(Circulation. 2000;101:524.)
© 2000 American Heart Association, Inc.
Basic Science Reports |
Characteristics of Coronary Smooth Muscle Cells and Adventitial Fibroblasts
From the Cardiovascular Research Center (S.P., Y.S., R.N., E.H.C., A.Z.), Department of Medicine (Cardiology), Thomas Jefferson University, Philadelphia, Pa; and Bryn Mawr Hospital (J.L.M.), Bryn Mawr, Pa.
Correspondence to Andrew Zalewski, MD, or Yi Shi, MD, PhD, Thomas Jefferson University, Division of Cardiology, Suite 410N, 1025 Walnut St, Philadelphia, PA 19107. E-mail andrew.zalewski@tju.edu or yi.shi{at}tju.edu
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Abstract |
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Background—Recent findings suggesting the involvement of adventitial cells in coronary repair have raised questions regarding the phenotypic "plasticity" of medial smooth muscle cells (SMCs). Accordingly, the aims of the present study were to examine the characteristics of coronary medial and adventitial cells and to compare the responses of coronary and noncoronary SMCs to stimulation.
Methods and Results—Enzymatically isolated coronary SMCs
(human and porcine) were distinct from noncoronary SMCs,
showing poor adhesion and spreading, as well as lower proliferation,
collagen synthesis, and LDL degradation. Several extracellular matrix
components (Matrigel, collagen I and IV, laminin,
vitronectin, fibronectin) or growth factors (epidermal
growth factor, platelet-derived growth factor-BB, insulin growth
factor-1, interleukin-1
) failed to augment the adhesion or
proliferation of coronary SMCs to the levels observed in
noncoronary SMCs. Unlike coronary SMCs,
coronary fibroblasts demonstrated high adhesion, proliferation,
collagen synthesis, and avid LDL metabolism. Limited
responses of coronary SMCs were associated with sustained
expression of differentiation markers (-smooth muscle actin,
h-caldesmon, and smooth muscle myosin heavy chain), whereas
noncoronary SMCs showed marked phenotypic
heterogeneity.
Conclusions—Coronary SMCs appeared to maintain highly differentiated phenotype in response to stimulation, whereas coronary adventitial fibroblasts demonstrated several characteristics that are essential during vascular repair. Coronary SMCs, however, were distinct from noncoronary medial cells, which displayed greater phenotypic heterogeneity and versatility in culture. We postulate that the mechanism of vascular repair may differ among vascular beds, pointing to the importance of coronary artery–specific investigations in vascular biology.
Key Words: remodeling • arteries • muscle, smooth • cells
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Introduction |
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Several mechanical, hemodynamic, or infectious factors contribute to the activation of resident vascular cells, resulting in the formation of intimal lesions.1 2 This process leads to changes in the cellular composition of the vessel wall, which accompany a broad spectrum of cardiovascular disorders.3 4 5 Although intimal cells share some similarities with medial smooth muscle (SM) cells (SMCs), they also display several distinct characteristics regarding morphology, gene expression, and synthetic properties (for a review, see Schwartz et al5 ). Several concepts have emerged regarding the mechanisms underlying the expansion of vascular cells in various disease states.6 7 8 The first paradigm is based on the assumption that the cells involved in vascular repair are primarily of SM origin. The most widely accepted view implies the "plasticity" of virtually all medial SMCs, which modulate their phenotype in response to stimulation. This has been based on the observed changes in cultured SMCs rapidly losing their contractile features and becoming "synthetic" cells.6 A second paradigm, which is based on more recent studies, however, suggests heterogeneity of the arterial media, with only a fraction of SMCs capable of a rapid response to stimulation. These "stem" SMCs, isolated from the rat aorta or carotid artery, differ in regard to morphology, gene expression, and behavior in culture compared with the remaining medial cells.9 10 The third paradigm implies that highly reactive "nonmuscle" cells, being able to proliferate, migrate, and synthesize extracellular matrix proteins, are also involved in the repair of the vessel wall. Although they are sparsely distributed within vascular media (eg, carotid artery, pulmonary artery, and saphenous vein), nonmuscle fibroblasts are abundant in the adventitia.11 12 13 After severe coronary injury (porcine model), nonmuscle cells of adventitial origin proliferate and migrate to the luminal region during neointimal formation.14 15 They also acquire
-SM actin (myofibroblast formation) and are capable of the synthesis
of several extracellular matrix proteins.16 17 18 Although these concepts remain the subject of ongoing debate, they are not mutually exclusive, because regional differences in the mechanisms of arterial repair may arise from the diverse lineages of vascular cells.19 In particular, the coronary vasculature demonstrates unique development, which differs from that of the aorta and its major tributaries.20 21 22 The results of the present study suggest that coronary SMCs are less responsive to stimulation and exhibit lower synthetic capability than other vascular SMCs. When the cellular constituents of the coronary arteries were analyzed, adventitial cells demonstrated particularly dynamic phenotypic characteristics. These findings suggest there are functional differences in the cellular constituents of coronary and noncoronary vascular beds, which may influence the mechanisms of vascular repair and lesion formation.
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Methods |
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Cell Isolation and Culture
Porcine arteries (epicardial coronary arteries, iliac artery, thoracic aorta) were harvested and opened, and their endothelial layers were removed. Human epicardial coronary arteries and samples of thoracic aorta, which were obtained from the recipients of heart transplantation, were prepared in an identical manner. For coronary arteries, the adventitia and media were separated with the assistance of surgical loops (magnification 2x). The separation of adventitial and medial layers was confirmed through immunostaining with
-SM actin
antibody of random samples (not shown). For other vascular beds, the
adventitial layer was stripped off, and only the media was used for
cell isolation. The tissues were minced into small pieces and
repeatedly incubated with collagenase type II (1 mg/mL;
Worthington) and elastase (0.5 mg/mL; Sigma) for 30 minutes at
37°C with rocking. The cells were collected after passage of the
digestion solution through a filter (pore size 70 µm; Becton
Dickinson). After the addition of 10% heat-inactivated FBS
and centrifugation, the cells were suspended in DMEM
supplemented with 10% FBS, 100 IU/mL penicillin, 100 µg/mL
streptomycin, and 2 mmol/L glutamine. The cells from all
digestions were pooled, and the viability was estimated in random
samples through trypan blue exclusion or MTT assay. Primary cells and
early passage cells (between 2 and 7) were used.
Adhesion and Growth Assays
Freshly isolated cells were plated at 20 000 cells/well in 10%
FBS with or without coatings (24-well plates). After 24 hours, the
wells were rinsed, and attached cells were trypsinized and counted in a
Coulter counter. The rate of adhesion was calculated as a percentage of
the initially plated cells. The following coatings were used to enhance
coronary SMC adhesion: Matrigel (10 µg/well; Becton
Dickinson), fibronectin (100 ng/well; Sigma), type I collagen (22
µg/well; Becton Dickinson), type IV collagen (22 µg/well; Becton
Dickinson), laminin (10 µg/well; Sigma), and vitronectin
(1 µg/well; Sigma). Briefly, the wells (24-well plates) were coated
with 200 µL of the solution at 4°C overnight with gentle rocking.
The excess of the coating solution was then removed, and the plates
were air dried for 10 minutes. In addition, the wells that were coated
with collagen, which was dissolved in 0.01 mol/L HCl, were rinsed twice
with DMEM. Each experiment was carried out in triplicate and repeated 3
times on different occasions with cells isolated from multiple donors.
Data represent mean±SD of 6 to 9 values from 3 separate
experiments.
For growth assay, the cells were plated at 20 000 or 100 000
cells/well (24-well plates) in DMEM supplemented with 10% FBS. At 2
days, cells were washed and fresh 10% FBS was added. Cells were then
trypsinized at 2, 4, 7, 9, and 14 days after plating and counted in a
Coulter counter. Because coronary SMCs did not adhere and grow
under these conditions, they were plated onto Matrigel-coated surfaces,
and various growth factors or their combinations were tested in
addition to 10% FBS: epidermal growth factor (EGF; 75 µg/mL; kindly
provided by Dr James San Antonio, Thomas Jefferson
University), platelet-derived growth factor (PDGF-BB; 100 ng/mL,
Sigma), insulin growth factor-1(IGF-1; 10 ng/mL; Sigma), and
interleukin (IL)-1 (1 ng/mL; Sigma). Each growth experiment was
carried out in triplicate for each time point and repeated 3 times on
different occasions with cells isolated from multiple donors. Data
represent mean±SD of 9 values from 3 experiments.
Protein Synthesis
For overall protein synthesis, cells were labeled with
35S-methionine (50 µCi/mL; DuPont-New England
Nuclear) for 40 hours (primary cells) or 24 hours (early passages). The
conditioned media and 2 washes were collected and subjected to TCA
(10%) precipitation. After being washed twice with 5% TCA,
radioactivity were determined with a beta scintillation counter
(Wallac). Protein synthesis was expressed as dpm per cell number. Data
represent mean±SD of 6 values derived from 3 experiments.
For collagen synthesis, subconfluent primary cells (200 000 cells) and confluent cells (passages 2 to 6) were labeled with 14C-proline (10 µCi/mL; DuPont-New England Nuclear) in DMEM containing 10% FBS and ascorbic acid (25 µg/mL) for 40 hours (primary cells) or 24 hours (confluent cells), respectively. The conditioned media and 2 washes were precipitated with 10% TCA at 4°C overnight. After centrifugation, the pellets were washed twice with 5% TCA and once with absolute ethanol. The pellets were dried and then hydrolyzed overnight with 0.45 mL of 0.2 mol/L NaOH. Radiolabeled collagen was quantified according to the collagenase digestion method.23 Briefly, hydrolyzed samples were divided into 2 aliquots (0.2 mL each) and neutralized by the addition of 0.2 mL of 0.08 N HCl. After the addition of N-ethylmaleimide (1.25 µmol/L), CaCl2 (0.25 µmol/L), and bacterial collagenases (25 µg/mL Clostridium histolyticum type III; Calbiochem), samples with a final volume of 0.5 mL were then incubated at 37°C for 5 hours. The control samples were treated without the addition of collagenases. The reaction was stopped by the addition of 0.5 mL of 20% TCA containing 0.5% tannic acid and BSA (100 µg/mL). Newly synthesized collagen was calculated as the difference between the radioactivity values of the samples treated, with or without collagenases (dpm/cell). The data represent mean±SD of 6 to 9 values derived from 3 experiments.
LDL Isolation, Modification, and Labeling
LDL (density, 1.020 to 1.063 g/mL) was isolated from fresh human
plasma through sequential
ultracentrifugation.24 25 LDL oxidation
(oxLDL) was achieved through the incubation of LDL (200 µg/mL) with
5 µmol/L CuSO4 at 37°C for 2 to 3 hours.
Oxidation was measured by monitoring the change of the diene absorption
(234 nm) and was stopped by the addition of 100 µmol/L EDTA and
40 µmol/L butylated hydroxytoluene. LDL was
iodinated with Na-125I (2 mCi;
DuPont-New England Nuclear) and eluted through PD-10 columns with 0.15
mol/L NaCl and 1 mmol/L EDTA.26 125I-LDL was
dialyzed for 48 hours at 4°C against 0.15 mol/L NaCl and 1
mmol/L EDTA. The activity of 125I-LDL ranged from
200 to 400 cpm/ng LDL protein with >98% of the
125I radioactivity precipitable by TCA and <5%
of the 125I radioactivity extractable with
chloroform-methanol. 125I-LDL was sterilized by
passage through a filter (pore size 0.45 µm), stored at 4°C
under argon, and used within 4 weeks.
For assessment of LDL degradation, freshly isolated cells were plated at 100 000 cells/well in DMEM supplemented with 10% human lipoprotein–deficient serum (for native LDL) or 10% FBS (for oxLDL) for 48 hours. Then, 125I-labeled native LDL or oxLDL was added to the cells for an additional 16 hours. The conditioned media were collected and subjected to TCA (10%) precipitation and chloroform extraction. The radioactivity values (125I-tyramine) in the aqueous phase were counted in a gamma counter (Wallac). The cells were lysed (0.1 mol/L NaOH), and cellular protein content was measured. Nonspecific LDL degradation (in the presence of 50-fold excess of unlabeled LDL and in cell-free wells) has been subtracted from all values. Data represent mean±SD of triplicate measurements. The experiments were repeated 3 times on separate occasions in both primary cells and cells at passages 2 to 7.
Immunostaining
Vascular cells were plated onto coverslips with or without
Matrigel coating. Coronary SMCs were grown on coverslips coated
with and without Matrigel. At different time points, the coverslips
were fixed with HistoChoice for 30 minutes and air dried. The
coverslips were stained with Vectastain Elite ABC system (Vector
Laboratories). They were incubated with primary antibodies for 1 hour
at room temperature, followed by biotinylated secondary horse
anti-mouse antibodies (1:2000; Vector Laboratories). Table 1
lists specific antibodies, the
concentrations used, and their sources. Negative controls included
either the omission of primary antibody or its replacement with
irrelevant mouse IgG (125 ng/mL). The immunostains were
visualized with the use of a diaminobenzidine tetrahydrochloride
substrate kit (Vector Laboratories) followed by hematoxylin
counterstaining.
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Statistical Analysis
Data are expressed as mean±SD values. One-way ANOVA was used to
compare the multigroup variables. If the F test results
were significant, Bonferroni’s analysis was carried out to
determine the differences among subgroups. A value of
P<0.05 was required to reject the null hypothesis.
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Results |
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Cell Morphology, Adhesion, and Growth
Enzymatic digestion yielded >80% viable cells regardless of vascular bed (trypan blue exclusion and MTT assay). As expected, medial cells isolated from the aorta and iliac artery displayed a spindle-shaped morphology, followed by a typical "hill-and-valley" pattern at confluence. In contrast, coronary SMCs demonstrated distinct morphology in culture (Figure 1
). Coronary adventitial cells
isolated from the same vessels, however, were spindle shaped after
plating and showed a hill-and-valley configuration at confluence, thus
displaying morphology similar to that of noncoronary SMCs
(Figure 1).
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10 days in culture. In contrast,
coronary adventitial cells and noncoronary medial cells
(aorta and iliac artery) displayed a spindle-shaped morphology. They
reached confluence at 7 to 10 days, when a typical hill-and-valley
pattern was observed (magnification 100x). FIB indicates
fibroblasts.
Porcine SMCs derived from noncoronary vascular media showed
rapid adhesion to plastic, which was comparable to that of
coronary adventitial fibroblasts (Figure 2A). This contrasted with
coronary medial SMCs, which showed significantly lower adhesion
(P<0.01). Because cell attachment is dependent on the
surrounding extracellular matrix proteins, we attempted to increase
coronary SM adhesion through the coating of culture surfaces
with several matrix proteins (Matrigel, collagen types I and IV,
laminin, vitronectin, fibronectin). As shown in Figure 2B, different extracellular matrix proteins, the use of EGF in
the culture medium, and the prolongation of adhesion time to 48 hours
produced only minimal improvement in the adhesion of coronary
SMCs.
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As expected, porcine noncoronary SMCs exhibited rapid growth in
the presence of 10% FBS, with exponential growth beginning at 2 days
(Figure 3A). These cells continued to
replicate after being subcultured for >10 passages and exhibited
similar growth on plastic or Matrigel-coated surfaces. In contrast, the
growth response of porcine coronary SMCs was markedly slower,
although coronary adventitial fibroblasts isolated from the
same vessels displayed a dynamic growth in culture (Figure 3A).
These differences were not species dependent, inasmuch as human
coronary SMCs also demonstrated lower growth capabilities
(Figure 3B). To determine whether specific growth factors are
required for coronary SMC growth, several growth factors (EGF,
PDGF-BB, IGF-1, or IL-1) were added to culture medium containing
10% FBS. Notwithstanding this stimulation, coronary SMCs
continued to demonstrate limited replication in culture (Figure 3C). To exclude the possibility that their slower growth was due
to low density, porcine coronary SMCs were also plated at high
density (100 000 cells/well), which failed to increase
coronary SMC growth or their ability to reach confluence after
>30 days in culture (not shown).
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Metabolic Labeling, Collagen Synthesis, and LDL
Metabolism
To compare metabolic activity in different vascular
cells, overall protein synthesis was assessed by
35S-methionine labeling. No major differences
were observed in total protein synthesis, which was consistent
with similar viability of primary porcine cultures (Figure 4A, P=NS). Because
"synthetic" phenotype of SMCs is marked by an increase in
extracellular matrix synthesis, de novo collagen synthesis was measured
with the use of 14C-proline incorporation,
followed by collagenase digestion. Noncoronary SMCs
from the aorta and iliac artery showed comparable collagen synthesis,
whereas coronary medial SMCs produced significantly less
collagen (Figure 4B, P<0.001). Coronary
adventitial fibroblasts from the same coronary arteries also
exhibited a higher ability to synthesize collagen than coronary
SMCs (P<0.001).
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The differences among vascular cells raised the question regarding
regional LDL metabolism. To this end, the ability of
vascular cells to degrade LDL was determined. Coronary SMCs
demonstrated significantly lower degradation of both native LDL
(P<0.01) and oxLDL (Figure 5A, P<0.001).
Coronary adventitial fibroblasts showed the most avid
metabolic processing of modified LDL, which exceeded values
observed in the coronary and noncoronary SMCs
(P<0.001). Differential LDL degradation was not species
dependent, because primary cultures of human vascular cells
demonstrated a similar pattern (Figure 5B). The avid degradation
of LDL by adventitial cells was maintained even in late passages
(passages 2 to 7; not shown).
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P<0.01
vs noncoronary SMCs by ANOVA with Bonferroni’s correction.
Abbreviations as in Figure 3
Expression of SM Differentiation Markers
To examine whether these characteristics of vascular cells are
related to their differentiation, several markers of SM differentiation
were assessed in primary cultures. There was pronounced
heterogeneity of -SM actin expression in
noncoronary SMCs (Table 2). As
illustrated in Figure 6, -SM actin
ranged from negative to strongly positive in subconfluent cells. The
markers of "late" SM differentiation, such as h-caldesmon and
SM-myosin heavy chain (MHC), also displayed
heterogenous distribution in noncoronary SMCs
(Figure 6). Likewise, a similar pattern was observed with SM1-
and SM2-MHC antibodies (not shown). In contrast, isolated
coronary SMCs demonstrated uniform expression of -SM actin
(Table 2). As shown in Figure 7, they lacked heterogenous distribution of cytoskeletal
markers (-SM actin, h-caldesmon, and SM-MHC) compared with
noncoronary SMCs. This highly differentiated phenotype
was present in cells cultured in 10% FBS for 1, 3, 5, and 10 days.
Figure 7 also illustrates the defective spreading of
coronary SMCs, which did not change in the presence of several
components of extracellular matrix (eg, vitronectin,
collagen types I and IV, or fibronectin; not shown).
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The vast majority of coronary adventitial fibroblasts were
devoid of -SM actin after isolation (Figure 8). This was followed by a dynamic
upregulation in -SM actin immunoreactivity in coronary
adventitial fibroblasts, which became almost uniformly positive for
-SM actin at 5 days, thereby acquiring the characteristics of
myofibroblasts.
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Discussion |
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Two major conclusions can be derived from the results of the present study. First, isolated coronary SMCs exhibit a highly differentiated phenotype, whereas noncoronary SMCs display greater phenotypic heterogeneity and more exuberant responses in culture (eg, proliferation, collagen synthesis, LDL degradation). Second, although coronary SMCs are less responsive than other SMCs, coronary adventitial nonmuscle cells demonstrate several phenotypic characteristics that enable them to react to pathophysiological stimuli.
Coronary Medial SMCs
Different morphology and gene expression, as well as distinct
proliferative and migratory capabilities, have been previously
identified among noncoronary SMCs.10 11 12 27 28 29
Whether the selective expansion of certain subtypes of SMCs or their
differential survival in culture accounted for phenotypic versatility
of noncoronary SMCs remains to be determined. Nevertheless,
coronary SMCs, isolated and cultured in an identical manner,
exhibited marked differences. Although differentiation and growth can
be dissociated in culture,30 31 coronary SMCs
continued to express differentiation markers and failed to replicate in
the presence of several growth factors (Figure 3
). To account
for possible species differences,32 33 34 we also examined
human coronary SMCs, which showed similar morphology and
proliferative activity as porcine coronary SMCs. It should be
underscored, however, that our findings do not necessarily contradict
previous reports describing the growth of -SM actin–positive cells
from human coronary arteries.35 36 The
particularly great ability of adventitial fibroblasts to migrate, to
overgrow medial SMCs, and to become myofibroblasts (Figure 8)
raises questions as to the origin of cells previously thought to be
SMCs.
The exact mechanism underlying differences between coronary and noncoronary SMCs remains to be determined. The development of coronary vessels in situ from the coelomic mesothelium, rather than from ectodermal or mesodermal portions of the aorta, may confer different cellular properties.19 22 In fact, cell type–specific transcriptional regulation of differentiation was recently emphasized by Owens et al.37 The extrinsic signals to be considered include extracellular matrix components (eg, laminin, heparan sulfate glycosaminoglycan) that modulate SMC differentiation.38 39 It is less likely, however, that regional variations in the composition of vascular extracellular matrix in situ were responsible for the observed differences after enzymatic cell isolation. We also emphasize that the effects of several other factors, which are absent in culture conditions, may promote dedifferentiation of coronary SM in vivo; these factors include age-, gender-, and environment-dependent phenomena (arterial retention of LDL or advanced glycosylation end products), which await clarifications in studies involving the coronary vasculature.40 41 42
Coronary Adventitial Nonmuscle Cells
The stimulation of coronary adventitial nonmuscle cells
resulted in phenotypic changes typically attributed to synthetic
SMCs.43 The expression of -SM actin by adventitial
nonmuscle cells exemplified the formation of myofibroblasts (Figure 8).44 45 The selective expansion of a small number
of SMCs in adventitial samples was unlikely, inasmuch as
coronary SMCs demonstrated poor attachment and growth (Figures 2 and 3). As shown in the present study, adventitial
nonmuscle cells displayed several properties necessary for
arterial repair (ie, high adhesion, proliferation, and
collagen synthesis). Recently, they have also been shown to possess
increased matrix-degrading activities and migration compared with
coronary SMCs, which constitutively express TIMP-1 and
-2.46 Notwithstanding these findings, we emphasize that
although the conditions for activation of nonmuscle cells likely exist
after acute mechanical medial injury in humans,47 the
migration of the cells during chronic medial damage is more difficult
to determine. The cytotoxic effects of modified LDL during
atherogenesis,42 however, often result in medial thinning,
which may enable nonmuscle cell translocation and their involvement in
lesion formation.48 49 The avid degradation of oxLDL by
activated coronary nonmuscle cells (Figure 5)
suggested a high expression of scavenger receptor and the possibility
of their involvement in foam cell formation on reaching the intima.
Clinical Implications
Disappointing results of several clinical studies that target
coronary restenosis have exemplified a low predictive
value of pharmacological testing in animal models of
noncoronary arterial injury. The presented
results and previous observations in vivo indicate greater
susceptibility of coronary adventitial fibroblasts to
mitogenic stimuli compared with coronary SMCs.
Whether the suggested involvement of adventitial cells will affect the
outcomes of antihyperplastic interventions applied in vivo (eg,
brachytherapy or pharmacological approaches) remains to be determined.
Interestingly, however, the adventitial delivery of some agents appears
to be more effective than endoluminal administration.50 We
postulate that a better understanding of the unique characteristics of
coronary vascular cells may provide the insight necessary for
future therapeutic interventions aimed specifically at a reduction in
coronary hyperplastic responses.
In conclusion, in the present study, we examined the characteristics of coronary medial SMCs and adventitial nonmuscle cells. Coronary SMCs, which differed from noncoronary SMCs, showed a highly differentiated phenotype with a limited ability to adhere, proliferate, and synthesize collagen. In contrast, adventitial nonmuscle cells displayed characteristics usually attributed to synthetic SMCs. We postulate that the mechanisms of vascular repair and lesion formation may differ among vascular beds, which points to the importance of future coronary artery–specific investigations in vascular biology.
Note Added in Proof
Since the submission of this manuscript, Christen et
al51 have also reported the presence of a highly
differentiated phenotype of coronary SMCs and the unique response of
these cells in culture.
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Acknowledgments |
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This study was supported by National Institutes of Health Grants HL-44150 and HL-60672 and the John S. Sharpe Foundation. The authors gratefully acknowledge Kenneth Margulies, MD, and Carol Fisher, BS (Temple University), for their assistance in the collection of human tissues. We thank Mary Ellen Henderson for her assistance in preparation of the manuscript.
Received May 14, 1999; revision received July 30, 1999; accepted August 13, 1999.
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