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Ann Thorac Surg 2002;74:90-95
© 2002 The Society of Thoracic Surgeons

Original article: cardiovascular

Platelet-derived growth factor-induced expression of c-fos in human vascular smooth muscle cells: implications for long-term graft patency

Dario F. Del Rizzo, MD, PhDa, Natalia Yurkova, PhDa, Michael C. Moon, MDa, Brenda Litchie, BSa, Peter Zahradka, PhD*a

a Institute of Cardiovascular Sciences, St. Boniface General Hospital Research Centre, University of Manitoba, Faculty of Medicine, Winnipeg, Manitoba, Canada

Accepted for publication March 18, 2002.

* Address reprint requests to Dr Zahradka, Institute of Cardiovascular Sciences, St. Boniface General Hospital Research Centre, 351 Taché Ave, Winnipeg, MB R2H 2A6, Canada
e-mail: peterz{at}sbrc.ca


   Abstract
Top
 Abstract
Introduction
 Material and methods
 Results
 Comment
 Acknowledgments
 References
 
Background. The internal mammary artery (IMA) has been shown to have a significantly superior long-term patency rate when compared with the saphenous vein (SV) graft. Cultured smooth muscle cells (SMCs) from the IMA are more resistant to the mitogenic effects of platelet-derived growth factor (PDGF), when compared with SMCs that are derived from the SV. The radial artery (RA) is currently being used as an alternative to the SV. However, no long-term patency data are available for the RA, and there is no information on the biological behavior of RA-derived SMCs in culture.

Methods. Smooth muscle cell cultures were taken from patients who underwent coronary artery bypass grafting with the IMA, RA, and SV. A quiescent state was induced by serum deprivation for 5 days. Thereafter cells were induced to proliferate by exposure to PDGF-BB. Levels of c-fos expression and 3H-thymidine incorporation were used as markers of cell proliferation.

Results. We found that even after serum deprivation, c-fos was still detectable; however, basal levels were higher in cells from the SV than cells from either the RA (p = 0.003) or IMA (p = 0.008). After stimulation with PDGF-BB, c-fos expression was greater in SMCs from the SV relative to the RA (p < 0.001 or the IMA (p = 0.02). Finally, relative to the SV, 3H-thymidine in the RA was 0.76 ± 0.22 (p < 0.05) and 0.39 ± 0.24 (p < 0.002) in the IMA, respectively.

Conclusions. The data indicate that SMCs from arterial conduits are more resistant to the mitogenic effects of PDGF-BB than those from venous conduits. Our results offer a mechanistic explanation of why arterial conduits might demonstrate patency superior to that of the SV.


   Introduction
Top
 Abstract
 Introduction
Material and methods
 Results
 Comment
 Acknowledgments
 References
 
The long-term benefits of coronary artery bypass grafting (CABG) surgery are in large part dependent on the fate of the conduits used to revascularize the myocardium. Saphenous vein (SV) grafts have a predictable failure rate such that 10 years after operation, only 50% to 60% of these grafts are patent [1]. The problem of vein graft attrition after CABG surgery is complex and multifactorial. However, smooth muscle cell (SMC) proliferation is considered to be a key component in this process [2–5]. Proliferation of SMCs results in intimal hyperplasia [6], also called neointimal proliferation or neointimal thickening, and is responsible for early graft failure. Late graft failure is the result of arteriosclerosis. However, the entire process can be considered to be part of the same disease spectrum the development of which is secondary to the uncontrolled proliferation of the SMCs within the vessel wall [7]. Under normal circumstances the SMCs in the walls of arteries and veins exhibit a nonproliferative contractile phenotype. However, as a result of injury, mitogenic stimulation, or altered metabolic states within the host, these cells begin to grow. Acquisition of the synthetic phenotype is associated with proliferation of these cells and occurs through a process that is referred to as phenotypic modulation.

Smooth muscle cells from SVs obtained postmortem from patients who had previously undergone CABG surgery demonstrate a decrease in levels of actin and myosin [8] that can be detected within days after operation [9]. The cells become motile and assume an appearance similar to that of fibroblasts. At this point they begin to secrete collagen and other proteins that eventually result in the formation of an extracellular matrix. The matrix then becomes the scaffold for calcification. These changes are characteristic of the synthetic phenotype. This process is known as vascular remodeling [5].

In contrast to the SV, the internal mammary artery (IMA) is more resistant to the development of arteriosclerosis [10]. This results in prolonged patency and superior long-term survival in patients undergoing CABG surgery with a left IMA (LIMA) versus a SV bypass to the left anterior descending coronary artery [11–14]. This knowledge was the rationale for the expanded use of alternate arterial conduits as bypass grafts that include the gastroepiploic (GEA), inferior epigastric (IEA), and radial artery (RA) [15–22]. Although it is presumed that these conduits will have superior patency rates than the SV, only limited long-term patency data exist to support this position [23].

The superior patency of the IMA over SV may be related to inherent differences in the proliferative capacity of SMCs from arterial versus venous conduits. Yang and colleagues [24] have shown that the SMCs from the IMA are relatively resistant to the mitogenic effects of platelet-derived growth factor (PDGF) when compared with SMCs from human SVs. These authors also showed that thrombin causes a much more pronounced mitogenic and contractile effect in SMCs originating from SV than in the IMA. These results suggest that biological differences in SMCs from arterial versus venous origin, particularly with respect to the degree of cell proliferation and phenotypic modulation, might be why patency of the IMA is superior to that of SV. However, no experimental data are available on the proliferative capacity of SMCs derived from the RA. In addition, as previously stated, no clinical studies have been published on the long-term (> 10 years) patency of the RA as a bypass conduit. The current study was undertaken to examine the mitogenic response to PDGF-BB of SMCs from RA, IMA, and SV conduits and to compare expression of markers of cell proliferation in these cells. The biological behavior of the RA may offer insight into its long-term performance as a bypass conduit.


   Material and methods
Top
 Abstract
 Introduction
 Material and methods
Results
 Comment
 Acknowledgments
 References
 
Patient demographics
Institutional ethics approval was obtained for the use of human tissue for research. After informed consent patients underwent elective CABG surgery. Any unused portion of the IMA, SV, and RA was placed in saline at 4°C and transferred to the laboratory.

There were 6 patients from whom we obtained sufficient material to culture SMCs from all three conduits (IMA, SV, and RA). The treating physicians provided laboratory personnel with demographic data on the patients. All were male with a mean age of 60.0 ± 10.2 years. One patient had non-insulin-dependent diabetes, 5 had hyperlipidemia, 2 had a previous smoking history, 4 had a positive family history of coronary artery disease (CAD), and 2 had underlying hypertension. All patients had at least two major risk factors for the development of CAD. Patient data are summarized in Table 1.


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  		Table 1. Demographic Data For Study Patients

 
Tissue culture materials and reagents
Tissue culture medium (Dulbecco’s Modified Eagle’s Medium [DMEM], high glucose), Nunc tissue culture plates, antibiotics, trypsin, fetal bovine serum (FBS), antibiotic-antimycotic (AB/AM 100X: 10,000 U/mL penicillin G, 10,000 µg/mL streptomycin sulfate, 25 µg/mL amphotericin), and TRIzol reagent were purchased from Gibco/BRL (Grand Island, NY). Linbro multiwell culture dishes were obtained from Flow Laboratories, Inc (McLean, VA). The PDGF-BB (ß-receptor binding isoform) was purchased from PeproTech Inc (Rocky Hill, NJ). Humulin-R (human insulin, [7.2 x 10-4 mol/L]) was obtained from Lilly (Eli Lilly Canada Inc, Toronto, ON). A 500-µL insulin stock solution was prepared by combining 69 µL of Humulin-R with 431 µL of sterile water in an autoclaved microfuge tube (final concentration 10-4 mol/L).

Transferrin, selenium, ascorbate, and sodium pyruvate were purchased from Sigma Chemical (St. Louis, MO). Supplemental stock of 100X PTSA (27.5 mg pyruvate, 12.5 mg transferring, 0.17 mg/mL selenium [2.5 µL], and 0.1 g ascorbate) was prepared in 25 mL of sterile water and stored in 5-mL aliquots at -80°C. Smooth muscle cell phosphate buffer saline (SMC-PBS) was prepared by the addition of 9% NaCl (50 mL) to 100 mL of 0.5 mol/L Na3PO4 (pH 7.1).

Standard (10X) culture medium contains 70 mL DMEM, 20 mL FBS, and 10 mL 10XAB/AM per 100 mL total volume. 2X culture medium contains 78 mL DMEM, 20 mL FBS, and 2 mL 10XAB/AM per 100 mL total volume. Serum-free medium contains 1 mL PTSA Supplement Stock, 1 mL 10XAB/AM Solution, 10 µL of Humulin-R (from a 10-4 mol/L stock solution), and 98 mL DMEM per 100 mL total volume. Radioisotopes for DNA synthesis ([methyl 3H]-thymidine, 2 Ci/mmol) studies were purchased from Amersham Canada (Oakville, ON, Canada).

Cell culture techniques
After transport to the laboratory, human conduits were immediately prepared for culture. Any adhering tissue (eg, perivascular fat from SVs, IMA pedicle), suture material, and surgical clips were removed by sharp dissection, avoiding stretching or puncturing of the vessels. They were then sectioned into rings 2 to 3 mm and completely immersed in 10X culture medium. The proximal and distal ends of each conduit were discarded. After 5 days vessel rings and cells were grown in 2X culture medium, which was changed every 2 to 3 days or whenever cells were seeded onto new plates.

Cell migration began after 7 to 14 days in culture. When cell clusters reached 20% to 30% confluency, the rings were transferred onto new plates. The first migration (or M1 cells) was allowed to grow until the clusters became dense. The cells were then trypsinized and plated onto progressively larger culture plates.

The new plates were watched for the beginning of the second migration, the M2 cells. When cultured cells (M1 or M2) reached 70% to 80% confluency, medium was removed and the plates were trypsinized for 5 minutes at 37°C. Gentle agitation facilitated the loosening of any residual adherent cells, and the trypsin solution was neutralized with an equal volume of culture medium and pipetted repeatedly to loosen cell clumps further. Cells were kept in serum-free medium for 5 days before experimental use. Previous work from our laboratory had demonstrated in porcine coronary artery SMCs that 100 hours of serum deprivation is required to induce a state of quiescence [25].

DNA synthesis
Cells were cultured as described above, rendered quiescent in 24-well dishes containing 1 mL of serum-free medium, and then stimulated by direct addition of PDGF-BB (0.1 µg/mL) without replacing the medium. Cells were incubated for 24 hours before addition of 1 µCi 3H-thymidine (0.1 mCi/mL stock). The cells were incubated for an additional 48 hours and lysed. The TCA precipitable fraction was captured on Whatman (GF/A) filters and counted in a liquid scintillation counter using a Scintiverse-II scintillation cocktail, as previously described [26].

RNA extraction
A quantity of 2 mL of medium was removed and 2 µL of PDGF-BB (0.1 µg/mL) added to each well. After 15 minutes culture medium was removed and the adherent cells were washed twice with PBS (pH 7.4) at 4°C. Total cellular RNA was extracted with TRIzol reagent. The RNA was quantified by measuring absorbance spectrophotometrically at 260 nm, and its integrity was assessed by electrophoresis in nondenaturing 1% agarose gels stained with ethidium bromide.

Semiquantitative reverse transcription polymerase chain reaction
Reverse transcription of 1 µg total RNA was performed using the Promega Access RT-PCR System (Promega Corporation, Madison, WI). Possible genomic DNA contamination was eliminated by incubating the RNA samples with DNase I for 15 minutes at room temperature before reverse transcription polymerase chain reaction (RT-PCR). The RT-PCR mixture, which was prepared according to the manufacturer’s protocol, contained 0.2 nmol/L dNTP, 1 µmol/L forward and reverse primers, 1 mmol/L MgSO4, 1X AMV/Tfl reaction buffer, 0.1 U/µL AMV reverse transcriptase, 0.1 U/µL Tfl polymerase. The temperature program for amplification was 25 cycles of 30 seconds per cycle at 94°C, followed by 1 minute at 62°C and 2 minutes at 68°C. The final extension was completed at 68°C for 7 minutes.

Levels of c-fos expression were normalized against GAPDH expression. Specific forward and reverse primers were designed accordingly:

c-fos: 258 bp: forward 5'-GGACCTATCTGGGTCCTTCTATG-3' reverse 5'-CGAGTCAGAGGAAGGCTCATT-3'.
GAPDH: 306 bp: forward 5'-CGCTGTGAACGGATTTGGCCGTAT-3' reverse 5'-AGCCTTCTCCATGGTGGTGAAGAC-3'.
RT-PCR products were analyzed by electrophoresis on 2% agarose gels stained with ethidium bromide. Relative band intensity was quantified by scanning densitometry (GS-670 Densitometer, Bio-Rad Laboratories, Mississauga, ON).

Statistical analyses
All statistical analyses were performed using Microsoft Excel 98 for MacIntosh. Data were analyzed by Student’s t test. Statistical significance was assumed for p values less than 0.05.


   Results
Top
 Abstract
 Introduction
 Material and methods
 Results
Comment
 Acknowledgments
 References
 
3H-thymidine incorporation
Levels of 3H-thymidine uptake after stimulation with PDGF-BB are summarized in Figure 1. For each tissue sample, base line and poststimulation counts were derived from six wells and the data were averaged. The absolute increase in thymidine incorporation was derived by subtraction of the average base line count from the poststimulation count. For each patient, thymidine uptake from SMCs of the SV was assigned the value 1, whereas uptake from the RA and IMA were expressed as a ratio (arterial conduit/venous conduit scintillation counts). We found that there was significantly less tracer uptake in SMCs from arterial versus venous conduits after stimulation with PDGF-BB. Relative to the SV, 3H-thymidine in the RA was 0.76 ± 0.22 (p < 0.05) and 0.39 ± 0.24 (p < 0.002) in the IMA, respectively. These data suggest that SMCs derived from arterial conduits are more resistant to mitogenic stimulation than those from venous conduits.



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  		Fig 1. 3H-thymidine uptake after stimulation by platelet-derived growth factor-BB (PDGF-BB). For each conduit the increase in isotope uptake was determined by subtraction of baseline counts (in cells rendered quiescent by serum deprivation) from poststimulation counts (after exposure to PDGF-BB). For each saphenous vein this number was assigned the value 1. Differences between the conduits are expressed as a ratio (radial artery/saphenous vein or internal mammary artery/saphenous vein). Relative to the saphenous vein graft (SVG), 3H-thymidine in the radial artery was 0.76 ± 0.22 (p < 0.05) and 0.39 ± 0.24 (p < 0.002) in the internal mammary artery (IMA), respectively.

 
Levels of c-fos expression before and after stimulation with PDGF-BB
Levels of c-fos expression in quiescent SMCs are summarized in Figure 2. Relative expression for each of the three conduits was determined with the aid of a densitometer. For each patient, c-fos expression in SMCs of the SV was assigned the value 1, whereas expression in the RA and IMA were expressed as a ratio (relative to the SV). We found that there was significantly less c-fos expression in the arterial conduits. Relative to the SV, c-fos expression in the RA was 0.51 ± 0.23 (p = 0.003) and 0.58 ± 0.24 (p = 0.008) in the IMA.



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  		Fig 2. Basal levels of c-fos expression in quiescent cells. After cells were rendered quiescent by serum deprivation, baseline levels of c-fos expression were determined by reverse transcription polymerase chain reaction. Relative band intensity was determined by densitometry. For each patient c-fos expression in smooth muscle cells of the saphenous vein (SV) graft (SVG) was assigned the value 1, whereas expression in the radial artery (RA) and internal mammary artery (IMA) was expressed as a ratio relative to the SV (RA/SV and IMA/SV). Relative to the SV, c-fos expression in the RA was 0.51 ± 0.23 (p = 0.003) and 0.58 ± 0.24 (p = 0.008) in the IMA.

 
In the next set of experiments we examined the effect of PDGF-BB on c-fos expression (Fig 3). Consistent with our previous findings, relative to the SV, c-fos expression in the RA was 0.69 ± 0.07 (p < 0.001) and 0.78 ± 0.16 (p = 0.02) in the IMA, respectively. Figure 4 illustrates relative band intensity before and after stimulation for three different patients. Lanes 1, 3, and 5 show basal c-fos for the SV, RA, and IMA respectively, whereas lanes 2, 4, and 6 illustrate the effect of PDGF-BB stimulation on c-fos expression for the same three conduits.



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  		Fig 3. Levels of c-fos expression after stimulation with platelet-derived growth factor-BB (PDGF-BB). Cells were rendered quiescent by serum deprivation and then exposed to PDGF-BB. Poststimulation levels of c-fos expression were determined by reverse transcription polymerase chain reaction. Relative band intensity was determined by densitometry. For each patient c-fos expression in smooth muscle cells of the saphenous vein (SV) graft (SVG) was assigned the value 1, whereas expression in the radial artery (RA) and internal mammary artery (IMA) was expressed as a ratio relative to the SV (RA/SV and IMA/SV). Relative to the SV, c-fos expression in the RA was 0.69 ± 0.07 (p < 0.001) and 0.78 ± 0.16 (p = 0.02) in the IMA, respectively.

 


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  		Fig 4. Levels of c-fos expression before and after stimulation with platelet-derived growth factor-BB (PDGF-BB). The RNA was extracted from cells rendered quiescent by serum deprivation and from cells after stimulation with PDGF-BB (0.1 µg/mL). Reverse transcription polymerase chain reaction was used to evaluate the expression of c-fos and the products were analyzed by electrophoresis on 2% agarose gels stained with ethidium bromide. Relative band intensity was quantified by scanning densitometry. The results from 3 separate patients (A, B, and C) are shown. Lane 1 = saphenous vein (SV)-PDGF-BB; lane 2 = SV + PDGF-BB; lane 3 = radial artery (RA)-PDGF-BB; lane 4 = RA + PDGF-BB; lane 5 = internal mammary artery (IMA)-PDGF-BB; and lane 6 = IMA + PDGF-BB.

 

   Comment
Top
 Abstract
 Introduction
 Material and methods
 Results
 Comment
Acknowledgments
 References
 
It is generally accepted that abnormal proliferation and migration of vascular SMCs plays an important role in the pathogenesis of atherosclerotic disease [2, 3, 27]. Stimulation by specific mitogens or growth factors induces expression of the c-fos gene product. The c-fos protein combines with the c-jun protein to form the AP-1 transcription factor, which in turn initiates downstream events that result in cellular proliferation [28, 29]. Under normal conditions, in the absence of proliferation, c-fos activity is very low or even undetectable.

Rivard and colleagues [7] recently demonstrated in New Zealand white rabbits that c-fos expression increased with age. They suggested that an age-dependent increase in the expression and DNA-binding activity of the c-fos protein might be responsible for the increase in vascular SMC proliferation associated with the development of atherosclerotic disease. These data offer a potential molecular explanation of why the incidence of atherosclerotic disease increases with advancing age.

We have examined c-fos expression in cultured SMCs from juvenile pig coronary arteries (data not shown). In these experiments, the cells were serum-deprived for 5 days to ensure entry into a quiescent state before determining levels of c-fos expression. After 5 days in serum-free medium, c-fos expression was not detectable in SMCs from these juvenile pigs. These experiments are consistent with the observation of Rivard and colleagues [7] that c-fos expression in vascular SMCs is regulated in an age-dependent manner.

The data presented herein demonstrate that even after serum deprivation for 5 days, a period of time sufficient to produce quiescence, c-fos activity could still be detected in human cultured SMCs of both arterial and venous origin. These data support the hypothesis that c-fos activity increases in an age-dependent manner, which in turn may be responsible for the genesis of arteriosclerosis in elderly individuals. Consistent with this notion is that the average age of the patients from which these cells were derived was 60.0 ± 10.2 years of age (Table 1).

When we compared the basal c-fos activity from different conduits, we noted that the level of expression was significantly greater in SMCs from SV when compared with SMCs from IMA or RA (Fig 2). In addition, basal activity levels were not significantly different in the IMA when compared with the RA. Overexpression of the AP-1 transcription factor c-fos can circumvent normal upstream negative regulators and can induce cell proliferation through interaction with the cAMP-responsive element [27]. Binding of AP-1 and cAMP-responsive element binding proteins signals downstream activation of cyclin-dependent kinase-2 and its regulatory subunits cyclin E and A, which are essential for cell cycle progression through G1 and S phase. Constitutively higher levels of c-fos in SMCs from SV suggest that the cells from these conduits have a greater propensity to enter into a proliferative state when compared with the SMCs originating in arterial grafts (IMA and RA).

In the next set of experiments we wanted to determine the response of arterial and venous vascular SMCs to mitogenic stimulation with PDGF. Yang and colleagues [24] recently demonstrated a significantly greater outgrowth of SMCs from SV versus IMA. Furthermore, SV-derived SMCs had a significantly greater proliferative response to PDGF than did IMA-derived cells. Our results are consistent with those reported by Yang and colleagues [24]. Stimulation with PDGF for 15 minutes induced a significant increase in c-fos expression of quiescent cells from all three sources. However, the absolute response was greater in cells from SV, whereas there were no differences in levels of expression between IMA-derived and RA-derived SMCs (Fig 3). These experiments indicate that, like the SMCs from the IMA, those of the RA are relatively resistant to PDGF-induced cell proliferation. As a second method to examine cell proliferation, we examined 3H-thymidine incorporation after stimulation of quiescent cells with PDGF-BB. These data are summarized in Figure 1. We found that 3H-thymidine incorporation was significantly higher in the SV when compared with that in the RA and IMA.

It is known that the IMA exhibits a low incidence of arteriosclerosis [10]. In contrast the SV, when placed in the coronary circulation, is subjected to significant atherosclerotic changes over time, resulting in a significant failure rate over a 10-year period [1]. It has been postulated that the resistance of IMA-derived SMCs to mitogenic stimulation by PDGF when compared with that of SV-derived SMCs may be responsible for the superior patency of an IMA graft versus an SV [24]. No long-term patency data are yet available for the RA, although early results are encouraging. One year after CABG, more than 90% of all RA grafts remain patent [19, 21, 22]. In our experiments, the biological behavior of cultured SMCs from RA grafts was similar to that of SMCs from IMA grafts.

A major limitation of this work is the assumption that the biological behavior of cultured cells will be consistent with how these cells behave in vivo. Although the results may be interesting, they do not necessarily reflect what occurs in the patient. Whether resistance to mitogenic stimulation of SMCs for the RA relative to the SV will result in superior long-term patency of the RA graft in comparison to the SV will require long-term angiographic follow-up studies.


   Acknowledgments
Top
 Abstract
 Introduction
 Material and methods
 Results
 Comment
 Acknowledgments
References
 
The authors wish to thank the Canadian Institutes for Health Research and the University of Manitoba Health Sciences Centre Research Foundation for their support of this work. We thank Mr Jason Voldeng for his technical assistance.


   References
Top
 Abstract
 Introduction
 Material and methods
 Results
 Comment
 Acknowledgments
 References
 

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J. Thorac. Cardiovasc. Surg., March 1, 2006; 131(3): 651 - 658.


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