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To increase the supply, many countries harvest allograft valves from explanted hearts of transplant recipients with ischaemic (ICM) or dilated cardiomyopathy (DCM). This study determines the structural integrity of valves from cardiomyopathic hearts.
Extracellular matrix (ECM) was examined in human valves obtained from normal, ICM, and DCM hearts. To confirm if ECM changes were directly related to the cardiomyopathy, we developed a porcine model of chronic ICM. Histology and immunohistostaining, as well as non-invasive multiphoton and second harmonic generation (SHG) imaging revealed marked disruption of ECM structures in human valves from ICM and DCM hearts. The ECM was unaffected in valves from normal and acute ICM pigs, whereas chronic ICM specimens showed ECM alterations similar to those seen in ICM and DCM patients. Proteins and proteinases implicated in ECM remodelling, including Tenascin C, TGFβ1, Cathepsin B, MMP2, were upregulated in human ICM and DCM, and porcine chronic ICM specimens.
Valves from cardiomyopathic hearts showed significant ECM deterioration with a disrupted collagen and elastic fibre network. It will be important to determine the impact of this ECM damage on valve durability and calcification in vivo if allografts are to be used from these donors.
Since their first introduction into clinical practice more than four decades ago, human allografts have been the valve of choice for women of child-bearing age, younger patients, patients with infective endocarditis, those with small aortic roots, and patients intolerant of anticoagulation with warfarin due to bleeding risks. They provide superior haemodynamic performance, freedom from anticoagulation, and relative resistance to infections, minimal thromboembolic complications, and good long-term durability.1–3 More widespread application of allograft valves is primarily restricted by availability. Currently, allograft valves are obtained mainly from cadaveric or non-heart beating donors; however, the number of intact cadaveric valves that are adequate for implantation is low and this donor type is restricted in some member states of the European Union.3,4 Thus, to increase the supply, many countries harvest allograft valves from the explanted hearts of transplant recipients with ischaemic (ICM) or dilated cardiomyopathy (DCM).4,5 Despite this common practice, the structural integrity of these valves remains unknown. Although the consequences of loss of normal extracellular matrix (ECM) structure on valve performance is not well understood, if ECM damage were present, these abnormalities could predispose to allograft degeneration and calcification that would affect long-term durability and survival leading to graft failure post-implant as has been suggested in some animal models.6,7 The aim of this study was therefore to determine the integrity of ECM structures in valves from cardiomyopathic hearts, with special focus on the aortic valve as it is the most widely used valve allograft. To confirm that any changes were directly related to the cardiomyopathy, we also developed a model of chronic ICM in pigs. Valve structures were examined using routine histology and immunohistostaining; however, one limitation of conventional histopathology is the risk that sample processing could introduce artefacts. To overcome this limitation, we utilized multiphoton-induced autofluorescence imaging and second harmonic generation (SHG) signal profiling for the minimal-invasive artefact-free three-dimensional imaging of endogenous ECM protein-containing elements.7–9 Normal valve leaflets typically display well-defined, highly autofluorescent elastin-containing fibres and collagen bundles that are three-dimensionally organized within the four tissue layers.9–12 These intra-tissue ECM structures can be visualized in native, unfixed tissues using wavelengths of 760 nm (cells and elastic fibres) and 840 nm (collagen structures) as excitation wavelengths.7–9 However, when ECM components lose their typical three-dimensional structure and native arrangement within a tissue they generate less autofluorescence and SHG signals. Therefore, diminished intrinsic fluorescence signal intensities are indicative of structural changes secondary to pathological ECM remodelling.7–9
Using both conventional and novel imaging technologies we have demonstrated that valves from cardiomyopathic hearts display a high degree of ECM structural remodelling accompanied by marked alterations of the collagen and elastic fibre network. Valves from the hearts of patients afflicted with a cardiomyopathy also displayed upregulated expression of a number of factors implicated in tissue remodelling including Tenascin C, transforming growth factor beta 1 (TGFβ1), Cathepsin B, and to a lesser extent matrix metalloproteinase 2 (MMP2). Our data suggest that it will be important to determine the impact of this ECM damage on valve durability, calcification, and premature degeneration in vivo if allograft valves from cardiomyopathic hearts will continue to be used as a source of allografts.
All studies involving human tissues were in accordance with institutional guidelines and were approved by the local research Ethics Committees. All research was carried out in compliance with the rules for investigation of human subjects, as defined in the Declaration of Helsinki. Animal handling and care followed the recommendations of the National Institutes of Health Guide for the Care and Use of Laboratory Animals. All animal protocols were approved by the institutional review board (IRB) of the David Geffen School of Medicine at UCLA, USA.
The valves used in this study were obtained from two different institutions. The demographic and main clinical characteristics of the study population are summarized in Table 1. After informed written consent was given, valves from hearts of patients with ICM and DCM that underwent heart transplantation were donated to the Homograft Bank located at the Department of Cardiothoracic and Vascular Surgery Friedrich Schiller University (FSU) Jena, Germany. All valves deemed not suitable as allografts between 2002 and 2004 at the FSU Jena were used in the current study after institutional ethics board approval. Valves from hearts with previous coronary artery bypass grafting (CABG) or any cardiac surgical procedures (e.g. implantation of a defibrillator or an assist device) were excluded to eliminate the risk that previous surgical manipulation may have caused tissue damage. In this study, we analysed aortic and pulmonary valves from seven ICM (three pulmonary valves, seven aortic valves) and seven DCM (four pulmonary valves, seven aortic valves) hearts. Archived paraffin-embedded aortic valve sections from 1 ICM and 1 DCM patient, and normal control specimens (pulmonary and aortic valves, each n = 6), obtained from post-mortem autopsy samples (expiration due to non-cardiac causes, with no history or evidence of cardiac disease on post-mortem inspection), were provided by the Department of Pathology, David Geffen School of Medicine at UCLA, USA.
Eleven Yorkshire-Duroc pigs, 3–4 months of age (35 ± 5 kg), were used in this study. The animals were divided into three groups: the first three pigs served as normal controls, the next four pigs were assigned to the acute ICM group, and the remaining four pigs were assigned to the chronic ICM group.
The acute ICM pig model has been described in great detail previously.13 Briefly, the animals were pre-medicated with intramuscular ketamine (15 mg/ kg) and diazepam (0.5 mg/ kg), at which point they were intubated and ventilated. Ventilation parameters were adjusted to maintain PaO2 (100–180 mmHg), PaCO2 (35–45 mmHg), and pH 7.36–7.44. General anaesthesia was maintained with inhaled 1.5–2.5% isoflurane and bolus injections of fentanyl (5–10 mcg/kg) and pancuronium (0.1 mg/kg). The body temperature was maintained at 36–37°C. The right carotid artery was cannulated for arterial pressure and blood gas measurements (IRMA Trupoint Blood Gas Analysis, Edison, NJ, USA). A 7.0 Fr pulmonary artery catheter (CCOmbo with Vigilance™, Edwards Lifesciences, Irvine, CA, USA) was inserted via the right external jugular vein to continuously measure mixed venous oxygen saturation, cardiac index, pulmonary artery, and central venous pressures. An AL1 guide catheter was placed via the femoral artery into the left main coronary artery. A 2.0–3.0 mm angioplasty balloon catheter was advanced over a guidewire and inflated in the mid-left anterior descending artery. The balloon was inflated for 30 min, followed by harvest of the heart and dissection of the aortic heart valves.
For the chronic ICM model, a closed-chest myocardial infarct (MI) model was established. After overnight fast, the animals were pre-medicated with 1.4 mg/kg Telazol, intubated, and ventilated. General anaesthesia was maintained with inhaled 2.5% isoflurane. Femoral arterial access was obtained, and an AL1 guide catheter was placed in the left main coronary artery. A 2.5–3.5 mm angioplasty balloon catheter was advanced and inflated in the left anterior descending artery (LAD). ECG and arterial pressure were monitored during infarction and recovery. Thirty seconds after balloon inflation, 5 mL suspension of sterile saline containing 0.5–1 mL polystyrene microspheres (Polybead® 90 µm, Polysciences, Inc., Warrington, PA, USA) were injected through the central lumen of the catheter. The site of occlusion in the LAD and the actual amount of microspheres injected were determined by a visual estimate of the size of the LAD and of the vascular bed that was subtended by the vessel distal to the point of occlusion. The balloon was deflated after 5 min and the catheter pulled out. The animals were housed for 8 weeks after which they underwent MRI assessment. Detailed information regarding MRI acquisition and analysis is provided in Supplementary material online. After MRI imaging, the pigs were sacrificed and aortic heart valves were excised and analysed.
For histological and immunofluorescence analysis, heart valve tissues were washed in sterile phosphate-buffered saline (PBS, Invitrogen, Carlsbad, CA, USA), fixed in 4% paraformaldehyde (Sigma-Aldrich, St Louis, MO, USA) and processed for paraffin embedding as previously described.14 A modified Movat pentachrome stain was used to demonstrate ECM components.12,14 Primary antibodies were a rabbit anti-Tenascin C (1:150; HPA004823, Sigma); and mouse anti-TGFβ1 (1:500; TB21, Abcam, Cambridge, MA, USA), anti-Cathepsin B (1:500; clone 3H2, Chemicon, Temecula, CA, USA), and anti-MMP2 (1:500; clone 42-5D11, Chemicon). After heat-mediated antigen retrieval using citrate buffer pH 6.0 (Sigma), sections were incubated in the primary antibody solution (antibody diluted in 1% bovine serum albumin, 0.1% cold fish skin gelatine, 0.1% Triton-X 100, 0.05% Tween 20 (all Sigma) in PBS) overnight at 4°C. Specimens treated using the buffer alone served as controls. Cell nuclei were counterstained using 4′-6-diamidino-2-phenylindole (DAPI, Sigma). Secondary antibodies included Alexa Fluor 594-conjugated goat anti-mouse IgG (H+L) and goat anti-rabbit IgG (H+L) (1:250; both from Molecular Probes, Eugene, OR, USA). All images were acquired using an inverted Zeiss Axiovert 200 microscope (Carl Zeiss MicroImaging Inc., Thornwood, NY, USA) and were processed with Adobe Photoshop CS3 (Adobe Systems Inc., San Jose, CA, USA).
Multiphoton-induced autofluorescence and SHG imaging were performed using a Zeiss LSM 510 META NLO femtosecond laser scanning system (Carl Zeiss), coupled to a software-tunable Coherent Chameleon titanium:sapphire laser (720–930 nm, 90 MHz; Coherent Laser Group, Santa Clara, CA, USA). All observations were made using unprocessed and untreated human and porcine tissues that had been transferred to the laboratory and analysed within 30 min after explantation. Following imaging, all samples were processed for histology and immunochemistry as described earlier. Extracellular matrix structure-dependent autofluorescence and SHG were induced using wavelengths of 760 nm (elastin) and 840 nm (collagen) as described previously.7–9,12,14 Non-invasive serial optical horizontal sections of six different areas of each of the specimens were taken in z-steps of 10 µm at depths of 20–250 µm.
To quantify the intrinsic fluorescence signals of collagen-containing structures within normal, acute, and chronic porcine ICM tissues, lambda stacks were ascertained at emission wavelengths of 390–500 nm (in 10 nm increments) using the two-photon Chameleon laser tuned to an excitation wavelength of 840 nm. Emission was collected using the Zeiss META detector (spectral separator) of the LSM 510 Meta NLO system, as previously described.7,9 Due to a reproducible laser excitation power and similar exposure times for all samples, mean intensities of the intrinsic SHG signals were calculated and used for quantification. Briefly, in each case six different areas of each tissue sample were scanned and intensities of the intrinsic SHG signals of the regions of interest (ROI) were detected. The SHG signal intensities were reflected by the grey values of all pixels within an ROI. Mean intensities were calculated from the six screened ROI areas.
To quantitate the ECM components of normal, acute, and chronic porcine ICM tissues, crude tissue lysates were prepared and all samples were normalized according to their wet weight in milligrams. Total collagen and proteoglycan/glycosaminoglycan contents were quantified using SIRCOL and BLYSCAN assays (Biocolor, Belfast, Northern Ireland) as per manufacturer's instructions. For the evaluation of gelatinase activity in normal, acute, and chronic porcine ICM tissues, total protein was extracted in 10 mm Tris, pH 7.4/150 mm NaCl/1 mm CaCl2/1 mm MgCl2/0.1% Triton X-100, in the presence of the protease inhibitor mixture Complete Mini (Roche Diagnostics GmbH, Mannheim, Germany). The protein content was determined by a colorimetric assay (Bio-Rad, Hercules/CA, USA). The MMP-Gelatinase Activity Assay Kit (Chemicon) was used according to the manufacturer's instructions.
For the statistical analyses, only aortic valve specimens were included. All data are presented as mean values±standard deviations (SD). Results were compared by analysis of variance and Fisher's PLSD (Tukey's) tests or two-sided, unpaired t-test as indicated, using significance at a P-value <0.05.
Outflow tract valves consist of three semilunar leaflets that separate the ventricles from the major arteries. Four well-defined tissue layers can be identified within the valve leaflets: arterialis, fibrosa, spongiosa, and ventricularis (Figure 1).2,9–12 To examine the integrity of the major ECM components (collagen, elastin, and glycosaminoglycans) within the aortic valve tissue regions schematically displayed in Figure 1, we performed Movat pentachrome staining on normal, ICM, and DCM human specimens (Figure 2). When compared with normal tissues (Figure 2A, D, G), valve leaflets isolated from human ICM (Figure 2B, E,H), and DCM (Figure 2C, F, I) hearts revealed a significant decrease of collagen within the arterialis and fibrosa layer (leaflet outflow side), increased interfibrillar spaces within the inner leaflet layer (spongiosa), as well as elastic fibre fragmentation within the inflow side layer (ventricularis); with a complete disappearance of the elastic fibre network in DCM specimens (Figure 2C). Abnormal ECM remodelling and turnover was also found in the muscle-leaflet junction area of human ICM and DCM tissues (Figure 2D–F), with evidence of cardiac muscle degradation in both ICM and DCM samples. However, in contrast to normal (Figure 2G) and ICM (Figure 2H) muscle samples, distinct interstitial fibrosis and irregular hypertrophy were exclusively seen within DCM muscle (Figure 2I).
Since manipulation and fixation of pathology samples can potentially introduce tissue artefacts, we directly assessed the tissue state and integrity of ECM structures in valves from normal, ICM, and DCM patients by multiphoton-induced autofluorescence microscopy (Figure 3). Elastic fibre autofluorescence was induced with laser pulses at 760 nm revealing the presence of a relatively preserved fibre network within outflow (Figure 3E and Q) and inflow (Figure 3G and S) sides of human aortic and pulmonary ICM valve leaflets, when compared with normal human tissues [Figure 3A, M (outflow side) and 33C, O (inflow side)]. However, using an excitation wavelength of 840 nm, degradation of collagen bundle structures was detectable predominantly within the collagen-rich outflow side of human aortic and pulmonary ICM leaflets (Figure 3F and R), which was more pronounced on the inflow side of the ICM leaflets (Figure 3H and T). Excitation of valve tissues isolated from human DCM hearts revealed marked disruption of ECM structures, including depletion and disintegration of elastin-containing structures, mainly within the outflow side of aortic and pulmonary leaflets (Figure 3I and U). Almost no autofluorescent collagen structures were detected within either side of aortic and pulmonary leaflets [Figure 3J, V (outflow side) and 33L, X (inflow side)]. Similar results were seen in all valves from human cardiomyopathic hearts imaged and were markedly different from normal valves including the absence of birefringed collagen fibres—histological hallmark of normal heart valve leaflet matrix (Figure 3A–D and M–P).
By the time patients with end-stage heart failure are transplanted, they represent a diverse population with variable disease aetiology, severity, duration, and treatment regimens. Thus, to determine more directly if the ECM damage that we observed within human valves from cardiomyopathic hearts was a consequence of the cardiomyopathy itself, we developed a model of ICM in pigs. This model specifically targets the left ventricular myocardium. Therefore, any changes observed within the aortic valves would presumably be secondary to the cardiomyopathic process. Compared with normal porcine valve tissues (Figure 4A, D, G, J), the leaflets, cardiac muscle, sinus wall, and aortic trunk of acute ICM specimens (Figure 4B, E, H, K) showed similar morphologic features without significant ECM changes. In contrast, the ECM structures of chronic ICM valves (Figure 4C, F, I, L) were fragmented and less organized, similar to human ICM and DCM samples. Accordingly, chronic ICM valve leaflets demonstrated less dense ECM with large disrupted spaces in the spongiosa layer, elastic fibre fragmentation within the ventricularis, and collagen remodelling within the fibrosa layer (Figure 4C). Severe degradation of collagen, elastin, and glycosaminoglycans was also seen within the muscle-leaflet junction of chronic ICM tissues (Figure 4F). Staining of the sinus wall region of chronic ICM specimens demonstrated pathologically increased ECM remodelling and turnover, as evidenced by a significant increase in fibrotic areas (intense red) and collagen deposition (yellow) (Figure 4I). Histomorphology and elastic fibre distribution within aortic trunk regions of chronic ICM specimens appeared to be largely unaffected; however, the content of glycosaminoglycans was slightly increased (Figure 4L).
To confirm these changes, we examined unprocessed tissues from normal, acute, or chronic ICM pigs using multiphoton-induced autofluorescence microscopy. No significant differences were detectable between normal and acute ICM leaflets (Figure 5A, C and E, G), cardiac muscle (Figure 6A and C), or aortic trunk (Figure 6G and I). Similar to the changes seen in valves from human ICM and DCM hearts, multiphoton imaging of the chronic ICM tissues demonstrated weak autofluorescence of collagen and elastic structures, indicating marked ECM remodelling (Figures 5I, K and 66E, K). The amount of collagen fibre autofluorescence, and hence ECM integrity, was quantified by analysing SHG signal profiles. Applying this novel technique, using the same laser powers for all specimens, we identified significant differences between intrinsic fluorescence signal intensity values reported in Table 2. Second harmonic generation signal patterning of normal, acute ICM, and chronic ICM leaflets, muscle and trunk regions are shown (Figure 5B, D, F, H, J, L; Figure 6B, D, F, H, J, L). There were no significant differences in SHG signal intensities of collagen structures in normal or acute ICM aortic specimens. However, SHG signal intensities from both were significantly higher than in all areas examined in chronic ICM tissues, indicating the substantial ultrastructural deterioration and disintegration of most collagen structures (Table 2). Second harmonic generation signals from normal leaflet outflow or inflow sides were 6.9-fold and 5.7-fold higher, respectively, than signal intensities measured in similar tissues from chronic ICM pigs (P < 0.001).
In addition to routine histology and qualitative and quantitative multiphoton imaging, we further determined collagen as well as proteoglycan/glycosaminoglycan contents in normal, acute, and chronic porcine ICM tissues. No differences were found in the amount of total collagen in normal valves vs. acute ICM or chronic ICM tissues suggesting the changes we saw in SHG signal were truly related to abnormalities in ECM architecture and not reductions in absolute collagen content (Table 3). In contrast, the proteoglycan/glycosaminoglycan content in valves from chronic ICM hearts was significantly decreased when compared with normal valves (P = 0.0061) or valves harvested from acute ICM hearts (P = 0.0078) (Table 3).
To explore the mechanism(s) underlying the ECM damage we observed, we examined the expression of a panel of factors implicated in regulating ECM remodelling and turnover (Figures 7 and 88). Immunofluorescence staining was performed using antibodies against Tenascin C (Figures 7A–C and 88A–C), TGFβ1 (Figures 7D–F and 88D–F), Cathepsin B (Figures 7G–I and 88G–I), and MMP2 (Figures 7J–L and 88J–L). In normal human as well as normal and acute ICM porcine valves, Tenascin C, a large ECM glycoprotein that has been implicated in the progression of aortic valvular disease,15 was predominantly expressed within the outflow side of the aortic valve leaflets (Figures 7A and 88A and B). In contrast, in human ICM and DCM, as well as porcine chronic ICM specimens, Tenascin C was expressed throughout the entire leaflets (Figures 7B and C and 88C). Similarly, TGFβ1, Cathepsin B, and MMP2, proteins and extracellular proteases associated with ECM remodelling, were not detectable within normal human control specimens (Figure 7D, G, J) or normal and acute ischaemic porcine valve leaflets (Figure 8D, E, G, H, J, K), but were detectable in human ICM tissues (Figure 7E, H, K). TGFβ1 was strongly expressed in human DCM tissues, whereas Cathepsin B (Figure 7I) and MMP2 (Figure 7L) were weakly expressed. In valves from porcine chronic ICM hearts, TGFβ1 (Figure 8F) and Cathepsin B (Figure 8I) were strongly expressed throughout the entire leaflets; and MMP2 expression was slightly upregulated (Figure 8L). To determine whether these low levels of MMP2 expression were associated with enhanced protease activity, we performed a gelatinase activity assay on lysates prepared from valvular tissue. This assay utilizes a biotinylated gelatinase substrate that is cleaved by active MMP2 or MMP9. There was a 71% increase in gelatinase activity in valves harvested from chronic ICM hearts when compared with normal valves or valves from acute ICM hearts (P < 0.0001).
Despite constant progress in treatment and expanded research efforts, valvular heart disease still accounts for significant mortality and morbidity.16,17 Unfortunately, there are no currently accepted treatments that reverse structural changes in diseased valves, although several have been proposed to delay the progression of established valvular disease.18 The only proven therapy is valve replacement and although the majority of valve replacements are performed with mechanical or bioprosthetic valves, heart valve allografts have become an attractive alternative in specific patient populations. However, the supply of allografts is limited and recent concerns over their propensity for degeneration, especially in pediatric recipients, has been a matter of debate.19,20
Since valves from patients with cardiomyopathy are often harvested to increase the supply of allografts, we sought to determine the integrity of ECM structures within these human valves. We used conventional histology and immunohistostaining, as well as novel non-invasive imaging technologies including multiphoton-induced autofluorescence microscopy and quantitative SHG signal profiling. Our studies revealed a significantly altered histoarchitecture and disintegration of major ECM proteins, including collagen, elastin, and glycosaminoglycans in human valves from ICM and DCM hearts when compared with normal control valves. Moreover, artefact-free multiphoton imaging of non-processed human aortic and pulmonary valve specimens demonstrated diminished intrinsic fluorescence signal intensities, indicating destructive remodelling of the collagen fibres within these valves. Interestingly, these degenerative changes were more pronounced in valve tissues from DCM patients when compared with ICM patients. Whether this represents a true biological difference vs. simply reflecting differences in patient populations with respect to disease duration and severity is unknown. In order to determine if the ECM changes we observed were directly related to the cardiomyopathic process, we developed a porcine model of chronic ICM by inducing left ventricular myocardial infarctions. Similar to the results seen in human valves from ICM and DCM hearts, histological analysis of aortic tissues from chronic porcine ICM tissues revealed pathological changes of the ECM, predominantly seen in the valve leaflets. Biochemical assays further revealed a significant decrease in proteoglycans/glycosaminoglycans in the chronic ICM tissues when compared with valves explanted from normal or acute control ICM hearts. Multiphoton-induced autofluorescence as well as quantitative SHG signal profiling confirmed marked alterations of the collagen and elastic fibre network. No significant differences were detectable between normal and acute ICM valve leaflets, cardiac muscle, or aortic trunk.
Little is known about the pathophysiology of structural valve deterioration or the mechanisms that regulate ECM remodelling in valve tissues. Identifying and quantifying the multi-factorial mechanisms that lead to degeneration and malfunction of graft tissues is therefore an important clinically relevant issue that will need to be addressed for the field to progress both on a potential preventative therapeutic as well as the research level. Although the effectors involved in pathological ECM remodelling are not well understood, ECM degradation appears to proceed the calcific changes in degenerating valves,21 and structurally altered elastic fibres as well as acidic glycoproteins may be a stimulus for valve calcification.6,22 Studies have also implicated MMPs and tissue inhibitors of metalloproteinases (TIMP) in ECM remodelling of native valves.23 Consistent with this data, we demonstrated that abnormal valve leaflet ECM remodelling and turnover in human ICM and DCM, as well as porcine chronic ICM specimens was associated with increased expression of Tenascin C, TGFβ1, Cathepsin B, and MMP2.
While it is speculative how cardiomyopathy could lead to pathological valve remodelling, marked haemodynamic alterations occur with the development of cardiomyopathy. These altered haemodynamic flow patterns could initiate ECM remodelling by activating expression of remodelling-associated proteins and proteases,24 and ultimately have structural and physiological consequences both for the endocardial endothelium and for the subjacent interstitial tissue, including the outflow tract valves.25 Indeed, it has been previously shown that collagen synthesis by valve interstitial cells can be stimulated by stretch and cyclic mechanical tension.26,27 Further, elevated TGFβ1 levels, in the presence of cyclic circumferential tension, resulted in synergistic increases in contractile and biosynthetic proteins in aortic valve interstitial cells, leading to enhanced collagen production.27 Tenascin C, which is expressed widely within heart valves during development, but is found only in restricted locations in the adult, can also be regulated by tissue biomechanics. Tenascin C has, in combination with MMP2, been implicated in the pathobiology of calcific aortic stenosis.15 In addition, cardiomyopathy has been associated with a generalized immune activation and previous studies have shown that inflammatory mediators can regulate ECM remodelling in calcific aortic valve stenosis.28
In summary, our data suggest that valves from cardiomyopathic hearts develop significant pathological ECM remodelling. These data suggest that it will be critically important to determine if the valve matrix damage that we observed has clinical consequences similar to the effect of ECM damage on survival of vascular grafts.29 If similar results are found it would imply the valves harvested from cardiomyopathic patients could be more predisposed to valve degeneration and calcification compromising valve function. This would have an important impact on current clinical practice with regard to utilization of allograft valves and choice of valve donors.
This work was supported by the Deutsche Forschungsgemeinschaft [to K.S.-L. Sche701/2-1], National Institutes of Health [5T32HL007895-10 to K.S.-L., P01 HL080111 and R01 HL70748 to W.R.M.], and the American Heart Association [0855177F to W.R.M.].
The authors would like to thank Yekaterina Butylkova, Derek Klarin, and the Translational Pathology Core Laboratory (TPCL) at UCLA for their excellent technical assistance. We are grateful to Jens Geiling (Institute of Anatomy, Friedrich-Schiller University Jena, Germany) for providing the heart valve illustration (Figure 1).
Conflict of interest: none declared.