Published online before print December 31, 2008, doi: 10.1161/CIRCULATIONAHA.108.773424
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
(Circulation. 2009;119:269-280.)
© 2009 American Heart Association, Inc.
Heart Failure |
Pressure Overload–Induced Alterations in Fibrillar Collagen Content and Myocardial Diastolic Function
Role of Secreted Protein Acidic and Rich in Cysteine (SPARC) in Post–Synthetic Procollagen Processing
From the Gazes Cardiac Research Institute, Division of Cardiology, Department of Medicine, RHJ Department of Veterans Affairs Medical Center, Medical University of South Carolina, Charleston.
Reprint requests to Amy D. Bradshaw, PhD, Medical University of South Carolina, Department of Medicine, Division of Cardiology, 114 Doughty St, Room 223, Gazes/Strom Thurmond Research Bldg, Charleston, SC, 29425. E-mail bradshad{at}musc.edu
Received February 14, 2008; accepted October 10, 2008.
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
Background— Chronic pressure overload causes myocardial hypertrophy, increased fibrillar collagen content, and abnormal diastolic function. We hypothesized that one determinant of these pressure overload–induced changes is the extracellular processing of newly synthesized procollagen into mature collagen fibrils. We further hypothesized that secreted protein acidic and rich in cysteine (SPARC) plays a key role in post–synthetic procollagen processing in normal and pressure-overloaded myocardium.
Methods and Results— To determine whether pressure overload–induced changes in collagen content and diastolic function are affected by the absence of SPARC, age-matched wild-type (WT) and SPARC-null mice underwent either transverse aortic constriction (TAC) for 4 weeks or served as nonoperated controls. Left ventricular (LV) collagen content was measured histologically by collagen volume fraction, collagen composition was measured by hydroxyproline assay as soluble collagen (1 mol/L NaCl extractable) versus insoluble collagen (mature cross-linked collagen), and collagen morphological structure was examined by scanning electron microscopy. SPARC expression was measured by immunoblot. LV, myocardial, and cardiomyocyte structure and function were assessed by echocardiographic, papillary muscle, and isolated cardiomyocyte studies. In WT mice, TAC increased LV mass, SPARC expression, myocardial diastolic stiffness, fibrillar collagen content, and soluble and insoluble collagen. In SPARC-null mice, TAC increased LV mass to an extent similar to WT mice. In addition, in SPARC-null mice, TAC increased fibrillar collagen content, albeit significantly less than that seen in WT TAC mice. Furthermore, the proportion of LV collagen that was insoluble was less in the SPARC-null TAC mice (86±2%) than in WT TAC mice (99±2%, P<0.05), and the proportion of collagen that was soluble was greater in the SPARC-null TAC mice (14±2%) than in WT TAC mice (1±2%, P<0.05) As a result, myocardial diastolic stiffness was lower in SPARC-null TAC mice (0.075±0.005) than in WT TAC mice (0.045±0.005, P<0.05).
Conclusions— The absence of SPARC reduced pressure overload–induced alterations in extracellular matrix fibrillar collagen and diastolic function. These data support the hypothesis that SPARC plays a key role in post–synthetic procollagen processing and the development of mature cross-linked collagen fibrils in normal and pressure-overloaded myocardium.
Key Words: collagen • proteins • heart failure • hypertrophy • diastole
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Chronic pressure overload, such as occurs with arterial hypertension, is a common cause of left ventricular (LV) remodeling and frequently leads to the development of chronic heart failure.1–3 Pressure overload–induced remodeling is characterized by the development of cardiomyocyte hypertrophy, an increase in extracellular matrix (ECM) fibrillar collagen content, and the development of abnormal diastolic function.3–6 However, the mechanisms by which pressure overload leads to cardiac remodeling, particularly a net increase in myocardial collagen content and the development of diastolic dysfunction, have not been completely defined.
Clinical Perspective p 280
Fibrillar collagen biosynthesis begins within a fibroblast with the synthesis of procollagen 
-chain monomeric proteins, which then form the triple-helical structure of a procollagen molecule7 (Figure 1). After synthesis, a procollagen molecule is secreted into the extracellular space, where it must undergo a series of ordered, time-sensitive, and location-sensitive processing steps to become a mature cross-linked insoluble structural collagen fibril. We hypothesized that one fundamental mechanism by which chronic pressure overload increases myocardial fibrillar collagen content and causes the development of abnormal diastolic function is an alteration in post–synthetic procollagen processing. We further hypothesized that one determinant of post–synthetic procollagen processing is the extracellular protein secreted protein acidic and rich in cysteine (SPARC).
|
Previous studies in noncardiovascular tissues have shown that SPARC is a procollagen-binding protein with counteradhesive activity and that it participates in the coordination of procollagen processing and facilitates the formation and assembly of mature cross-linked insoluble structural collagen fibrils.8,9 Studies in dermal tissue and dermal fibroblasts support the hypothesis that when SPARC binds to newly secreted procollagen, it chaperones the procollagen molecule through processing steps within the extracellular space (Figure 1) and limits binding of procollagen to cell-surface receptors.10–13 By diminishing procollagen engagement of cell surface receptors, SPARC facilitates the optimal sequence and timing of the processing steps and prevents procollagen from being degraded prematurely or processed improperly.10 SPARC is not absolutely required to process procollagen, but SPARC enhances and facilitates procollagen processing. In transgenic mice that do not express SPARC (SPARC-null mice), there was a substantial decrease in dermal collagen concentration, collagen fibrils were significantly smaller, and the dermis had a significantly decreased tensile strength (and decreased stiffness) compared with dermis from wild-type (WT) mice.11
However, post–synthetic procollagen processing has not been studied extensively in myocardial tissue. It is not known whether SPARC is an important determinant of post–synthetic procollagen processing in the myocardium or whether procollagen processing is altered in the myocardium of the pressure-overloaded LV. It is also not known whether a change in SPARC expression is a fundamental mechanism by which pressure overload increases myocardial fibrillar collagen content and causes diastolic dysfunction. Accordingly, the purpose of the present study was to test the hypothesis that (1) SPARC is an important determinant of myocardial fibrillar collagen content in normal myocardium, (2) SPARC is significantly increased in pressure-overloaded myocardium, and (3) the absence of SPARC significantly alters the effects of pressure overload on myocardial fibrillar collagen content and diastolic function.
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Protocols
Four groups of mice were studied (WT control, WT transverse aortic constriction [TAC], SPARC-null control, and SPARC-null TAC) with 4 protocols (A through D; Figure 2). Transgenic mice that do not express SPARC (SPARC-null mice, produced by targeted gene deletion) were generated on a C57Bl6/SV129 background. SPARC-null mice were compared with their WT littermate mice (C57Bl6/SV129) both in the control state and after TAC. To determine whether the absence of SPARC resulted in a change in cardiac phenotype, nonoperated WT and SPARC-null control mice underwent echocardiography and then were randomly assigned to 1 of 4 experimental protocols (A through D). To determine whether the absence of SPARC altered the phenotypic response to TAC-induced pressure overload, WT mice and SPARC-null mice underwent TAC at 4 weeks, followed by echocardiography and random assignment to 1 of 3 experimental protocols (A, B, or C). The methods used to perform echocardiography14 and TAC have been described previously and are presented in detail in the online-only Data Supplement.
|
Twenty-nine nonoperated control SPARC-null mice (16 males, 13 females) and 29 WT mice (15 males, 14 female) were studied at 12 weeks of age and were entered into protocols A (n=10 SPARC null, 10 WT), B (n=7 SPARC null, 7 WT), C (n=6 SPARC null, 6 WT), and D (n=6 SPARC null, 6 WT). Forty-three SPARC-null (22 male, 21 female) and 42 WT (22 male, 20 female) mice were subjected to TAC surgery at 12 weeks of age. Twenty-six SPARC-null (14 male, 12 female) and 18 WT (10 male, 8 female) mice survived 4 weeks of TAC and were entered into protocols A (n=14 SPARC null, 8 WT), B (n=6 SPARC null, 5 WT), and C (n=6 SPARC null, 6 WT). All procedures performed were approved by the Medical University of South Carolina Institutional Animal Care and Use Committee in accordance with National Institutes of Health guidelines.
Protocol A
Mice were anesthetized with inhalation isoflurane and given heparin 200 U IP. The LV was isolated, weighed, divided into 2 pieces, and frozen in liquid nitrogen. Two sets of analyses were performed: (1) Collagen composition was determined by biochemistry, and (2) SPARC abundance was determined by immunoblot.
Extraction and Biochemical Quantification of Collagen
Frozen LV tissue was lyophilized, weighed (dry weight), pulverized, resuspended in 1 mol/L NaCl with protease inhibitors, tumbled overnight at 4°C, and centrifuged. The supernatant then contained the salt-soluble collagen (ie, largely non–cross-linked collagen); the pellet contained the salt-insoluble collagen (fully mature cross-linked fibrillar collagen). Each collagen fraction was processed separately. Collagen fractions underwent complete acid hydrolysis with 6N HCl for 18 hours at 120°C, and then each was neutralized to pH 7 with 4N NaOH. One milliliter of chloramine T was added to 2-mL volumes of collagen sample and incubated at room temperature for 20 minutes. One milliliter of Ehrlich’s reagent (60% perchloric acid, 15 mL of 1-propanol, 3.75 g of p-dimethyl-aminobenzaldehyde in 25 mL) was added, and samples were incubated at 60°C for 20 minutes. Absorbance at 558 
was read on a spectrophotometer. Collagen was quantified as micrograms of hydroxyproline per milligram of dry weight LV myocardium. Total collagen was calculated as the sum total of the salt-soluble and -insoluble fractions. Thus, 3 collagen measurements were made by this hydroxyproline quantification of differentially isolated collagen fractions: Salt-insoluble collagen (mature, fully processed cross-linked collagen) and salt-soluble collagen (not processed or incompletely processed but non–cross-linked collagen) and total collagen (the sum of soluble plus insoluble collagen).
Western Blot Analysis
Immunoblots were performed on LV tissue samples extracted with 1 mol/L NaCl to obtain the salt-soluble fraction or LV tissue samples treated with Triton extraction buffer. SDS-PAGE gels were probed with murine anti-SPARC polyclonal antibodies (1:20 000 dilution, R&D Systems, Minneapolis, Minn).
Protocol B
Mice were anesthetized and given heparin 200 U IP. The LV was isolated, the aorta cannulated, the LV perfused with 2,3-butanedione monoxime, and the papillary muscle isolated to determine in vitro myocardial systolic and diastolic function.
Papillary Muscle Preparation and Myocardial Function Measurements
Previously, our laboratory performed isolated papillary muscle studies in normal and pressure-overloaded cats and rats.15–17 The methods used in these studies were modified for the study of murine papillary muscles (Data Supplement). Passive diastolic stiffness was examined in 2 ways: (1) by defining rest stress at the length (Lmax) at which maximum active tension was developed and 2) by performing a muscle stretch at a very slow stretch rate (1 mm/min) beginning from near slack length (very lightly preloaded muscle at 0.1 g) to a muscle length of 15% greater than that at slack length (equivalent to Lmax preload). The myocardial stress-versus-strain relationship during this muscle stretch was used to calculate the passive stiffness constant, β, as stress=Ae(βStrain)+C, where A and C are curve-fitting constants. Myocardial stress was calculated from muscle force divided by muscle cross-sectional area, and strain was calculated as L–L0/L0, where L=muscle length during stretch and L0=muscle length at 0.1 g of preload.
Protocol C
Mice were anesthetized and given heparin 200 U IP. The LV was isolated and the aorta cannulated, and the LV was divided in half. One half of the LV was fixed in 4% paraformaldehyde and stored in 4% paraformaldehyde at 4°C to undergo quantitative collagen content analysis with light microscopy studies. The other half of the LV was fixed in 2% glutaraldehyde to undergo morphometric analysis of fibrillar collagen with scanning electron microscopy studies.
Collagen Content by Light Microscopy
LV sections were stained with picrosirius red (PSR) to detect collagen fibers and were viewed with polarized light under dark field optics to detect birefringence of collagen fibers. Quantitative analysis of PSR-stained images captured with polarized light was performed. Five fields chosen at random from each mouse were scanned with SigmaScan software. Fields with large blood vessels were excluded from the analysis. Areas examined were distributed throughout the myocardium from the subendocardium to the subepicardium and excluded the epicardial surface. Collagen volume fraction (CVF) was calculated as the area stained by PSR divided by the total area of interest using previously published techniques.18 PSR birefringent staining was used to identify and quantify mature, fully processed cross-linked insoluble fibrillar collagen within the myocardium.19,20 Additional studies confirmed that PSR birefringent staining does not detect procollagen and is specific for mature collagen fibrils; these are presented in the Data Supplement.
Collagen Morphological Structure by Scanning Electron Microscopy
LV samples were processed according to previously published techniques, which included immersion fixation in 2% cacodylate glutaraldehyde, postfixation in 2% osmium tetroxide, dehydration with 100% alcohol, and drying with hexamethyldisilazane with a critical point dryer.21 Samples were mounted on scanning electron microscopy stubs, sputter-coated with gold palladium, and imaged with a JEOL JSM-5410 scanning microscope (JEOL Ltd, Tokyo, Japan) at 15 kV, with lens current set at 46 and the working distance set at 14.
Protocol D
Tail-cuff arterial blood pressure was measured, then LV cardiomyocytes were isolated by collagenase techniques, and cardiomyocyte systolic and diastolic function was assessed. Mice blood pressure was obtained by a noninvasive tail cuff (BP2000 system, VisiTech, Sunderland, United Kingdom).22 Blood pressure measurements were used together with echocardiographically determined LV stroke volume to calculate effective arterial elastance (Ea).23 Briefly, arterial elastance=systolic pressure/stroke volume, where stroke volume=end-diastolic volume–end-systolic volume. After the blood pressure and echocardiographic studies, mice underwent cardiomyocyte isolation.
Cardiomyocyte Isolation and Function Measurements
Cardiomyocyte isolation was performed by collagenase digestion methods (Data Supplement).24 Using methods described below, 20 cardiomyocytes from each mouse underwent study to examine their systolic and diastolic properties by use of the IonOptix system (IonOptix, Milton, Mass), and 10 cardiomyocytes from each mouse underwent study to examine passive stiffness of cardiomyocytes by use of our custom stretch system.
Cardiomyocyte systolic and diastolic relaxation properties were measured in isolated cardiomyocytes with the IonOptix system (Data Supplement).25 The passive stiffness properties of isolated cardiomyocytes were determined by a passive stretch method (Data Supplement).26
Statistical Analysis
Data are presented as mean±SEM. Differences in continuous variables between the WT control, WT TAC, SPARC-null control, and SPARC-null TAC groups were determined by 1-way ANOVA followed by Tukey testing of pairwise analysis. Although data comparing WT control versus SPARC-null control are presented in the Results section separately from data comparing WT TAC versus SPARC-null TAC, all comparisons were done in the context of a 4-group comparison (WT control, WT TAC, SPARC-null control, SPARC-null TAC) by the statistical methods stated above. Differences in dichotomous variables were determined by 
2 test, and adjustment for multiple comparisons was made with Bonferroni methods for testing of pairwise differences, which has a relatively conservative approach similar to that used with the continuous variables. The survival data presented in Figure 3 utilized a standard Kaplan-Meier analysis. P<0.05 was considered significant. The primary outcome variable for the present study was the concentration of insoluble collagen measured by biochemically determined hydroxyproline levels. The other measurements presented are secondary end points that are included as an exploratory analysis to examine pathophysiological mechanisms.
|
The authors had full access to and take full responsibility for the integrity of the data. All authors have read and agree to the manuscript as written.
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
SPARC-Null Mouse Cardiac Phenotype
Fibrillar Collagen Content by Light Microscopy
There was a significant decrease in PSR staining in SPARC-null LV myocardium compared with that seen in the WT control LV myocardium (Figure 4). CVF (Figure 4) was decreased in the SPARC-null control (0.46±0.04%) compared with the WT control (0.92±0.10%, P<0.05).
|
Collagen Composition by Biochemistry
In SPARC-null control mice, there was a decrease in total collagen and a decrease in the relative proportion of collagen that was insoluble compared with WT control mice (Figure 4). The relative proportion of insoluble collagen fell from 91±1% in WT to 87±1% in SPARC-null mice (P<0.05; Table 1). By contrast, the relative proportion of collagen that was soluble increased from 9±1% in WT to 13±1% in SPARC-null mice (P<0.05).
|
Collagen Morphology by Scanning Electron Microscopy
Morphological structure and distribution of fibrillar collagen was examined qualitatively in LV samples from WT and SPARC-null control mice (Figure 4). Fibrillar collagen struts and weave were reduced in the SPARC-null mice.
LV Structure and Function
There were no significant differences in LV volume, mass, mass/body weight ratio, volume/mass ratio, or ejection fraction in SPARC-null mice compared with WT mice (Table 2). Thus, the absence of SPARC did not alter LV volume, geometry, mass, or systolic function.
|
Vascular Function
The absence of SPARC did not significantly affect blood pressure or effective arterial elastance. Systolic blood pressure was 102±7 mm Hg in WT mice versus 104±6 mm Hg in SPARC-null mice, and effective arterial elastance was 2.3±0.3 mm Hg/µL in WT mice versus 2.4±0.2 mm Hg/µL in SPARC-null mice. There were no significant differences in any of these measurements between WT and SPARC-null control mice.
Myocardial Function
The effects of the absence of SPARC on myocardial systolic properties, diastolic relaxation, and passive stiffness were assessed by isolated papillary muscle studies. As shown in Table 3, myocardial systolic properties were not significantly different in SPARC-null control mice compared with WT control mice. Similarly, there were no significant differences in myocardial relaxation rates. By contrast, myocardial passive stiffness was decreased in SPARC-null control mice compared with WT control mice (Figure 5). The myocardial stiffness constant (β) was decreased significantly in papillary muscles isolated from SPARC-null mice compared with WT mice (Figure 5).
|
|
Cardiomyocyte Function
Cardiomyocyte systolic properties as measured by the percent and rate of shortening were not significantly different in SPARC-null versus WT control cardiomyocytes (Table 3). In addition, cardiomyocyte diastolic properties as measured by the rate of lengthening and cardiomyocyte stiffness were not significantly different in SPARC-null versus WT control cardiomyocytes.
Effects of TAC
The 28-day Kaplan–Meier survival curves for the 4 groups of mice studied (WT control, WT TAC, SPARC-null control, and SPARC-null TAC) are presented in Figure 3. This analysis showed that the procedural and periprocedural death rate after TAC was similar in WT and SPARC-null mice; late survival after TAC was also similar in WT and SPARC-null mice. There were no deaths in the control groups during the 28-day period.
SPARC Immunoblot
There was a significant increase in SPARC expression in WT mice 4 weeks after TAC measured in both salt-soluble and detergent-soluble LV myocardial tissue samples (Figure 6). As expected, there was no SPARC expression in SPARC-null mice.
|
Fibrillar Collagen Content by Light Microscopy
There was a significant increase in PSR staining in WT mice after TAC (Figures 4C and 4D). There was also an increase in PSR staining in SPARC-null mice after TAC. However, the increase in PSR staining in SPARC-null TAC mice was less than that which occurred after TAC in WT mice. CVF was increased in WT TAC mice (4.8±0.8%, P<0.05 versus WT control). CVF was also increased in SPARC-null TAC mice (1.49±0.13%, P<0.05 versus SPARC-null control); however, this TAC-induced increase in CVF was less in the SPARC-null TAC mice than in the WT TAC mice (P<0.05). The coefficient of variation between LV regions in the 2 groups of mice was low.
Collagen Composition by Biochemistry
In WT mice, TAC increased total collagen and increased the relative proportion of collagen that was insoluble (Figure 4). Relative insoluble collagen increased to 99±2% in WT TAC mice (P<0.05 versus control WT mice; Table 1); however, in WT mice, TAC decreased the relative proportion of collagen that was soluble. Relative soluble collagen decreased to 1±2% in WT TAC mice (P<0.05 versus WT control mice). In SPARC-null mice, TAC resulted in an increase in total collagen compared with SPARC-null control mice, and this increase was similar in extent to the increase in total collagen that occurred in WT mice after TAC (Figure 4). In SPARC-null mice, TAC increased both insoluble and soluble collagen concentrations compared with SPARC-null control mice. In addition, the relative proportion of collagen that was insoluble (86±2%) and the relative proportion of collagen that was soluble (14±2%) were not changed after TAC in SPARC-null mice compared with SPARC-null control mice (Table 1). However, after TAC, the relative proportion of collagen that was insoluble was smaller and the relative proportion of collagen that was soluble was larger in the SPARC-null mice than in the WT control mice (both P<0.05, SPARC-null TAC versus WT TAC mice).
Collagen Morphology by Scanning Electron Microscopy
Morphological structure and distribution of fibrillar collagen were qualitatively examined in LV samples from WT and SPARC-null TAC mice (Figure 4). In both WT and SPARC-null TAC mice, the fibrillar collagen struts and weave were increased compared with control mice; however, these changes were less pronounced in SPARC-null mice than in WT mice.
LV Structure and Function
TAC-induced pressure overload resulted in an equivalent increase in LV mass in WT and SPARC-null mice (Table 2). LV mass, wall thickness, and relative wall thickness increased in both WT and SPARC-null mice as a result of TAC for 4 weeks. The aortic pressure gradient created by the TAC was increased to an equivalent degree in both WT and SPARC-null mice after TAC. The aortic gradient reflects the pressure overload stimulus created by TAC. This stimulus was similar in both WT and SPARC-null mice that underwent TAC.
Myocardial Function
Papillary muscle cross-sectional area was not affected by the absence of SPARC in control mice (cross-sectional area=0.41±0.07 mm2 in WT versus 0.43±0.04 in SPARC-null mice) or TAC mice (cross-sectional area=0.53±0.04 in WT versus 0.58±0.04 in SPARC-null mice); however, the cross-sectional area was larger in TAC mice than in control mice (P<0.05). Minimum myocardial papillary muscle stress at Lmax was lower in SPARC-null mice than in WT mice both in control (4.88±0.2 mN/mm2 in WT control versus 3.05±0.35 mN/mm2 in SPARC-null control, P<0.05) and after TAC (7.06±0.44 mN/mm2 in WT TAC versus 5.26±0.81 mN/mm2 in SPARC-null TAC, P<0.05); however, minimum myocardial papillary muscle stress at Lmax was higher in TAC versus control mice (P<0.05). Myocardial diastolic stiffness (β) was increased in WT TAC mice compared with control WT mice. Myocardial diastolic stiffness also increased in TAC SPARC-null mice compared with control SPARC-null mice; however, this increase was significantly less than that which occurred in WT mice after TAC (Figure 5).
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
The present studies demonstrated the following: (1) SPARC-null mice had a significant change in cardiac phenotype compared with WT mice, which was characterized by a decrease in mature cross-linked insoluble collagen fibrils and a decrease in myocardial diastolic stiffness; (2) TAC-induced pressure overload in WT mice increased SPARC expression, insoluble collagen, and myocardial stiffness; and (3) TAC-induced pressure overload in SPARC-null mice resulted in a significantly smaller increase in insoluble collagen and myocardial stiffness than it did in WT mice. These data support the hypothesis that SPARC is an important determinant of post–synthetic procollagen processing and the development of mature cross-linked collagen fibrils in the normal myocardium and that SPARC is one fundamental determinant by which chronic pressure overload increases myocardial fibrillar collagen content and contributes to the development of diastolic dysfunction.
Procollagen Processing in Noncardiovascular Tissues
After synthesis, procollagen molecules are secreted into the extracellular space, where they must undergo a series of ordered, time-sensitive, and location-sensitive processing steps to become a mature cross-linked insoluble structural collagen fibril27 (Figure 1). These procollagen processing steps appear to be most efficient and effective in leading to formation of or incorporation into a mature cross-linked insoluble structural collagen fibril when they occur in the presence of matricellular proteins such as SPARC.8,9,27 The hypothesis that SPARC coordinates procollagen processing and facilitates collagen fibril assembly and formation9 is based largely on studies performed in the dermis from normal WT and SPARC-null mice. These experiments demonstrated that SPARC-null mice had substantial decreases in dermal collagen concentrations (half that of WT dermis). Collagen fibrils from the dermis of SPARC-null mice were significantly smaller and had a decreased range of fibril diameters compared with the dermis from WT mice.11 In addition, these structural changes in collagen fibrils were associated with changes in the mechanical and functional properties of the dermis, including decreased tensile strength (and decreased stiffness).11
In experimental models of dermal injury and fibrosis, SPARC-null mice had a significantly blunted response to the fibrotic stimulus, with significantly less fibrillar collagen deposition than WT mice.12,13 For example, when foreign material is implanted in the skin, a fibrous capsule is synthesized around the foreign material. In SPARC-null mice, the fibrillar collagen surrounding the foreign material exhibited fewer and smaller diameter fibrils than in WT mice.13
Therefore, there is convincing evidence that SPARC is an important determinant of post–synthetic procollagen processing and the development of mature cross-linked collagen fibrils in the ECM of dermal, pulmonary, skeletal, and renal tissues, particularly in response to injury. On the basis of these studies in noncardiovascular tissues, we hypothesized that procollagen processing and the development of mature cross-linked collagen fibrils would be altered in myocardial tissues of SPARC-null mice and that the response to a stimulus that increases myocardial fibrillar collagen content would be altered in SPARC-null mice.
Proposed Mechanisms by Which SPARC Affects Procollagen Processing
The mechanism by which SPARC influences procollagen processing has been studied in vitro with primary dermal fibroblast cultures from WT and SPARC-null mice.10 Procollagen, collagen intermediates, and soluble and insoluble collagen were measured in the conditioned media and in the "cell associated layer." In the absence of SPARC, procollagen secreted from fibroblasts had a greater tendency to associate with and bind to the fibroblast cell surface and to undergo either degradation or premature, disordered processing, and they did not efficiently or effectively develop into mature cross-linked collagen fibrils. If recombinant SPARC was added to SPARC-null primary fibroblast cultures, procollagen processing was restored toward normal. When recombinant SPARC was added to SPARC-null fibroblasts, procollagen binding to or association with the fibroblast cell surface was decreased, and effective procollagen processing into mature collagen fibrils was returned to levels comparable to the WT fibroblasts.10
Therefore, these in vitro results suggest that SPARC limits procollagen binding to cell surface receptors and promotes processing of procollagen into mature collagen fibrils. In the absence of SPARC, the regulation of procollagen processing is disrupted, and collagen interaction with receptors is enhanced, which leads to increased degradation of procollagen at the expense of incorporation of processed collagen into insoluble collagen fibrils.
Myocardial Collagen: Effects of the Absence of SPARC
LV myocardial collagen was examined in the present study by 3 independent methods: (1) Collagen content was examined with light microscopy PSR birefringent staining and quantified by measuring CVF; (2) collagen composition was examined with a biochemically determined hydroxyproline assay to quantify insoluble versus soluble collagen; and (3) collagen morphology was examined by scanning electron microscopy. These data showed that the absence of SPARC reduced the pressure overload–induced increase in myocardial collagen content, changed the composition of myocardial collagen in favor of more soluble and less insoluble collagen, and altered the collagen morphology. Although the differences in CVF were directionally similar to the hydroxyproline analysis, the magnitude of the change in fibrillar collagen as measured by CVF differed somewhat from the biochemically derived collagen measurements made by hydroxyproline analysis. When collagen content was assessed by CVF, there was a more robust response to TAC-induced pressure-overload hypertrophy in mice than was measured by the hydroxyproline analysis. We attribute this difference in collagen measurement to at least 3 factors. First, the hydroxyproline analysis included measurement of all of the collagen in the LV, including that found in association with large blood vessels and the epicardial surface. We do not believe that the absence of SPARC changes collagen associated with large blood vessels and the epicardial surface, and therefore, the effects of the absence of SPARC on myocardial fibrillar collagen would be somewhat minimized (or perhaps underestimated) by the hydroxyproline analysis. By contrast, CVF measurements were restricted to changes in interstitial collagen and excluded large blood vessels and the epicardial surface. Second, the hydroxyproline assay measured all collagen types, not just those that contribute to interstitial fibrillar collagen. By contrast, CVF primarily quantified collagen types I and III, which constitute interstitial fibrillar collagen. Third, CVF measurements were likely influenced by proteoglycan incorporation into collagen fibrils. Myocardial proteoglycans are known to be increased in response to pressure-overload hypertrophy but are not likely to be affected by the absence of SPARC. Therefore, the inclusion of proteoglycans by the PSR assay is likely to increase the apparent collagen fiber diameter as assessed by PSR, increase the measured CVF, and somewhat maximize (or perhaps overestimate) fibrillar collagen. By contrast, hydroxyproline is specific to collagen molecules only. These 3 factors help explain why there were differences between CVF and hydroxyproline assessment of mature, fully processed cross-linked myocardial fibrillar collagen and why the percent change in CVF was larger than that of the hydroxyproline assay. The hydroxyproline assay minimized (or perhaps underestimated) and the PSR assay maximized (or perhaps overestimated) the effects of the absence of SPARC on myocardial fibrillar collagen content. Nonetheless, we believe that the measurements of both CVF and hydroxyproline are complementary, that each measurement contributes to the overall understanding of collagen homeostasis in pressure-overload hypertrophy and confers an ability to examine the effects of SPARC on procollagen processing in pressure-overload hypertrophy. Importantly, both collagen measurements support the hypothesis that the absence of SPARC influences post–synthetic procollagen processing and alters collagen concentrations in the LV both in the control state and after pressure-overload hypertrophy induced by TAC.
In previous studies, there has been some variability in the effects of pressure overload on percent interstitial fibrosis, CVF, and collagen content.28–30 Variability in these measurements may be technique-dependent, based in part on whether PSR staining was performed with birefringence (polarized light) or without birefringence, whether Masson’s trichrome staining was used, and the exact morphological quantitation program used. PSR staining with birefringence stains only mature, fully processed cross-linked fibrillar collagen. PSR staining without birefringence is less specific for these collagen fibers. Masson’s trichrome stain binds to many types of ECM proteins and is not specific for fibrillar collagen. In addition, previous studies did not always state whether vasculature and epicardial surfaces were excluded from CVF measurements. For example, in a study by Zhang et al,28 CVF measured by quantification of LV myocardial sections stained with Masson’s trichrome increased from 
1% in WT C57Bl6 mice to 12% in pressure-overload hypertrophic mice. Zeisberg et al30 reported 20% fibrotic area in banded WT C57Bl6 mice, and Barrick et al29 reported an increase from 1% in normal mice to 5% in pressure-overload hypertrophic mice in C57Bl6 mice. The measurements of CVF using PSR-stained images in the present study are in close agreement to that of Barrick et al.29 It appears likely that the differences in ECM quantification as visualized by Masson’s trichrome versus PSR account for the range of values reported. Therefore, interpretation of the present results and comparison with previously published results are dependent on a clear understanding of these methodological differences.
Myocardial Stiffness: Effects of the Absence of SPARC
The increase in LV passive stiffness that occurred during pressure-overload hypertrophy was likely to be caused by changes in both the cardiomyocyte and the ECM. When ECM properties are altered, the remaining abnormalities in cardiomyocytes may still result in impaired LV diastolic function. When cardiomyocyte properties are altered, the remaining abnormalities in ECM may still result in impaired LV diastolic function. The data presented in previous studies and in the present study support this conclusion.16,17 The most important comparisons in the present study were those between WT control versus WT TAC and between SPARC-null control versus SPARC-null TAC. Examined in this fashion, SPARC-null TAC mice had an increase in insoluble collagen compared with SPARC-null controls; however, this increase was significantly less than in WT mice. The same pattern was seen in the myocardial passive stiffness measurements. Therefore, the absence of SPARC did not completely normalize insoluble collagen or myocardial stiffness when SPARC-null TAC mice were compared with SPARC-null controls. These data support the conclusion that the increase in LV passive stiffness that occurred during pressure-overload hypertrophy was caused by abnormalities in both the cardiomyocyte and the ECM.
Systolic Function and Arterial Stiffness: Effects of the Absence of SPARC
Data from the present study support the conclusion that the absence of SPARC did not affect arterial blood pressure or vascular compliance. These results are concordant with previous studies that demonstrated that systolic and diastolic blood pressure and arterial elastance were not significantly affected by the absence of SPARC.31 In addition, the absence of SPARC did not affect myocardial systolic properties, myocardial relaxation, or cardiomyocyte function. These data suggest that the effects of the absence of SPARC on myocardial fibrillar collagen primarily relate to myocardial passive stiffness.
Study Limitations
Fibrillar collagen homeostasis is influenced by at least 3 regulatory control mechanisms: procollagen biosynthesis, post–synthetic procollagen processing, and collagen degradation.32,33 The balance between these 3 regulatory control mechanisms determines the total fibrillar collagen content present in a given pathological state at a specific time. The present study focused specifically on the role that SPARC plays in post–synthetic procollagen processing and facilitation of the formation and assembly of mature cross-linked insoluble structural collagen fibrils. However, we recognize that experiments examining the effects of a change in SPARC on fibrillar collagen content must be interpreted in light of potential simultaneous changes in the other regulatory control mechanisms.
In the TAC experiments, neither procollagen synthesis nor mature cross-linked collagen fibril degradation was measured directly; however, some of the data obtained in the present study and data from experiments performed in dermal fibroblasts suggest that the absence of SPARC did not change either procollagen synthesis or mature cross-linked collagen fibril degradation. For example, TAC-induced pressure overload increased total collagen to a similar extent in both WT and SPARC-null mice; the primary substantial differences were in the insoluble versus soluble collagen fractions, which resulted in an increase in collagen content as measured by CVF and a change in fibrillar collagen morphology as evidenced in the scanning electron microscopy photomicrographs. In addition, data obtained from previous studies examining collagen synthesis and degradation in dermis and dermal fibroblasts showed no significant changes in collagen synthesis or degradation rates in SPARC-null versus WT mice. Although this conclusion will have to be supported by future studies that include direct in vivo measurements, data in the present study support the hypothesis that SPARC plays an important role in post–synthetic procollagen processing and facilitation of the formation and assembly of mature cross-linked insoluble structural collagen fibrils.
The myocardial response to stress is dependent on the type of stress imposed (pressure overload versus volume overload versus myocardial infarction), the strain of animal used, and the length over which the stress is imposed and the response monitored. Therefore, the effects of the absence of SPARC seen in the present study with TAC-induced pressure-overload hypertrophy may differ if any of these variables are changed. One genetic background strain was used both in the SPARC-null and WT mice: C57Bl6/SV129. Thus, variability in at least this determinant of the response to the imposed stress was eliminated. The response to pressure overload will produce a different response than volume overload or myocardial injury from a myocardial infarction. Whether procollagen processing and SPARC play an equivalent and important role in each type of stimulus is not fully known at this time. The present report selectively examined the response to pressure overload. One previous study examined the response of SPARC-null mice to the imposition of a myocardial infarction.34 In that study, there was an 80% mortality rate after myocardial infarction in SPARC-null mice, whereas there was a 15% mortality rate in WT mice after myocardial infarction. In addition, after myocardial infarction, SPARC-null mice developed misaligned and disorganized ECM. To date, no other published studies have examined the effect of the absence of SPARC in response to pressure overload, volume overload, or myocardial infarction. Future studies must be designed to fully evaluate the function of SPARC in post–synthetic procollagen processing in response to these different types of stimuli and the others factors listed above.
Whether there is a differential gender response to pressure-overload hypertrophy or the absence of SPARC has not been examined. In general, when the data were divided into male and female, the response appeared to be similar between genders. In particular, there appeared to be no gender differences in response to the absence of SPARC. However, the present study was not powered with a sufficient sample size to perform a statistically valid comparison between genders. This question will require future studies.
Conclusions
Data from the present study support the following conclusions: (1) SPARC plays an important role in post–synthetic procollagen processing in normal myocardium; (2) SPARC is significantly increased in pressure-overloaded myocardium; and (3) the absence of SPARC significantly alters the effects of pressure overload on myocardial fibrillar collagen content and diastolic function. SPARC is one fundamental mechanism by which chronic pressure overload increases myocardial fibrillar collagen content and contributes to the development of diastolic dysfunction. The regulation of SPARC and post–synthetic procollagen processing may provide important mechanism-based targets for the development of novel therapies for patients with chronic heart failure.
|
Acknowledgments |
|---|
Sources of Funding
This study was supported by the Research Service of the Department of Veterans Affairs (Drs Zile and Bradshaw) and grants from the National Heart, Lung, and Blood Institute (PO1-HL-48788 to Dr Zile).
Disclosures
None.
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
1. Chobanian AV, Bakris GL, Black HR, Cushman WC, Green LE, Izzo JL Jr, Jones DW, Materson BJ, Oparil S, Wright JT Jr, Roccella EJ; National High Blood Pressure Education Program Coordinating Committee. The seventh report of the Joint National Committee on Prevention, Detection, Evaluation, and Treatment of High Blood Pressure. JAMA. 2003; 289: 2560–2572.
2. Lloyd-Jones DM, Larson MG, Leip EP, Beiser A, D'Agostino RB, Kannel WB, Murabito JM, Vasan RS, Benjamin EJ, Levy D. Lifetime risk for developing congestive heart failure: the Framingham Heart Study. Circulation. 2002; 106: 3068–3072.
3. Ahmed SH, Clark LL, Pennington WR, Webb CS, Bonnema DD, Leonardi AH, McClure CD, Spinale FG, Zile MR. Matrix metalloproteinases/tissue inhibitors of metalloproteinases: relationship between changes in proteolytic determinants of matrix composition and structural, functional, and clinical manifestations of hypertensive heart disease. Circulation. 2006; 113: 2089–2096.
4. Diez J, Querejeta R, Lopez B, Gonzalez A, Larman M, Martinez Ubago JL. Losartan-dependent regression of myocardial fibrosis is associated with reduction of left ventricular chamber stiffness in hypertensive patients. Circulation. 2002; 105: 2512–2517.
5. Solomon SD, Janardhanan R, Verma A, Bourgoun M, Daley WL, Purkayastha D, Lacourciere Y, Hippler SE, Fields H, Naqvi TZ, Mulvagh SL, Arnold JMO, Thomas JD, Zile MR, Aurigemma GP; for the Valsartan in Diastolic Dysfunction (VALILDD) Investigators. Effect of angiotensin receptor blockade and antihypertensive drugs on diastolic function in patients with hypertension and diastolic dysfunction: a randomized trial. Lancet. 2007; 369: 2079–2087.[CrossRef][Medline] [Order article via Infotrieve]
6. Martos R, Baugh J, Ledwidge, M. O'Loughlin C, Conlon C, Patle A, Donnelly SC, McDonald K. Diastolic heart failure: evidence of increased myocardial collagen turnover linked to diastolic dysfunction. Circulation. 2007; 115: 888–895.
7. Bishop JE, Laurent GJ. Collagen turnover and its regulation in the normal and hypertrophying heart. Eur Heart J. 1995; 16: 38–44.[Abstract]
8. Bornstein P, Sage EH. Matricellular proteins: extracellular modulators of cell function. Curr Opin Cell Biol. 2002; 14: 608–616.[CrossRef][Medline] [Order article via Infotrieve]
9. Bradshaw AD, Sage EH. SPARC, a matricellular protein that functions in cellular differentiation and tissue response to injury. J Clin Invest. 2001; 107: 1049–1054.[Medline] [Order article via Infotrieve]
10. Rentz TJ, Poobalarahi F, Bornstein P, Sage EH, Bradshaw AD. SPARC regulates processing of procollagen I and collagen fibrillogenesis in dermal fibroblasts. J Biol Chem. 2007; 282: 22062–22071.
11. Bradshaw AD, Puolakkainen P, Dasgupta J, Davidson JM, Wight TN, Sage EH. SPARC-null mice display abnormalities in the dermis characterized by decreased collagen fibril diameter and reduced tensile strength. J Invest Dermatol. 2003; 120: 949–955.[CrossRef][Medline] [Order article via Infotrieve]
12. Bradshaw AD, Reed MJ, Sage EH. SPARC-null mice exhibit accelerated cutaneous wound closure. J Histochem Cytochem. 2002; 50: 1–10.
13. Puolakkainen P, Bradshaw AD, Kyriakides TR, Reed M, Brekken R, Wight T, Bornstein P, Ratner B, Sage EH. Compromised production of extracellular matrix in mice lacking secreted protein, acidic and rich in cysteine (SPARC) leads to a reduced foreign body reaction to implanted biomaterials. Am J Pathol. 2003; 162: 627–635.
14. Lang RM, Bierig M, Devereux RB, Flachskampf FA, Foster E, Pellikka PA, Picard MH, Roman MJ, Seward J, Shanewise JS, Solomon SD, Spencer KT, St John Sutton M, Stewart WJ. Recommendations for chamber quantification: a report from the American Society of Echocardiography’s Guidelines and Standards Committee and the Chamber Quantification Writing Group, developed in conjunction with the European Association of Echocardiography, a branch of the European Society of Cardiology. J Am Soc Echocardiogr. 2005; 18: 1440–1463.[CrossRef][Medline] [Order article via Infotrieve]
15. Zile MR, Koide M, Sato H, Ishiguro Y, Conrad CC, Buckley JM, Morgan JP, Cooper G IV. Role of microtubules in the contractile dysfunction of hypertrophied myocardium. J Am Coll Cardiol. 1999; 33: 250–260.
16. Harris TS, Baicu CF, Conrad CH, Zile MR. Constitutive properties of hypertrophied myocardium: cellular contribution to changes in myocardial stiffness. Am J Physiol. 2002; 282: H2173–H2182.
17. Stroud JD, Baicu CF, Barnes MA, Spinale FG, Zile MR. Viscoelastic properties of pressure overload hypertrophied myocardium: effect serine protease treatment. Am J Physiol. 2002; 282: H2324–H2335.
18. Blom AS, Mukherjee R, Pilla JJ, Lowry AS, Yarbrough WM, Mingoia JT, Hendrick JW, Stroud RE, McLean JE, Affuso J, Gorman RC, Gorman JH III, Acker MA, Spinale FG. Cardiac support device modifies left ventricular geometry and myocardial structure after myocardial infarction, Circulation. 2005; 112: 1274–1283.
19. Junqueira LC, Bignolas G, Brentani RR. Picrosirius staining plus polarization microscopy, a specific method for collagen detection in tissue sections. Histochem J. 1979; 11: 447–455.[CrossRef][Medline] [Order article via Infotrieve]
20. Whittaker P, Kloner RA, Boughner DR, Pickering JG. Quantitative assessment of myocardial collagen with picrosirius red staining and circularly polarized light. Basic Res Cardiol. 1994; 89: 397–410.[CrossRef][Medline] [Order article via Infotrieve]
21. Spinale FG, Tomita M, Zellner JL, Cook JC, Crawford FA, Zile MR. Collagen remodeling and changes in ventricular function during the development and regression of supraventricular tachycardia induced cardiomyopathy in swine. Am J Physiol. 1991; 261: H308–H318.[Medline] [Order article via Infotrieve]
22. Krege JH, Hodgin JB, Hagaman JR, Smithies O. A noninvasive computerized tail-cuff system for measuring blood pressure in mice. Hypertension. 1995; 25: 1111–1115.
23. Kelly RP, Ting CT, Yang TM, Liu CP, Maughan WL, Chang MS, Kass DA. Effective arterial elastance as index of arterial vascular load in humans. Circulation. 1992; 86: 513–521.
24. Ishibashi Y, Takahashi M, Isomatsu Y, Qiao F, Iijima Y, Shiraishi H, Simsic JM, Baicu CF, Robbins J, Zile MR, Cooper G IV. Role of microtubules versus myosin heavy chain isoforms in contractile dysfunction of hypertrophied murine cardiocytes. Am J Physiol Heart Circ Physiol. 2003; 285: H1270–H1285.
25. Lim CC, Apstein CS, Colucci WS, Liao R. Impaired cell shortening and relengthening with increased pacing frequency are intrinsic to the senescent mouse cardiomyocyte. J Mol Cell Cardiol. 2000; 32: 2075–2082.[CrossRef][Medline] [Order article via Infotrieve]
26. Zile MR, Richardson K, Cowles MK, Buckley JM, Koide M, Cowles BA, Gharpuray V, Cooper G IV. Constitutive properties of adult mammalian cardiac muscle cells. Circulation. 1998; 98: 567–579.
27. Prockop DJ, Kivirkko KI. Collagens: molecular biology, diseases, and potentials for therapy. Ann Rev Biochem. 1995; 64: 403–434.[CrossRef][Medline] [Order article via Infotrieve]
28. Zhang P, Xu X, Hu X, van Deel ED, Zhu G, Chen Y. Inducible nitric oxide synthase deficiency protects the heart from systolic overload-induced ventricular hypertrophy and congestive heart failure. Circ Res. 2007; 100: 1089–1098.
29. Barrick CJ, Rojas M, Schoonhoven R, Smyth SS, Threadgill DW. Cardiac response to pressure overload in 129S1/SvImJ and C57BL/6J mice: temporal- and background-dependent development of concentric left ventricular hypertrophy. Am J Physiol Heart Circ Physiol. 2007; 292: H2119–H2130.
30. Zeisberg EM, Tarnavski O, Zeisberg M, Dorfman AL, McMullen JR, Gustafsson E, Chandraker A, Yuan X, Pu WT, Roberts AB, Neilson EG, Sayegh MH, Izumo S, Kalluri R. Endothelial-to-mesenchymal transition contributes to cardiac fibrosis. Nat Med. 2007; 13: 952–961.[CrossRef][Medline] [Order article via Infotrieve]
31. Socha MJ, Manhiani M, Said N, Imig JD, Motamed K. Secreted protein acidic and rich in cysteine deficiency ameliorates renal inflammation and fibrosis in angiotensin hypertension. Am J Pathol. 2007; 171: 1104–1112.
32. Spinale FG. Myocardial matrix remodeling and the matrix metalloproteinases: influence on cardiac form and function. Physiol Rev. 2007; 87: 1285–342.
33. Nimni ME. Fibrillar collagens: their biosynthesis, molecular structure, and mode of assembly. In: Zern MA, Reid LM, eds. Extracellular Matrix. New York, NY: Marcel Dekker; 1993: 121–148.
34. Schellings MW, van Loon M, Sage H, Pinto YM, Heymans S. Loss of SPARC results in increased cardiac rupture and heart failure after acute myocardial infarction. Circulation. 2006; 114 (suppl II): II-274. Abstract.
|
Footnotes |
|---|
The online-only Data Supplement is available with this article at http://circ.ahajournals.org/cgi/content/full/CIRCULATIONAHA.108.773424/DC1.
Related Article:
Circulation 2009 119: 201-203.
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||