufacturer. The concentration of cRNA was determined spectrophotometrically by absorbance at lambda  = 260 nm, and analysis by agarose electrophoresis revealed a single band. cRNA was stored at -80°C.

For accurate quantitation, the internal standard must compete in a linear relationship with the native eNOS cDNA. Therefore, a standard curve was generated by incubating decreasing concentrations of eNOS cDNA (1.3-166 amol) with a fixed concentration of eNOS-C cDNA (21 amol). PCR analysis was performed as described previously, using the eNOS primers in Table 1 under the following conditions for 35 cycles: 45 s at 94°C denaturing, 61°C annealing, and 72°C extension. Conversely, decreasing concentrations of eNOS-C (1.3-332 amol) were competed against a fixed concentration of eNOS (21 amol) cDNA. To examine sample mRNA levels, linear competition must also exist between full-length eNOS mRNA and the truncated eNOS-C. Decreasing amounts of AEC total RNA (2-0.0625 g) were incubated with a fixed quantity of cRNA (10 pg). The amount of eNOS mRNA per mass of total RNA was derived from comparison with a standard curve generated from the competition between decreasing concentrations of eNOS cDNA and a fixed concentration of cRNA (10 pg). To standardize the addition of competitor between samples and the standard curve, cRNA was added to the reverse transcriptase master mix. Reverse transcription was performed as described above, except for the inclusion of 2.5% acetylated BSA. PCR products were resolved on a 1.2% agarose gel as described above, photographed, and digitized with an HP scanner. Respective bands of eNOS cDNA and eNOS-C were analyzed with National Institutes of Health image software (1.61), and peak areas were quantitated by integration.

Purification of Microvessels

Four mongrel dogs (body wt 20-25 kg) were anesthetized with pentobarbital sodium (50 mg/kg), and a thoracotomy was performed. The heart and a segment (3-5 cm) of aorta were excised and immersed in ice-cold PBS (0.1% BSA, pH 7.4). The circumflex, left anterior descending (LAD), and right coronary artery (RCA) were dissected free from the heart and together with the aorta were cleaned of periadventitial fat and superficial connective tissue. The vessels were snap-frozen in liquid nitrogen and were stored at -80°C until used. Coronary microvessels were prepared from the left ventricle as previously described (11, 31). To determine the effect of the purification procedure on RNA integrity, total RNA was extracted from the left ventricle, crude microvessel extract with associated myocytes, and enriched coronary microvessels, representing the start, midpoint, and conclusion of the procedure. The piece of ventricle, crude microvessel extract, and terminal microvessels were snap-frozen in liquid nitrogen. Frozen blood vessels and myocardium were pulverized in a mortar and pestle, immersed in TRIZOL, and homogenized with a Polytron. RNA (1 µg) was reverse transcribed, and 10% of the resultant cDNA was amplified by PCR using the primers listed in Table 1. Denaturation was at 95°C, annealing at 61°C, and extension at 72°C for a total for 45 s each.

Quantitation of eNOS mRNA in Canine Blood Vessels

Total RNA (500 ng) from aorta, circumflex, LAD, RCA, and microvessels was incubated with a fixed amount of eNOS-C cRNA (50 fg, 0.2 amol) and reverse transcribed, and the cDNA (10%) was amplified by PCR with eNOS primers (Table 1) for 35 cycles as described above. Before reverse transcription, RNA concentrations were normalized, and the ribosomal RNAs, 28S and 18S, were analyzed by agarose gel electrophoresis. The number of copies of eNOS mRNA was estimated based on comparison with a standard curve generated in parallel. To normalize eNOS mRNA levels to a gene that is expressed in all vascular cells and to a gene that is an endothelial cell marker, residual cDNA (10%) from the same reverse transcriptase reactions was amplified by PCR, using primers for canine GAPDH and vWF, respectively.

Statistics

Results are expressed as means ± SE. Regression lines were calculated by the least-squares regression method (Prism; GraphPad Software). Comparisons between blood vessels were made by one-way ANOVA, and significance was calculated with the Newman-Keuls post hoc test. A value of P < 0.05 was considered significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cloning of NOS cDNA Fragments

A 1.86-kb fragment of eNOS cDNA was cloned from canine AEC (accession number, AF068681), corresponding to nucleotides 64-1927 of the human eNOS cDNA. Canine eNOS was ~85% similar to human (M95296), bovine (M95674), murine (U53142), and porcine (U59924) eNOS cDNAs (Table 2). However, canine eNOS shares only 44-51% homology with iNOS sequences from rat (L12562), mouse (M87039), human (L24553), and guinea pig (AF027180) and 57% homology to mouse (D14552) and human (U17327) nNOS cDNAs.

                              
View this table:
[in this window]
[in a new window]
  		Table 2.   Interspecies comparison of cloned NOS isoforms with canine NOS cDNA fragments

With the use of LPS-treated splenic vein endothelial cells, a 1.96-kb segment of canine iNOS cDNA was cloned (AF068682). Canine iNOS cDNA was ~80% similar to published sequences for human, mouse, rat, and guinea pig (Table 2). Canine iNOS was 44-51% identical to human, mouse, and porcine eNOS sequences and 36-48% homologous to mouse and human nNOS sequences.

With the use of canine myocardium as a source of nNOS, a 1.16-kb fragment of canine nNOS cDNA was isolated (AF068683). Interspecies comparison of the canine nNOS cDNA sequence with neuronal isoforms from human, mouse, and rat (X59949) is shown in Table 2. Canine nNOS was only 54-57% similar to eNOS sequences from human, bovine, porcine, and mouse and 55-56% homologous with iNOS from guinea pig, mouse, human, and rat iNOS sequences. Within-species comparison of cloned cDNA fragments for the canine NOS indicated homology of only 43-54% between eNOS, iNOS, and nNOS.

Specificity of Primers

Because of significant homology between canine eNOS, iNOS, and nNOS, the specificity of primers for each isoform was confirmed by PCR. As shown in Fig. 2A, primers specific for eNOS selectively amplified eNOS cDNA but did not amplify regions of iNOS and nNOS that overlap the eNOS sequence. Primers designed to recognize canine iNOS and nNOS templates amplified their respective targets only.

Enrichment of eNOS During Isolation of Cardiac Microvessels

Next, we examined the expression of several genes, including eNOS, iNOS, nNOS, vWF, Cav-1, and GAPDH in canine left ventricle, a crude mixture of microvessels and myocytes, and purified microvessels. As seen in Fig. 2B, eNOS, vWF, and Cav-1 were enriched by the purification procedure, whereas GAPDH was expressed equally. Although substantially less abundant than eNOS, iNOS and nNOS mRNAs were also enriched. Also shown in Fig. 2B, the integrity of ribosomal RNAs was maintained throughout the isolation procedure.

Amplification Kinetics

The ability of the competitor to compete in a linear relationship with native eNOS was determined from either 1) decreasing concentrations of eNOS cDNA in the presence of a fixed concentration of eNOS-C (Fig. 3A) or 2) decreasing concentrations of eNOS-C cDNA against a fixed concentration of eNOS cDNA (Fig. 3B). Regardless of how the curves were constructed, the relationship between the competitor and the full-length eNOS was always linear, with correlation coefficients (r2) of 0.996 (P < 0.001; Fig. 3A) and 0.9981 (P < 0.001; Fig. 3B, n = 3). Next, we examined the relationship between eNOS-C RNA (cRNA) and full-length eNOS mRNA. Quantitation of eNOS mRNA from serial dilutions (2-0.0625 µg) of AEC total RNA also resulted in a linear relationship with an r2 of 0.989 (P < 0.001; n = 3; data not shown). Extrapolating from this curve, 1 µg of AEC total RNA contains ~1.74 fmol of eNOS mRNA.


View larger version (25K):
[in this window]
[in a new window]
  		Fig. 3.   Competitive PCR for eNOS is linear with respect to native template and competitor. A: fixed concentration of eNOS cDNA (21 amol) was incubated in the presence of decreasing concentrations (332-2.6 amol) of competitor eNOS (eNOS-C) cDNA and were amplified by PCR for 35 cycles. The relative densities of eNOS (411 bp) and eNOS-C (332 bp) bands were determined by integration and were plotted as a ratio vs. the concentration of input eNOS-C cDNA (n = 4). B: converse relationship between decreasing concentrations of eNOS cDNA (332-2.6 amol) and a constant concentration of eNOS-C (21 amol).

Quantitation of eNOS mRNA Levels

The number of copies of eNOS mRNA transcripts was estimated in total RNA (500 ng) from the aorta, RCA, LAD, left circumflex, and coronary microvessels. The highest amount of eNOS was found in canine microvessels followed by circumflex, RCA, LAD, and aorta (n = 4 dogs, Fig. 4A). PCR amplification of total RNA without reverse transcription did not result in a defined product, excluding the amplification of genomic DNA. Within-assay coefficients of variation were <10%. The sensitivity of the assay under stated conditions was 0.2 amol. The expression of vWF and GAPDH was also examined in the various vessels. As seen in Fig. 4B, the ratio of vWF to GAPDH was greatest in coronary microvessels, followed by the RCA, circumflex, and the LAD. The aorta had the lowest expression of vWF relative to that of GAPDH. The amount of eNOS mRNA normalized to the relative abundance of vWF from each blood vessel was greatest in the circumflex artery, closely followed by the microvessels, intermediate in the LAD and RCA, and the least in the aorta (Fig. 4C).


View larger version (13K):
[in this window]
[in a new window]
  		Fig. 4.   Relative levels of eNOS mRNA in the canine cardiac vasculature. A: total RNA (500 ng) from the aorta, right coronary artery (RCA), left anterior descending (LAD), circumflex artery, aorta, and microvessels was incubated with competitor (50 fg), reverse transcribed, and amplified by PCR for 35 cycles (n = 4). Quantitation was achieved via comparison with a standard curve. B: densitometric ratio of vWF and GAPDH mRNA from cardiac blood vessels. With the use of primers for respective genes, cDNA from A was amplified by PCR for 25 cycles. C: eNOS mRNA levels normalized to vWF. ** Significantly different from all other vessels (P < 0.01). + Significantly different from aorta (P < 0.05).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Canine cardiovascular models are well-established correlates of the human conditions they emulate. However, there is little available information on how canine genes involved in cardiovascular function are regulated due to the lack of specific molecular probes. In the present study, using a PCR-based approach, we have cloned and sequenced cDNA fragments for canine eNOS, iNOS, and nNOS. The sequences of canine eNOS, iNOS, and nNOS are closely related to respective isoforms of other species (80-90% homology), which is in agreement with existing interspecies variation. Comparison between different isoforms indicates that canine NOS are also as dissimilar as that reported for other species (40-50% homology).

Previous studies have measured changes in canine eNOS mRNA expression via Northern analysis (24, 27). However, the sensitivity of this technique is such that large amounts of RNA (10-12 µg) must be obtained from endothelial cells scraped from the lumen of the thoracic aorta. We therefore designed a competitive RT-PCR assay, based on standard curve methodology, to quantify eNOS expression in canine blood vessels. The inherent sensitivity of PCR obviates the need to separate endothelial cells before RNA extraction. A PCR-based gene-fusion strategy was employed to generate a competitor with an internal deletion, enabling one to distinguish native eNOS and competitor by agarose gel electrophoresis. Optimal primers were designed, based on the sequence of canine eNOS, and were demonstrated not to recognize related sequences on iNOS and nNOS. Competition between the internal deletion mutant and native eNOS followed a linear relationship, a necessity for accurate quantitation, and the relationship remained linear regardless of whether the competing entities were of DNA or RNA origin.

The isolation of coronary microvessels from the left ventricle has been shown to result in the appearance of what morphologically resembles a mixture of arterioles, capillaries, and collagen fibers (11). The procedure is time intensive, and initially we were concerned about the ability to extract reasonable amounts of nondegraded RNA. Although the 18S and 28S ribosomal RNA bands appeared intact and nondegraded, we cannot rule out the possibility that there was some degree of degradation during the isolation procedure. One potential caveat of RNA isolation is that, in the absence of obvious degradation, it is possible that the extent of degradation for individual transcripts may differ. The comparative analysis of several genes isolated from microvessels in this study makes the assumption that the rate of degradation, if any, is similar between the products quantified. However, not only were we able to isolate quality RNA from microvessels, but we have also demonstrated that the purification procedure is associated with an enrichment of genes that are recognized endothelial cell markers, including vWF, eNOS, and Cav-1, consistent with their increased expression in vascular endothelial cells.

Quantitation of eNOS mRNA levels in cardiac blood vessels revealed that, per 500 ng of total RNA, purified microvessels contain the highest concentration of eNOS, which was 14.9 times greater than that found in the aorta, which had the lowest concentration of eNOS. The circumflex artery had the next highest level of eNOS, with 3.1-fold less than in microvessels, followed by the RCA with 4.7-fold and the LAD with 6.8-fold less eNOS per 500 ng of RNA. Given the monolayer nature of the vascular endothelium, the differences in eNOS expression most likely reflect changes in the ratio of the number of cells in the intima versus the media. Indeed, the relative ratios of vWF mRNA, an endothelial cell marker to GAPDH, a gene expressed in all vascular cells, mirrored the expression profile for eNOS mRNA, with the greatest levels detected in microvessels and the least from the aorta. Furthermore, cultured canine AECs possess 123 times more copies of eNOS per 0.5 µg of RNA than do canine microvessels. However, quantitation of the expression of eNOS mRNA in canine blood vessels does not reflect the number of copies or density of eNOS mRNA per endothelial cell. As an estimate of that determinant, we normalized eNOS expression to vWF, which was coamplified from the same pool of cDNA. The ratio of eNOS to vWF was greatest in the circumflex artery, closely followed by the microvessels, and the smallest ratio was seen in the aorta. Normalization relies on the uniform expression of vWF mRNA in the endothelium of canine blood vessels, and a potential caveat to an estimate of density per endothelial cell is that expression of vWF may not be homogeneously distributed within the cardiac vasculature (10, 28). There are numerous reports in the literature of the greater capacity of large vessels to generate NO and exhibit endothelium-dependent relaxations that are NO mediated (9, 20, 24) relative to the microvasculature. Thus our estimates of eNOS mRNA levels across the cardiac vasculature may not parallel the ability of individual vessels to produce NO due to the many constraints on the enzymatic production of NO, which may also vary regionally and segmentally throughout the vasculature (1).

The standard curve approach to quantitation of mRNA (29) offers several advantages over other methods. With a titration assay, a serial dilution must be generated for each sample, requiring an enormous number of reactions per assay. Single tube competitive PCR requires that the competitor and the native gene have identical amplification kinetics and that kinetics remain identical in both the exponential and plateau phases of PCR. With the use of a standard curve approach, many samples can draw from a single standard curve. Regardless of the magnitude of the ratio between the native gene and the competitor, there is a corresponding ratio that is identical at some point on the standard curve, and because the standard curve is constructed with the same concentration of RNA competitor the amplification kinetics and formation of heteroduplexes must be identical. The sensitivity of the current assay was 0.2 amol. This is the quantity of native eNOS beyond which the ratio of competitor (50 fg) to native eNOS is no longer discernible. Depending on the requirements, the sensitivity of the assay can be adjusted by titration such that a decrease in the quantity of competitor added to RT reactions would result in increased sensitivity. The lower limits of the assay have not been defined. In addition, because the reverse transcriptase is primed with random hexamers, many genes can be evaluated simultaneously from the same pool of cDNA.

In summary, we have determined partial cDNA sequences for the three canine NOS isoforms. Based on these sequences, we have developed a selective, sensitive, and accurate nonradioactive method to measure eNOS mRNA expression in all canine cardiac blood vessels. The ability to accurately measure eNOS expression from many different blood vessels, especially coronary microvessels, which determine coronary resistance and thus blood flow, will provide a greater perspective in our understanding of the molecular changes in eNOS that may occur in canine models of cardiovascular conditions.


    ACKNOWLEDGEMENTS

This work was supported by National Heart, Lung, and Blood Institute Grants HL-PO-1 43023, HL-50142, and HL-53053 (T. H. Hintze) and HL-57665 and HL-51948 (W. C. Sessa). D. Fulton was supported by a consortium arrangement on HL-PO-143023. W. C. Sessa is an Established Investigator of the American Heart Association.


    FOOTNOTES

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. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: W. C. Sessa, Yale Univ. School of Medicine, BCMM 436D, 295 Congress Ave., New Haven, CT 06536-0812 (E-mail: william.sessa{at}yale.edu).

Received 23 March 1999; accepted in final form 15 September 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Andries, L. J., D. L. Brutsaert, and S. U. Sys. Nonuniformity of endothelial constitutive nitric oxide synthase distribution in cardiac endothelium. Circ. Res. 82: 195-203, 1998[Abstract/Free Full Text].

2.   Cardillo, C., C. M. Kilcoyne, A. A. Quyyumi, R. O. Cannon III, and J. A. Panza. Selective defect in nitric oxide synthesis may explain the impaired endothelium-dependent vasodilation in patients with essential hypertension. Circulation 97: 851-856, 1998[Abstract/Free Full Text].

3.   Drexler, H., D. Hayoz, T. Munzel, H. Just, R. Zelis, and H. R. Brunner. Endothelial dysfunction in chronic heart failure. Experimental and clinical studies. Arzneimittelforschung 44: 455-458, 1994[Medline].

4.   Feron, O., F. Saldana, J. B. Michel, and T. Michel. The endothelial nitric-oxide synthase-caveolin regulatory cycle. J. Biol. Chem. 273: 3125-3128, 1998[Abstract/Free Full Text].

5.   Garcia-Cardena, G., R. Fan, V. Shah, R. Sorrentino, G. Cirino, A. Papapetropoulos, and W. C. Sessa. Dynamic activation of endothelial nitric oxide synthase by Hsp90. Nature 392: 821-824, 1998[Medline].

6.   Garcia-Cardena, G., R. Fan, D. F. Stern, J. Liu, and W. C. Sessa. Endothelial nitric oxide synthase is regulated by tyrosine phosphorylation and interacts with caveolin-1. J. Biol. Chem. 271: 27237-27240, 1996[Abstract/Free Full Text].

7.   Garcia-Cardena, G., P. Martasek, B. S. Masters, P. M. Skidd, J. Couet, S. Li, M. P. Lisanti, and W. C. Sessa. Dissecting the interaction between nitric oxide synthase (NOS) and caveolin. Functional significance of the nos caveolin binding domain in vivo. J. Biol. Chem. 272: 25437-25440, 1997[Abstract/Free Full Text].

8.   Gardiner, S. M., A. M. Compton, T. Bennett, R. M. Palmer, and S. Moncada. Control of regional blood flow by endothelium-derived nitric oxide. Hypertension 15: 486-492, 1990[Abstract/Free Full Text].

9.   Garland, C. J., F. Plane, B. K. Kemp, and T. M. Cocks. Endothelium-dependent hyperpolarization: a role in the control of vascular tone. Trends Pharmacol. Sci. 16: 23-30, 1995[Medline]. (published erratum appears in Trends Pharmacol. Sci. 16: 177, 1995)

10.   Gebrane-Younes, J., L. Drouet, J. P. Caen, and L. Orcel. Heterogeneous distribution of Weibel-Palade bodies and von Willebrand factor along the porcine vascular tree. Am. J. Pathol. 139: 1471-1484, 1991[Abstract].

11.   Gerritsen, M. E., and M. P. Printz. Sites of prostaglandin synthesis in the bovine heart and isolated bovine coronary microvessels. Circ. Res. 49: 1152-1163, 1981[Abstract/Free Full Text].

12.   Griffith, O. W., and D. J. Stuehr. Nitric oxide synthases: properties and catalytic mechanism. Annu. Rev. Physiol. 57: 707-736, 1995[Web of Science][Medline].

13.   Kaiser, L., R. C. Spickard, and N. B. Olivier. Heart failure depresses endothelium-dependent responses in canine femoral artery. Am. J. Physiol. Heart Circ. Physiol. 256: H962-H967, 1989[Abstract/Free Full Text].

14.   Kichuk, M. R., X. Zhang, M. Oz, R. Michler, G. Kaley, A. Nasjletti, and T. H. Hintze. Angiotensin-converting enzyme inhibitors promote nitric oxide production in coronary microvessels from failing explanted human hearts. Am. J. Cardiol. 80: 137A-142A, 1997[Medline].

15.   Liu, J., G. Garcia-Cardena, and W. C. Sessa. Palmitoylation of endothelial nitric oxide synthase is necessary for optimal stimulated release of nitric oxide: implications for caveolae localization. Biochemistry 35: 13277-13281, 1996[Medline].

16.   Mayer, B., and B. Hemmens. Biosynthesis and action of nitric oxide in mammalian cells. Trends Biochem. Sci. 22: 477-481, 1997[Web of Science][Medline].

17.   Michel, T., G. K. Li, and L. Busconi. Phosphorylation and subcellular translocation of endothelial nitric oxide synthase. Proc. Natl. Acad. Sci. USA 90: 6252-6256, 1993[Abstract/Free Full Text].

18.   Miller, V. M., and P. M. Vanhoutte. Enhanced release of endothelium-derived factor(s) by chronic increases in blood flow. Am. J. Physiol. Heart Circ. Physiol. 255: H446-H451, 1988[Abstract/Free Full Text].

19.   Nadaud, S., M. Philippe, J. F. Arnal, J. B. Michel, and F. Soubrier. Sustained increase in aortic endothelial nitric oxide synthase expression in vivo in a model of chronic high blood flow. Circ. Res. 79: 857-863, 1996[Abstract/Free Full Text].

20.   Nagao, T., S. Illiano, and P. M. Vanhoutte. Heterogeneous distribution of endothelium-dependent relaxations resistant to NG-nitro-L-arginine in rats. Am. J. Physiol. Heart Circ. Physiol. 263: H1090-H1094, 1992[Abstract/Free Full Text].

21.   Pieper, G. M., D. A. Mei, P. Langenstroer, and S. T. O'Rourke. Bioassay of endothelium-derived relaxing factor in diabetic rat aorta. Am. J. Physiol. Heart Circ. Physiol. 263: H676-H680, 1992[Abstract/Free Full Text].

22.   Saenz de Tejada, I., I. Goldstein, K. Azadzoi, R. J. Krane, and R. A. Cohen. Impaired neurogenic and endothelium-mediated relaxation of penile smooth muscle from diabetic men with impotence. N. Engl. J. Med. 320: 1025-1030, 1989[Abstract].

23.   Sessa, W. C., G. Garcia-Cardena, J. Liu, A. Keh, J. S. Pollock, J. Bradley, S. Thiru, I. M. Braverman, and K. M. Desai. The Golgi association of endothelial nitric oxide synthase is necessary for the efficient synthesis of nitric oxide. J. Biol. Chem. 270: 17641-17644, 1995[Abstract/Free Full Text].

24.   Sessa, W. C., K. Pritchard, N. Seyedi, J. Wang, and T. H. Hintze. Chronic exercise in dogs increases coronary vascular nitric oxide production and endothelial cell nitric oxide synthase gene expression. Circ. Res. 74: 349-353, 1994[Abstract/Free Full Text].

25.   Shaul, P. W., E. J. Smart, L. J. Robinson, Z. German, I. S. Yuhanna, Y. Ying, R. G. Anderson, and T. Michel. Acylation targets emdothelial nitric-oxide synthase to plasmalemmal caveolae. J. Biol. Chem. 271: 6518-6522, 1996[Abstract/Free Full Text].

26.   Shesely, E. G., N. Maeda, H. S. Kim, K. M. Desai, J. H. Krege, V. E. Laubach, P. A. Sherman, W. C. Sessa, and O. Smithies. Elevated blood pressures in mice lacking endothelial nitric oxide synthase. Proc. Natl. Acad. Sci. USA 93: 13176-13181, 1996[Abstract/Free Full Text].

27.   Smith, C. J., D. Sun, C. Hoegler, B. S. Roth, X. Zhang, G. Zhao, X. B. Xu, Y. Kobari, K. Pritchard, Jr., W. C. Sessa, and T. H. Hintze. Reduced gene expression of vascular endothelial NO synthase and cyclooxygenase-1 in heart failure. Circ. Res. 78: 58-64, 1996[Abstract/Free Full Text].

28.   Smith, J. M., J. H. Meinkoth, T. Hochstatter, and K. M. Meyers. Differential distribution of von Willebrand factor in canine vascular endothelium. Am. J. Vet. Res. 57: 750-755, 1996[Web of Science][Medline].

29.   Tsai, S. J., and M. C. Wiltbank. Quantification of mRNA using competitive RT-PCR with standard-curve methodology. Biotechniques 21: 862-866, 1996[Web of Science][Medline].

30.   Wang, J., N. Seyedi, X. B. Xu, M. S. Wolin, and T. H. Hintze. Defective endothelium-mediated control of coronary circulation in conscious dogs after heart failure. Am. J. Physiol. Heart Circ. Physiol. 266: H670-H680, 1994[Abstract/Free Full Text].

31.   Zhang, X., Y. W. Xie, A. Nasjletti, X. Xu, M. S. Wolin, and T. H. Hintze. ACE inhibitors promote nitric oxide accumulation to modulate myocardial oxygen consumption. Circulation 95: 176-182, 1997[Abstract/Free Full Text].

Am J Physiol Heart Circ Physiol 278(2):H658-H665
0363-6135/00 $5.00 Copyright © 2000 the American Physiological Society


This article has been cited by other articles:


Home page
 Physiol. Rev.Home page
J.-L. Balligand, O. Feron, and C. Dessy
eNOS Activation by Physical Forces: From Short-Term Regulation of Contraction to Chronic Remodeling of Cardiovascular Tissues
Physiol Rev, April 1, 2009; 89(2): 481 - 534.
[Abstract] [Full Text] [PDF]

Home page
 Circ. Res.Home page
R. D. Rudic, D. Brinster, Y. Cheng, S. Fries, W.-L. Song, S. Austin, T. M. Coffman, and G. A. FitzGerald
COX-2-Derived Prostacyclin Modulates Vascular Remodeling
Circ. Res., June 24, 2005; 96(12): 1240 - 1247.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
K. Funabiki, K. Onishi, K. Dohi, T. Koji, K. Imanaka-Yoshida, M. Ito, H. Wada, N. Isaka, T. Nobori, and T. Nakano
Combined angiotensin receptor blocker and ACE inhibitor on myocardial fibrosis and left ventricular stiffness in dogs with heart failure
Am J Physiol Heart Circ Physiol, December 1, 2004; 287(6): H2487 - H2492.
[Abstract] [Full Text] [PDF]

Home page
 Physiol. Rev.Home page
D. L. Brutsaert
Cardiac Endothelial-Myocardial Signaling: Its Role in Cardiac Growth, Contractile Performance, and Rhythmicity
Physiol Rev, January 1, 2003; 83(1): 59 - 115.
[Abstract] [Full Text] [PDF]

Home page
 HypertensionHome page
M. Fujii, A. Wada, T. Tsutamoto, M. Ohnishi, T. Isono, and M. Kinoshita
Bradykinin Improves Left Ventricular Diastolic Function Under Long-Term Angiotensin-Converting Enzyme Inhibition in Heart Failure
Hypertension, May 1, 2002; 39(5): 952 - 957.
[Abstract] [Full Text] [PDF]

Home page
 Circ. Res.Home page
R. D. Rudic, M. Bucci, D. Fulton, S. S. Segal, and W. C. Sessa
Temporal Events Underlying Arterial Remodeling After Chronic Flow Reduction in Mice : Correlation of Structural Changes With a Deficit in Basal Nitric Oxide Synthesis
Circ. Res., June 9, 2000; 86(11): 1160 - 1166.
[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 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 Web of Science (7)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Fulton, D.
Right arrow Articles by Sessa, W. C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Fulton, D.
Right arrow Articles by Sessa, W. C.

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