(Hypertension. 1995;25:219-226.)
© 1995 American Heart Association, Inc.
Articles |
Cellular Distribution of Angiotensin-Converting Enzyme After Myocardial Infarction
From the Department of Pharmacology, University of Kiel (Germany) (M.F., Y.-C.Z., H.M.S., T.U.); the Department of Pathology, University of Giessen (Germany) (F.F., R.M.B.); the Department of Anatomy and Cell Biology, University of Heidelberg (Germany) (S.B.); and the National Cardiology Research Center, Moscow, Russia (S.D.).
Correspondence to Mechthild Falkenhahn, Department of Pharmacology, University of Kiel, Hospitalstr. 4, 24105 Kiel, FRG.
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Abstract |
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Abstract We studied the cellular distribution of angiotensin-converting enzyme (ACE) in the heart related to the cell types involved in left ventricular repair and remodeling before and after myocardial infarction by immunohistochemical techniques using monoclonal and polyclonal antibodies. In noninfarcted myocardium of both human and rat, ACE expression was confined to endothelial cells and subendocardial cell layers of the aortic valve. ACE was prominent in endothelia of small arteries and arterioles, whereas only half the coronary capillaries were immunoreactive and venous vessels were almost completely devoid of the enzyme. In a rat model of myocardial infarction, ACE distribution was determined 1, 3, and 7 days and 2, 3, and 6 weeks after coronary occlusion. Three and 7 days after infarction, endothelial cells of sprouting capillaries and macrophages in the marginal zone of necrosis revealed ACE expression. In both human and rat with the onset of fibrosis, intense staining of the enzyme was found in the marginal zone of the repair tissue. In situ hybridization for collagen type I in the rat revealed that zones with high collagen content had almost no ACE immunoreactivity. Vascular smooth muscle cells and cardiomyocytes revealed no ACE expression throughout the study. We conclude that endothelial cells are the principal source for the expression of ACE after myocardial infarction. The observed induction of ACE with the onset of fibrosis suggests a role of this enzyme that is related to tissue repair and remodeling.
Key Words: angiotensin-converting enzyme • heart • myocardial infarction • endothelium • macrophages
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Introduction |
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Angiotensin-converting enzyme (ACE) is a key element of the renin-angiotensin and kallikrein-kinin systems. ACE takes part in the various functions of both systems by generating angiotensin II (Ang II) and degrading bradykinin. Advances in understanding the structure and regulation of somatic ACE have been facilitated by the molecular cloning of the ACE gene1 2 3 4 and by the recently reported functional analysis of the human and rabbit ACE gene promoter sequence.5 6 The presence of ACE in the rat heart is well documented. With the use of quantitative autoradiography, the binding of radioligands revealed ACE expression in different areas of the heart, including cardiac valves, coronary vessels, atria, and myocardium.7 8 Several earlier experimental studies in pressure-overloaded left ventricular hypertrophy and experimental heart failure showed an induction of ACE activity and ACE mRNA synthesis.9 10 11 12 This tissue-specific induction of the ACE gene may contribute to harmful effects of the renin-angiotensin system in cardiac diseases. Clinical studies have corroborated this hypothesis. Thus, administration of ACE inhibitors after myocardial infarction (MI) not only reduced the development of heart failure but also the risk of recurrent MI and improved the survival of patients with heart failure.13 The time of initial ACE treatment after MI seems to be critical because a study with early treatment within 24 hours after MI could not confirm the beneficial effects of ACE inhibitors.14 Moreover, experimental results showed that cardiac function improved in rats treated with captopril 3 to 5 weeks after MI, whereas cardiac function of rats treated 1 to 21 days after MI deteriorated.15
Intense efforts have been made to find out whether the beneficial effects of ACE inhibitors in heart failure can be attributed to local rather than systemic effects of the renin-angiotensin and kallikrein-kinin systems. Evidence for both bradykinin- and Ang II–dependent local effects has been obtained.16 17 18 These and other studies revealed that the functions of cardiac ACE go beyond blood pressure control and may be associated with hypertrophy, fibrosis, and inflammatory response.
The cells responsible for ACE induction associated with MI have not yet been identified. Therefore, we studied the cellular distribution of ACE in human and rat heart with immunohistochemical staining techniques and then analyzed the time-dependent changes in ACE distribution at different times during ventricular remodeling after MI. Using cell type–specific markers, we further characterized the cell types involved in ACE induction. Our results demonstrate that endothelial cells are mainly responsible for an increase in ACE generation after MI.
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Methods |
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Rat MI Model
MI was induced in rats by permanent ligation of the left descending coronary artery with a modified technique described by Johns and Olson.19 Briefly, after induction of ether anesthesia, male Wistar rats (Dr Karl Thomae GmbH, Biberach, Germany) weighing 250 to 300 g were intubated, artificially respirated, and connected to an electrocardiogram recorder. The left descending coronary artery was ligated with sterile suture material (Ethibond 6-0, Ethicon) under a stereomicroscope. In sham-operated rats, ligations were placed beside the coronary artery. Successful ligation of the coronary artery was verified by the occurrence of arrhythmia in the electrocardiogram and visually by the color change of the ischemic area. Mortality within 24 hours was less than 40%.
Preparation of Tissue Samples
Rats were killed 24 hours, 3 days, 7 days, 2 weeks, 3 weeks, and
6 weeks after MI (n=3 at each time). Infarcted and sham-operated hearts
were removed, snap-frozen, and stored in liquid nitrogen. Corresponding
horizontal cross sections of whole rat hearts with coronary occlusion
and of the sham-operated hearts were stained. Comparable cross sections
inferior to the level of cardiac valves were analyzed. These sections
allowed the analysis of both left and right ventricular walls
(infarcted and noninfarcted). With very little interindividual
variability, the necrotic area was transmural; thus, the margin of the
infarcted area was sharply demarcated from the intact
myocardium.18 Sham-operated hearts showed no areas of
necrosis (not shown).
ACE staining persistence was tested in six myocardial autopsy specimens between 6 and 48 hours postmortem. ACE reactivity was found to be stable for up to 48 hours postmortem. No diffusion artifacts appeared within the media of the arteries. Infarcted human myocardium was taken from eight patients who had not been treated with ACE inhibitors; noninfarcted human myocardium was taken from eight patients without a history of hypertension or cardiovascular diseases. Tissue samples were dissected at autopsy within 24 hours postmortem (16 women and men; 28 to 65 years old), snap-frozen, and stored in liquid nitrogen. Serial cryostat sections (5 µm) of freshly frozen tissue were mounted on slides, air-dried, and stored at -20°C.
Monoclonal Antibodies and Antisera
Six monoclonal antibodies (MAbs) against different epitopes of
the N-domain of human ACE were tested for immunohistochemical staining
of ACE in human and rat myocardial sections (clones: 9B9, 0.5 µg/mL;
i2H5, 1 µg/mL; 3G8, 5 µg/mL; 5F1, 10 µg/mL; 3A5, 7 µg/mL; i1A8,
30 µg/mL; all mouse IgG1).20 21 We also
stained human myocardial sections with a polyclonal rabbit anti-human
ACE antiserum (Y4, 1.7 µg/mL; kindly provided by
Dr F. Alhenc-Gelas)22 and rat myocardial sections with a
polyclonal rabbit anti-mouse ACE antiserum diluted 1:200 (kindly
provided by Dr K.E. Bernstein).4 ACE-positive cells were
characterized by double-staining techniques using cell type–specific
markers: CD31 (1:100), mouse IgG1 anti-human endothelial
cell (Dako); MRC OX-43 (1:1000), mouse IgG1 anti-rat
endothelial cell (Serotec)23 ; CD68 (1:50), mouse
IgG3 anti-human macrophage (Dako); and Ki-M2R (1:50), mouse
IgG1 anti-rat macrophage (Dianova).24 For
staining of smooth muscle cells and cardiac
fibroblasts,25 26 we used mouse anti-human 
-actin IgG
(1:400) (Sigma Chemical Co) and polyclonal goat anti-human collagen
types I, II, and III antibodies (1:50) (Southern Biotechnology
Associates, Inc). For negative control, we used mouse anti-rabbit
immunoglobulin (clone: MR12/52, 1 µg/mL; Dako).
Immunohistochemistry Techniques
Immunoenzymatic detection was mainly performed by the highly
sensitive alkaline phosphatase anti-alkaline phosphatase (APAAP)
technique, using a slightly modified method of Cordell et
al.27 In brief, 5-µm frozen tissue sections were
air-dried and fixed in acetone for 10 minutes at room temperature.
Sections were incubated with the primary MAb and then with rabbit
anti-mouse immunoglobulin (rabbit-"link," 1:40) (Dako)
supplemented with either 12.5% pooled human serum or 25% pooled rat
serum to inhibit unspecific cross-reactivity. This was followed by an
incubation with the APAAP complex (1:50) (Dako). Each step lasted 30
minutes at room temperature. Samples were thoroughly washed in
Tris-HCl–buffered saline (pH 7.6) between steps. The
rabbit-"link" and APAAP complex steps were then repeated for 10
minutes each at room temperature. Alkaline phosphatase substrate
reaction with new fuchsin (100 µg/mL) and levamisole (400 µg/mL)
was performed for 20 minutes at room temperature. Sections were
counterstained with hematoxylin and mounted in gelatin.
The APAAP technique was combined either with labeled streptavidin biotin (LSAB) or with silver intensified immunogold for cell type characterization. The LSAB technique was applied according to the manufacturer's protocol (Dako). The silver intensified immunogold technique was performed as previously described.28 For double-staining immunofluorescence, sections were incubated with a mixture of monoclonal mouse and polyclonal rabbit primary antibodies for 45 minutes at 37°C (concentrations 10 to 20 times higher than in APAAP), rinsed in phosphate-buffered saline, and incubated for 30 minutes at 37°C with a mixture of fluorescein-conjugated goat anti-mouse immunoglobulins (FITC, Cappel) and rhodamine-conjugated pig anti-rabbit immunoglobulins (TRITC, Dako). Sections were rinsed in phosphate-buffered saline and mounted in Mowiol (E Merck).
In Situ Hybridization
For preparation of the cRNA probe, a pGEM3Z transcription vector
containing a 360 bp fragment of the human collagen type I cDNA (kindly
provided by Dr K. von der Mark) was used. The sense and anti-sense cRNA
probes were synthesized and labeled by in vitro transcription using
digoxigenin (DIG)–11-UTP (Boehringer). The in situ hybridization was
performed as previously described.29 Briefly, after
prehybridization steps, each section was covered with 20 µL
hybridization mixture (appropriate concentration of labeled probe, 0.5
to 5 ng/µL). After hybridization at 41°C for 16 to 18 hours, slides
were washed in 2x sodium saline citrate (SSC) for 30 minutes at room
temperature followed by washes in SSC containing 50% formamide at
41°C (1x SSC, 1 hour; 0.5x SSC, 1 to 2 hours; and 0.1x SSC, 1
hour) and a final wash in 0.1x SSC at room temperature for 20 minutes.
Detection of the signal was performed using a Boehringer DIG detection
system. For a combination of in situ hybridization and indirect
immunofluorescence, a combined labeling protocol was established on
cryostat sections that first went through the regular in situ
hybridization protocol as described above. Both sheep anti-DIG
conjugate (1:500) and rabbit anti-mouse ACE antiserum (1:50) diluted in
blocking medium were administered to the sections for 1 hour at room
temperature followed by 12 to 18 hours at 4°C. Slides were rinsed in
buffer 1 (100 mmol/L Tris-HCl, 150 mmol/L NaCl, pH 7.5) for 30 minutes
and then incubated with fluorescein-conjugated anti-rabbit
immunoglobulins (1:100) for 2 hours at room temperature. Sections were
rinsed in buffer 1 for 30 minutes and then processed as in the regular
in situ hybridization protocol29 for visualization of
anti–DIG-phosphatase color reaction.
Reagents and Chemicals
Restriction enzymes were obtained from GIBCO and RNA polymerase
from Promega. All other chemicals were obtained from Sigma Chemical
Co.
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Results |
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We determined the cellular distribution of ACE in human and rat myocardium by APAAP immunohistochemistry. All MAbs against human ACE revealed identical reactivity patterns on human heart sections in the concentrations used (not shown). Only one of the MAbs (9B9) reacted with rat ACE. Both polyclonal antibodies and MAbs against ACE showed identical reaction patterns (Fig 1A, 1B, and 1C). ACE expression was confined to endothelial cells in noninfarcted myocardium; however, staining intensity of endothelial cells differed remarkably with respect to vessel type. Endothelia of nearly all small arteries and arterioles revealed strong ACE staining (Fig 1B), whereas only some capillaries were ACE positive, and venous vessels showed almost no staining (see arrows, Fig 1A, 1C, 1E, and 1F). The subendothelial cells of both rat and human cardiac valves were stained ACE positive, whereas most of the valvular endothelial cells were obviously ACE negative (Fig 1D). Although these ACE-positive cells had fibroblast morphology and were negative for endothelial cell markers, they failed to demonstrate a conclusive coexpression of
-actin and
collagen.
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To study ACE expression in the progress of left ventricular repair and remodeling, we analyzed the cellular ACE distribution at different times after MI. Twenty-four hours after MI, no ACE was found in areas of necrosis. This probably resulted from the destruction of capillaries and vascular structures within the infarcted area. Immigrated granulocytes and macrophages were ACE negative. Three days after MI, few ACE-positive cells appeared within the granulation tissue demarcating the zone of necrosis (Fig 2A). Further organization of the necrotic zone and the influx of sprouting capillaries 7 days after MI were associated with an intensified ACE staining (Fig 2B). ACE could mainly be localized in endothelial cells. Double-staining technique for ACE and Ki-M2R macrophage antigen revealed that some of the rat macrophages in this area also expressed ACE (Fig 2C and 2D). Similar results were found in human infarcted myocardium (Fig 2E).
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Three weeks after MI, we found an induction of ACE expression within
the newly formed fibrous repair tissue (Fig 3A and 3B).
Areas with intense ACE staining had a high density of reconstituted
cardiac vessels and capillaries and some ACE-positive macrophages (Fig 3B, upper left). Some of the ACE-positive cells looked like typical
fibroblasts with tapering cell extensions. To further characterize
these cells, we stained serial sections with anti-ACE, anti–-actin,
and anti-collagen types I, II, and III, respectively. Although
anti–collagen II and III staining was negative, areas moderately
stained for both -actin and collagen type I also showed some diffuse
staining for ACE (Fig 3B, 3C, and 3D), suggesting that not only
endothelial cells but also some cardiac fibroblasts in these areas of
fibrosis expressed ACE. However, a combination of in situ hybridization
and immunohistochemical staining revealed that fibroblasts with a
strong expression of collagen type I mRNA within the infarcted tissue
were ACE negative (Fig 4). Interestingly, smooth muscle
cells of cardiac vessels in the fibrous scar tissue showed no ACE
expression throughout our study (Fig 3C). A comparable ACE distribution
pattern was seen 6 weeks after MI in rat hearts and in corresponding
human MI tissue (not shown). At no time before and after MI was ACE
staining found in cardiomyocytes.
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Discussion |
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In recent years, intensive efforts have been made to better define the physiological role of cardiac ACE. To localize ACE expression in the heart, several investigators have used autoradiographic methods.7 8 However, these methods fall short in specifying the cellular site where ACE is synthesized and acting. Our results were obtained using highly sensitive immunohistochemical staining techniques, so we were able to study ACE expression after MI in relation to specific cell types.
In noninfarcted myocardium of both human and rat, ACE was confined to endothelial cells. ACE expression was clearly accentuated in endothelia of small arteries and arterioles, whereas only about half the coronary capillaries were ACE positive and endothelial cells of venous vessels were almost completely ACE negative. Since MAbs often show a low antigen affinity, we also tested a polyclonal antibody against ACE and found identical staining for both MAbs and polyclonal antibodies against ACE. These results demonstrate striking quantitative differences with respect to ACE expression in cardiac endothelial cells.
Our findings are consistent with the previous observation that ACE is differentially expressed within the vascular tree, with high levels of the enzyme in the precapillaric coronary microcirculation and decreasing concentrations toward capillaries and the venous system30 (unpublished observations). Endothelial cells are able to modify their ACE expression. Various stimuli that activate ACE synthesis in bovine aortic endothelial cells are known.4 31 Furthermore, positive and negative regulatory elements of the human ACE gene promoter have been shown to determine ACE regulation in endothelial cells in vitro.5 6 The coronary autoregulation might give an explanation for the heterogeneous pattern of ACE expression observed in cardiac capillaries. Under resting conditions, autoregulation results in the perfusion of only about half the available capillaries.32 Resting endothelial cells of capillaries, which are temporarily excluded from the circulating bloodstream, have a minimized supply of ACE substrates and for economic reasons might reduce their ACE expression by minimizing their protein synthesis on the maintenance of basal turnover. ACE expression in vascular endothelial cells thus would be related to their functional status.
In the rat model of MI, we found a few ACE-positive macrophages 3 to 7 days after MI in the marginal zone of necrosis and also later, 3 to 6 weeks after MI, in the fibrous repair tissue. Although mononuclear cells synthesize none or very little ACE, the expression of this enzyme is induced when the cells transform to macrophages.33 It has also been reported that serum ACE and ACE localized in intracellular secretory compartments are involved in the processing of antigens major histocompatibility complex class I–restricted T lymphocytes.34 35 This raises the interesting possibility that ACE may be involved in mechanisms of activation and development of inflammatory cells. Since Ki-M2R recognizes activated macrophages,24 it appears that the few macrophages that were found to be ACE positive were at a certain stage of maturation in which they express ACE.
It is presently under discussion whether treatment with ACE inhibitors can affect the inflammatory response after MI. Inhibition of local ACE in capillaries of granulation tissue and surrounding myocardium will lead to decreased breakdown of bradykinin and to reduction of Ang II concentration, resulting in an improved blood supply to the marginal zone of necrosis.18 Bradykinin is thought to play an important role in the inflammation response soon after coronary occlusion and in cardiac ischemia in isolated hearts, when the local kinin system in the heart has been shown to be activated.36 Several studies have demonstrated that the reduction of infarction size by ACE inhibitors can be abolished by a specific bradykinin (B2) receptor antagonist.18 37 Moreover, even low doses of ACE inhibitors have been found to improve cardiac blood flow in hypertrophied myocardium, an effect that may be related to an increased capillary length density.38 39
Apart from its role in the regulation of coronary flow, ACE might be involved in mechanisms of tissue repair. In contrast to our results, Rakugi et al40 have demonstrated recently that vascular smooth muscle cells are responsible for an induction of ACE expression after intima injury. Neointima formation might also be related to elevated Ang II levels, because ACE inhibitors have been shown to prevent neointima proliferation.41 Thus, Ang II is no longer considered solely to be a potent vasoconstrictor but also to be an effective agent in the regulation of cell proliferation and extracellular matrix production.42 43 44 Thus, Ang II might play an important role during structural changes in cardiac disease.
In our model of MI, high levels of ACE expression in the repairing scar
tissue appeared with the onset of fibrosis. An induction of cardiac ACE
can be expected to lead to considerably enhanced Ang II formation.
Stimulation of growth in myocytes and fibroblasts through direct
Ang II effects has been discussed as a basis for compensatory
ventricular remodeling.17 In cardiac fibroblasts, elevated
Ang II levels have been shown to increase both the mRNA expression and
protein synthesis of type I collagen.45 Therefore, high
Ang II concentrations in the fibrous scar tissue after MI might
stimulate fibroblast growth and collagen synthesis. Indeed, it has been
reported that treatment with the angiotensin type 1 receptor antagonist
losartan resulted in a reduction of heart weight and collagen content
after MI.16 Our findings using a double-staining technique
show that ACE induction 3 and 6 weeks after MI was mainly caused by an
increase of the microvascular endothelial compartment of the infarcted
zone. Double staining for anti–collagen I and anti-ACE revealed that
areas of high collagen content were low in ACE immunoreactivity. The
combined detection of collagen type I mRNA and ACE immunoreactivity
demonstrated that collagen-generating fibroblasts did not contain ACE.
However, these fibroblasts lie close to ACE-positive vascular
endothelia. We cannot rule out at present the possibility that
certain fibroblasts not detected by collagen types I, II, or III or
-actin staining (known to be synthesized by mature cardiac
fibroblasts) might participate in ACE expression.25 26
In experimental heart failure, tissue-specific ACE induction has been observed not only in scar tissue after infarction but also in unaffected atria and ventricles of the rat heart. In addition, the participation of cardiomyocytes in ACE induction associated with cardiac disease such as left ventricular hypertrophy has been discussed by some investigators.9 46 In our model of MI, ACE staining in the unaffected myocardium was confined to the vascular endothelial cells and was clearly not present in cardiomyocytes. ACE induction after MI was principally localized in the vascular endothelium. Further investigation is needed to clarify whether the ACE induction in hypertrophied myocardium reported by other authors is based on an overall induction of ACE in cardiac endothelial cells or whether the recruitment of capillaries due to a higher oxygen demand32 is responsible for the ACE upregulation.
In conclusion, our data demonstrate that the expression of cardiac ACE under normal conditions is restricted to endothelial cells and that the level of ACE activity in the heart after MI is mainly regulated by this cell type. Apart from some activated macrophages and some non–collagen-producing fibroblastic cells, all other cell types failed to show evidence of ACE synthesis. The observed ACE induction with the onset of fibrosis indicates a role of ACE that goes beyond blood pressure control and may be related to tissue repair and remodeling. It remains to be investigated which mechanisms are involved in the ACE-related structural changes after MI.
Received June 28, 1994; first decision September 8, 1994; accepted October 19, 1994.
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