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Research Article
Skeletal Muscle Resident Progenitor Cells Coexpress
Mesenchymal and Myogenic Markers and Are Not Affected by
Chronic Heart Failure-Induced Dysregulations
R. I. Dmitrieva ,
1
T. A. Lelyavina,
2
M. Y. Komarova,
3
V. L. Galenko,
2
O. A. Ivanova,
4
P. A. Tikanova,
5
N. V. Khromova,
1
A. S. Golovkin ,
1
M. A. Bortsova,
2
A. Sergushichev,
4
M. Yu. Sitnikova,
2
and A. A. Kostareva
1
1
Institute of Molecular Biology and Genetics, National Almazov Medical Research Centre, Saint Petersburg, Russia
2
Heart Failure Department, National Almazov Medical Research Centre, Saint Petersburg, Russia
3
Peter the Great St. Petersburg Polytechnic University, Saint Petersburg, Russia
4
ITMO University, Saint Petersburg, Russia
5
Saint Petersburg State University, Saint Petersburg, Russia
Correspondence should be addressed to R. I. Dmitrieva; renata.i.dmitrieva@gmail.com
Received 4 July 2018; Revised 6 October 2018; Accepted 7 November 2018; Published 3 January 2019
Guest Editor: Zhaoping Ding
Copyright © 2019 R. I. Dmitrieva et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Background and Purpose. In heart failure (HF), metabolic alterations induce skeletal muscle wasting and decrease of exercise
capacity and quality of life. The activation of skeletal muscle regeneration potential is a prospective strategy to reduce muscle
wasting; therefore, the aim of this project was to determine if functional properties of skeletal muscle mesenchymal progenitor
cells (SM-MPC) were affected by HF-induced functional and metabolic dysregulations. Methods. Gastrocnemius muscle biopsy
samples were obtained from 3 healthy donors (HD) and 12 HF patients to purify mRNA for further analysis and to isolate
SM-MPC. Cells were expanded in vitro and characterized by immunocytochemistry and flow cytometry for expression of
mesenchymal (CD105/CD73/CD166/CD146/CD140b/CD140a/VIM) and myogenic (Myf5/CD56/MyoG) markers. Cells were
induced to differentiate and were then analyzed by immunostaining and Q-PCR to verify the efficiency of differentiation. The
expression of genes that control muscle metabolism and development was compared for HD/HF patients in both muscle
biopsy and in vitro-differentiated myotubes. Results. The upregulation of MYH3/MYH8/Myf6 detected in HF skeletal muscle
along with metabolic alterations indicates chronic pathological activation of the muscle developmental program. SM-MPC
isolated from HD and HF patients represented a mixed population that coexpresses both mesenchymal and myogenic markers
and differs from AD-MMSC, BM-MMSC, and IMF-MSC. The functional properties of SM-MPC did not differ between HD
and HF patients. Conclusion. In the present work, we demonstrate that the metabolic and functional alterations we detected in
skeletal muscle from HF patients do not dramatically affect the functional properties of purified and expanded in vitro
SM-MPC. We speculate that skeletal muscle progenitor cells are protected by their niche and under beneficial
circumstances could contribute to muscle restoration and prevention and treatment of muscle wasting. The potential new
therapeutic strategies of HF-induced skeletal muscle wasting should be targeted on both activation of SM-MPC regeneration
potential and improvement of skeletal muscle metabolic status to provide a favorable environment for SM-MPC-driven
muscle restoration.
1. Introduction
In heart failure (HF), functional and metabolic alterations
are detected not only in cardiac muscle [1, 2] but also in
skeletal muscle tissue. Oxidative stress, systemic inflamma-
tion, chronic hypoxia, and decreased fatty acid oxidation
coupled with mitochondrial dysfunction are the factors
contributing to HF-induced muscle damage that include
Hindawi
Stem Cells International
Volume 2019, Article ID 5690345, 11 pages
https://doi.org/10.1155/2019/5690345
a shift in fiber type, induction of atrophy, development of
insulin resistance, dysregulation of lipid metabolism, and
ectopic fat depositions in the skeletal muscles. Additionally,
chronic activation of adrenergic and natriuretic peptide
systems in HF results in sustained lipolysis in adipocytes
resulting in the accumulation of toxic and neutral lipid
species in adipose and skeletal muscle that also contrib-
utes to skeletal muscle damage [3–9]. Impairments in
skeletal muscle stem cell function have also been sug-
gested as an important factor causing the loss of muscle
mass with increasing age [10] and could similarly be con-
sidered as a factor contributing to HF-induced skeletal
muscle wasting.
The development of preventive and therapeutic strategies
against muscle wasting disorders remains an unresolved
challenge. By now, exercise training, either alone or in
combination with nutritional support, is the most proven
strategy to reduce skeletal muscle wasting in HF patients
and is recommended by treatment guidelines [7, 11]. Conse-
quently, the activation of skeletal muscle developmental,
growth, and regeneration potential is an essential mechanism
to treat/prevent skeletal muscle wasting. Thus, the skeletal
muscle progenitor cells that contribute to skeletal muscle
regeneration and growth might be a prospective therapeutic
target, and the analysis of the functional properties of skeletal
muscle stem cells derived from heart failure patients has
become a crucial issue.
Identification and characterization of myogenic progeni-
tors in postnatal tissues are important for the evaluation of
regeneration potential. In our recent work [12], we have
demonstrated that bone marrow multipotent mesenchymal
stromal cells (BM-MMSC) derived from heart failure
patients are affected by heart failure in multiple ways: (1) in
HF-derived cultures, we detected the upregulation of genes
that control regeneration and fibrosis, including the Tgf-β
pathway, synthesis of ECM, remodeling enzymes, and
adhesion molecules; (2) during in vitro expansion, BM-
MMSC from HF patients demonstrated early development
of replicative senescence and decrease of proliferative
activity; and (3) altered differentiation potential was also
observed in HF-derived samples. However, when culturing
conditions were modified, we have achieved the predomi-
nant purification and expansion of the highly proliferative
nonprofibrotic CD146+/SMAαfraction that proves the
potential efficacy of HF-derived BM-MMSC in regeneration
processes [12].
Multipotent mesenchymal stromal cells are tissue-
committed progenitors that preferentially contribute to
the regeneration of certain types of tissue. For skeletal
muscle, the role of nonsatellite resident myogenic progen-
itors including multipotent mesenchymal stromal cells in
tissue regeneration was also reported [13–17]. In the cur-
rent work, we sought to investigate whether or not HF-
induced metabolic dysregulations affect the functional
properties of resident skeletal muscle mesenchymal progen-
itor cells (SM-MPC) in order to determine if these cells
could respond sufficiently to therapeutic interventions
aimed at activating muscle regeneration and growth,
including physical rehabilitation programs focused on the
stabilization of muscle metabolism and prevention of skeletal
muscle wasting.
2. Methods
2.1. Study Design and Ethical Issues. The current work is part
of a complex ongoing project focused at evaluating the
efficiency of aerobic physical training and developing person-
alized physical rehabilitation programs for heart failure
patients. The first results that demonstrate the physiological
response to aerobic physical training were published recently
[18, 19]. During this project, the skeletal muscle biopsies
should be taken from a selected group of patients enrolled
in the program before and after a course of exercise training
in order to examine the regenerative potential of muscle
progenitor cells in HF patients as presented in this work;
RNA-Seq analysis will be employed to reveal the global
response of skeletal muscle to exercise training and deter-
mine potential specific targets when sample collection will
be completed. In the current work, the first portion of the
biopsy samples was used. All samples were collected under
the agreement of the Institutional Ethics Committee at the
Almazov National Medical Research Centre. All patients
and donors entering the program agreed to and signed an
institutional review board-approved statement of informed
consent. The study was conducted in compliance with cur-
rent Good Clinical Practice standards and in accordance with
the principles under the Declaration of Helsinki (1989).
2.2. Human Subjects and Gastrocnemius Muscle Biopsy
Samples. Only male donors were recruited into this study.
A total of 3 healthy adult donors (HD) and 12 chronic heart
failure patients (HF) were enrolled. HF patients have the
following characteristics: NYHA II-III functional class, age
54 ±12.5 years, body mass index (BMI) 26.5 ±6.4 kg/m
2
,
and left ventricle ejection fraction (LV EF) 26.4 ±1.4%. The
NYHA II : III patient ratio is 67% : 33%.
Inclusion criteria for the study are as follows: age 18–65
years, LV EF <40% (Simpson), stable CHF NYHA II-III
functional class, informed consent signing, ability to perform
CPET, and optimal drug and electrophysiological therapy
(ICD, CRT-D).
Exclusion criteria for the study are as follows: myocardial
infarction and myocardial revascularization less than 3
months, stroke and CRT-D implantation less than 6 months,
expressed cognitive impairment, any chronic disease decom-
pensation, and high-gradation ventricular arrhythmias with
no implanted cardioverter-defibrillator (ICD).
Gastrocnemius muscle biopsy samples were collected
from each donor/patient at baseline and after 3–6 months
of follow-up. Biopsy samples were divided into two portions.
One was immediately transferred to liquid nitrogen for
further mRNA purification. The second portion was used
immediately for skeletal muscle progenitor cell purification.
Since our study was limited by the small number of HD
enrolled into the project, we demonstrate the consistent
response of HD-derived cells to stimulation of myogenic
differentiation and similarities in immunophenotypes in
supplemental Figure S1.
2 Stem Cells International
2.3. Purification and Separation of Skeletal Muscle
Mesenchymal Progenitor Cells (SM-MPC) and Mesenchymal
Stromal Cells from Intermuscular Fat (IMF-MSC). SM-MPC
were isolated enzymatically according to the protocols
described previously [20, 21] with minor changes. In brief,
isolated muscles were placed into an enzyme solution,
mechanically disrupted with scissors, and digested for
60 min at 37
°
Cin5mLfiltered 0.1% collagenase I (C0130,
Sigma-Aldrich, Germany). To remove collagenase and cell
debris after digestion, the cell suspension was centrifuged
for 5 min at 1000 × g and the supernatant was discarded. To
release the stem cells from the fibers, the pellet was resus-
pended using sterile pipette tips in 2.5 mL of washing media
(DMEM supplemented with 10% horse serum (HS) (Gibco,
USA)). After the resuspension, the fibers were allowed to
settle for 5 min and then the supernatant containing stem
cells was transferred to a fresh tube. To increase the yield, this
step was repeated twice. The double-collected supernatant
was filtered through a 40 μm nylon cell strainer and centri-
fuged for 10 min at 1000 × g in order to discard debris. Then,
the resultant supernatant was discarded and the pellet of cells
was placed in a proliferation media (DMEM supplemented
with 10% FCS) on cell culture dishes and cultured until
80% confluence.
IMF-MSC samples were obtained from intermuscular
adipose tissue located in biopsy material. IMF-MSC cultures
were prepared as described in [22]. The separated sample of
adipose tissue was washed with phosphate-buffered saline
(PBS) and suspended in an equal volume of DMEM supple-
mented with 0.1% collagenase type III, prewarmed to 37
°
C.
The tissue was placed in a shaking water bath at 37
°
C with
continuous agitation for 30 min and centrifuged for 5 min
at 300 ×g at room temperature; then, the tissue sample was
resuspended in culture media (DMEM supplemented with
10% FBC) and plated in a culture dish for expansion.
Bone marrow mesenchymal multipotent stromal cells
(BM-MMSC) and subcutaneous adipose mesenchymal mul-
tipotent stromal cells (AD-MMSC) were collected, character-
ized, and saved as in our previous projects [22]. For this
project, they were obtained from the biobank of the Almazov
National Medical Research Centre, cultured as IMF-MSC,
and used as control samples where appropriate.
2.4. Differentiation Protocols. Fusion of some cells without
external stimuli usually was observed in subconfluent
SM-MPC cultures and served as a reliable indicator, after
which we induced skeletal muscle differentiation. To induce
differentiation, the proliferation media was removed and
replaced with differentiation media that was renewed after
every other day. The DMEM media was supplemented with
2% of horse serum. Cultures were taken for experiments at
day five and day seven after induction when myotubes were
clearly visualized. Adipose tissue differentiation was stimu-
lated as described earlier [22] by replacing the culture media
with adipocyte induction medium composed of culture
medium supplemented with 1 μM insulin, 1 μM dexametha-
sone, and 0.5 μM 3-isobutyl-1-methylxanthine. Differenti-
ated adipocytes were fixed and stained with Oil Red O at
day 9 after induction.
2.5. Immunocytochemistry. The nature of the isolated cells
was confirmed by immunocytochemical staining. Cells
seeded onto cover glasses were fixed in 4% paraformaldehyde
for 10 min at 4
°
C and then permeabilized with 0.02% Triton
X-100 for 5 min. Nonspecific binding was blocked by incu-
bation in 15% FCS for 30 min, followed by one-hour incu-
bation with the following primary antibodies: anti-MyoG
(R&D Systems, USA), anti-MYF5 (R&D Systems, USA),
anti-vimentin (Sigma-Aldrich, USA), anti-CD146 (Sigma-
Aldrich, USA), anti-desmin (D33, Dako, Denmark), myo-
sin heavy chain (MF20, MAB4470, R&D Systems, USA),
anti-myosin (skeletal fast; human MYH1/MYH2) (M4276,
Sigma-Aldrich, USA), anti-myosin (skeletal slow; human
MYH7) (M8421, Sigma-Aldrich, USA), and anti-myogenin
(MAB6686, R&D Systems, USA). The secondary antibodies
conjugated with Alexa Fluor 546/Alexa-488 (Molecular
Probes, USA) were applied for 45 min at room tempera-
ture. Nuclei were counterstained with DAPI (Molecular
Probes, USA).
2.6. Flow Cytometry Analysis. The immunophenotype of
stem cells was evaluated by flow cytometry analysis per-
formed on CytoFLEX (Beckman Coulter). Сells were resus-
pended in 100 μL of PBS containing 1% of bovine serum
albumin (Sigma-Aldrich, Saint Louis, MO, USA) and
incubated for 20 min at 20
°
C in the dark with the following
monoclonal antibodies (Ab): anti-CD56 PC7 (Beckman
Coulter, USA, A21692), anti-CD146 PE (Beckman Coulter,
USA, A07483), anti-CD166 PE (Beckman Coulter, USA,
A22361), anti-CD73 PE (BD Pharmingen, USA, 550257),
anti-CD105 APC (R&D Systems, USA, FAB1097A-100),
anti-CD45 PC5 (Beckman Coulter, USA, A07785), anti-
PDGFRβAPC (BD Pharmingen, USA, FAB1263A), and
anti-CD140a PE (BioLegend, USA, 323506). Data were
analyzed using the CytExpert 2.0 (Beckman Coulter).
2.7. Cell Sorting. All sorting procedures were performed on a
BD FACSAria™III (Becton Dickinson, USA) flow cytometer
using BD FACSDiva (Becton Dickinson, USA) software.
Flow calibration was performed using Acudrop Beads (BD
FACS™) with following stabilization for at least 25 minutes.
The nozzle size was 100 mm, and the sheath pressure was
set at 17 psi. During sorting, the flow rate was restricted to
<800 events/sec to ensure minimal contamination. Addition-
ally, a “4-way purity”sort option was used and is sufficient to
gain a 99% pure sample. Before sorting commenced, appro-
priate settings were determined for all parameters.
Cell cultures were stained with antiCD56 PE (Beckman
Coulter, USA, A07788) monoclonal antibodies according to
the manufacturer’s protocols. Primarily cells were detected
in logarithmical scales in forward scattering (FS) and side
scattering (SC). Sorting was performed according to the
electronic gating strategy. Two target cell populations
(CD56+ and CD56-, respectively) were collected in 15 mL
falcon tubes containing PBS supplemented with 2% of FBS.
Sorting efficiency was controlled using additional flow
cytometry analysis of sorted samples. Detected purity was
no less than 95%.
3Stem Cells International
2.8. RNA Isolation, cDNA Synthesis, and Q-PCR. Sequences
for Q-PCR primers can be found in supplemental Table S1.
Total RNA was isolated using the ExtractRNA reagent
(Evrogen, cat. no. BC032, Russia). cDNA was synthesized
from 500 ng of total RNA using a Moloney Murine
Leukemia Virus Reverse Transcriptase MMLV RT kit
(Evrogen, SK021, Russia). A quantitative evaluation of gene
expression was performed with qPCR mix-HS SYBR+ROX
(Evrogen, cat. no. PK156, Russia). Q-PCR data are presented
as arbitrary units of mRNA expression normalized to
GAPDH expression and to expression levels in the
reference sample.
2.9. Statistical Methods. Statistical analysis was performed
using GraphPad Prism 7 software. All data were analyzed
with at least three biological replicates and presented as
mean ±SEM. See figure legends for details for each spe-
cific experiment.
3. Results
3.1. Pathological Upregulation of Genes That Regulate
Developmental/Regeneration Program and Metabolism
Detected in Skeletal Muscle from HF Patients. In order to
detect markers of HF-induced functional and metabolic
alterations in skeletal muscles, the expression analysis was
performed in HD- and HF-derived biopsy samples for
markers and regulators of skeletal muscle development,
maturation and function (Myf6, Myh3,Myh8, Myh1, Myh4,
Myh9, Myh10, Myh7, TNNI2, and TTNC1), and energy
metabolism, including the expression of genes that regulate
lipids and glucose handling (Pgc1a, HIF1a, GLUT1, GLUT4,
aP2, PLIN2, PLIN3, PPARg, ATGL, SCD1, GOS, CGI58,
CD36, NPRA, NPRB, and NPRC). A few genes from this
panel demonstrated significantly altered expression in
HF-derived samples (Figure 1(a)).
We have found that the balance between expression
levels of slow oxidative skeletal muscle fiber MHC isoform
MYH7 and fast glycolytic isoform MYH1 was notably down-
regulated in HF muscle (Figure 1(b)), along with the down-
regulation of the expression of Pgc1a (Figure 1(c)) that
indicates an impairment in the regulation of the transcrip-
tional program for mitochondrial biogenesis and oxidative
metabolism in HF [23]. Furthermore, the expression levels
of both developmental myosins, embryonic (MYH3) and
neonatal (MYH8), were substantially upregulated in HF-
derived skeletal muscle biopsies, which along with the
upregulation of myogenic regulatory factor Myf6 detected
in HF-derived samples (Figures 1(d)–1(f)) may indicate the
chronic shift to developmental program and pathological
stimulation of muscle regeneration in HF [24, 25].
We have also detected in HF skeletal muscle the alter-
ations in the expression of the NPRA/B-to-NPRC ratio
(Figures 1(g)–1(i)) that controls the biological activity of
the natriuretic peptide system (NP) at the target tissue level
[26], including control of sensitivity to insulin and lipid
oxidative capacity through a Pgc1a-dependent pathway [27]
and cardiac progenitor cell proliferation and differentiation
into cardiomyocytes [28].
3.2. The Characterization of In Vitro Expanded SM-MPC
Cells and Comparison with Mesenchymal Multipotent Cells
from Different Sources. Purified and expanded in vitro
populations of SM-MPC derived from HD and HF muscle
biopsies demonstrated a mixed phenotype that may change
dynamically during in vitro expansion. Cells did not differ
significantly from sample to sample and between HF
patients and HD in both flow cytometry analysis and immu-
nocytochemistry analysis. BM-MMSC, IMF-MSC, and AD-
MMSC were used to demonstrate differences and similarities
between muscle-derived stem cells and mesenchymal multi-
potent stromal cells from other sources. The representative
images are presented in Figure 2.
Most of the SM-MPC cells were CD73+/CD105+/
CD166+/CD140b+, and about 30–40% of the cells in the
sample expressed NCAM/CD56, known as a reliable molec-
ular marker of satellite cells and myoblasts in human skeletal
muscle, as well as myotubes, and muscle fibers during devel-
opment and/or regeneration [29] (Figure 2(a)). Interestingly,
we detected two distinct subpopulations of CD140b
dim/bright
cells in some samples, but none of these populations was
CD56+ (Figure 2(b)). Furthermore, practically all SM-MPC
cells demonstrated no CD140a expression and only about
15% of them were CD146
dim
. Markedly, a substantial fraction
of CD146
dim
cells was CD56+ (Figure 2(b)).
BM-MMSC demonstrated, as expected, the CD73+/
CD166+/CD140a+/CD140b+/CD146+ phenotype and, sur-
prisingly, expressed NCAM/CD56. The expression of CD56
on bone marrow-derived MSC is not common, but was
reported previously at both mRNA and protein levels and
was donor-specific with the CD56+ fraction ranging from
24 to 88.5% [30]. In our experiments in all 3 samples, we
detected quite a big fraction of CD56+ cells; however, the
fraction of CD56
bright
cells in these samples was less than
15%, while in SM-MPC samples this fraction was more
than 30% as indicated on Figure 2(a). Interestingly, unlike
SM-MPC, in BM-MMSC samples all CD56+ cells were
CD140a+/CD140b+.
The immunophenotypes of both fat-derived samples,
IMF-MSC and AD-MMSC, were very similar and differ sub-
stantially from the ones of the SM-MPC and BM-MMSC
samples. Virtually all cells in the IMF-MSC and AD-MMSC
samples were CD56 negative and expressed mesenchymal
markers CD73, CD105, CD166, and CD140b (PDGFRb),
but they demonstrated little or no CD140a (PDGFRa) and
CD146 expression.
The immunostaining analysis also revealed the coexpres-
sion of mesenchymal markers and markers of myogenic cells
in SM-MPC (Figure 2(c)). Virtually all cells were expanded
in vitro; however, the cells were not stimulated to differenti-
ate into cells expressing early myogenic regulatory factor
Myf5 [25], and some cells, presumably those that will
undergo spontaneous fusion, expressed myogenin (MYOG)
that regulates the fusion of myocytes and the formation of
myotubes [25]. Furthermore, in vitro expanded skeletal
muscle-derived stem cells expressed vimentin (Figure 2(c)),
known to be expressed not only in mesenchymal cells [31]
but also in myoblasts and in myotubes during early stages
of embryonic development [32, 33]. The promyogenic nature
4 Stem Cells International
of cells that express the melanoma cell adhesion molecule
(MCAM, or CD146) was demonstrated recently [16], and
we also have detected the CD146+ population in our samples
(Figure 2(c)). We did Myf5 and MyoG immunostaining
simultaneously for the same cultures that were stained for
vimentin, and these data in combination with FACS analysis
provide evidence of the coexpression of mesenchymal lineage
cell markers with myogenic markers.
3.3. Both CD56+ and CD56- Fractions of SM-MPC
Demonstrate Myogenic Potential. In order to determine if
both major subpopulations of SM-MPC (CD56-/CD56+)
possess the myogenic potential and may therefore be impor-
tant for the maintenance of skeletal muscle regeneration,
we employed FACS sorting to separate these subpopula-
tions from SM-MPC; purified CD56-/CD56+ samples were
expanded in vitro and induced to differentiate into adipose
and muscle tissue (Figure 3). IMF-MSC samples were used
as a control.
As we expected, IMF-MSC samples did not respond to
the stimulation of myogenesis but were differentiated actively
into adipocytes (Figure 3(b)). On the contrary, both subpop-
ulations of SM-MPC did not differentiate into adipocytes
under adipogenic stimuli but demonstrated the ability to dif-
ferentiate into myotubes (Figure 3(c)). The fusion coefficient
was slightly but not significantly higher in the CD56+
subpopulation (30 ± 3% vs 24 ± 35) (Figure 3(d)), which
confirmed the potential significance of both fractions of
SM-MPC for muscle regeneration/development. Taking
into account all information mentioned above, we choose
to use the whole unsorted HD- and HF-derived SM-MPC
cellular samples in further work.
3.4. Immunohistochemical Analysis and Gene Expression
Analysis during HD- and HF-Derived SM-MPC Differentiation
In Vitro Did Not Reveal Differences between Groups. HD-
and HF-derived SM-MPC demonstrated a similar ability
to respond to the stimulation of myogenesis in vitro. We
Fold change (log2)
SCD1
GOS
MYH3
MYH8
MYH1
aP2
HIF1a
CGI58
MYF6
GLUT1
PPARg
CD36
PLIN2
TTNC1
MYH7
NPRB
LXRA
CPT1b
MYH9
GLUT4
MYH10
TNNI2
ATG L
PLIN3
MCAD
ERRA
NPRA
DGAT1
Pgc1a
NPRC
−1.5 5
(a)
HD HF
0
1
2
3
4
5
Relative MYH7/MYH1 mRNA
expression (arbitrary units)
p < 0.01
(b)
HD HF
0
2
4
6p < 0.01
Pgc1c mRNA expression
(arbitrary units)
(c)
HD HF
0
2
4
6
8
10
p < 0.01
MYH3 mRNA expression
(arbitrary units)
(d)
HD HF
0
10
20
30
40
p < 0.02
MYH8 mRNA expression
(arbitray units)
(e)
HD HF
0
5
10
15 p < 0.05
Myf6 mRNA expression
(arbitrary units)
(f)
HD HF
0
5
10
15
NS
NPRA mRNA expression
(arbitrary units)
(g)
HD HF
0
2
4
6
NS
NPRB mRNA expression
(arbitrary units)
(h)
HD HF
0
10
20
30
40
p < 0.05
NPRC mRNA expression
(arbitrary units)
(i)
Figure 1: The expression of genes that regulate skeletal muscle development and metabolism is altered in HF patients. (a) Results of Q-PCR
screening of key regulators and markers of skeletal muscle development and metabolism. Green bars: upregulation in HF; red bars:
downregulation in HF; n=3 (HD) and n=12 (HF). (b–i) Results of Q-PCR analysis of mRNA expression for genes that demonstrated
significant differences in between HD and HF. n=3(HD) and n=12 (HF).
5Stem Cells International
have done immunocytochemical staining at early and late
steps of differentiation. At day five after induction, the
formation of myotubes was observed and immunocyto-
chemical staining detected the expression of myogenin,
CD146, and vimentin. It is known that vimentin is the
most abundant intermediate filamentproteininimmature
myoblasts/muscle progenitors. During the early steps of
muscle development, desmin and vimentin are coexpressed.
Upon further differentiation into mature muscle cells,
desmin is strongly upregulated, while the expression of
CD56
SM-MPC IMF-MSC AD-MMSC BM-MMSC
41% CD56 0.7% CD56 4% CD56 92%
CD73 88% CD73 45% CD73 96% CD73 100%
CD105 100% CD105 99% CD105 99% CD105 24%
CD166 79% CD166 95% CD166 92% CD166 100%
CD146 16% CD146 8% CD146 1% CD146 75%
CD140a 8% CD140a 3% CD140a 0% CD140a 96%
CD140b 77% CD140b 90% CD140b 92% CD140b 100%
CD45 0% CD45 0% CD45 0% CD45 0%
35% 15%
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(a)
CD56 + /CD140b−
CD56 + /CD140a−
CD56 + /CD146b+
CD56
CD140b
CD56
CD140a
CD56
CD146
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(b)
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MYF5
VIMCD146
DAPI DAPI
DAPI DAPI
(c)
Figure 2: Characterization of SM-MPC, IFM-MSC, BM-MMSC, and AD-MMSC cultures expanded in vitro. (a) Representative histograms
demonstrate the results of a comparative FACS surface marker analysis of SM-MPC, IFM-MSC, BM-MMSC, and AD-MMSC. Green
histograms indicate the stained samples, and grey ones indicate the negative controls. The red ring indicates the CD56
bright
subpopulations
in SM-MPC and BM-MMSC samples (n=3–5 in each group). (b) Visualization of FACS analysis in SM-MPC samples: CD56+/CD140a-,
CD56+/CD140b-, and CD56+/CD146+ subpopulations are indicated in a rectangle gate; quadrant gates specify the negative/positive
populations. Unstained cells were used as a negative control for each sample. (c) Immunocytological phenotyping of skeletal muscle
precursor cells at early steps of expansion in vitro. Cells were stained for the expression of MYF5 (red; ~100% of positive), MYOG
(red; 25±4% of positive), vimentin (VIM, green; ~100 of positive), or CD146 (green; 20±4.5% of positive). The insert demonstrates
the enlarged region with a CD146+ cell. Nuclei were labelled with DAPI (blue). Scale bars represent 50 μm.
6 Stem Cells International
vimentin completely ceased [34]. In our cultures, we
detected vimentin in progenitor cells (Figure 2(c)) and in
myotubes at the early steps of differentiation (Figure 4),
while desmin expression was detected in all tested condi-
tions(Figures3and4).
Staining for desmin (Figure 4) and MF20 (data not
shown) at day 5 did not demonstrate a cross-striated pattern
specific for mature skeletal muscle fiber (Figure 4). At day
seven of immunocytochemical staining with antibodies
against MF20, slow and fast MyHCs confirmed that differ-
entiated myotubes demonstrate a cross-striated pattern,
similar to that seen in adult muscle fibers. Both HD- and
HF-derived cells developed myotubes that were positive
for both slow (MYH7) and fast (MYH1/MYH2) myosins,
and the fractions of nuclei incorporated into myosin-
positive myotubes did not differ significantly between
HF and HD samples (Figure 5(a)). The fusion coefficient
did not differ significantly between HD and HF samples
(28 ± 4% vs 32 ± 5%) (Figure 5(b)).
The gene expression analysis of markers and regulators of
myogenic differentiation of SM-MPC derived from HD and
HF patients confirmed the results of immunohistochemistry:
the expression of slow skeletal muscle fiber MHC isoform
MYH7 and fast isoform MYH1 did not differ significantly
between HD- and HF-derived differentiated myotubes, and
the expression of embryonic (MYH3) and neonatal (MYH8)
myosins did not differ significantly as well. The expression
of myogenic regulatory factor Myf6, Pgc1a, and of atrial
natriuretic peptide receptor C in differentiated myotubes also
did not differ significantly between groups (Figure 5(c)).
4. Discussion
Heart failure is a multiorgan syndrome affecting different cell
types, including skeletal muscle. The development of preven-
tive and therapeutic strategies against muscle wasting disor-
ders in HF remains an unresolved challenge, and activation
of developmental/regeneration programs in skeletal muscle
could be considered as a prospective approach. Therefore,
the activation of skeletal muscle progenitor cells would be
beneficial for the restoration of skeletal muscle structure
and performance. In this work, we aimed to determine if
HF-induced skeletal muscle alterations affect SM-MPC
developmental/regeneration potential.
IFM-MSC
Stimulation of
myogenesis and adipogenesis
SM-MPC
(a)
IFM-MSC
Adipogenesis Myogenesis
No response
(b)
Adipogenesis Myogenesis
No response
CD56-PC7
CD73-PE
0 10
3
10
2
10
3
10
4
10
5
10
4
10
5
CD56−
CD56+
Desmin
DAPI
DEsmin
DAPI
SM-MPC
(c)
50
40
30
20
10
0CD56−CD56+
Fusion coecient (%)
(d)
Figure 3: Comparative analysis of the functional properties of subpopulations of skeletal muscle progenitor cells derived from muscle tissue
(SM-MPC) and from intermuscular fat (IMF-MSC). (a) The design of the experiment is as follows: SM-MPC and IMF-MSC were purified
from muscle biopsy, expanded in vitro, characterized, and induced to differentiate. (b) IMF-MSC under appropriate stimulation undergo
adipogenesis but do not respond to promyogenic stimulation. (c) Both CD56+ and CD56- fractions of SM-MPC demonstrate the ability
to differentiate into myotubes but do not respond to adipogenic stimuli; scale bars represent 100 μm. (d) Fusion coefficient is calculated as
a percent of nuclei incorporated in MF20+ myotubes, and it does not differ between the CD56+ and CD56- subpopulations.
HD HDHD
HF HF
HF
DAPI
DAPI DAPI
DAPI
DAPI
DAPI Desmin
Desmin
VIM
VIM
CD146
CD146
MYOG
MYOG
Figure 4: Immunocytological phenotyping of differentiated myotubes at early steps of myogenesis. At day 5 after stimulation, myotubes
coexpress myogenin (MYOG) and CD146. All cells in culture express vimentin (VIM), and desmin expression is specifically associated
with multinucleated myotubes. Scale bars represent 50 μm.
7Stem Cells International
We have detected a number of chronic dysregulations in
the skeletal muscle tissue from our patients. The most
important one was the chronic activation of the develop-
mental program: the expression of mRNA of MYH3/
MYH8 myosins and myogenic regulatory factor Myf6 was
detected in an HF-derived biopsy. These alterations in com-
bination with changes in slow/fast fiber composition may
severely influence skeletal muscle metabolism, structure,
and performance. We suggest that in HF, the regeneration
process is stimulated in response to HF-induced damage:
skeletal muscle progenitor cells progress successfully through
activation, proliferation, differentiation into myoblasts, and
fusion steps; however, instead of advancing to the fiber mat-
uration stage, they get “stuck”at the developmental phase
presumably due to chronic metabolic alterations observed
in HF skeletal muscle. The reexpression of developmental
myosins in adult skeletal muscle was detected in different
pathological conditions that involve muscle degeneration/
regeneration, such as trauma, chronic denervation, muscu-
lar dystrophy, and different types of myopathies (reviewed
in [24]); however, to our knowledge none was previously
reported for HF.
The shift from oxidative fiber type I to glycolytic fiber
type II and reduced oxidative enzyme activities is the best-
described HF-induced metabolic alteration in skeletal muscle
[3, 5, 8, 35, 36]. We have also detected the shift in the expres-
sion ratio between fiber type I and fiber type II in skeletal
muscle from HF patients, as well as the downregulation of
expression of Pgc1a (Figure 1), which is an important medi-
ator of mitochondrial metabolic properties in skeletal muscle
and is downregulated in various types of atrophying muscle
[37] including skeletal muscle of rats with HF [38–41].
Furthermore, HF is a state of chronic activation of adrenergic
and NP systems, which besides their well-documented role in
the cardiovascular system also plays a role in favoring fat
oxidative capacity in human skeletal muscle cells [42] via
the activation of cGMP signaling, induction of PGC1a, and
enhancement of mitochondrial respiration [27]. There are
also recent data showing that the NP system is involved in
the regulation of cardiac progenitor cell proliferation via
NPR-A and differentiation into cardiomyocytes via NPR-B
[28] contributing to heart development and regeneration.
The switch in expression balance from NPR-A/B to NPRC
detected in our work indicates an increase of NP system
MF20
MF20
Slow
Slow
DAPI
DAPI
DAPI
DAPI
DAPI
DAPI
Fast
Fast
HF HF HF
HD HD HD
(a)
HD HF
0
20
40
60
Fusion coecient (%)
(b)
1
10
100
1000
HD
MYH1
MYH7
MYH3
MYH8
MYH6
NPRA
NPRB
NPRC
Pgc1a
HF
mRNA expression
(arbitrary units, log)
(c)
Figure 5: Differentiated myotubes do not differ significantly between HD- and HF-derived skeletal muscle progenitor cells. (a) At day 7 after
stimulation, myotubes were stained for the expression of MyHC with an antibody that recognizes the heavy chain of myosin II (MF20) and
markers of slow MYH7 and fast MYH1/MYH2 fibers. Nuclei were labelled with DAPI (blue). Representative images are given for both HF-
and HD-derived samples. Scale bars represent 50 μm. (b) Fusion coefficient is calculated as a percent of nuclei incorporated in MF20+
myotubes at day 7 after stimulation, and it does not differ between HD- and HF-derived samples. (c) mRNA expression analysis was
performed for key markers of muscle development and metabolism for both HF- and HD-derived samples.
8 Stem Cells International
activity [9] that could also impact on the upregulation of
developmental signaling in HF skeletal muscle. Together,
we have detected a number of alterations in HF-derived
skeletal muscle that would affect developmental, metabolic,
structural, and functional properties of skeletal muscle.
Next, we purified SM-MPC from HD- and HF-derived
biopsies and investigated if the functional properties of
HF-derived cells were affected by these alterations. The
SM-MPC that we isolated from skeletal muscle biopsy
samples from HD and HF patients represented a mixed
population of cells that express both mesenchymal and
myogenic markers. Samples were characterized by FACS
analysis (Figures 2(a)–2(c)) and immunocytochemistry
(Figure 2(c)), and they were also investigated for the abil-
ity to differentiate in vitro (Figure 3). In order to better
evaluate the myogenic potential of muscle progenitor cells
derived from HD and HF subjects based on obtained
results, we concluded using the whole unsorted samples
in differentiation experiments in order to retain in cultures
all subpopulations that could possibly contribute to stimu-
lated in vitro myogenesis either via paracrine signaling
mechanisms and/or direct cell-cell interaction.
Indeed, there is a lot of evidence in the literature that
describes different subpopulations of skeletal muscle progen-
itor cells that support skeletal muscle development, growth,
and regeneration (reviewed in [13]). The best characterized
myogenic progenitors in postnatal muscle are satellite cells
that are activated in response to injury or stimulation to
growth, which then start to proliferate and generate a pool
of myoblasts able to fuse into newly formed myofibers.
CD56 is considered as the most reliable satellite cell surface
marker [38]. However, human satellite cells are not easy to
isolate, purify, and expand in culture: most of the studies with
satellite cells were done on mice, but not on humans [16].
Furthermore, in recent years reports of myogenic cells
distinct from satellite cells have accumulated, and not all
of these cells are reported to be CD56+. For example,
PW1+/Pax7–interstitial cells (PICs) that do not express
CD56 but demonstrate bipotential behavior in vitro, gen-
erating both smooth and skeletal muscles, were isolated
and characterized [17]; the coexpression of mesenchymal
(CD90, CD73, CD166, and CD105) and myogenic (CD56)
markers was reported on several cell populations with myo-
genic potential including human embryonic mesodermal
progenitors [43], expanded in vitro muscle-derived primary
cultures [14], and myogenic mesenchymal progenitors
derived from hES and iPSC [29]. It was also shown that sub-
endothelial- (mural) cultured CD146+ cells (also known as
“mesenchymal stem cells”) purified from various groups of
skeletal muscle were able to spontaneously generate myo-
tubes in vitro and myofibrils in vivo, and the expression of
CD146 and CD56 was mutually exclusive in distinct myo-
genic cell subsets, with no coexpression [16]. Importantly,
in this work the authors report that sorted and cultured
CD146+ human muscle-derived cells progressively turn on
the expression of myogenic markers PAX7, PAX3, Myf5,
CD56, desmin, and MyHC, as they were verified by fluores-
cent immunocytochemistry [16]. Finally, in some protocols
CD146 is recommended as a positive selection marker in cell
sorting to obtain a human fetal myoblast population [44].
Together, all these previous findings support observations
made in our work. Firstly, in expanded in vitro adherent
cells purified from gastrocnemius muscle biopsy we also
detected substantial subpopulations of cells that coexpress
mesenchymal and myogenic markers (Figure 2). Secondly,
we have found that not only the sorted CD56+ but also
the sorted CD56- subpopulation demonstrates myogenic
potential (Figure 3). Because of the limited volumes of
samples, we were not able to monitor the dynamics of marker
expression during in vitro expansion of sorted CD56+/CD56-
populations; however, we can speculate that the myogenic
potential of the CD56- subpopulation can be related to the
CD146+ subpopulation of cells and those cells could turn
on the expression of myogenic markers in the course of
expansion as described by Persichini et al. [16]. The data
presented on Figures 2 and 3 support this speculation pretty
well: the coexpression of CD146 and CD56 on certain sub-
populations of expanded in vitro SM-MPC (Figures 2(a)
and 2(c)) and coexpression of myogenic regulatory factor
myogenin (MyoG) with CD146 at early steps of myogenic
differentiation (Figure 3) confirm the importance of the
CD146+ SM-MPC fraction for myogenic differentiation.
Also, as indicated on Figure 3, both major fractions of
SM-MPC (CD56+/CD56-) demonstrated a similar ability to
differentiate into myotubes but not into adipocytes. This is
an important observation: dysregulation of lipid metabolism
in the skeletal muscles of HF patients is a well-described
metabolic disorder [9], and fatty degeneration of skeletal
muscle is often associated with metabolic dysregulation
[45, 46]. In our experiments, neither HD nor HF-derived
SM-MPC demonstrated in vitro adipogenic potential. Our
data fit well the observations described by Uezumi et al.
[45] who demonstrated that only the CD140a+ mesenchymal
progenitor population of muscle-derived cells show efficient
adipogenic differentiation both in vitro and in vivo. In
our work, SM-MPC demonstrated a CD140a- phenotype.
Interestingly, both adipose tissue-derived AD-MMSC and
IMF-MSC, but not BM-MMSC samples that all differentiate
into adipocytes (Figure 2, our previous data [22]), demon-
strated a CD140a- phenotype.
Interestingly, we have detected myotube formation not
only upon canonical differentiation conditions (2% horse
serum), but even under adipogenic stimulation (data not
shown). Similar observations were made previously by
others: myogenesis in skeletal muscle-derived progenitor
cells under adipogenic stimulation was mentioned earlier
by Persichini et al. [16] and by Uezumi et al. [45]; those data,
however, did not get much attention and deserve further
investigation. Together, these observations allow us to con-
clude that SM-MSC samples purified from both HD- and
HF-derived skeletal muscle were restricted/committed to
myogenic differentiation and, in general, do not differ
between healthy donors and heart failure patients.
5. Conclusion
In the present work, we demonstrate that the metabolic and
functional alterations we detected in skeletal muscle from
9Stem Cells International
HF patients do not dramatically affect the functional prop-
erties of purified and expanded in vitro skeletal muscle
mesenchymal progenitor cells (SM-MPC).
These findings allow us to speculate that skeletal muscle
progenitor cells are quite well protected by their niche from
HF-induced metabolic stress, and they could, under benefi-
cial circumstances, contribute to damaged muscle restora-
tion, prevention, and treatment of muscle wasting; the exact
mechanisms behind alterations in the muscle regeneration
program in HF remain to be investigated, and the potential
new therapeutic strategies of HF-induced skeletal muscle
wasting should be targeted on both activation of skeletal
muscle stem cell regeneration potential and improvement
of skeletal muscle metabolic status in order to provide a
favorable environment for SM-MPC-driven improvement
of muscle structure and performance.
Data Availability
The data used to support the findings of this study are
available from the corresponding author upon request.
Conflicts of Interest
Dmitrieva R.I., Lelyavina T.A., Komarova M.Y., Ivanova
O.A., Galenko V.L., Tikanova P.O., Khromova N.V.,
Golovkin A.S., Bortsova M.A., Sergushichev A., Sitnikova
M.Yu., and Kostareva A.A. declare that they have no
conflict of interest.
Acknowledgments
Cell sorting experiments were performed on the FACSAria
III Cell Sorter (Becton, Dickinson and Company, USA) at
the Research Resource Centre “Molecular and Cell Tech-
nologies”, Saint Petersburg State University; http://www.
biomed.spbu.ru/en/. Work was funded by Russian Science
Foundation grant #16-15-10178 (RD).
Supplementary Materials
Table S1: list of primers used in Q-PCR experiments. Figure
S1: SM-MPC from HD demonstrate the consistent response
to stimulation of myogenic differentiation and similarities
in immunophenotypes. Upper panel: immunocytological
visualization of differentiated myotubes in HD-derived sam-
ples: myotubes were stained for expression of MyHC with an
antibody that recognizes the heavy chain of myosin II
(MF20). Nuclei were labelled with DAPI (blue). Lower panel:
histograms demonstrate results of FACS surface marker
analysis of SM-MPC derived from HD. (Supplementary
Materials)
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