Nonmyocyte ERK1/2 signaling contributes to load-induced cardiomyopathy in Marfan mice

Abstract

Among children with the most severe presentation of Marfan syndrome (MFS), an inherited disorder of connective tissue caused by a deficiency of extracellular fibrillin-1, heart failure is the leading cause of death. Here, we show that, while MFS mice (Fbn1C1039G/+ mice) typically have normal cardiac function, pressure overload (PO) induces an acute and severe dilated cardiomyopathy in association with fibrosis and myocyte enlargement. Failing MFS hearts show high expression of TGF-β ligands, with increased TGF-β signaling in both nonmyocytes and myocytes; pathologic ERK activation is restricted to the nonmyocyte compartment. Informatively, TGF-β, angiotensin II type 1 receptor (AT1R), or ERK antagonism (with neutralizing antibody, losartan, or MEK inhibitor, respectively) prevents load-induced cardiac decompensation in MFS mice, despite persistent PO. In situ analyses revealed an unanticipated axis of activation in nonmyocytes, with AT1R-dependent ERK activation driving TGF-β ligand expression that culminates in both autocrine and paracrine overdrive of TGF-β signaling. The full compensation seen in wild-type mice exposed to mild PO correlates with enhanced deposition of extracellular fibrillin-1. Taken together, these data suggest that fibrillin-1 contributes to cardiac reserve in the face of hemodynamic stress, critically implicate nonmyocytes in disease pathogenesis, and validate ERK as a therapeutic target in MFS-related cardiac decompensation.

Full text

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RESEARCH ARTICLE
Authorship note: E.G. MacFarlane
and E. Takimoto contributed equally
to this work.
Conflict of interest: The authors have
declared that no conflict of interest
exists.
Submitted: November 3, 2016
Accepted: June 29, 2017
Published: August 3, 2017
Reference information:
JCI Insight. 2017;2(15):e91588.
https://doi.org/10.1172/jci.
insight.91588.
Nonmyocyte ERK1/2 signaling contributes
to load-induced cardiomyopathy in Marfan
mice
Rosanne Rouf,1 Elena Gallo MacFarlane,2 Eiki Takimoto,1 Rahul Chaudhary,1 Varun Nagpal,2
Peter P. Rainer,1 Julia G. Bindman,2 Elizabeth E. Gerber,2 Djahida Bedja,1 Christopher Schiefer,1
Karen L. Miller,1 Guangshuo Zhu,1 Loretha Myers,2 Nuria Amat-Alarcon,1 Dong I. Lee,1
Norimichi Koitabashi,1 Daniel P. Judge,1 David A. Kass,1 and Harry C. Dietz2,3
1Division of Cardiology, Department of Medicine, and 2McKusick-Nathans Institute of Genetic Medicine, Johns Hopkins
University School of Medicine, Baltimore, Maryland, USA. 3Howard Hughes Medical Institute, Bethesda, Maryland, USA.
Introduction
Heterozygous mutations in the gene encoding fibrillin-1 (FBN1) cause Marfan syndrome (MFS), a system-
ic connective tissue disorder characterized by manifestations in the ocular, skeletal, and cardiovascular
systems, including aortic root aneurysm (1). Heart failure is the leading cause of death in young children
with MFS (2, 3). While the prevailing view is that cardiac decompensation is a consequence of hemody-
namic load imposed by associated mitral valvular regurgitation, some clinical studies of MFS patients
have reported ventricular enlargement or cardiac dysfunction out of proportion to the severity of valvu-
lar disease (4–7), suggesting either a concomitant primary dilated cardiomyopathy (DCM) or myocardial
vulnerability to mechanical stress (8). Elucidation of the mechanism of cardiac dysfunction in MFS has
important implications for the determination of surgical thresholds for valve repair and for development of
novel medical therapies for heart failure.
Fibrillin-1–rich microfibrils are a critical component of the extracellular matrix (ECM) in many tis-
sues (9). The Fbn1C1039G/+ mouse model of MFS harbors a heterozygous cysteine substitution in an epider-
mal growth factor–like domain in fibrillin-1 representative of the most common class of mutation in MFS
patients (1). Such mutations result in normal mRNA expression but decreased fibrillin-1 deposition in
the ECM (1). Studies in Fbn1C1039G/+ mice have suggested that failed matrix sequestration of latent TGF-β,
with consequent increased cytokine activity, drives many manifestations of MFS, including emphysema,
skeletal muscle myopathy, aortic aneurysm, and myxomatous valvular disease (10–13). All of these man-
ifestations are attenuated in Fbn1C1039G/+ mice by pharmacological antagonism of the canonical TGF-β
signaling (Smad2/3) pathway with either TGF-β–neutralizing antibody (NAb) (10–14) or the angiotensin
Among children with the most severe presentation of Marfan syndrome (MFS), an inherited
disorder of connective tissue caused by a deficiency of extracellular fibrillin-1, heart failure is the
leading cause of death. Here, we show that, while MFS mice (Fbn1C1039G/+ mice) typically have normal
cardiac function, pressure overload (PO) induces an acute and severe dilated cardiomyopathy in
association with fibrosis and myocyte enlargement. Failing MFS hearts show high expression of
TGF-β ligands, with increased TGF-β signaling in both nonmyocytes and myocytes; pathologic
ERK activation is restricted to the nonmyocyte compartment. Informatively, TGF-β, angiotensin II
type 1 receptor (AT1R), or ERK antagonism (with neutralizing antibody, losartan, or MEK inhibitor,
respectively) prevents load-induced cardiac decompensation in MFS mice, despite persistent PO. In
situ analyses revealed an unanticipated axis of activation in nonmyocytes, with AT1R-dependent
ERK activation driving TGF-β ligand expression that culminates in both autocrine and paracrine
overdrive of TGF-β signaling. The full compensation seen in wild-type mice exposed to mild PO
correlates with enhanced deposition of extracellular fibrillin-1. Taken together, these data suggest
that fibrillin-1 contributes to cardiac reserve in the face of hemodynamic stress, critically implicate
nonmyocytes in disease pathogenesis, and validate ERK as a therapeutic target in MFS-related
cardiac decompensation.
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II type 1 receptor (AT1R) blocker (ARB), which suppresses expression of TGF-β receptors and ligands
(12, 14). AT1R-dependent signaling cascades also converge on the ERK1/2 pathway (15), which has also
been shown to be activated in MFS aortas; selective inhibition of ERK1/2 activation using RDEA119
(refametinib) rescues aortic growth and wall architecture in MFS mice (16). Intriguingly, tissue analysis
in non-MFS rodent and human heart failure reveals evidence for increased TGF-β signaling (17–20) as
well as increased deposition of collagens type I and III (21, 22), which are encoded by TGF-β–responsive
genes. Furthermore, in myocardial cells, TGF-β expression and secretion can be induced by angiotensin
II (AngII) signaling (23) and mechanical stretch (24), respectively, and have been shown to drive the tran-
scriptional changes that accompany the transition from compensated load-induced hypertrophy to heart
failure (25–27).
In the myocardium, fibrillin-1 is a predominantly nonmyocyte-derived ECM protein (28, 29) that is
localized in perivascular and perimysial spaces, closely paralleling the spatial arrangement of collagen
(20, 30). Recent studies in Fbn1C1039G/+ mice have shown late-onset development of DCM, which uniformly
associates with valvular regurgitation and increased myocardial ERK1/2 signaling (31); however, these
studies do not segregate the effects of perturbed hemodynamic load from intrinsic defects in the myocar-
dium. Similarly, Fbn1mgR/mgR mice, in which a homozygous hypomorphic allele of Fbn1 causes severe defi-
ciency of fibrillin-1 throughout development, develop DCM associated with severe valvular regurgitation
(32). Conditional deletion of Fbn1, using homozygous Fbn1lox/lox mice in combination with a cardiomyo-
cyte-specific αMHC-Cre-recombinase driver (Fbn1αMHC–/–), also associates with structural and functional
changes consistent with DCM (32). Although this latter study seems to point to an intrinsic defect in
cardiomyocytes as the primary driver of DCM in the context of profound deficiency of fibrillin-1 present
throughout development, the relevance of these findings to the broad clinical spectrum of MFS patients
with heterozygous mutations that impose a relative and postnatally progressive deficiency of extracellular
fibrillin-1 remains unclear.
To address these issues, we studied young Fbn1C1039G/+ mice (prior to valvular or myocardial dysfunc-
tion) in combination with a conditional provocation (transverse aortic constriction [TAC]) to impose acute
hemodynamic load. Comprehensive assessment of cardiac function, gross morphology, and cellular sig-
naling events in tight temporal sequence with TAC allowed a precise definition of gene-by-environment
interactions in the pathogenesis of MFS-associated DCM.
Results
Fbn1C1039G/+ mice are predisposed to load-induced heart failure. We performed serial echocardiography and pres-
sure-volume (PV) loop analysis in Fbn1C1039G/+ mice. Echocardiography of Fbn1C1039G/+ mice showed that the
presence of valvular regurgitation (mitral and/or aortic) associated directly with increased end-diastolic
(r = 0.82, P < 0.001) and end-systolic (r = 0.84, P < 0.0005) diameters and inversely with fractional short-
ening (r = –0.7792, P < 0.005) (Supplemental Figure 1A; supplemental material available online with this
article; https://doi.org/10.1172/jci.insight.91588DS1). However, in mice that did not manifest valvular
regurgitation, as assessed by color flow Doppler, cardiac dimensions and function were indistinguishable
between WT and Fbn1C1039G/+ mice (Supplemental Figure 1A). PV loop analysis, which isolates assessment
of cardiac function from loading conditions on the ventricle (33), also showed that systolic and diastolic
parameters were identical between WT and Fbn1C1039G/+ hearts in the absence of valvular regurgitation at
4 or 12 months of age (Supplemental Figure 1, B and C). Finally, up to 12 months of age, heart weights
(Supplemental Figure 1D) and left ventricular activation of Smad2 (phosphorylated [pSmad2]) or ERK1/2
(phosphorylated [pERK1/2]) was unaltered in Fbn1C1039G/+ mice (Supplemental Figure 2, A–C), as were
expression levels of prototypical TGF-β target genes Ctgf and Serpine1 (Supplemental Figure 2D). Impor-
tantly, these findings suggest that, in the unstressed heart, cardiac structure and function is normal in the
Fbn1C1039G/+ mouse model of MFS.
To study whether the cardiac response to increased hemodynamic load is altered in Fbn1C1039G/+ hearts, we
exposed WT and littermate Fbn1C1039G/+ mice without valvular regurgitation to TAC (referred to as WT:TAC
and Fbn1C1039G/+:TAC mice), using a magnitude of aortic constriction titrated to cause a mild pressure over-
load (PO). Baseline cardiac dimensions and function were indistinguishable between Fbn1C1039G/+ and WT
control mice (Supplemental Figure 3A). However, after 1 week of TAC, Fbn1C1039G/+ mice uniquely showed
marked ventricular dilatation and dysfunction, which progressed in the ensuing 3 weeks (Figure 1, A and B).
Rare Fbn1C1039G/+ and WT mice that had developed valvular regurgitation were excluded from all analyses.
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Postmortem analysis after 4 weeks of TAC revealed a profound increase in heart size and mass in Fbn1C1039G/+
mice compared with WT mice subjected to equivalent TAC (Figure 1C). Marked lung congestion (wet lung
weight) (Figure 1C) occurred as an expected correlate with left heart failure in Fbn1C1039G/+:TAC mice. More-
over, histological analysis disclosed a marked increase in cardiomyocyte cross-sectional area and interstitial
fibrosis in Fbn1C1039G/+:TAC hearts when compared with WT animals subjected to TAC (Figure 1, D and E,
and Supplemental Figure 3, B and C). The expression of myocyte-specific hypertrophy-related genes, atrial
Figure 1. Fbn1C1039G/+ mice develop marked heart failure acutely from pressure overload. (A) Weekly temporal changes in cardiac dimensions and function
after TAC. EDD, end-diastolic diameter; ESD, end-systolic diameter; FS, fractional shortening; LV mass, left ventricular mass; BL, baseline. Comparison
pairs shown in brackets on right side of panels. n = 4–8 per group. (B) Representative M-mode echocardiogram 4 weeks after TAC. White arrow, end-di-
astolic diameter; yellow arrow, end-systolic diameter. (C) Representative whole hearts and summary results for heart weight normalized to tibia length
(HW/TL) in Fbn1+/+ and Fbn1C1039G/+ mice subjected to 4 weeks of TAC. Wet lung weight (LW) was normalized to TL. n ≥ 5 per group. *P < 0.05, **P < 0.01,
***P < 0.001; ##P < 0.01 vs. Fbn1+/+:sham; ††P < 0.01 vs. Fbn1C1039G/+:sham, 2-way ANOVA, Bonferroni’s correction. (D and E) Representative hematoxylin
and eosin and Masson’s trichrome staining of formalin-fixed heart sections from Fbn1+/+ and Fbn1C1039G/+ mice subjected to 4 weeks of TAC. Scale bars: 75
μm (D); 150 μm (E). White dotted line (in D) outlines myocyte circumference. Blue stain (in E) indicates fibrosis.
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natriuretic peptide (Nppa), brain natriuretic peptide (Nppb), and β–myosin heavy chain (Myh7) (26, 34), was
significantly increased in Fbn1C1039G/+ hearts when compared with WT hearts subjected to TAC (Supplemen-
tal Figure 3D) but was comparable between sham groups. We found that though cardiac expression of Fbn1
mRNA was significantly increased in Fbn1C1039G/+ mice and WT controls in response to TAC (Figure 2A),
deposition of fibrillin-1 protein in the interstitial pericapillary and perivascular spaces adjacent to cardiac
Figure 2. Fibrillin-1 protein fails to deposit in myocardium in load-induced heart failure in Fbn1C1039G/+ mice. (A) mRNA expression of Fbn1 normalized to
Gapdh and then to Fbn1+/+:sham data, assessed by real-time RT-PCR. n ≥ 4 per group. (B) Representative images of immunofluorescent fibrillin-1 staining
(red) of heart sections from Fbn1+/+ and Fbn1C1039G/+ mice subjected to 4 weeks of TAC. Red, fibrillin-1 (interstitial space); blue, DAPI (nuclei); green, lipofus-
cin (myocytes). Scale bar: 20 μm. (C) Quantitative representation of fibrillin-1 normalized to unit area. n = 4–7 per group. (D and E) Representative Western
blot and summary quantification for phosphorylated/total (p/t) Smad2 and ERK1/2 using left ventricular tissue lysates. Lanes were run on the same gel
but were noncontiguous. n ≥ 4 per group. (F) mRNA expression of TGF-β targets, normalized to 18S and then to Fbn1+/+ data, assessed by real-time RT-PCR
from left ventricular tissue. Col1a1, collagen type 1 a1; Fn1, fibronectin; Serpine1, plasminogen activator inhibitor, type 1; Postn, periostin; Ctgf, connective
tissue growth factor; Spp1, osteopontin. n = 3–6 per group. *P < 0.05, **P < 0.01, ***P < 0.001, 1-way ANOVA, Tukey’s correction. In box-and-whisker plots,
the lower and upper margins of each box define the 25th and 75th percentiles, respectively; the internal line defines the median, and the whiskers define
the range. Values outside 1.5 times the interquartile distance are shown.
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nonmyocytes was increased in the compensated WT heart but not in decompensated Fbn1C1039G/+ hearts
exposed to TAC (Figure 2, B and C, and Supplemental Figure 4). Thus, while MFS hearts have no demon-
strable cardiac dysfunction at baseline, they do display an abnormal response to mechanical load, revealing
that fibrillin-1 plays a critical role in the adaptive response to hemodynamic stress.
Canonical TGF-β signaling contributes to load-induced cardiac decompensation in Fbn1C1039G/+ mice. To gain insight
into the mechanism of load-induced heart failure in Fbn1C1039G/+ mice, we measured changes in the activation
of the Smad2 and ERK1/2 signaling pathways, which are known to mediate the progression of aortic aneu-
rysm in MFS mice (12, 16). Immunoblots showed increased levels of pSmad2 and pERK1/2 in Fbn1C1039G/+
Figure 3. Smad2 and ERK1/2 activation are dierentially increased in the myocyte and nonmyocyte compartments of failing Fbn1C1039G/+ hearts. (A)
Representative sections of pSmad2 (red) immunostaining. Scale bar: 20 μm. White arrowheads point to examples of positively stained myocyte nuclei.
(B) Summary quantification of total intensity normalized to Fbn1+/+:sham and labeling index (percentage positive cells) of pSmad2 in the nonmyocyte and
myocyte compartments. (C and D) Representative sections and summary quantification of pERK1/2 (red) immunostaining. Scale bar: 20 μm. n = 6 fields
per group. **P < 0.01, ***P < 0.001, 1-way ANOVA, Tukey’s correction.
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hearts when compared with WT controls after 4 weeks of TAC (Figure 2, D and E). Increased expression of
prototypical TGF-β–driven gene products, such as Col1a1, Postn, Serpin1, Ctgf, Spp1, and Fn1 (Figure 2F), was
also observed in Fbn1C1039G/+:TAC hearts when compared with WT:TAC controls. In situ analyses of heart
sections by immunofluorescence showed increased pSmad2 and pERK1/2 in Fbn1C1039G/+:TAC hearts when
compared with WT:TAC controls (Figure 3). Furthermore, in Fbn1C1039G/+:TAC hearts, pSmad2 colocalized
Figure 4. Treatment with TGF-β NAb, losartan, or MEKi improves load-induced cardiac decompensation in Fbn1C1039G/+ hearts, despite persistent
load. (A) Temporal changes of cardiac dimensions and function of Fbn1C1039G/+ hearts after TAC, with treatment arms. Fbn1+/+, blue circle with solid line;
Fbn1C1039G/+, red square with solid line; Fbn1C1039G/+ with TGF-β NAb treatment, orange triangle with dotted line; Fbn1C1039G/+ with losartan (Los) treatment,
green diamond with dotted line; Fbn1C1039G/+ with MEK1/2 inhibitor (MEKi) treatment, purple circle with dotted line. Early time point, 1 week after TAC; mid
time point, 2–4 weeks after TAC; end time point, 3–5 weeks after TAC. EDD, end-diastolic diameter; ESD, end-systolic diameter; FS, fractional shortening;
LV mass, left ventricular mass; BL, baseline. Comparison pairs shown in brackets on right side of respective panel. n ≥ 5 per group. (B) Representative
M-mode echocardiograms at end time point, Fbn1C1039G/+ sham vs. TAC groups, with and without treatment. (C) Summary quantification of heart weight
normalized to tibia length (HW/TL). n ≥ 5 per group. *P < 0.05, **P < 0.01, ***P < 0.001, 1-way ANOVA, Tukey’s correction.
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with both nonmyocyte and myocyte compartments (Figure 3, A and B; Supplemental Figure 5, A and B, top;
and Supplemental Figure 5, C and D, left), whereas enhanced pERK1/2 colocalized predominantly with the
nonmyocyte compartment (Figure 3, C and D; Supplemental Figure 5, A and B, bottom; and Supplemental
Figure 5, C and D, right). Further characterization of pSmad2-positive and pERK1/2-positive nonmyocytes
showed positive costaining of vimentin in a subset of cells (Supplemental Figure 5D), suggesting nonmyo-
cytes to be of fibroblast, endothelial, or myeloid origin (35).
Figure 5. Functional improvement of mechanically loaded Fbn1C1039G/+ hearts correlates with reduced fibrosis. (A) Representative Masson’s trichrome–
stained heart sections of Fbn1C1039G/+ sham vs. TAC groups, with and without treatment, showing fibrosis stained in blue. Scale bars: 750 μm (top); 100 μm
(bottom). (B) Quantification of fibrosis, quantified by total number of blue pixels normalized to whole tissue area and then to Fbn1+/+:sham data. n = 4–7
per group. (C) mRNA expression of TGF-β–responsive fibrogenic genes, normalized to Gapdh and then to Fbn1+/+:sham data, assessed by real-time RT-PCR.
Col1a1, collagen type 1 a1; Spp1, osteopontin. n = 4–6 per group. *P < 0.05, **P < 0.01, ***P < 0.001, 1-way ANOVA, Tukey’s correction. In box-and-whisker
plots, the lower and upper margins of each box define the 25th and 75th percentiles, respectively; the internal line defines the median, and the whiskers
define the range. Values outside 1.5 times the interquartile distance are shown.
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We next assessed for contributions of TGF-β, AT1R, and ERK1/2 signaling by pretreating Fbn1C1039G/+
mice systemically with a pan-specific TGF-β–neutralizing polyclonal antibody (NAb) (12), an ARB (losar-
tan) (12), or a selective MEK1/2 inhibitor (MEKi, PD98059 or RDEA119) (16, 36), respectively. Cultured
cardiac fibroblasts were used to show that MEKi antagonizes phosphorylation of ERK1/2 but not Smad2
(Supplemental Figure 6) as expected (16, 37, 38).
Serial echocardiography showed that NAb, losartan, or MEKi treatment substantially improved end-di-
astolic and end-systolic dimensions, fractional shortening, and left ventricular mass in Fbn1C1039G/+:TAC
hearts when compared with vehicle-treated counterparts (Figure 4). Therapeutic effects correlated with
a significant reduction in fibrotic area (Figure 5, A and B). Furthermore, TAC-induced upregulation of
TGF-β–responsive fibrogenic genes, collagen (Col1a1) and osteopontin (Spp1) (39–41), was attenuated by
NAb treatment and abrogated by either losartan or MEKi (Figure 5C). Curiously, myocyte size, which
is increased in Fbn1C1039G/+:TAC hearts, was normalized upon treatment with losartan or MEKi but not
with NAb (Supplemental Figure 7, A–C). Paralleling these observations, the hypertrophic gene program
induced by TAC in Fbn1C1039G/+ mice was unaltered by NAb treatment but was significantly suppressed by
either losartan or MEKi (Supplemental Figure 7D). Thus, the comparable therapeutic efficacy of NAb,
losartan, and MEKi associate with a reduction of fibrosis but not with myocyte hypertrophy.
We next examined Smad2 and ERK1/2 signaling in Fbn1C1039G/+ hearts subjected to TAC and treated
with NAb, losartan, or MEKi in an attempt to define the pathogenic sequence that culminates in cardiac
decompensation. The therapeutic efficacies of NAb, losartan, and MEKi correlated with reduced levels
of pSmad2 (Figure 6, A–D), supporting the notion that proximal events in TGF-β signaling contribute to
load-induced heart failure. However, with NAb treatment, pERK1/2 levels were unaltered (Figure 6, A and
B), suggesting that ERK1/2 activation occurs independently or upstream of TGF-β signaling. Informa-
tively, MEKi treatment normalized both pERK1/2 and pSmad2 levels, suggesting that ERK1/2 signaling
contributes directly or indirectly to induction of TGF-β canonical signaling. Losartan also normalized both
pERK1/2 and pSmad2 levels, suggesting an AT1R:ERK1/2:Smad2 axis of activation (Figure 6, C–E). In
further support of an AT1R:ERK1/2:Smad2 axis of activation, we show that AngII treatment of murine
cardiac fibroblasts after 24 hours results in increased activation of ERK1/2 but not Smad2 (Supplemental
Figure 8). Whether or not these events represent cell-autonomous relationships remained to be determined.
Losartan or MEKi potently suppresses Smad2 activation in both nonmyocyte and myocyte compartments. To
determine relationships of signaling pathways in nonmyocyte and myocyte compartments, we performed
in situ analysis of pSmad2 and pERK1/2 in Fbn1C1039G/+:TAC hearts treated with NAb, losartan, or MEKi.
NAb treatment achieved a marked decrease in nonmyocyte Smad2 activation, whereas myocyte Smad2
activation largely persisted (Figure 7A and Supplemental Figure 9A). In contrast, treatment with both
losartan and MEKi resulted in full normalization of pSmad2 in both cellular compartments. Losartan or
MEKi achieved a marked decrease in nonmyocyte ERK1/2 activation; however, NAb had no effect (Figure
7B and Supplemental Figure 9B). All three treatments reduced vimentin reactivity (Figure 7) and prevented
TAC-induced expansion of the nonmyocyte compartment in Fbn1C1039G/+ mice (Supplemental Figure 9C).
The failure of NAb to suppress myocyte Smad2 signaling likely reflects a bioavailability issue as previously
discussed (42). Taken together, these data are consistent with a model that invokes a primary increase in
AT1R-mediated ERK1/2 activation in nonmyocytes that is required for subsequent Smad2 activation in
both nonmyocytes and myocytes.
ERK1/2 activation is pivotal in the progression of load-induced heart failure and upregulates TGF-β ligand expres-
sion. We postulated that ERK1/2 signaling may induce nonmyocyte TGF-β ligand production, thereby driv-
ing canonical TGF-β signaling in both an autocrine and paracrine manner. In keeping with this hypothesis,
we found that TAC increased expression of Tgfb2 and Tgfb3 in Fbn1C1039G/+ hearts that was normalized with
losartan or MEKi but not NAb (Figure 8). Furthermore, in situ analysis demonstrated that the expression
of TGF-β3 mRNA localized predominantly to the nonmyocyte compartment (Supplemental Figure 10, A
and B), similarly to that of vimentin, a mesenchymal cell marker that is upregulated in fibroblast-to-myofi-
broblast transformation (43) (Supplemental Figure 10C). Cultured cardiac fibroblasts demonstrate a 12-fold
higher expression of Tgfb3 compared with cultured cardiomyocytes (Supplemental Figure 10D). AngII treat-
ment of cardiac fibroblasts after 12 hours resulted in ERK1/2 activation but not direct Smad2 activation,
while TGF-β3 treatment showed intact Smad2 activation (Supplemental Figure 11). Furthermore, cardiac
fibroblasts cultured from Fbn1C1039G/+ hearts demonstrated even greater sensitivity to ERK1/2 activation in
response to AngII treatment than WT hearts (Supplemental Figure 11). Taken together, these data identify a
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mechanism by which ERK1/2 activation may promote the deleterious effects of TGF-β signaling. This may
be particularly important in the fibrillin-1–deficient myocardium of the hemodynamically loaded MFS heart.
Discussion
While prior reports have suggested a tendency for cardiac decompensation in MFS, it has been difficult
to determine with confidence if this relates to an intrinsic myocardial predisposition, the influence of
valve dysfunction, or a combination of the two. This ambiguity relates to the high frequency of mitral
Figure 6. ERK1/2 activation is upstream of Smad2 activation in Fbn1C1039G/+ hearts. (A–D) Representative Western blot and summary quantification for
Smad2 and ERK1/2 phosphorylation from left ventricular tissue lysates. n = 4–6 per group. *P < 0.05, **P < 0.01, ***P < 0.001, 1-way ANOVA, Tukey’s
correction. (E) Diagram illustrating the pathogenic sequence of load-induced heart failure in the fibrillin-1–deficient myocardium of Fbn1C1039G/+ mice. NM,
nonmyocytes; M, myocytes. In box-and-whisker plots, the lower and upper margins of each box define the 25th and 75th percentiles, respectively; the
internal line defines the median, and the whiskers define the range. Values outside 1.5 times the interquartile distance are shown.
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and/or aortic valve regurgitation in people and mice with MFS that can be difficult to quantitate and the
use of mouse models with targeted disruption of both Fbn1 alleles that show accelerated cardiovascular
disease (32) but uncertain mechanistic relevance to Marfan patients with heterozygous mutations and a
deficiency of extracellular fibrillin-1 that progresses postnatally. In keeping with a critical contribution
of hemodynamic stress, heart failure and infantile death in children with the most severe presentation of
MFS uniformly associate with significant volume overload (44, 45) and can be delayed or even prevented
by aggressive surgical intervention for valve dysfunction (46–48). In this study, we utilized a standardized
PO (via TAC) as a conditional provocation in young mice heterozygous for a typical MFS-associated
missense mutation in fibrillin-1. Importantly, these animals had no abnormality of heart structure or
function at the onset of the study, and only mice that did not have any valvular regurgitation during
Figure 7. Inhibition of ERK1/2 activation suppresses Smad2 activation in both nonmyocyte and myocyte compartments. Representative coimmu-
nostaining of (A) pSmad2 (red, row 1) and vimentin (pink, row 2) and of (B) pERK1/2 (red, row 1) and vimentin (pink, row 2) of myocyte-enriched sec-
tions of Fbn1C1039G/+ hearts subjected to sham vs. TAC, with treatment groups. Scale bars: 20 μm; zoom inset: ×2 of nonmyocyte-enriched areas. Blue,
DAPI (nuclei); green, lipofuscin (myocytes).
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the entire follow-up period were analyzed. MFS mice
exposed to TAC showed acute and profound DCM and
heart failure, while their WT counterparts remained ful-
ly compensated. Furthermore, decompensation in MFS
associated with aberrant activation of cellular signal-
ing in the myocardium, including both the TGF-β and
ERK1/2 pathways. Pharmacologic attenuation of aber-
rant signaling with TGF-β, AT1R, or ERK1/2 antago-
nists preserved heart structure and function, despite a
fixed PO, suggesting that predisposition does not simply
relate to a chronic deficiency in the structure and hence
mechanical integrity of the ECM, but rather reflects
dynamic but modifiable compensatory cellular signaling
events that attend myocardial remodeling. These data
suggest a number of conclusions with strong potential
for clinical relevance. First, the concept that the myocar-
dium is sensitized to acute hemodynamic stress in MFS
supports consideration of more frequent cardiovascular
surveillance for individuals with volume overload and
aggressive avoidance of hypertension for indications
beyond suppression of aneurysm progression. Second,
the observation that mild cardiovascular stress selec-
tively tips the MFS heart into profound failure supports
the earlier application of methods to more accurately
quantify valvular dysfunction (such as cardiac MRI) in
patients with MFS. The thresholds used to determine
when valve repair or replacement should be performed
should be reconsidered in this context. Finally, as dis-
cussed below, a refined understanding of the molecular
pathogenesis of heart failure in MFS may allow the test-
ing and implementation of novel medical therapies for
this and related disorders.
Prior work performed using a mouse model that is
homozygous for a hypomorphic Fbn1 allele that expresses
normal but reduced levels of Fbn1, diminishing the total
amount of produced fibrillin to about 15% of normal
from the time of conception, attributed the relatively mild
cardiac dysfunction that was observed to altered mecha-
nosensing and ultimately enhanced ERK1/2 signaling by
cardiomyocytes (32). The focus on this cell type stemmed from the observation of DCM in a mouse line in
which both copies of a conditional (flanked by loxP) Fbn1 allele were deleted through expression of Cre-re-
combinase using a cardiac myosin heavy chain promoter element (αMHC-Cre). These experiments were
difficult to interpret with certainty; however, the use of this Cre driver was also associated with a substantial
(~50%) reduction in fibrillin-1 expression in cardiac fibroblasts that was equivalent to that observed in mice
Figure 8. Inhibition of ERK1/2 activation suppresses expres-
sion of all three TGF-β isoforms. mRNA expression of Tgfb1
(TGF-β1), Tgfb2 (TGF-β2), and Tgfb3 (TGF-β3) isoforms, nor-
malized to Gapdh and then to Fbn1+/+:sham data, assessed by
real-time RT-PCR. n = 4–6 per group. *P < 0.05, **P < 0.01, ***P
< 0.001, 1-way ANOVA, Tukey’s correction. In box-and-whisker
plots, the lower and upper margins of each box define the 25th
and 75th percentiles, respectively; the internal line defines the
median, and the whiskers define the range. Values outside 1.5
times the interquartile distance are shown.
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with germline haploinsufficiency for Fbn1 (32). Furthermore, isolated expression of Cre-recombinase in the
myocardium has previously been associated with predisposition for cardiomyopathy (49, 50). While control
experiments by Cook and colleagues suggested that neither isolated Fbn1 haploinsufficiency nor Cre-recom-
binase expression was sufficient to induce cardiomyopathy in their hands, the potential pathogenic influence
of these events in combination was not tested (32). Our study now documents overt pathogenic significance
for ERK1/2 signaling in load-induced heart failure in Marfan mice, given that specific antagonism with
a MEKi prevented all perturbations of myocardial architecture and function. A number of observations,
however, focus attention on the nonmyocyte compartment. First, application of PO in WT animals was
associated with pronounced upregulation of fibrillin-1 expression, with increased deposition in the nonmyo-
cyte compartment; failure of this event in MFS mice correlated with adverse cardiac remodeling and failure.
Second, our application of in situ–based methods revealed that excessive ERK1/2 activation is specific to
nonmyocytes in the failing hearts of MFS mice. Finally, the attenuation of disease seen upon treatment with
TGF-β NAb was associated with inhibition of canonical Smad signaling in nonmyocytes but not myocytes,
presumably due to limited bioavailability in the latter, as has previously been reported (42).
The pathogenic sequence for load-induced heart failure in nonmyocytes of MFS mice appears to ini-
tiate with increased ERK1/2 activation through AT1R signaling. In theory, this could relate to increased
expression of factors along the angiotensin-converting enzyme/AngII/AT1R axis in response to the com-
bined influence of an altered matricellular environment and mechanical stress. What is both clear and
surprising is that ERK1/2 signaling drives TGF-β ligand production (largely TGF-2 and TGF-β3) by
inference in nonmyocytes and increases canonical TGF-β signaling in both the nonmyocyte and myocyte
compartments. This is in contrast to previous studies in the aorta of MFS mouse models that have shown
that excessive ERK1/2 activation is, at least in part, dependent upon and therefore downstream of TGF-β
(14, 16, 51). Of greater importance, this work shows that, similar to findings in mice with hypertrophic
cardiomyopathy (52), load-induced nonmyocyte expansion, fibrosis, and heart failure in MFS mice are
greatly attenuated by specific antagonism of TGF-β using NAb; the substantial residual Smad2 activa-
tion in myocytes but not nonmyocytes (likely on the basis of differential bioavailability of NAb) infers
specific pathogenic significance in the progression of heart failure for the latter. Prevention of myocyte
hypertrophy parallels the attenuation of myocyte Smad2 signaling seen with treatment with either ARB
or MEKi (but not with TGF-β NAb) and suggests that AT1R- and ERK1/2-dependent nonmyocyte secre-
tion of TGF-β ligands drives hypertrophy via paracrine mechanisms, while driving fibrosis and failure in
an autocrine fashion. These data support a primary role for extracellular fibrillin-1 in compensating for
ventricular injury and stress and help to establish the nonmyocyte microenvironment as a critical factor
in the loaded MFS heart. These findings may also inform mechanisms relevant to other presentations of
heart failure beyond MFS.
Methods
Mice. Fbn1C1039G/+ mice were developed as previously described (1) and bred on a pure C57BL/6 background
(backcrossed for >9 generations).
TAC and drug treatments. PO was performed by surgical placement of suture around the transverse
aorta as described previously (53). A 7-0 ligature was placed around the vessel using a 26-gauge needle to
ensure consistent occlusion. This needle size was chosen to elicit a mild response, as initial studies using
our standard TAC model (27-gauge needle) led to increased mortality in Fbn1C1039G/+ mice. Size-, age-,
and sex-matched (male) Fbn1C1039G/+ mice and littermate Fbn1+/+ mice were randomized to receive TAC
versus sham surgery (same procedure but without constriction).
For drug treatment studies, size-, age-, and sex-matched (male) Fbn1C1039G/+ mice were randomized to
receive drug treatment versus vehicle as follows: rabbit polyclonal TGF-β NAb (pan-specific against TGF-β1,
TGF-β2, and TGF-β3) and control IgG (R&D Systems) were reconstituted in PBS and administered via i.p.
injection at a dose of 10 mg/kg every 10 days. The first dose was given 2 days prior to TAC. The MEKi,
PD98059 (Calbiochem), was reconstituted in 25% DMSO/50% ethanol in PBS and administered daily i.p.
at a dose of 3 mg/kg, or placebo (DMSO) by was administered by i.p. injection, 5 days a week (36). The
first dose was given 1 day prior to TAC. RDEA119 (NIH/National Human Genome Research Institute,
Selleck Chemicals) was reconstituted in 10% 2-hydroxypropyl-β-cyclodextrin (Sigma-Aldrich) in PBS and
administered twice daily by oral gavage at a dose of 25 mg/kg. The first dose was given 1 day prior to TAC.
Losartan was administered daily in drinking water at a dose of 0.6 g/l as previously described (12). Mice
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were euthanized at 3 weeks (TGF-β NAb, PD98059, and losartan) or 5 weeks (RDEA119) for tissue analy-
sis. The studies and analyses were performed blinded as to experimental group.
Echocardiography. All transthoracic echocardiography was performed in conscious mice. To assess the
presence of valvular regurgitation, high-frame rate color Doppler imaging was performed before TAC and
at the study end (Vevo 2100, 40-MHz mechanical transducer, VisualSonics). Rare mice that developed
valvular regurgitation after TAC were excluded from all after TAC analyses. M-mode echocardiography
followed reported methods (42).
In vivo hemodynamics. Cardiac function and arterial loading were assessed by PV analyses as described
previously (33, 53). Briefly, mice were anesthetized, intubated, and mechanically ventilated. The LV apex
was exposed between the seventh and eighth ribs, and a 1.4-Fr PV catheter (SPR 839; Millar Instruments
Inc.) was advanced through the apex to lie along the longitudinal axis. Absolute volume was determined by
saline calibration and aortic ultrasound, and data were assessed at steady state and during preload reduc-
tion. Data were analyzed with custom software.
Molecular analyses. Protein electrophoresis and immunoblot assays followed standard procedures, using
cell lysis buffer (Cell Signaling) containing protease inhibitor and phosphatase inhibitor cocktail (Milli-
pore). Assays were run on 10% Bis-Tris gels (Bio-Rad), subjected to SDS-PAGE, and transferred to PVDF
membranes. Membranes were probed for pSmad2 (3101, Cell Signaling), pERK1/2 (4377, Cell Signaling),
GAPDH (ab9483, Abcam), tSmad2 (3103, Cell Signaling), and tERK1/2 (4695, Cell Signaling), followed
by labeling with anti-rabbit/goat secondary antibody and visualizing by either enhanced chemilumines-
cence (SuperSignal West Femto, Life Technologies) or fluorescence detection (LI-COR Odyssey). Blots
were quantified using ImageJ software (NIH) or the LI-COR Odyssey system.
Quantitative PCR using RT-PCR by amplification (Applied Biosystems) was used to assess RNA
expression using standard procedures. The following TaqMan probes (Applied Biosystems) were used:
Mm00514908_m1 (Fbn1), Mm00600555_m1 (Myh7), Mm01255747_g1 (Nppa), Mm00435304_g1 (Nppb),
Mm99999915_g1 (Gapdh), Mm01178820_m1 (Tgfb1), Mm00436955_m1 (Tgfb2), Mm00436960_m1
(Tgfb3), Mm00801666_g1 (Col1a1), Mm01256744_m1 (Fn1), Mm00436767_m1 (Spp1), Mm00435858_m1
(Serpine1), Mm01192933_g1 (Ctgf), and Mm00450111_m1 (Postn). Relative quantification for each tran-
script was obtained by normalizing against housekeeping control transcript.
Histology. Myocardium was fixed with 10% formaldehyde, paraffin embedded, and sectioned into
5-μm-thick slices. Masson’s trichrome staining was used to visualize fibrosis. Quantification of fibrosis
content was performed in whole heart sections using the pen tool in Aperio ImageScope (Leica Bio-
systems) to exclude large vessels, epicardium, or pericardium and automated morphometric analysis of
digital images and a previously validated modified positive pixel algorithm to quantify the amount of Tri-
chrome (blue) stain (54) (Aperio ImageScope). Quantification of myocyte cross-sectional area, when per-
formed in hematoxylin and eosin–stained sections, was performed with ImageJ. For wheat germ aggluti-
nin (WGA) staining, slides were deparaffinized, rehydrated, and subjected to citrate-based heat-mediated
antigen retrieval as previously described in (55). Sodium borohydride solution (1 mg/ml in PBS) was used
for quenching. Slides were incubated with Alexa Fluor 647–preconjugated WGA (W32466, Invitrogen)
overnight at 4°C and mounted using Prolong Gold mounting medium (P36934, Thermofisher). Image
acquisition was performed on an EVOS epifluorescence microscope (Life Technologies). Quantification
of myocyte cross-sectional area was performed using an automated algorithm with Image J, analyzing
1,040 ± 70 cells from 6–16 areas per mouse heart.
Immunohistochemistry. Slides with 5-μm-thick sections were deparaffinized with xylene and hydrated in
a graded alcohol series. Slides were immersed in boiling antigen retrieval solution (10 mM sodium citrate
buffer, pH 6.0) for 10 minutes and then set aside to cool in the same solution at room temperature for 30
minutes. Slides were incubated in freshly prepared sodium borohydride solution (10 mg/ml PBS) for 20
minutes, incubated in 100 mM glycine TBT (TBS containing 0.1% Triton-X 100) for 20 minutes, and then
treated with Fc block (NB309) and Background-buster (NB306) from Innovex Biosciences according to
the manufacturer’s instructions. Slides were incubated in a humid chamber overnight at 4°C in the fol-
lowing primary antibodies, as applicable: anti-fibrillin-1 (1:100, ab53803, Abcam), anti-pSmad2 (1:100,
4-953, Millipore), and anti-pERK1/2 (1:100, 4370, Cell Signaling) diluted in TBT. After washing, slides
were then incubated in a humid chamber with Alexa Fluor 594–conjugated donkey anti-rabbit IgG sec-
ondary antibody (1:500, R37119, Life Technologies) at room temperature for 30 minutes, washed, and
then mounted in VECTASHIELD HardSet Mounting Medium with DAPI (H-1500, Vector Laboratories).
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When applicable, following staining with the secondary antibody, slides were further incubated with Alexa
Fluor 647–preconjugated anti-vimentin antibody (1:100, Cell Signaling, 9856). Image acquisition was per-
formed on a Zeiss AxioExaminer with 710NLO-Meta multiphoton confocal microscope at magnifications
of ×20 and ×40.
High-resolution images of immunostaining (red fluorescence) were quantified using ImageJ. After
background correction, fibrillin-1–positive staining was quantified by signal intensity (red) per unit area.
The quantification of pSmad2 and pERK1/2 (red fluorescence) staining in myocytes and nonmyocytes
was performed using ImageJ. Myocytes, known to exhibit robust lipofuscin autofluorescence and sarco-
meric Z-lines in sodium borohydride-treated formaldehyde-fixed sections (56), were defined by the fol-
lowing characteristics: (a) cells with circular or oval-shaped nuclei (blue channel) centrally located with-
in myocytes (green channel), (b) relatively large cell size, and (c) presence of sarcomeric Z-lines (56).
Nonmyocytes were otherwise defined as all remaining cells with identifiable nuclei located peripheral
to myocytes, with minimal autofluorescence, in addition to being located in the interstitial space. The
number of positive cells (staining red for pSmad2 or pERK1/2) per total cells was manually counted in
each compartment by two blinded observers in 10–15 areas per treatment condition.
In situ RNA hybridization. RNAscope technology (Advanced Cell Diagnostics) was used to perform in situ
mRNA hybridization on formalin-fixed paraffin-embedded slides following the manufacturer’s instructions.
Briefly, 5-μm-thick tissue sections were hybridized with Tgfb3 (406211) and Vim (457961) from RNAscope
probes (Advanced Cell Diagnostics). A negative control probe and a positive control probe served as techni-
cal quality controls. Multiple images were acquired in Z-stack mode using a Zeiss 780 Laser Scanning Con-
focal Microscope. The in situ hybridization images are maximum intensity projections of acquired images.
Isolation of cardiac fibroblasts. Neonatal ventricles from 3-day-old mice or rats were separated and minced
in ice-cold balanced salt solution, as described previously with minor modifications (57). To isolate cardiac
cells, the tissues were incubated in a balanced salt solution containing 0.2% collagenase type 2 (Worthing-
ton Biochemical) for 6 minutes at 37°C with agitation. The digestion buffer was replaced 7 times, at which
point the tissues were completely digested. The dispersed cells were incubated in T25 flasks for 120 minutes
to plate nonmyocytes. Nonmyocytes that attached to the dishes were cultured in DMEM supplemented
with 10% FBS and allowed to grow to confluence; then they were trypsinized and passaged at 1:4. This
procedure yielded cell cultures that were almost exclusively fibroblasts by the first passage. Experiments
were carried out after 1 additional passage.
Statistics. Data are presented as dot plots, with mean ± SEM, or as box-and-whisker plots, with the
lower and upper margins of each box defining the 25th and 75th percentiles, respectively; the internal line
defining the median; and the whiskers defining the range. Values outside 1.5 times the interquartile distance
are shown. Comparisons of multiple groups were performed using either 1-way or 2-way ANOVA, as
appropriate, followed by Tukey’s or Bonferroni’s post-hoc correction (GraphPad Prism). Two-group anal-
ysis used an unpaired 2-tailed Student’s t test. Statistically significant differences (defined as P ≤ 0.05) and
trends (defined as 0.05 < P < 0.1) between genotypes (WT vs. CH) and respective treatment groups (sham
vs. TAC vs. TAC plus drug treatment) are shown.
Study approval. All mice were cared for and protocols were performed under strict compliance with and
with approved by the Animal Care and Use Committee of the Johns Hopkins University School of Medicine.
Author contributions
RR designed, directed, performed, and analyzed experiments. ET and GZ performed expert TAC and PV
loop analysis. RR and DB acquired and analyzed all echocardiographic images. RR, EGM, RC, VN, PPR,
JGB, EEG, CS, KLM, LM, NAA, and DIL performed experiments and analyzed data. NK provided tis-
sue for analysis. DPJ, DAK, and HCD provided valuable guidance and expertise. RR and HCD wrote the
paper. All authors approved the content and submission of this manuscript.
Acknowledgments
We thank the Johns Hopkins Institute Biomedical Sciences Core Microscope Facility, funded by NIH
grants S10RR024550 and S10OD016374, for assistance in acquisition of immunofluorescence images.
This research was supported by the Sarnoff Cardiovascular Research Foundation (to RR), the William
S. Smilow Foundation for Marfan Syndrome Research (to HCD), and the Howard Hughes Medical
Institute (to HCD).
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Address correspondence to: Rosanne Rouf, Division of Cardiology, Department of Medicine, Johns Hop-
kins University School of Medicine, 720 Rutland Avenue, Ross Building 809, Baltimore, Maryland 21224,
USA. Phone: 410.502.2857; Email: rrouf1@jhmi.edu.
Or to: Harry C. Dietz, Institute of Genetic Medicine, Johns Hopkins University School of Medicine, MRB
539, 733 N. Broadway, Baltimore, Maryland 21205, USA. Phone: 410.614.0701; Email: hdietz@jhmi.edu.
RC’s present address is: Department of Medicine, Sinai Hospital of Baltimore, Baltimore Maryland, USA.
PPR’s present address is: Division of Cardiology, Medical University of Graz, Graz, Austria.
NK’s present address is: Department of Cardiovascular Medicine, Gunma University Graduate School of
Medicine, Maebashi, Japan.
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