FASEB J. Avanti Polar Lipids
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Published as doi: 10.1096/fj.05-5517fje.
This Article
Right arrow Abstract Freely available
Right arrow Summary
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
fj.05-5517fjev1
20/8/1266    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Petermann, I.
Right arrow Articles by Reinheckel, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Petermann, I.
Right arrow Articles by Reinheckel, T.
(The FASEB Journal. 2006;20:1266-1268.)
© 2006 FASEB

Lysosomal, cytoskeletal, and metabolic alterations in cardiomyopathy of cathepsin L knockout mice

Ivonne Petermann*,1, Christian Mayer*,1, Jörg Stypmann{dagger}, Martin L. Biniossek*, Desmond J. Tobin{ddagger}, Markus A. Engelen{dagger}, Thomas Dandekar§, Tilman Grune||, Lorenz Schild, Christoph Peters*,2 and Thomas Reinheckel*

* Institut für Molekulare Medizin und Zellforschung, Albert-Ludwigs-Universität Freiburg, Freiburg, Germany;

{dagger} Medizinische Klinik und Poliklinik C (Kardiologie und Angiologie), Zentrale Projektgruppe Kleintierdiagnostik des Interdisziplinären Zentrums für Klinische Forschung Münster, Universitätsklinikum Westfälische Wilhelms-Universität Münster, Münster, Germany;

{ddagger} Medical Biosciences, School of Life Sciences, University of Bradford, Bradford, West Yorkshire, UK;

§ Lehrstuhl für Bioinformatik, Biozentrum, Am Hubland, Würzburg, Germany;

|| Research Institute for Environmental Medicine at the Heinrich Heine University Duesseldorf, Molecular Aging Research, Duesseldorf, Germany; and

Institut für Klinische Chemie und Pathologische Biochemie, Bereich Pathologische Biochemie, Medizinische Fakultät der Otto-von-Guericke-Universität Magdeburg, Germany

2Correspondence: Institut für Molekulare Medizin und Zellforschung, Albert-Ludwigs-Universität Freiburg, Freiburg D-79104, Germany. E-mail: christoph.peters{at}mol-med.uni.freiburg-de

ABSTRACT

Although lysosomal proteases are expressed in the heart at considerable levels, their specific functions in this organ remain elusive. Mice deficient for the lysosomal cysteine protease cathepsin L (CTSL) develop a late onset dilated cardiomyopathy (DCM) that is characterized by cardiac chamber dilation, fibrosis, and impaired cardiac contraction at 12 month of age. Investigation of the pathogenic sequence of DCM in ctsl–/– mice revealed numerous dysmorphic lysosome-like structures in heart muscle as early as 3 days after birth, whereas skeletal muscle was not affected. Labeling of the acidic cell compartment of neonatal cardiomyocytes and detection of lysosomal markers after subcellular fractionation confirmed increased lysosome content in CTSL deficient myocardium; however, specific storage materials were not detected. The myocardium of ctsl+/+ and ctsl–/– mice revealed no differences in incidence of cell death, proliferation, and capillary density during DCM progression. However, an observed increase in mRNA expression of natriuretic peptides in young adult mice indicates the activation of the adaptive "fetal" gene program, while proteome analysis revealed decreased levels of the sarcomere-associated proteins {alpha}-tropomyosin, desmin, and calsarcin 1, as well as considerable changes of metabolic enzymes. Bioinformatic pathway analysis suggested a switch to anaerobic catabolism and impairment of mitochondrial respiration. This interpretation was supported by a 50% reduction in resting state oxygen consumption and impaired respiration capacity in ctsl–/– myocardial homogenates. In summary, the data indicate an essential role of CTSL in maintaining the structure of the endosomal/lysosomal compartment in cardiomyocytes. Lysosomal impairment in ctsl–/– hearts results in metabolic and sarcomeric alterations that promote DCM development.—Petermann, I., Mayer, C., Stypmann, J., Biniossek, M. L., Tobin, D. J., Engelen, M. A., Dandekar, T., Grune, T., Schild, L., Peters, C., Reinheckel, T. Lysosomal, cytoskeletal, and metabolic alterations in cardiomyopathy of cathepsin L knockout mice.


Key Words: protease • cardiomyopathy • lysosome • proteome

CATHEPSIN L (CTSL) BELONGS to the papain-like family of lysosomal cysteine proteases (1) . The cysteine cathepsins comprise 11 human and 19 murine enzymes (2) . Because of their principal endosomal/lysosomal localizations and redundant substrate specificities, cysteine cathepsins have been initially thought to cooperate in terminal degradation of proteins that were delivered to the lysosomes by endocytosis or autophagy (3) . However, over the past few years cysteine cathepsins have been detected in the cytosol and nucleus, as well as in the extracellular space, and multiple specific functions have been assigned to individual cathepsins (4) . CTSL is a potent endoprotease that is able to perform limited proteolysis in the acidic cellular compartment of specific cell types. Recently, established functions of CTSL include the catalysis of a distinct step of myosin heavy chain (MHC) II invariant chain processing in thymic cortex epithelial cells, the maturation of enkephalin in chromaffin granules of neuroendocrine cells, and the degradation and recycling of growth factors and their receptors in epidermal keratinocytes (5 6 7) . In the cytosol, CTSL has been suggested to regulate apoptosis by activation of tBid and it has been shown to process the nuclear transcription factor CDP/Cux (8 , 9) . In addition, CTSL promotes invasion of tumor cells and endothelial progenitor cells by degradation of extracellular matrix molecules (10) . CTSL knockout mice exhibit a prominent skin phenotype characterized by epidermal hyperproliferation, acanthosis, and hyperkeratosis, as well as by abnormalities in development and cycling of hair follicles that result in periodic hair loss (11 , 12) . Recently, we identified a marked dilated cardiomyopathy (DCM) occurring in 1-yr-old CTSL-deficient mice (13) . DCM describes a group of myocardial diseases characterized by cardiac dilation, decreased left ventricular contraction, cardiac remodeling, and finally congestive heart failure (14) . Currently, known causes of DCM include viral infections, ischemia, and mutations in genes encoding sarcomeric and structural proteins essential for the generation and transmission of contractile forces within the cardiomyocyte; however, these etiologies cause complex adaptive and pathogenic responses on biochemical and cell biological levels (15 16 17) .

Since CTSL could be involved in multiple cellular processes the alterations of which might finally cause a cardiomyopathy, the present study was initiated to investigate the pathogenic sequence that occurs in the heart of ctsl–/– mice. Specifically, we aimed to establish the time course of disease progression and to identify the cell biological processes that are altered by CTSL deficiency in the myocardium.

MATERIALS AND METHODS

Animals and tissues
CTSL-deficient mice were generated by gene targeting in mouse embryonic stem cells as described previously (11) . Expression of CTSL mRNA, CTSL protein, and activity was completely abolished in CTSL-deficient mice (11) . The generation, maintenance, and breeding of the mice as well as animal experiments were performed in accordance with our institutional regulations. Echocardiographic measurements were performed as described previously (18) . Hearts were dissected after perfusion with 25 ml PBS via the left ventricle. The atria and heart valves were removed, and ventricular myocardium was prepared for histological analyses or immediately stored at –80°C for biochemical and molecular biology methods.

High-resolution light microscopy and transmission electron microscopy
Tissues were immediately fixed in half-strength Karnovsky’s fixative, postfixed in 2% osmium tetroxide, and embedded in araldite resin. Semithin and ultrathin sections were cut; the former were stained with the metachromatic stain, toluidine blue/borax, and examined by light microscopy and photographed (Leitz), while the latter were stained with uranyl acetate and lead citrate and examined and photographed using a Jeol 1200EX transmission electron microscope (TEM; Jeol). Multiple blocks were examined from each heart, and 3 mM long and 1 µm thick sections were examined by light microscopy from each block. Tissue blocks were further examined by electron microscopy (EM) with each ultrathin section {approx}1 mM across.

Histology
Myocardium was washed in PBS and subsequently fixed in 4% buffered formalin for 24 h. After being embedded in paraffin, 5 µm thick serial sections were cut, deparaffinated, and rehydrated, and interstitial fibrosis was stained with Masson’s Trichrome. Carbohydrates were detected by the with periodic acid Schiff’s reagent (PAS staining) reaction (19) . Glucosaminoglycanes and lipids were detected by Mowry and Sudan III stainings, respectively (20) .

Immunohistochemistry and terminal dUTP nick-end labeling
For detection of the cell proliferation marker Ki67, slides were boiled in citrate buffer (0.01 M) for 15 min and endogenous peroxidase activity was blocked with 3% H2O2 in methanol for 20 min at room temperature. After being blocked with rabbit serum in PBS-Tween 20 (0.2% Tween 20 in PBS), the slides were incubated with 130 µl rat anti-mouse Ki67 (DakoCytomation) antibody (Ab; 1:20 in PBS-Tween 20) overnight at 4°C. Ki67-positive cells were detected by the Vectastain Elite ABC Rat IgG system (Vector Laboratories). Stained slides were observed with a 200-fold magnification. Ki67-positive nuclei were counted in nine defined fields of view equally distributed within the cross-section (3 covering the left ventricle, 3 covering the right ventricle, 3 covering the septum).

Apoptotic cells were detected by usingan established, commercially available, terminal dUTP nick-end labeling (TUNEL) kit (ApopTag; Oncor). Briefly, 5 µm sections were deparaffinized, treated with proteinase K for 15 min at room temperature, and then incubated with digoxigenin-dUTP in the presence of terminal desoxyriboxyl-transferase. Subsequently, TUNEL-positive cells were visualized by antidigoxigenin peroxidase conjugated F(ab)2 fragments. TUNEL-positive and TUNEL-negative cells were counted as described for Ki67, respectively. Three hearts were analyzed per genotype and age group.

For detection of CD31 (PECAM-1) and laminin, cryostat sections (5 µm) were washed in PBS and blocked with serum for 1 h at 37°C. The slides were incubated with 100 µl rat anti-mouse CD31 (BD Pharmingen) and rabbit anti-mouse laminin (Abcam) antibodies (1:250 in PBS-Tween 20) for 2 h at 37°C. Subsequently, the sections were incubated with 100 µl Alexa Fluor 647 rat IgG (BD Pharmingen) and Alexa Fluor 488 goat anti-rabbit (Invitrogen/Molecular Probes) secondary antibodies for 2 h at 37°C. Nuclei were stained after being washed three times with PBS-Tween 20 with 100 µl Hoechst 33342 (82 ng/ml) for 5 min.

Labeling of the acidic compartment in primary murine cardiomyocytes
Hearts from 3-day-old mice were washed in ice-cold ADS buffer (116 mM NaCl, 20 mM HEPES (pH 7.4), 1 mM Na2H2PO4, 5.4 mM KCl, 0.8 mM MgSO4 x7 H2O, and 5.5 mM glucose, sterile filtered). Neonatal ventricles were digested in ADS buffer containing 127 U/ml collagenase II (PAA) and 1 mg/ml pancreatin (Sigma) in a shaker (10,000 rpm) at 37°C for 15 min. After digestion, crude material in the sample was allowed to sediment for 1 min and was digestion was repeated twice. The cells in the supernatants of the three digestion steps were washed by centrifugation (800 rpm) in a 3:1 mixture of DMEM and Medium 199 supplemented with 10% horse serum, 5% FBS, 2 mmol/l L-glutamine, 10 mg/ml streptomycin, and 100 UI/ml penicillin (all cell culture materials were purchased from Invitrogen/Life Technologies). Cardiomyocytes and nonmyocyte cells (NMCs) were separated by differential plating (45 min, 37°C, 10% CO2) and plated at a density of 1 x 105 cells/cm2 on gelatin-coated wells with poly-L-lysine coated coverslips and noncoated wells, respectively. After 24 h, cardiomyocytes were switched to serum-free medium and washed in PBS. The acidic compartment was stained with 500 nM Lysotracker (Invitrogen/Molecular Probes) in a 3:1 mixture of DMEM and Medium 199 with supplements and without serum for 1 h (37°C, 10% CO2) and washed for 2 h in fresh media. The cells on the coverslips were fixed with 4% paraformaldehyde in 0.1 M PIPES (pH 6.8) for 5 min and washed in PBS and blocked with 1% BSA in PBS for 1 h. After permeabilization with 0.05% saponin in 0.1 M PIPES, pH 6.8, for 5 min and being washed two times, the cells were fixed with ice-cold acetone for 10 min. Cells were stained with 65 µl goat anti-mouse troponin I-C Ab (1:200 in PBA, Santa Cruz) for 90 min at room temperature. After being washed three times with PBS-Tween 20, the cells were incubated with 65 µl rhodamine conjugated secondary Ab (1:200 in PBA) at room temperature.

Subcellular fractionation
Fresh ventricle myocardium sample (100 mg) was minced in 500 µl homogenization buffer (250 mM sucrose, 10 mM Tris, and 5 mM MgCl2, pH 7.4) using a Dounce Homogenizer and centrifuged at 1000 g for 10 min. The resulting supernatants were adjusted to equal protein content (10 mg/ml) and centrifuged for 20 min at 17,000 g. The resulting organelle pellet was dissolved in 250 µl homogenization buffer and separated on a 3.25 ml Percoll-gradient ranging from 1.04–1.12 g/ml for 45 min at 50,000 g (21) . Fractions of 250 µl were collected and used for further enzyme activity determination and Western blotting.

Measurement of ß-hexosamidase activity
ß-Hexosamidase hydrolytic activity was determined by measuring the degradation of p-nitrophenyl-N-acetyl-ß-D-glucoseaminide by photometry at 405 nM, and 10 µl of density gradient fractions were added to 100 µl of substrate solution (100 mM sodium citrate, 10 µM p-nitrophenyl-N-acetyl-ß-D-glucoseaminide, pH 4.6) 15 min at 37°C. The reaction was terminated by adding 500 µl stop solution (400 mM glycine, pH 10.4), and enzyme activity was calculated by using the formula: activity ß-hex(mU/ml) = {Delta}E405 nM*220 mU/ml.

Detection of the lysosome-associated membrane protein 1 by Western blotting
Density fractions were applied to 10% SDS-PAGE under reducing conditions and subsequently electrotransferred onto a PVDF membrane. For detection of the lysosome-associated membrane protein 1 (Lamp1), the monoclonal rat-antimouse Lamp1 Ab (Abcam) was used in a 1:38 dilution. The binding of secondary Ab (antirat IgG-POD; Abcam; 1:2500 dilution) was detected by the SuperSignal Chemiluminescent Substrate (Pierce).

Two-dimensional electrophoresis
Proteins were extracted from freeze-dried myocardial samples based on their differential solubility; a sample of 20 mg was Dounce homogenized in 1 ml detergent free extraction buffer I (40 mM Tris, 1 mM EDTA, 1 mM pefablock, 10 µM pepstatin A, and 100 µM leupeptin, pH 7.4). After centrifugation (16,000 g, 12 min, 4°C), the supernatant was saved for two-dimensional electrophoresis (2DE) as hydrophilic protein fraction. The pellet was dissolved in 1 ml detergent containing extraction buffer II (40 mM Tris, 5 M urea, 2 M thiourea, 2% (w/v) CHAPS, 2% (w/v) SB 3–10, 1 mM EDTA, 1 mM pefablock, 1 µM pepstatin, and 100 µM leupeptin, pH 7.4). After centrifugation (16,000 g, 12 min, room temperature), the supernatant was saved for 2DE as detergent-soluble protein fraction. Fractions of 1.5 mg protein were precipitated with 10% (w/v) trichloroacetic acid (final concentration) and stored at –20°C. For 2DE, samples were thawed and centrifuged at 12,000 g at 4°C for 10 min. Pellets were washed with acetone at –20°C, reprecipitated by centrifugation, and dissolved in 375 µl sample buffer [6 M urea, 2 M thiourea, 4% (w/v) CHAPS, 2% carrier ampholytes, pH 3–10. 1% (w/v) dithiothreitol]. Samples were centrifuged at 12,000 g at room temperature for 5 min and subsequently applied to isoelectric focusing (IEF) in precast Immobiline dry strips (18 cm, pH 3–7 linear pH-gradient) on an IPGphor unit (AmhershamPharmacia, Germany). IEF was performed at 20°C for a total of 50,000 V-hours. Subsequently, IEF strips were equilibrated with in 8 M urea, 20% (v/v) glycerol, 1% (w/v) SDS, and 150 mM Tris-HCl (pH 6.8) and placed on top of a 14% SDS-polyacrylamide gel (20 cm x 20 cm x 1.5 mM). SDS-PAGE was performed using the Laemmli buffer system at 80 V for 16 h. Proteins were stained by colloidal Coomassie G-250 dye (22) , and 2DE gels were scanned at a resolution of 200 dots per inch (dpi) by a scanner calibrated for a linear signal detection range of at least 2 optical density. The resulting images were evaluated with the Z3-software package (Compugen, Israel), and differentially expressed proteins (t test, P<0.05) were selected for identification by mass spectrometry.

Mass spectrometry
Protein spots were excised from the gel and "ingel" digested by trypsin (Promega, Mannheim, Germany; ref 23 ). As a modification of this protocol, peptides were eluted only once by 100% acetonitrile, dried, and, for mass spectrometry, dissolved in 2% TFA (trifluoroacetic acid 99.9%, SDS, Peypin; water: Lichrosolv, Merck). MALDI-TOF mass spectra were acquired on a Reflex III mass spectrometer (Bruker Daltonik) in the reflector mode (positively charged ions) with external calibration. Samples were prepared with the thin-layer technique (24) with {alpha}-cyano-4-hydroxy cinnamic acid (97%, Aldrich, recrystallized from ethanol)/nitrocellulose (Bio-Rad, Trans-Blot Transfer Medium) as matrix. The sample was washed with 0.1% TFA (2°C). HPLC-mass spectrometry (MS)/MS was performed on a QSTAR Pulsar i mass spectrometer (Applera GmbH, Germany) coupled to an Ultimate micro pump (Dionex, Germany). HPLC-column tips (fused silica) with 75 µm inner diameter (New Objective) were self-packed (23) with Reprosil-Pur 120 ODS-3 (Dr. Maisch, Ammerbuch, Germany). A gradient of A [0.5% acetic acid (ACS Reagent, Sigma) in water] and B [0.5 acetic acid in 80% acetonitrile (HPLC gradient grade, ACS; Peypin)] with increasing organic proportion was used for peptide separation. The mass spectrometer was operated in the data-dependent mode and switched automatically between MS and MS/MS. For peptide mapping with MALDI-TOF, the MASCOT-Software (25 ; Matrixscience) in combination with the NCBInr database (National Center for Biotechnology Information) was used for protein identification [monoisotopic m/z, mass accuracy 100 ppm or better, up to one missed cleavage, fixed modification: carbamidomethyl (-Cys), variable modification: oxidized methionine]. In selected cases, nonsignificant MALDI-TOF hits were confirmed by additional LC-MS/MS sequence information.

Reverse transcription and quantitative RT-PCR
Total RNA from myocardium was prepared using the RNeasy Mini kit (Qiagen). Five micrograms of total RNA were reverse transcribed by the SuperScript first-strand synthesis system for RT-PCR (Invitrogen). PCR amplification of the reverse transcribed cDNA was performed using the intercalating SYBR-green dye, cDNA/RNA, Taq-polymerase, and specific primers under the following conditions: 1 cycle for 1 min at 72°C, 50 cycles (94°C for 15 s, 60°C for 30 s, 72°C for 30 s), and 1 cycle at 72°C for 7 min in the MyiQTM single-color real-time PCR detection system (Bio-Rad).

Malondialdehyde measurement
Malondialdehyde measurement (MDA) was determined according to Wong et al. (26) with modifications of Sommerburg et al. (27) . MDA was determined as the thiobarbituric acid (TBA) derivative. The tissue was homogenized in PBS containing 0.5 mM butylated hydroxytoluene (BHT). The modification reaction contained phosphoric acid (440 mM), sample or MDA standard, and TBA solution (42 mM). The reaction mixture was incubated at 100°C for 60 min, and then the samples and standards were cooled on ice. To neutralize the phosphoric acid and to precipitate the proteins before the sample was injected into the HPLC system, the samples and standards were diluted 1:1 (v/v) with NaOH (0.1 M) in methanol. Afterward all samples were centrifuged at 10,000 g for 2 min, and aliquots of supernatants were injected into the reversed phase HPLC and separated by isocratic elution with phosphate buffer (50 mM, pH 6.8) containing 40% (v/v) methanol. The TBA-MDA complex was detected by means of fluorescence using an excitation wavelength of 525 nM and emission of 550 nM.

Protein carbonyl determination
For determination of protein carbonyl groups, the method of Buss et al. (28) with modifications of Sitte et al. was used (28) . After the samples were homogenized and the protein concentration was determined, the samples were diluted to the same concentration of protein (1 mg/ml) and derivatized with dinitrophenylhydrazine (DNPH) solution. Sample loading and washing of ELISA plates were performed as described by Buss et al. (28) . Development was performed using a detection system described by Sitte et al. (29) . Absorbances were determined at 492 nM. A standard curve of oxidized BSA was included in each plate. Blanks of PBS without protein were subtracted from standards and samples absorbances. Oxidized BSA was prepared by modifying solved BSA with hypochlorite. The carbonyl content of the oxidized BSA was determined according to Buss et al. (28) . Reduced BSA was obtained as described by Buss et al. (28) .

Determination of mitochondrial respiration
Ventricular myocard (200 mg) was homogenized in 1 ml of a buffer containing 180 mM KCl and 10 mM EDTA, pH 7.4. The resultant homogenate (50 µl) was preincubated in 2 ml of O2-saturated medium containing 110 mM mannitol, 60 mM Tris, 10 mM NaH2PO4, 1 mM MgCl2, and 0.5 mM EGTA, pH 7.4 at 30°C for 2 min. The functional integrity of mitochondria was controlled by determination of the respiratory control index with and without addition of ADP and assessment of the oligomycine sensitivity of respiration. For the calibration of the Clark-type oxygen electrode, the O2 content of air-saturated incubation medium was taken to be 217 nmol O2/ml (30) . O2 consumption was measured after addition of 5 mM glutamate, 5 mM malate, and 10 µmol oligomycine or after addition of 5 mM succinate, 5 µmol rotenone, and 10 µmol oligomycine. Subsequently, the uncoupling agent carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone (FCCP) was titrated until maximal O2 consumption was reached.

Statistical analysis
Data are arithmetic mean and SE. Comparisons of groups were done by t test (2-sided) or ANOVA for experiments with more than two subgroups. Post hoc range tests and pair wise multiple comparisons were performed with t test (2-sided). Proportion data were analyzed by Pearson’s Chi-Square test. P values ≤ 0.05 were considered as statistically significant.

RESULTS

Morphology and functional impairment of CTSL-deficient hearts
CTSL-deficient mice are fertile and show normal breeding behavior. Pups devoid of CTSL exhibit a higher mortality than their wild-type littermates on weaning (15% ctsl–/– vs. 6% wild type), but thereafter spontaneous mortality of both groups does not show any significant difference during the observation period of up to 1 yr of age (11) . At 2 months of age, hearts of CTSL knockout mice show no gross morphological alteration ex vivo and in vivo (Fig. 1 A, B). However, at 1 yr of age ctsl–/– mice show increased left-ventricular diameters by echocardiographic assessment (Fig. 1B ). Approximately 25% of the hearts will develop pronounced dilation of the left and right ventricles with only little hypertrophy at 12 months of age (arrow, Fig. 1C ). This dilation is usually linked to the development of valve insufficiencies or stenoses causing volume and pressure stress that cannot be not compensated by the CTSL-deficient myocardium (13) . The percentage of fractional shortening is an echocardiographic measure of myocardial contraction. At 2 months of age, ctsl–/– mice already show a small but significant impairment of fractional shortening (Fig. 1D ). This functional impairment progressed during aging, reaching a 50% reduced contraction of hearts from 1-yr-old CTSL knockout mice (Fig. 1D ), even in the absence of extreme dilation (arrow, Fig. 1C ). Transcriptional activation of the natriuretic peptides ANP and BNP in left-ventricular myocardium reflects an increased tension in heart walls (31) . Consistent with the slightly reduced fractional shortening (Fig. 1D ), CTSL-deficient mice show increased mRNA levels of ANP and BNP already in ventricular myocardium of 2-month-old mice as compared with age matched controls (Fig. 1E ).


Figure 1
View larger version (33K):
[in this window]
[in a new window]
  		Figure 1. Morphology and functional impairment of CTSL-deficient (ctsl–/–) hearts. A) Cross-sections (below heart valves) of wild-type (WT; ctsl+/+) and CTSL-deficient (ctsl–/–) mouse ventricles from 2-month-old mice. B) Length of left ventricle (LV) in vivo determined by echocardiography in ctsl+/+ and ctsl–/– hearts at 2 and 12 months of age. C) Cross-sections (below heart valves) of ctsl+/+ and ctsl–/– mouse ventricles from 12-month-old mice. Arrow marks a representative example of the 25% of hearts that develop pronounced dilation of LV and right ventricle (RV). D) Echocardiographic assessment of fractional shortening, a parameter for heart myocardial contraction in 2- and 12-month-old ctsl–/– mice. E) mRNA expression of the natriuretic peptides ANP and BNP in ventricular myocardium of 2-month-old mice determined by quantitative, real-time RT-PCR. n.s. = not significant, **P < 0.01, ***P < 0.001 by Student’s t test.

Fibrosis and ultrastructural alterations in CTSL-deficient myocardium
Development of extensive interstitial fibrosis is a hallmark of cardiac remodeling in the pathogenesis of cardiomyopathies (32) . We used a grading system on Masson’s Trichrome stained ventricular sections for semiquantitative assessment of age-dependent fibrosis in the hearts of ctsl+/+ and ctsl–/– mice (Fig. 2 A). On aging, the wild-type mice show a slight increase in myocardial fibrosis (Fig. 2B ). CTSL-deficient hearts do not reveal increased fibrosis at 2 months of age as compared with the wild type. However, starting at an age of ~4 months ctsl–/– hearts develop increased fibrosis that can be severe (i.e., grade 4, at 1 yr of age). In contrast to fibrosis that appears relatively late in the pathogenesis of the cardiomyopathy, an aberrant myocardial ultrastructure, characterized in part by abnormal accumulations of enlarged lysosomes, is already observed in hearts of 3-day-old ctsl–/– animals (Fig. 2C ). At this age, wild-type myocardium exhibited normal muscle fibers, normal mitochondrial morphology, and only a few small electron-dense lysosomes. In CTSL-knockout mice, the cardiomyocytes contained some large vacuolar lysosomes next to numerous smaller and compact lysosomes. By 2 months of age, ctsl+/+ hearts showed normal tissue ultrastructure, with no vacuolation or accumulations of lysosomes apparent (Fig. 2D ). By contrast, high resolution light microscopy of CTSL-deficient myocardium revealed abundant cytoplasmic vacuolation, often distributed in linear arrays. TEM of longitudinally cut section of cardiac muscle fibers revealed extensive vacuolation, and numerous end-stage lysosomes/residual bodies (pale-staining lysosomes) in addition to active lysosomes (dark-staining lysosomes). Some of the vacuolated structures also revealed features of mitochondria and were closely associated with muscle fibers. Regions of cytoplasmic cell debris located between muscle fibers were also detected. In addition, marked nuclear pleomorphism with clumping of heterochomatism was found in the myocardium of 2-month-old CTSL knockout mice (Fig. 2D ). Identical ultrastructural findings were observed in hearts of 12-month-old CTSL knockout mice (data not shown). Interestingly, these alterations appeared to be specific for the myocardium of ctsl–/– mice, since skeletal muscle of 1-yr-old wild-type and CTSL-deficient mice exhibited identical and normal ultrastructures (Fig. 2E ).


Figure 2
View larger version (89K):
[in this window]
[in a new window]
  		Figure 2. Myocardial fibrosis and ultrastructure in CTSL-deficient myocardium. A) Heart sections were stained with Masson’s Trichrome and scored for grade of fibrosis. B) Interstitial fibrosis during aging of WT (ctsl+/+, closed circles) and CTSL-deficient (ctsl–/–, open circles) hearts *P < 0.05, **P < 0.01 by Pearson’s Chi-Square test. C-E) HRLM (blue images) and TEM (grayscale images) of ctsl+/+ and ctsl–/– myocardium from 3-day (C) and 8-wk-old mice (D), as well as from skeletal muscle of 12-month-old animals (E). Normal cardiac muscles from ctsl+/+ mice show a rich plexus of blood capillaries (Cap) located between individual cardiomyocytes. No vacuolation or accumulations of lysosomes (Ly) is apparent. TEM of ctsl+/+ myocardium (D) shows centrally located oval nuclei (Nu) and intercalated disks (interdigitating intercellular junctions) joining individual cells (ID). Cardiac muscle cell contains numerous mitochondria (Mt), and cell cytoplasm is not vacuolated and contains few distinct lysosomes. HRLM of cardiac muscle fibers from ctsl–/– mice show abundant cytoplasmic vacuolation (Vac), often distributed in linear arrays. Large deeply stained accumulations of lysosomes are visible in perinuclear region of some cardiomyocytes. TEM ctsl–/– cardiomyocytes exhibits extensive vacuolation and numerous end-stage lysosomes/residual bodies (pale-staining lysosomes) in addition to active lysosomes (dark-staining lysosomes). In D a region of cytoplasmic cell debris (CD) is located between 2 muscle fibers marked nuclear pleomorphism, with some regions of this highly extended nucleus exhibiting intense heterochromatin clumping (He). In contrast to findings in ctsl–/– heart muscle, both genotypes show typical normal skeletal muscle architecture in HRLM and TEM (E).

Alterations of the acidic cellular compartment of CTSL-deficient myocardium
The myocardial ultrastructure and transcriptome analysis of ctsl–/– hearts suggested a pathological alteration of the acidic cellular compartment. Labeling of primary cardiomyocytes isolated from newborn ctsl+/+ and ctsl–/– mice with an acidophilic dye (Lysotracker) revealed strong staining in cardiomyocytes of both genotypes compared with "contaminating" cardiac fibroblasts also present in these cell cultures (Fig. 3 A). In contrast to wild-type cardiomyocytes, which show a reticular distribution of acidic organelles, ctsl–/– cardiomyocytes show a patchy pattern of acidophilic staining that most likely correlates with the increased number of lysosome-like organelles seen by EM (Fig. 2C ). These findings were further supported by sucrose-density fractionation of myocardial postnuclear supernatants, with identical total protein content for both genotypes (Fig. 3B ). Activity of the lysosomal/endosomal enzyme ß-hexosaminidase was increased approximately twofold in the density fractions from 1.04 to 1.09 g/ml. Lamp1 was detected by Western blotting in the fractions from 1.08 to 1.10 g/ml in both genotypes, and, in comparison to the wild-type, Lamp1 was more abundant in the ctsl–/– fractions (Fig. 3B ). To test if this alteration of the acidic cellular compartment reflected a lysosomal storage disease, we stained myocardial histological sections of 1-yr-old mice with PAS staining; for the detection of carbohydrates), with Sudan III (intracellular lipid staining), and by the Mowry method for detection of glucosaminoglycan storage (Fig. 4 ). Only the latter revealed any difference between ctsl+/+ and ctsl–/– myocardium, with an associated increase in the expression of glucosaminoglycan storage observed in CTSL knockout mice. However, this increased glucosaminoglycan staining in ctsl–/– hearts was restricted to the extracellular space and therefore highly likely to be associated with the myocardial fibrosis and remodeling detected in older CTSL-deficient animals (Fig. 2A, B ).


Figure 3
View larger version (43K):
[in this window]
[in a new window]
  		Figure 3. Alterations of acidic cellular compartment of CTSL-deficient myocardium. A) Cultured primary cardiomyocytes from neonatel ctsl+/+ and ctsl–/– mice were stained with the acidophilic dye Lysotracker (green), cardiac specific Troponin I (red), and the nuclei with Hoechst #33342 (blue). Note reticular lysosomal staining in ctsl+/+ compared with lysosomal patches in ctsl–/–. B) Equal amounts of total protein from postnuclear supernatants of myocardium from 4-month-old ctsl+/+ and ctsl–/– mice were subjected to sucrose gradient centrifugation. Activity of ß-hesosaminidase (ß-Hex) and Lamp1 was detected by Western blot (WB) in the ctsl+/+ (wt) and ctsl–/– (ko) gradient fractions.


Figure 4
View larger version (180K):
[in this window]
[in a new window]
  		Figure 4. Examination of intracellular storage material in 12-month-old ctsl+/+ and ctsl–/– myocardium. A) Carbohydrate staining. Both genotypes show absence of PAS-positive structures in myocardium. B) Lipid staining Sudan III. Ctsl+/+ and ctsl–/– show several distinct lipid droplets but no difference could be observed between both genotypes. C) Mowry’s colloidal staining (blue color) reveals glucosaminoglycans in extracellular space, which correlates with myocardial fibrosis (as shown in Fig. 2 , A and B). Nuclei stained blue in A and B and pink in C.

Cell death, cell proliferation, and vascular structure in CTSL-deficient myocardium
There are some examples of cardiomyopathies for which an imbalance of apoptotic cell death and cell proliferation in the myocardium has been shown to be an essential pathogenic factor (33) . Furthermore, CTSL knockout mice show hyperproliferation of epidermal keratinocytes and reduced rates of keratinocyte cell death during regression of hair follicles in the catagen phase of the hair cycle (11 , 12) . However, ctsl–/– myocardium did not reveal any difference in the incidence of TUNEL-positive dying cells or Ki67-positive proliferating cells compared with wild-type myocardium in mice at 2–12 months of age (Fig. 5 A, B). CTSL also mediates the invasion of endothelial progenitor cells during postnatal neovascularization, a process contributing to the formation of capillaries in ischemic or hypoxic tissues (10) . However, in the current study, myocardium of CTSL knockout and wild-type mice revealed identical capillary densities at 2 months (data not shown) and 12 months of age (Fig. 5C ).


Figure 5
View larger version (41K):
[in this window]
[in a new window]
  		Figure 5. Cell death, cell proliferation, and vascular structure in CTSL-deficient myocardium. A) DNA fragmentation in cardiac tissue sections was detected by TUNEL method. For each sample, TUNEL-positive and TUNEL-negative nuclei were counted in 9 defined fields at x200 magnification. Two hearts were analyzed per genotype and per age group. Statistical analysis did not reveal any significant difference between groups. B) Proliferation marker Ki67 was detected by immunohistochemistry in cardiac tissue sections. For each sample, Ki67-positive and Ki67-negative nuclei were counted in 9 defined fields at 200-fold magnification. Three hearts were analyzed per genotype and per age group. Statistical analysis did not reveal any significant difference between groups. C) Sections of myocardium (5 µm) from 12-month-old mice were immunostained for the basement membrane glycoprotein Laminin (green) and for platelet endothelial cell adhesion molecule CD31 (red). Ctsl+/+ and ctsl–/–myocardium show identical capillary density and architecture. Arrow indicates a larger blood vessel. Representative for 3 hearts analyzed.

Proteome analysis of ctsl+/+ and ctsl–/– myocardium
To obtain further data on the molecular pathogenesis of the DCM in CTSL knockout mice, we screened the myocardial protein pattern of ctsl+/+ and ctsl–/– mice at ages of 2, 6, 8, and 12 months for differentially expressed proteins by 2DE and identified these proteins by MS (Table 1 , see Supplementary Tables for details on mass spectrometric protein identification). Remarkably, some of these proteins could be detected by our proteome screen in several age groups in which these proteins show similar ratios of protein levels for ctsl–/–/ctsl+/+ (with the exception of the NADH-dehydrogenase Fe-S protein 1; Table 1 ). After the differentially expressed proteins were classified according to their functions, the data reveal reduced levels of proteins involved in myocardial contraction, i.e., tropomyosin 1, calsarcin-1, and desmin, in the myocardium of ctsl–/– mice. Strikingly, the levels of many proteins involved in cellular and mitochondrial catabolism as well as in oxidative phosphorylation are altered in CTSL-deficient hearts (Table 1) . In addition, increased levels of peroxiredoxin 2 and oxidized peroxiredoxin 6 were associated with decreased levels of reduced peroxiredoxin 6, suggesting the presence of significant oxidative stress in the myocardium of CTSL-deficient mice (34) . Such a proteomic comparison of ctsl+/+ and ctsl–/– myocardium permits us to generate several hypotheses on cellular processes underlying DCM development.


View this table:
[in this window]
[in a new window]
  		Table 1. Results of differential proteome analysis of ctsl+/+ and ctsl–/– ventricular myocardiuma

Parameters of oxidative damage and mitochondrial respiration in CTSL-deficient myocardium
The proteome of ctsl–/– hearts revealed up-regulation of some peroxiredoxins that are antioxidant proteins (34) . To assess the extent of damage caused by free oxygen radicals in ctsl–/– hearts, we measured MDA and protein-bound carbonyl-groups that represent stable end-products of lipid peroxidation and oxidative protein damage, respectively (35 , 36) . There was no significant difference in either parameters of oxidative damage in samples from ctsl–/– vs. wild-type myocardium (Fig. 6 A, B), indicating that the antioxidant system in ctsl–/– hearts is able to prevent extensive oxidative damage. Mitochondria are the major source of ATP in the heart muscle (37) . Screening the proteome of CTSL-deficient hearts revealed reduced levels of respiratory chain proteins (Table 1) . Thus, we compared oxygen consumption in myocardial homogenates of 5-month-old ctsl–/– and ctsl+/+ mice (Fig. 6C, D ). Measurement of citrate-synthase activity indicated similar amounts of mitochondria in ctsl–/– and ctsl+/+ homogenates (Fig. 6C , inset). However, with the use of glutamate in combination with malate as respiratory substrates, a reduction of 50% in the oxygen consumption rates in resting and FCCP-uncoupled respiration was observed in ctsl–/– homogenates (Fig. 6C ) compared with homogenates from wild-type mice. Respiration with succinate as substrate was revealed significant impairment in the ctsl–/– group, whereas this effect did not reach significance in presence of FCCP (Fig. 6D ). The impairment of mitochondrial respiration is highly likely to impair ATP production and, therefore, to contribute to the pathogenesis of the cardiomyopathy in CTSL-deficient mice.


Figure 6
View larger version (22K):
[in this window]
[in a new window]
  		Figure 6. Parameters of oxidative damage and mitochondrial respiration. A) Malondialdehyde as a marker for lipidperoxidation was measured in myocardial samples from 2- and 12-month-old ctsl+/+ (column light gray) and ctsl–/– (column dark gray) mice by HPLC and was normalized to total protein in the sample. No significant difference was detected between both genotypes and age groups (n=5). B) Protein-bound carbonyls as an indicator for protein peroxidation were detected by ELISA in ctsl+/+ (column light gray) and ctsl–/– (column dark gray) myocardium. Amount of protein-bound carbonyls normalized to total protein content showed no significant difference between both genotypes at 2 and 12 months (n=5). C–D) Oxygen consumption in myocardial homogenates of 4-month-old mice (n=10) was determined by Clark-electrode measurements with glutamate/malate (C) or succinate (D) as substrates. In measurements for D, rotenone was added for complex I inhibition. FCCP served as uncoupling agent. Rates of respiration are presented in % of air saturation; 100% of air saturation correspond to 217 nmol O2/ml/secxmg protein of homogenate. Inset shows activity of TCA-cycle enzyme citrate synthease in ctsl+/+ (column light gray) and ctsl–/– (column dark gray) myocardial homogenates. **P < 0.01 by Student’s t test

DISCUSSION

This study reports the pathogenesis of DCM in mice with a deficiency for the lysosomal protease CTSL. The essential function of CTSL in the heart is most likely located in the endosomal/lysosomal compartment, because an altered structure of lysosomes in ctsl–/– hearts can be detected already in myocardium and cardiomyocytes of 3-day-old mice. A phase of functional compensation is characteristic for the initial pathogenesis of a cardiomyopathy (14) . However, already during functional compensation, myocardial wall stress is increased and results in the activation of an adaptive "fetal" gene program, as demonstrated in our model for the myocardial expression of the natriuretic factors ANP and BNP in 2-month-old ctsl–/– mice. At this age the morphology and histology of the ctsl–/– hearts were not affected and myocardial contraction showed only small, but significant, impairment. By contrast, myocardial ultastructure was severely impaired in 2-month-old ctsl–/– mice showing numerous end-stage lysosomes/residual bodies and active lysosomes. Interestingly skeletal muscle ultrastructure was not affected, indicating a heart-specific function of CTSL. Increased production of lysosomes is also indicated by a higher abundance of the lysosomal markers ß-hexosaminidase and Lamp1 in ctsl–/– myocardium. Accumulation and abnormal ultrastucture of lysosome-like organellles have been reported in ctsl–/– keratinocytes and in neurons of mice with double deficiency for cathepsins B and L (12 , 38) . This finding is supported by treatment of cultured hypocampal neurons with broad spectrum inhibitors of cysteine cathepsins (39) . Thus, CTSL appears to be critical for the homeostasis of the lysosomal/endosomal compartment in some, but not all, cell types. For CTSL-deficient keratinocytes, we have shown that the recycling of epidermal growth factor (and possibly other growth factors) is enhanced, resulting in sustained mitogenic growth factor signaling and keratinocyte hyperproliferation in the epidermis (12) . In analogy, the impairment of the lysosomal compartment in CTSL-deficient myocardium could cause distinct changes in cardiomyocyte growth factor signaling that results in the complex disease process described by the present data.

Furthermore, the hearts of young adult mice show significant quantitative changes in protein pattern. Loss of function of sarcomeric and cytoskeletal components involved in cardiac force generation and force transmission has been frequently shown to cause cardiomyopathies (17) . For instance, specific point mutations in {alpha}-tropomyosin have been shown to result in either in hypertrophic or dilated cardiomyopathies in humans and mice. Interestingly, we found {alpha}-tropomyosin down-regulated synergistically at mRNA (data not shown) and protein levels. Over the past years, the sarcomeric Z-disc has been proven to be a focal point in the pathogenesis of human cardiomyopathies (40) . Two Z-disc associated proteins, desmin and calsarcin 1, were detected at significantly reduced levels in the hearts of CTSL knockout mice compared with wild-type mice. Interestingly, degeneration of cardiomyocytes and myocardial fibrosis can also be seen in desmin knockout mice (41) . Mice with deficiency for calsarcin-1, a negative regulator of calcineurin, reveal an excess of slow skeletal muscle fibers and an activated hypertrophic gene program, despite the absence of hypertrophy (42) . Thus, reduction of {alpha}-tropomyosin, calsarcin-1, and desmin levels are likely to contribute to impaired cardiac contraction and cardiac remodeling in our mouse model.

Reactive oxygen species (ROS) are known for their ability to damage cellular DNA, phospholipids, and proteins. Therefore, ROS have been extensively studied in the failing heart and evidence for promotion of human heart failure by ROS has been obtained (43) . Two-dimensional electrophoresis-based proteome analyses in our model revealed increased levels of antioxidative peroxiredoxins and heat shock proteins, which are induced under oxidative stress conditions by oxidation-dependent activation of the heat shock factor 1 transcription factor (44) . Most interestingly, the oxidized form of peroxiredoxin 6 was found at increased concentration in CTSL-deficient myocardium, whereas the corresponding reduced form was decreased (34) . These findings indicate an increased ROS generation in the hearts of CTSL null mice. However, the measurement malondialdehyde and protein carbonyls (indicators of ROS induced damage of phospholipids and proteins, respectively) revealed no differences between wild-type and CTSL-deficient myocardium. This indicates that the antioxidative system in ctsl–/– hearts, including the peroxiredoxins, is sufficient to detoxify ROS and to prevent oxidative damage of cell structures. Major intracellular sources of ROS are dysfunctional mitochondria with impaired electron transport and respiration (45) . Remarkably, proteome analysis of CTSL-deficient myocardium revealed a significant reduction in the levels of respiratory chain components compared with wild-type controls. Consistently, a decreased oxidative capacity was detected in myocardial homogenates of ctsl–/– mice. Functional impairment of mitochondria has frequently been demonstrated in animal models with exogenous induction of heart failure (e.g., by aortic banding, myocardial infarction, or pacing) (46 47 48) . Furthermore, tissue specific deletion of the mitochondrial transcription factor A (Tfam) in the mouse myocardium results in decreased respiratory complex activities and DCM (49) . Interestingly, these mice show a metabolic shift from fatty acid oxidation to glycolysis on development of cardiomyopathy (50) , a finding that is consistent with increased levels of glycolytic enzymes (i.e., phosphopyruvate-hydratase (alpha-enolase) and lactate-dehydrogenase), in ctsl–/– myocardium. Thus, it appears that mitochondrial impairment and metabolic alterations are important pathophysiological factors for the progression of cardiomyopathy in CTSL-deficient mice.

We conclude that deficiency of CTSL in the heart primarily affects the lysosomal system, namely by increasing the number and changing the morphology, of acidic organelles. Since lysosomal storage materials were not observed, the cardiomyopathy of CTSL knockout mice can be classified as a lysosomal cardiomyopathy without specific storage. This is an interesting contrast to the accumulation of lysosomal storage materials in other cardiomyopathies caused by defects of lysosomal proteins (e.g., Lamp2 deficiency and glycogen storage in Danon disease) (51 , 52) . The primary defects in the acidic cellular compartment of CTSL-deficient hearts result in complex biochemical and cellular alterations that are hallmarks of cardiomyopathies. These characteristics include the induction of an adaptive gene program, loss of cytoskeletal proteins, and mitochondrial impairment. Together, these interdependent cellular alterations lead to impaired cardiomyocyte function, to cardiac remodeling, and to progressive DCM. Our observation that CTSL plays an essential role in the structure and function of the lysosomal compartment in myocardium, but not apparently in skeletal muscle, requires further investigation.

ACKNOWLEDGMENTS

We thank Susanne Dollwet-Mack and Heidi Bräuner (Institut für Molekulare Medizin und Zellforschung, Freiburg) for excellent technical assistance. This study was supported by a grant of the Deutsche Forschungsgemeinschaft (Re1584/2–1/2–2).

FOOTNOTES

1 These authors contributed equally to this work.

Received for publication November 30, 2005. Accepted for publication January 27, 2006.

REFERENCES

  1. Rawlings, N. D., Barrett, A. J. (2000) MEROPS: the peptidase database. Nucleic Acids Res. 28,323-325[Abstract/Free Full Text]
  2. Puente, X. S., Sanchez, L. M., Overall, C. M., Lopez-Otin, C. (2003) Human and mouse proteases: a comparative genomic approach. Nat. Rev. Genet. 4,544-558[CrossRef][Medline]
  3. Ciechanover, A. (2005) Proteolysis: from the lysosome to ubiquitin and the proteasome. Nat. Rev. Mol. Cell. Biol. 6,79-87[CrossRef][Medline]
  4. Bromme, D., Kaleta, J. (2002) Thiol-dependent cathepsins: pathophysiological implications and recent advances in inhibitor design. Curr. Pharm. Des. 8,1639-1658[CrossRef][Medline]
  5. Yasothornsrikul, S., Greenbaum, D., Medzihradszky, K. F., Toneff, T., Bundey, R., Miller, R., Schilling, B., Petermann, I., Dehnert, J., Logvinova,, et al (2003) Cathepsin L in secretory vesicles functions as a prohormone-processing enzyme for production of the enkephalin peptide neurotransmitter. Proc. Natl. Acad. Sci. U. S. A. 100,9590-9595[Abstract/Free Full Text]
  6. Nakagawa, T., Roth, W., Wong, P., Nelson, A., Farr, A., Deussing, J., Villadangos, J. A., Ploegh, H., Peters, C., Rudensky, A. Y. (1998) Cathepsin L: critical role in Ii degradation and CD4 T cell selection in the thymus. Science 280,450-453[Abstract/Free Full Text]
  7. Reinheckel, T., Hagemann, S., Dollwet-Mack, S., Martinez, E., Lohmuller, T., Zlatkovic, G., Tobin, D. J., Maas-Szabowski, N., Peters, C. (2005) The lysosomal cysteine protease cathepsin L regulates keratinocyte proliferation by control of growth factor recycling. J. Cell Sci. 118,3387-3395[Abstract/Free Full Text]
  8. Cirman, T., Oresic, K., Mazovec, G. D., Turk, V., Reed, J. C., Myers, R. M., Salvesen, G. S., Turk, B. (2004) Selective disruption of lysosomes in HeLa cells triggers apoptosis mediated by cleavage of Bid by multiple papain-like lysosomal cathepsins. J. Biol. Chem. 279,3578-3587[Abstract/Free Full Text]
  9. Goulet, B., Baruch, A., Moon, N. S., Poirier, M., Sansregret, L. L., Erickson, A., Bogyo, M., Nepveu, A. (2004) A cathepsin L isoform that is devoid of a signal peptide localizes to the nucleus in S phase and processes the CDP/Cux transcription factor. Mol. Cell. 14,207-219[CrossRef][Medline]
  10. Urbich, C., Heeschen, C., Aicher, A., Sasaki, K., Bruhl, T., Farhadi, M. R., Vajkoczy, P., Hofmann, W. K., Peters, C., Pennacchio, L. A., et al (2005) Cathepsin L is required for endothelial progenitor cell-induced neovascularization. Nat. Med. 11,206-213[CrossRef][Medline]
  11. Roth, W., Deussing, J., Botchkarev, V. A., Pauly-Evers, M., Saftig, P., Hafner, A., Schmidt, P., Schmahl, W., Scherer, J., Anton-Lamprecht, I., et al (2000) Cathepsin L deficiency as molecular defect of furless: hyperproliferation of keratinocytes and pertubation of hair follicle cycling. FASEB J. 14,2075-2086[Abstract/Free Full Text]
  12. Tobin, D. J., Foitzik, K., Reinheckel, T., Mecklenburg, L., Botchkarev, V. A., Peters, C., Paus, R. (2002) The lysosomal protease cathepsin L is an important regulator of keratinocyte and melanocyte differentiation during hair follicle morphogenesis and cycling. Am. J. Pathol. 160,1807-1821[Abstract/Free Full Text]
  13. Stypmann, J., Glaser, K., Roth, W., Tobin, D. J., Petermann, I., Matthias, R., Monnig, G., Haverkamp, W., Breithardt, G., Schmahl, W., Peters, C., Reinheckel, T. (2002) Dilated cardiomyopathy in mice deficient for the lysosomal cysteine peptidase cathepsin L. Proc. Natl. Acad. Sci. U. S. A. 99,6234-6239[Abstract/Free Full Text]
  14. Francis, G. S. (2001) Pathophysiology of chronic heart failure. Am. J. Med. 110(Suppl),7A, 37S-46S
  15. Mason, J. W. (2003) Myocarditis and dilated cardiomyopathy: an inflammatory link. Cardiovasc. Res. 60,5-10[Abstract/Free Full Text]
  16. Nian, M., Lee, P., Khaper, N., Liu, P. (2004) Inflammatory cytokines and postmyocardial infarction remodeling. Circ. Res. 94,1543-1553[Abstract/Free Full Text]
  17. Morita, H., Seidman, J., Seidman, C. E. (2005) Genetic causes of human heart failure. J. Clin. Invest. 115,518-526[CrossRef][Medline]
  18. Strauch, O. F., Stypmann, J., Reinheckel, T., Martinez, E., Haverkamp, W., Peters, C. (2003) Cardiac and ocular pathologies in a mouse model of mucopolysaccharidosis type VI. Pediatr. Res. 54,701-708[CrossRef][Medline]
  19. Mc Manus, J. F. A. (1948) Histological and histochemical uses of periodic acid. Stain Technol. 23,99-108[Medline]
  20. Mowry, R. W. (1958) Improved procedure for the staining of acidic polysaccharides by Muller’s colloidal (hydrous) ferric oxide and its combination with the Feulgen and the periodic acid-Schiff reactions. Lab. Invest. 7,566-576[Medline]
  21. Pertoft, H., Laurent, T. C., Laas, T., Kagedal, L. (1978) Density gradients prepared from colloidal silica particles coated by polyvinylpyrrolidone (Percoll). Anal. Biochem. 88,271-282[CrossRef][Medline]
  22. Neuhoff, V., Arold, N., Taube, D., Ehrhardt, W. (1988) Improved staining of proteins in polyacrylamide gels including isoelectric focusing gels with clear background at nanogram sensitivity using Coomassie Brilliant Blue G-250 and R-250. Electrophoresis 9,255-262[CrossRef][Medline]
  23. Shevchenko, A., Wilm, M., Vorm, O., Mann, M. (1996) Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68,850-858[Medline]
  24. Kussmann, M. (1997) Matrix-assisted laser desorption/ionisation mass specrtometry sample preparation techniques designed for various peptide and protein analytes. J. Mass. Spectrom. 32,593-601[CrossRef]
  25. Perkins, D. N., Pappin, D. J., Creasy, D. M., Cottrell, J. S. (1999) Probability-based protein identification by searching sequence databases using mass spectrometry data. Electrophoresis 20,3551-3567[CrossRef][Medline]
  26. Wong, S. H., Knight, J. A., Hopfer, S. M., Zaharia, O., Leach, C. N., Jr, Sunderman, F. W., Jr (1987) Lipoperoxides in plasma as measured by liquid-chromatographic separation of malondialdehyde-thiobarbituric acid adduct. Clin. Chem. 33,214-220[Abstract/Free Full Text]
  27. Sommerburg, O., Grune, T., Klee, S., Ungemach, F. R., Siems, W. G. (1993) Formation of 4-hydroxynonenal and further aldehydic mediators of inflammation during bromo-trichloromethane treatment of rat liver cells. Mediators Inflamm. 2,27-31[Medline]
  28. Buss, H., Chan, T. P., Sluis, K. B., Domigan, N. M., Winterbourn, C. C. (1997) Protein carbonyl measurement by a sensitive ELISA method. Free Radic. Biol. Med. 23,361-366[CrossRef][Medline]
  29. Sitte, N., Merker, K., Grune, T. (1998) Proteasome-dependent degradation of oxidized proteins in MRC-5 fibroblasts. FEBS Lett. 440,399-402[CrossRef][Medline]
  30. Reynafarje, B., Costa, L. E., Lehninger, A. L. (1985) O2 solubility in aqueous media determined by a kinetic method. Anal. Biochem. 145,406-418[CrossRef][Medline]
  31. Suttner, S. W., Boldt, J. (2004) Natriuretic peptide system: physiology and clinical utility. Curr. Opin. Crit. Care 10,336-341[CrossRef][Medline]
  32. Maisch, B. (1996) Ventricular remodeling. Cardiology 81(Suppl),1, 2-10
  33. Foo, R. S., Mani, K., Kitsis, R. N. (2005) Death begets failure in the heart. J. Clin. Invest. 115,565-571[CrossRef][Medline]
  34. Chevallet, M., Wagner, E., Luche, S., van Dorsselaer, A., Leize-Wagner, E., Rabilloud, T. (2003) Regeneration of Peroxiredoxins during Recovery after Oxidative Stress: only some overoxidized peroxiredoxins can be reduced during recovery after oxidative stress. J. Biol. Chem. 278,37146-37153[Abstract/Free Full Text]
  35. Levine, R. L., Williams, J. A., Stadtman, E. R., Shacter, E. (1994) Carbonyl assays for determination of oxidatively modified proteins. Methods Enzymol. 233,346-357[Medline]
  36. Draper, H. H., Hadley, M. (1990) Malondialdehyde determination as index of lipid peroxidation. Methods Enzymol. 186,421-431[Medline]
  37. Huss, J. M., Kelly, D. P. (2005) Mitochondrial energy metabolism in heart failure: a question of balance. J. Clin. Invest. 115,547-555[CrossRef][Medline]
  38. Felbor, U., Kessler, B., Mothes, W., Goebel, H. H., Ploegh, H. L., Bronson, R. T., Olsen, B. R. (2002) Neuronal loss and brain atrophy in mice lacking cathepsins B and L. Proc. Natl. Acad. Sci. U. S. A. 99,7883-7888[Abstract/Free Full Text]
  39. Bednarski, E., Ribak, C. E., Lynch, G. (1997) Suppression of cathepsins B and L causes a proliferation of lysosomes and the formation of meganeurites in hippocampus. J. Neurosci. 17,4006-4021[Abstract/Free Full Text]
  40. Perriard, J. C., Hirschy, A., Ehler, E. (2003) Dilated cardiomyopathy: a disease of the intercalated disc?. Trends Cardiovasc. Med. 13,30-38[CrossRef][Medline]
  41. Milner, D. J., Weitzer, G., Tran, D., Bradley, A., Capetanaki, Y. (1996) Disruption of muscle architecture and myocardial degeneration in mice lacking desmin. J. Cell Biol. 134,1255-1270[Abstract/Free Full Text]
  42. Frey, N., Barrientos, T., Shelton, J. M., Frank, D., Rutten, H., Gehring, D., Kuhn, C., Lutz, M., Rothermel, B., Bassel-Duby, R., et al (2004) Mice lacking calsarcin-1 are sensitized to calcineurin signaling and show accelerated cardiomyopathy in response to pathological biomechanical stress. Nat. Med. 10,1336-1343[CrossRef][Medline]
  43. Giordano, F. J. (2005) Oxygen, oxidative stress, hypoxia, and heart failure. J. Clin. Invest. 115,500-508[CrossRef][Medline]
  44. Ahn, S. G., Thiele, D. J. (2003) Redox regulation of mammalian heat shock factor 1 is essential for Hsp gene activation and protection from stress. Genes Dev. 17,516-528[Abstract/Free Full Text]
  45. Nohl, H., Kozlov, A. V., Gille, L., Staniek, K. (2003) Cell respiration and formation of reactive oxygen species: facts and artefacts. Biochem. Soc. Trans. 31,1308-1311[Medline]
  46. Garnier, A., Fortin, D., Delomenie, C., Momken, I., Veksler, V., Ventura-Clapier, R. (2003) Depressed mitochondrial transcription factors and oxidative capacity in rat failing cardiac and skeletal muscles. J. Physiol. 551,491-501[Abstract/Free Full Text]
  47. Ide, T., Tsutsui, H., Hayashidani, S., Kang, D., Suematsu, N., Nakamura, K., Utsumi, H., Hamasaki, N., Takeshita, A. (2001) Mitochondrial DNA damage and dysfunction associated with oxidative stress in failing hearts after myocardial infarction. Circ. Res. 88,529-535[Abstract/Free Full Text]
  48. Marin-Garcia, J., Goldenthal, M. J., Moe, G. W. (2001) Abnormal cardiac and skeletal muscle mitochondrial function in pacing-induced cardiac failure. Cardiovasc. Res. 52,103-110[CrossRef][Medline]
  49. Li, H., Wang, J., Wilhelmsson, H., Hansson, A., Thoren, P., Duffy, J., Rustin, P., Larsson, N. G. (2000) Genetic modification of survival in tissue-specific knockout mice with mitochondrial cardiomyopathy. Proc. Natl. Acad. Sci. U. S. A. 97,3467-3472[Abstract/Free Full Text]
  50. Hansson, A., Hance, N., Dufour, E., Rantanen, A., Hultenby, K., Clayton, D. A., Wibom, R., Larsson, N. G. (2004) A switch in metabolism precedes increased mitochondrial biogenesis in respiratory chain-deficient mouse hearts. Proc. Natl. Acad. Sci. U. S. A. 101,3136-3141[Abstract/Free Full Text]
  51. Tanaka, Y., Guhde, G., Suter, A., Eskelinen, E. L., Hartmann, D., Lullmann-Rauch, R., Janssen, P. M., Blanz, J., von Figura, K., Saftig, P. (2000) Accumulation of autophagic vacuoles and cardiomyopathy in LAMP-2-deficient mice. Nature 406,902-906[CrossRef][Medline]
  52. Nishino, I., Fu, J., Tanji, K., Yamada, T., Shimojo, S., Koori, T., Mora, M., Riggs, J. E., Oh, S. J., Koga, Y., Sue, C. M., et al (2000) Primary LAMP-2 deficiency causes X-linked vacuolar cardiomyopathy and myopathy (Danon disease). Nature 406,906-910[CrossRef][Medline]



This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
D. Spira, J. Stypmann, D. J. Tobin, I. Petermann, C. Mayer, S. Hagemann, O. Vasiljeva, T. Gunther, R. Schule, C. Peters, et al.
Cell Type-specific Functions of the Lysosomal Protease Cathepsin L in the Heart
J. Biol. Chem., December 21, 2007; 282(51): 37045 - 37052.
[Abstract] [Full Text] [PDF]

Home page
 HypertensionHome page
X. W. Cheng, K. Obata, M. Kuzuya, H. Izawa, K. Nakamura, E. Asai, T. Nagasaka, M. Saka, T. Kimata, A. Noda, et al.
Elastolytic Cathepsin Induction/Activation System Exists in Myocardium and Is Upregulated in Hypertensive Heart Failure
Hypertension, November 1, 2006; 48(5): 979 - 987.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Summary
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
fj.05-5517fjev1
20/8/1266    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Petermann, I.
Right arrow Articles by Reinheckel, T.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Petermann, I.
Right arrow Articles by Reinheckel, T.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS