(2022) 12:24
Yamanouchi et al. Skeletal Muscle
https://doi.org/10.1186/s13395-022-00307-7

Open Access

RESEARCH

Macroglossia and less advanced dystrophic
change in the tongue muscle of the Duchenne
muscular dystrophy rat
Keitaro Yamanouchi1*, Yukie Tanaka1, Masanari Ikeda1, Shizuka Kato2, Ryosuke Okino3, Hiroki Nishi3,
Fumihiko Hakuno3, Shin‑Ichiro Takahashi3, James Chambers2, Takashi Matsuwaki1 and Kazuyuki Uchida2

Abstract
Background: Duchenne muscular dystrophy (DMD) is an X-linked muscle disease caused by a complete lack ofdystrophin, which stabilizes the plasma membrane of myofibers. The orofacial function is affected in an advanced
stage of DMD and this often leads to an eating disorder such as dysphagia. Dysphagia is caused by multiple etiologies
including decreased mastication and swallowing. Therefore, preventing the functional declines of mastication andswallowing in DMD is important to improve the patient’s quality of life. In the present study, using a rat model of DMD
we generated previously, we performed analyses on the masseter and tongue muscles, both are required for proper
eating function.
Methods: Age-related changes of the masseter and tongue muscle of DMD rats were analyzed morphometrically,
histologically, and immunohistochemically. Also, transcription of cellular senescent markers, and utrophin (Utrn), afunctional analog of dystrophin, was examined.
Results: The masseter muscle of DMD rats showed progressive dystrophic changes as observed in their hindlimb
muscle, accompanied by increased transcription of p16 and p19. On the other hand, the tongue of DMD rats showed
macroglossia due to hypertrophy of myofibers with less dystrophic changes. Proliferative activity was preserved in
the satellite cells from the tongue muscle but was perturbed severely in those from the masseter muscle. While Utrn
transcription was increased in the masseter muscle of DMD rats compared to WT rats, probably due to a compensa‑
tory mechanism, its level in the tongue muscle was comparable between WT and DMD rats and was similar to that in
the masseter muscle of DMD rats.
Conclusions: Muscular dystrophy is less advanced in the tongue muscle compared to the masseter muscle in the
DMD rat.
Keywords: Duchenne muscular dystrophy, Masseter muscle, Tongue, Macroglossia, Rat*Correspondence: akeita@g.ecc.u-tokyo.ac.jp
1
Laboratory of Veterinary Physiology, Graduate School of Agricultural and Life
Sciences, The University of Tokyo, 1‑1‑1 Yayoi, Bunkyo‑ku, Tokyo 113‑8657,
Japan
Full list of author information is available at the end of the article

Background
Duchenne muscular dystrophy (DMD) is an X-linked
muscle disease caused by out-of-frame mutations in thedystrophin gene that results in the complete loss of dystrophin protein [9]. Dystrophin is a structural proteinthat stabilizes the plasma membrane of the myofibers,
and the lack of dystrophin protein causes myofibers to
become fragile [9]. Skeletal muscle myofibers normally© The Author(s) 2022. Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which
permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to theoriginal author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images orother third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line
to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory
regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of thislicence, visit http://​creat​iveco​mmons.​org/​licen​ses/​by/4.​0/. The Creative Commons Public Domain Dedication waiver (http://​creat​iveco​
mmons.​org/​publi​cdoma​in/​zero/1.​0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Yamanouchi et al. Skeletal Muscle(2022) 12:24

have a remarkable regenerative capacity [11]. Once
myofibers are damaged, the myogenic stem/progenitor
cells, satellite cells, are activated and proliferate, theneventually fuse each other to form multinucleated myotubes. The myotubes mature to constitute newly regenerated myofibers. DMD is characterized by repeatedcycles of degeneration and regeneration of myofibers,
which finally result in a loss of regenerative capacity due
to exhaustion of the satellite cell pool [21]. DMD patientsshow progressive weakness and premature death due to aloss of ambulation [9].
In an advanced stage of DMD, an orofacial function isaffected [3] and eating disorders such as dysphagia areoften a problem [37]. Dysphagia is caused by multiple
etiologies. Decreased mastication is one of the causesof dysphagia. Masticatory movement involves masseter,
temporalis, medial, and lateral pterygoid muscles [17].
Masticated foods are then swallowed using the tongue
muscle in the oral cavity. Swallowing is initiated by thecontraction of digastric and tongue muscles against thepalatine to move the bolus into the oropharynx, followed
by a complex coordination of pharyngeal and esophageal
muscles. The tongue muscles consist of the extrinsic muscles, which move the tongue from the outside, and the
intrinsic muscles, which change the shape of the tongue.
The former consists of four types of tongue muscles: thegenioglossus, hyoglossus, styloglossus, and palatoglossus
muscles, and the latter consists of four types: the superior
longitudinal, inferior longitudinal, transverse, and vertical muscles. Several reports demonstrated that dysphagia
in DMD patients is associated with dystrophic changes inmuscles related to mastication [36]. Macroglossia, whichis common in DMD patients [28, 37], is defined as anenlarged tongue and is often associated with dysphagia[5]. Therefore, preventing the deterioration of mastication and swallowing functions in DMD is important toimprove the patient’s quality of life.
We previously reported the generation of a rat modelof DMD (DMD rat) [22, 33]. DMD rats show a progressive muscle pathology in their hindlimb muscle
and diaphragm like human DMD [22, 33]. The pathological changes seen in the muscle of DMD rats include
decreased strength, increased fibrosis, and fatty infiltration [22, 33]. We further demonstrated the occurrence ofpremature cellular senescence in the muscle of DMD rats
and suggested that this could be one of the causes of satellite cell depletion, which leads to a loss of regenerative
capacity [33].
In the above our studies using DMD rats [22, 33], we
analyzed only the muscle of the hindlimbs and diaphragm. During breeding and maintenance of DMD rats,
we had experienced a decreased food intake in accordance with the decrease in the body weight in their later

Page 2 of 13

life, and this prompted us to examine whether the muscles related to food intake are also affected. Thus, in thisstudy, we focused on the masseter and tongue muscles as
the muscles involved in mastication and swallowing andinvestigated their development and histological changes,
as well as the dynamics of satellite cells.

Methods
Animals

We previously generated and characterized a strain of
DMD rat [22, 33]. In this strain of DMD rat, a deletionof 329 bp around the splice site in intron2 leads to exon3
skipping, and an insertion of 1 bp in exon16 causes thegeneration of a stop codon, resulting in a loss of dystrophin protein [22, 33]. Adult ­XDmdX female rats weremated with wild-type (WT) male rats to generate maleWT (XY) and DMD ­(XDmdY) rats. They were maintained
under controlled environmental conditions, at 23 °C
with a light/dark cycle (lights on 0800–2000). Laboratory chow (Labo MR Standard, Nihon Nousan Co., Yokohama, Japan) and water were given ad libitum. All animal
experiments performed in this study were in accordancewith the Guide for the Care and Use of Laboratory Animals of the University of Tokyo and were approved by
the Institutional Animal Care and Use Committee of the
University of Tokyo (P18-125).
Measurement of food intake

At 3 or 7 months old, the rats were transferred individually to a plastic cage and their amount of food intake wasmeasured for two consecutive days. The measured values
were divided by 2 and expressed as food intake per day.
Histological and immunohistochemical analyses

At indicated age, the rats were killed by inhalation of carbon dioxide gas. After removal of facial skin, the area corresponding to the masseter muscle was measured. The
masseter muscle was considered as an ellipse (Fig. 1B),
and its long and short diameters were measured withcalipers to calculate the area. Then, the lower half part ofthe masseter muscle was removed. The tongue was cut
out at the root. Collected tissues were fixed in 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS)
for paraffin-embedded sections or snap-frozen in liquid
nitrogen-cooled isopentane for cryosections.
The paraffin-embedded sections were used for hematoxylin–eosin (HE) or Masson’s trichrome staining. The
sections were observed and photographed using a microscope (BX51, Olympus, Tokyo, Japan) equipped with adigital camera (DP73, Olympus). The HE-stained tongue
sections were used for measurement of the width andarea of the tongue (Fig. 1D). Masson’s trichrome-stained
sections were used for quantitative analysis of the fibroticYamanouchi et al. Skeletal Muscle(2022) 12:24

area. Two fields were randomly selected in the section
using a 4 × objective and the area occupied by fibrotic tissues stained blue and the total area of sections were calculated using ImageJ software (v1.47; National Institutesof Health, Bethesda, MD, USA).
The cryosections (7 μm) were used for immunohistochemistry. The sections were fixed with 4% PFA in PBS
for 15 min, followed by blocking with 5% normal donkey
serum (NDS) in PBS. The sections were incubated withprimary antibodies overnight at 4 °C, then secondary
antibodies for 1 h.
For identification of satellite cells, vascular endothelial
cells, and mesenchymal progenitor cells, anti-Pax7 (1:200
with 5% NDS in PBS, mouse, clone P3U1; Developmental
Studies Hybridoma Bank (DSHB), Iowa City, IA, USA),anti-CD31 (1:400 with 5% NDS in PBS, rabbit, NB1002284, Novus Biological, Centennial, CO, USA), and antichondroitin sulfate proteoglycan 4 (CSPG4) (1:50 with
5% NDS in PBS, mouse, clone 5C12 [34]) were used as
primary antibodies, respectively, followed by AlexaFluorlabeled donkey anti-mouse IgG (1:500 with 5% NDS in
PBS, Jackson ImmunoResearch, West Grove, PA, USA).
For identification of necrotic myofibers, anti-laminin(1:400 with 5% NDS in PBS, rabbit, L9393 Sigma, St
Louis, MO, USA) was used as the primary antibody,and AlexaFluor-labeled goat anti-rat IgG and donkey
anti-rabbit IgG (1:500 with 2.5% NDS/2.5% normal goat
serum (NGS) in PBS, Jackson ImmunoResearch, West
Grove) were used as secondary antibodies.
For identification of newly formed regenerated myofibers, anti-embryonic myosin heavy chain (eMHC) (1:400
with 5% NDS in PBS, mouse, clone F1.652, DSHB) andanti-laminin were used as primary antibodies, andAlexaFluor-labeled donkey anti-mouse IgG and donkey anti-rabbit IgG (1:500 with 5% NDS in PBS, Jackson
ImmunoResearch) were used as secondary antibodies.
After the reaction with secondary antibodies, nuclei werecounterstained with Hoechst 33258. The sections
were observed and photographed using a microscope
equipped with a digital camera.
For quantification of Pax7-positive cells, IgG-positive
necrotic myofibers, and embryonic myosin heavy chain(eMHC)-positive myofibers, several numbers of the field
were photographed using × 4 or × 10 objectives, andthe number of positive cells or myofibers were counted.

Page 3 of 13

Myofibers were identified by double-staining with ananti-laminin antibody. Data were expressed as percent
positive myofibers or number of positive cells per area.
For quantification of myofiber size and the number of nuclei per myofiber, the sections were immunostained with anti-laminin, followed by labeling with
AlexaFluor-labeled secondary antibody. The nuclei werecounterstained with Hoechst 33258. The sections were
photographed as described above. The myofibers wereautomatically identified by Cellpose Google Colab script(https://​colab.​resea​rch.​google.​com/​drive/​1958U​QIH-​
XAYog​Kvbxn​aUHAL​YvR73​KLj2) [38], and their diameters (minimum Feret diameters) and numbers were
measured using ImageJ software. The number of nuclei
per myofiber was calculated by dividing the total numberof nuclei by the number of myofibers.
The area occupied by CD31-positive cells and CSPG4positive cells was calculated by ImageJ.
Cell culture and immunocytochemistry

Mononuclear cells were isolated from skeletal muscle as
described previously [39]. In brief, the rats were killed byinhalation of carbon dioxide gas, and their tongue andmasseter muscles were extirpated. As described above,the lower half part of the masseter muscle and tongue
cut out at the root were used for cell isolation. They wereminced with scissors and digested with 1.25 mg/mL protease (from Streptomyces griseus, type XIV; Sigma) at
37 °C for 1 h. Cells were separated from myofiber fragments through differential centrifugation and plated onto
poly-l-lysine- and fibronectin-coated 48-well plates.
Cells from each muscle piece were divided into two equal
portions (one for immunocytochemistry of Pax7 andanother for MyoD) and plated. Cells were cultured in
Dulbecco’s modified Eagle medium (Gibco, Life Technologies, Palo Alto, CA, USA) containing 10% fetal bovine
serum, 100 U/mL penicillin, 100 μg/mL streptomycin,
and 50 μg/mL gentamicin for 4 days.
The cells were fixed with 4% PFA/PBS for 15 min andthen blocked with 5% NGS/PBS containing 0.1% Triton
X-100 for 10 min. The cells were incubated with antiPax7 (1:200 with 5% NGS/PBS) or anti-MyoD (1:200,
with 5% NGS/PBS, mouse, clone 5.8A, Novocastra, Newcastle upon Tyne, UK) overnight at 4 °C. After washing,
they were incubated with AlexaFluor-labeled secondary(See figure on next page.)
Fig. 1 Food intake and morphometrical analyses of masseter and tongue muscles of WT and DMD rats. A The amount of food intake per day
was measured at 3 and 7 months old. The data are expressed as mean ± SE. **, p < 0.01 by unpaired Student’s t-test. B The masseter muscle wasconsidered as an ellipse (dotted white circle) and its long and short diameters were measured with calipers to calculate the area. C Changes in
the area of the masseter muscle at 1, 3, 6, and 8 months old. The data are expressed as mean ± SE. **, p < 0.01 by unpaired Student’s t-test. D The
tongue was cut at the root (dotted black line) (left) and the width (dotted black line) and area (dotted red circle) were measured. Scale bar = 1 cm. E
Changes in the width and cross-sectional area of the tongue at 1, 3, and 8 months old. The data are expressed as mean ± SE. **, p < 0.01 by unpaired
Student’s t-test

Yamanouchi et al. Skeletal Muscle(2022) 12:24

Fig. 1 (See legend on previous page.)

Page 4 of 13

Yamanouchi et al. Skeletal Muscle(2022) 12:24

antibody for 1 h. Nuclei were counterstained with Hoechst 33,258. The Pax7- and MyoD-positive cells werecounted in randomly selected five fields at 20 × objective of a fluorescence microscope (BX50, Olympus). The
total numbers of nuclei were also counted. The data wereexpressed as percent positive cells.
Quantitative real‑time PCR

Total RNA was extracted from 100 cryosections (7 μm)
with FastGene RNA Basic kit (Nippon Genetics Co., Ltd.,Tokyo, Japan) and reverse transcribed to cDNA using
Super Script II kit (Invitrogen). Quantitative real-time
PCR was performed on a Light Cycler 2.0 (Roche Diagnostics, Roche, Basel, Switzerland) with the Thunderbird
SYBR qPCR Mix (TOYOBO, Osaka, Japan). The following primer sets were used: p16: forward, 5′-TTC ACC
AAA CGC CCC GAA CA-3′; reverse, 5′-CAG GAG
AGC TGC CAC TTT GAC-3′; p19: forward, 5ʹ-GTG
TTG AGG CCA GAG AGG AT-3ʹ; reverse, 5ʹ-TTG CCC
ATC ATC ATC ACC T-3ʹ; p21: forward, 5ʹ-GAC ATCTCA GGG CCG AAA-3ʹ; reverse, 5ʹ-GGC GCT TGG
AGT GAT AGA AA-3ʹ; p53: forward, 5ʹ-AGA GAG CACTGC CCA CCA-3ʹ; reverse, 5ʹ-AAC ATC TCG AAG
CGC TCA C-3ʹ; Utrn: forward, 5ʹ‐TAG AGC AAT ACG
CCA CAC GA‐3ʹ; reverse, 5ʹ‐ACG CTC TTC CTT CTC
CAC AG‐3ʹ; MuRF1: forward, 5ʹ‐AGG ACT CCT GCC
GAG TGA C-3′; reverse, 5′-TTG TGG CTC AGT TCC
TCC TT-3′; Atrogin1: forward, 5′-GAA GAC CGG CTA
CTG TGG AA-3′; Atrogin1: reverse, 5′-ATC AAT CGC
TTG CGG ATC T-3′, and HPRT: forward, 5ʹ‐GAC CGG
TTC TGT CAT GTC G‐3ʹ; reverse, 5ʹ‐ACC TGG TTC
ATC ATC ACT AAT CAC‐3ʹ. The expression of eachgene was analyzed using the crossing‐point method andexpressed after normalization with that of HPRT.
Western blotting

Two hundred cryosections (7 μm) of the tongue muscle were homogenized using radioimmunoprecipitation assay (RIPA) buffer [10 mM ­NaH2PO4, 150 mM
NaCl, 2 mM ethylenediaminetetraacetic acid (EDTA),
0.1% sodium deoxycholate, 1% Nonidet P-40, 10 μg/mL
leupeptin, 5 μg/mL pepstatin, 1.84 g/L N­ a3VO4, 10 mg/
mL p-nitrophenylphosphate (PNPP), 100 KIU/mL aprotinin, 20 μg/mL phenylmethanesulfonylfluoride (PMSF)],and protein concentration of the lysates was determinedusing Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA). Lysates were, then, diluted
with RIPA buffer to the same concentrations of protein.
The samples were subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) andwestern blotting as described in the previous study [1].
Antibodies used were as follows: anti-phospho-Akt antibody (Ser473, Cell Signaling Technology, Danvers, MA,

Page 5 of 13

USA, #9271), anti-Akt antibody (Cell Signaling Technology, #9272), anti-phospho-p70 S6K antibody (Thr389,
Cell Signaling Technology, #9234), anti-p70 S6K antibody(Santa Cruz Biotechnology, C-18, sc-230), anti-phospho-S6 ribosomal protein antibody (Ser240/244, Cell
Signaling Technology, #2215), anti-S6 ribosomal protein
antibody (Cell Signaling Technology, 54D2, #2317), anti4EBP1 antibody (Cell Signaling Technology, #9452), antiα-tubulin antibody (Sigma Aldrich, T9026), anti-rabbit
IgG, horseradish peroxidase (HRP)-linked whole antibody from donkey (GE healthcare, NA934V), and antimouse IgG, HRP-linked whole antibody from sheep (GEhealthcare, NA931V). Band intensities of each blot were
quantified using ImageJ.
Statistical analyses

Except for an analysis of myofiber diameters, grapheddata are expressed as means ± SE. Unpaired Student’s
t-test (between two groups) and one-way analysis of variance (ANOVA) followed by a Tukey–Kramer test (formultiple group comparison) were used to evaluate statistical differences. For the experiment to assess the difference of myofiber diameters, the p-value was determinedusing the Wilcoxon rank sum test. p-values less than 0.05
were considered statistically significant.

Results
We previously reported that the body weight of DMD
rats decreased progressively after the age of 6 months[33]. Suspecting that eating disorder occurs in DMD
rats, we measured the amount of food intake at the agesof 3 months (before the onset of body weight decrease)and 7 months (after the onset of bodyweight decrease).
The amount of food intake at 3 months old was comparable between WT and DMD rats (Fig. 1A). However,at 7 months old, significantly decreased food intake wasobserved in DMD rats, indicating the occurrence of aneating disorder (Fig. 1A).
The above results suggest that muscles involved in foodintake are affected by the progress of the disease. We thenperformed morphometric analyses of muscles involved infood intake. We chose the masseter and tongue muscles
because these two muscles are to be approached easily and distinguishable from other muscles. As shown in
Fig. 1B, the area corresponding to the masseter muscle
was considered as an ellipse, and its long and short diameters were measured with calipers to calculate the area.
The area of the masseter muscle was measured from 1 to
8 months old. At 1 month old and after 6 months old, thearea of the masseter muscle was significantly reduced in
DMD rats compared to WT rats. Especially, the greater
reduction was observed at 6 and 8 months old (Fig. 1C).

Yamanouchi et al. Skeletal Muscle(2022) 12:24

The appearance of the tongue at 8 months old is shownin Fig. 1D. The cross-section of the posterior part of the
tongue was larger in DMD rats than in WT rats (Fig. 1D).
Then, we measured the area and width of the upper halfof the tongue sections. Quantitative analysis showed thatthe width of the tongue of DMD rats was significantly
larger than that of WT rats after 3 months old (Fig. 1E).
In addition, the area of the tongue was significantly
larger in DMD rats at the age of 8 months than in WT
rats (Fig. 1F). These results indicate that macroglossia is
occurring in DMD rats.
To further gain insight into the age-related changes ofthe masseter and tongue muscles of DMD rats, histological analyses were performed. For the tongue muscles, theintrinsic and extrinsic muscles were examined separately.
The masseter muscle of DMD rats showed myofiber
degeneration and necrosis with infiltration of neutrophils and macrophages at 1 month old (Fig. 2B) andPage 6 of 13

regenerative myofibers with enlarged centralized nuclei
and basophilic cytoplasm at 3 months old (Fig. 2D). At
8 months old, in addition to inflammatory and degenerative changes, severe fibrosis and fatty infiltration wereobserved (Fig. 2F, H). No significant pathological changes
were observed in the masseter muscle of WT rats at allages (Fig. 2A, C, E, G).
In contrast, in the intrinsic tongue muscle of DMD rats,
regenerative myofibers with enlarged centralized nuclei
and basophilic cytoplasm were observed at 1 month old(Fig. 2J), and these regenerative changes were accompanied by mild fibrosis at 3 months old (Fig. 2L). At
8 months old, mild fibrosis, also confirmed by Masson’s
trichrome stain (Fig. 2P), was persisted (Fig. 2N). The
extrinsic tongue muscles generally showed similar findings as to the intrinsic tongue muscles, although regenerative changes were observed in some parts of thegenioglossus muscles of DMD rats at 3 and 8 months

Fig. 2 Histological analyses of masseter and tongue muscles of WT and DMD rats. A–F HE- and G, H Masson’s trichrome-stained sections of themasseter muscle of WT and DMD rats. In the masseter muscle of DMD rats, myofiber degeneration and necrosis with infiltration of neutrophils
and macrophages at 1 month old, regenerative myofibers with enlarged centralized nuclei and basophilic cytoplasm at 3 months old, and severe
fibrosis, confirmed by Masson’s trichrome staining, at 8 months old were observed, respectively. No significant pathological changes were observedin the masseter muscle of WT rats at all ages. I–N HE- and O, P Masson’s trichrome-stained sections of the tongue muscle of WT and DMD rats. In
the tongue muscle of DMD rats, regenerative myofibers with enlarged centralized nuclei and basophilic cytoplasm at 1 month old, the regenerativechanges accompanied by mild fibrosis at 3 months old, and persistence of mild fibrosis, confirmed by Masson’s trichrome staining, at 8 months old
were observed, respectively. No significant pathological changes were observed in the tongue muscle of WT rats at all ages

Yamanouchi et al. Skeletal Muscle(2022) 12:24

old (Supplementary Fig. 1). No significant pathologicalchanges were observed in both the intrinsic and extrinsic
tongue muscle of WT rats at all ages (F ig. 2I, K, M, O).
To further examine whether myofiber necrosis andmuscle regeneration are ongoing at 6 months old whenmacroglossia is occurring in the tongue muscle of DMD
rats, immunohistochemical analyses of endogenous IgG(myofiber necrosis) and embryonic MHC (muscle regeneration) were performed. The masseter muscle of DMD
rats, though there is variation among individuals, showedmarked myofiber necrosis (Fig. 3A, E) and muscle regeneration (Fig. 3C, F), while the intrinsic tongue muscle
showed none of these (Fig. 3B, D–F). These results indicate that, although the masseter and tongue muscles of
DMD rats undergo pathological changes that are notseen in WT rats, the changes seen in the tongue muscle
are much milder than in the masseter muscle of DMD
rats.
To investigate the factors that are responsible for macroglossia seen in DMD rats, we measured the myofiber
diameters of the intrinsic tongue muscle of WT and
DMD rats at 6 months old. The myofiber diameters of
DMD rats were larger than those of WT rats (Fig. 3G,
H; p < 0.01 by the Wilcoxon rank sum test). The ratio of
myofibers with central nuclei was significantly higher
in DMD rats than in WT rats, indicating that muscle
regeneration had occurred in the past (F ig. 3I). In addition, quantitative analysis of Masson’s trichrome-stained
sections as shown in Fig. 2 reveals that the fibrotic areain the tongue muscle of DMD rats is about twice that of
WT rats (Fig. 3J). Thus, these results suggest that boththe hypertrophic response of myofibers and the accumulation of collagen between myofibers were combined to
produce macroglossia. Considering the possibility thatcells present in the stromal area are affected, the numberof vascular endothelial cells and mesenchymal progenitor
cells in the tongue muscle was compared between WTPage 7 of 13

and DMD rats at 6 months old. The area occupied by
CD31-positive (endothelial) and CSPG4-positive (mesenchymal progenitor) cells were comparable (Supplementary Fig. 2).
Hypertrophy of myofibers occurs by two mechanisms: increased protein synthesis through activation of
the PI3K-Akt-mTOR pathway and the addition of new
myonuclei by fusion of satellite cells [12]. To gain moreinsight into the mechanism of hypertrophy of myofibers
observed in the tongue muscle of DMD rats, we investigated the amount and phosphorylation of molecules that
act in the downstream of PI3K-Akt-mTOR pathway as
well as the number of nuclei per myofiber in 6-month-old
tongue muscle. No differences were observed between
WT and DMD rats in the amount and phosphorylation of S6K, S6, and 4E-BP1 (Fig. 3K–R; Supplementary
Fig. 3). Neither the expression nor the phosphorylation of
Foxo1, which is thought to be involved in protein degradation, could be detected (data not shown). In addition,
no significant differences were observed in the transcription of two skeletal muscle-specific ubiquitin ligase
genes, Atrogin 1 and MuRF1, which are involved in protein degradation [2] (Fig. 3S). When the number of nuclei
per myofiber in transverse sections was calculated, atrend toward a higher number of nuclei was observed in
the DMD rats, although this was not statistically significant (Fig. 3T). These results suggest that, at least at theage of 6 months, there is neither an increase in protein
synthesis nor a suppression of protein degradation andthat an increased number of fused satellite cells is more
likely involved in the hypertrophy of myofibers.
We reported previously that cellular senescence withincreased transcription of p16 and p19 occurs in thehindlimb muscles of DMD rats and that the proliferation of muscle satellite cells is markedly suppressed inculture [33]. Therefore, we examined the transcription ofsenescence-associated genes in the masseter and tongue(See figure on next page.)
Fig. 3 Analyses of necrotic and regenerating myofibers in the masseter and tongue muscle of WT and DMD rats (A–F). A, B Representative
immunohistochemistry of endogenous IgG (red) and laminin (green) in the masseter (A) and tongue (B) muscle of DMD rats at 6 months old. Nuclei
were stained with Hoechst 33258 (blue). Arrowheads (white) indicate necrotic myofibers. C, D Representative immunohistochemistry of embryonicmyosin heavy chain (eMHC) (red) and laminin (green) in the masseter (A) and tongue (B) muscle of DMD rats at 6 months old. Nuclei were stainedwith Hoechst 33258 (blue). Arrowheads (white) indicate regenerating myofibers. E Quantitative analysis of necrotic and regenerating myofibers inmasseter and tongue muscle of WT (n=5) and DMD (n=6) rats at 6 months old. The data are expressed as mean±SE. *, p<0.05 vs others by Tukey–
Kramer test. Analyses of tongue muscle of WT and DMD rats (G–J). G, H Distribution of myofiber diameters at 6 months old. The left panel (G)depicts the relative distribution of myofibers with certain diameters. The right panel (H) depicts relative cumulative plots based on the data shownin the left panel. The myofiber diameters of DMD rat (n=6) were larger than those of WT rats (n=5). p<0.01 by Wilcoxon rank sum test (number ofmyofibers analyzed, WT=3112 and DMD=2662). I Ratio of myofibers with central nuclei at 6 months old. The data are expressed as mean±SE. **,p<0.01 by unpaired Student’s t-test. J Relative fibrotic area at 6 months old. The data are expressed as mean±SE. **, p<0.01 by unpaired Student’s
t-test. Quantitative analyses of molecules involved downstream of Akt-mTOR pathway in the tongue muscle of WT (n=5) and DMD rats (n=6) at 6
months old (K–R). K–N Ratio of phosphorylated Akt, S6K (S6 kinase), S6 (S6 ribosomal protein), and 4E-BP1. O, P Total amount of Akt, S6K (S6 kinase),
S6 (S6 ribosomal protein), and 4E-BP1 normalized to that of α-tubulin. The data are expressed as mean±SE. Quantitative analyses of expression of
two muscle-specific E3 ubiquitin ligases (Atrogin1 and MuRF1) in the tongue muscle of WT (n=5) and DMD rats (n=6) at 6 months old (S). The data
are expressed as mean±SE. Quantitative analysis in the number of nuclei per myofiber of the tongue muscle of WT (n=5) and DMD rats (n=6) at 6
months old (T). The data are expressed as mean±SEYamanouchi et al. Skeletal Muscle(2022) 12:24

Fig. 3 (See legend on previous page.)

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Yamanouchi et al. Skeletal Muscle(2022) 12:24

muscles at 9 months old. p16 and p19 were markedly
upregulated in the masseter muscle of DMD rats compared to WT rats (Fig. 4A, B). On the other hand, both
p16 and p19 were slightly upregulated in the tongue
muscle of DMD rats but the degrees were significantly
lower than those in the masseter muscle (Fig. 4A, B). p21
transcription was significantly upregulated only in themasseter muscle of DMD rats (Fig. 4C), and there wasno difference in the p53 transcription in both the muscles between WT and DMD rats (Fig. 4D). There was no
significant difference in the number of Pax7-positive satellite cells in the sections, although there was a large variation between individuals (Fig. 4E). On the other hand,when cultured in vitro for 4 days, the number of Pax7- or
MyoD-positive satellite cells derived from the massetermuscle of DMD rats was significantly lower than that of
WT rats (Fig. 4F, G, Supplementary Fig. 4A and B), while
the number of satellite cells derived from the tongue
muscle of DMD rats was only slightly lower than that of
WT rats (Fig. 4F, G, Supplementary Fig. 4A and B). These
results indicate that cellular senescence, possibly throughp16- and p19-dependent pathway, occurs extensively inPage 9 of 13

the masseter muscle of DMD rats as was seen in theirhindlimb muscles [33], while it is significantly less severein the tongue muscle.
The above results suggest that the dystrophic changes
are less advanced in the tongue muscle of DMD rats.
Utrophin, known as dystrophin-related protein, had beenshown to compensate for the function of dystrophin [7,13, 20, 26]. In a mouse model of DMD, mdx mouse, theskeletal muscle phenotype is milder than that seen in
DMD patients [20], and this had been once thought to be
due to an upregulation of utrophin [20]. Thus, we examined the Utrn transcription in the masseter and tongue
muscles of WT and DMD rats at 9 months old. In themasseter muscle, Utrn transcription was significantly
upregulated by the absence of dystrophin (Fig. 4H). Interestingly, in the tongue muscle, Utrn transcription was
also upregulated but its basal expression in the presenceof dystrophin (WT rats) was comparable to that in themasseter muscle of DMD rats (Fig. 4H). This result suggests that the tongue muscle is intrinsically expressing ahigher level of Utrn transcription even in the presence ofdystrophin, and this might be related to the resistance ofFig. 4 Quantitative analyses of gene expression and satellite cells in muscles of WT and DMD rats. A p16, B p19, C p21, and D p53 transcriptionsin the masseter and tongue muscle at 9 months old. The data are expressed as mean ± SE. * and **, p < 0.05 and p < 0.01 by unpaired Student’s
t-test, respectively (WT, n = 5; DMD, n = 6). E Number of Pax7-positive satellite cells in the sections at 9 months old. The data are expressed asmean ± SE. There was no significant difference among all groups by Tukey–Kramer test (WT, n = 5; DMD, n = 6). F, G Relative number of Pax7- and
MyoD-positive satellite cells from 9-month-old rats after 4-day culture in vitro. The data are expressed as mean ± SE. * and **, p < 0.05 and p < 0.01 by
unpaired Student’s t-test, respectively (WT, n = 3; DMD, n = 3). H Utrn transcription in the masseter and tongue muscle at 9 months old. The data areexpressed as mean ± SE. *, p < 0.05 by unpaired Student’s t-test (WT, n = 5; DMD, n = 6)

Yamanouchi et al. Skeletal Muscle(2022) 12:24

this tissue against degenerative changes in the absence ofdystrophin.

Discussion
In the present study, we demonstrated that the tongue of
DMD rats shows macroglossia and degenerative changes
were observed transiently only in the early period of life.
On the other hand, chronic degenerative and necroticchanges were seen in the masseter muscle of DMD rats.
Cellular senescence is caused by oxidative stress due
to chronic inflammation [6]. We previously reported
progressive myofiber degeneration and necrosis, alongwith cellular senescence with increased transcription of
p16 and p19, in the tibialis anterior (TA) and diaphragmof DMD rats [33]. p16 transcript was mainly present insatellite cells and mesenchymal progenitor cells, and their
proliferative activity was severely suppressed in vitro. The
phenotype of the masseter muscle of DMD rats seen inthis study is similar to our previous reports; the masseter muscle showed progressive myofiber degenerationand necrosis, and accordingly, p16 and p19 transcription
was upregulated. In contrast, the transcription of p16 andp19 in the tongue muscle of DMD rats was much lower
than those in the masseter muscle, and the proliferative
activity of the satellite cells was relatively well-preservedin vitro. This is in agreement with the histological observation that myofiber necrosis and inflammation were notobserved in the tongue muscle of DMD rats, suggesting
the absence of oxidative stress that could induce cellularsenescence.
A similar phenotype to the tongue muscle of DMD rats
had been observed in the hindlimb muscle of mdx mice
[10, 19, 35]. In the tongue muscle of DMD rats, regeneration of myofibers was transiently observed at 1 monthold, and at later periods of their life, myofibers with central nuclei persisted without any degenerative changes. In
the skeletal muscle of mdx mice, necrotic regeneration isobserved only up to 1 to 2 months of age, and regenerated
myofibers with central nuclei remain for a long time [10,19, 35]. At present, it is not clear why myofiber necrosis
occurs only transiently in mdx mice, but it has been suggested that one reason is that compensatory increases
in utrophin, a functional homolog of dystrophin, may
replace the function of dystrophin [20]. In the present
study, the Utrn transcription was higher in the tongue
muscle of WT rats than in their masseter muscle. Therefore, one of the possibilities is that some mechanism may
be responsible for the increased transcription of Utrn inthe tongue muscle of the rat, and myofiber necrosis may
not occur even under dystrophin deficiency. However,this study only showed differences in transcription levelsof utrophin by quantitative RT-PCR, and immunohistochemical analysis to determine the localization, as well as

Page 10 of 13

quantitative analysis of protein levels by western blotting,is needed to draw a precise conclusion.
The degree of skeletal muscle regeneration after injury
varies depending on the body part; it has been reported
that the diameter of myofibers and the degree of fibrosis in masticatory muscles of 100-day-old mdx mice vary
according to the muscles [31]. In addition, Yoshioka et al.
reported that when injury and subsequent regenerationof skeletal muscle were induced by cardiotoxin injection in WT and mdx mice, there were differences in therate of regeneration and the degree of increase in muscle
weight after regeneration [40]. In particular, the masseter muscle had a longer time to recover its muscle weightafter injury than the TA muscle, and no hypertrophy ofregenerated myofibers occurred [40]. They suggested
that these differences were due to a distinct nature of thesatellite cells present in each skeletal muscle. Regarding
this, the origin of the satellite cells varies among different muscles [27]. The tongue muscle including satellite
cells is thought to be composed of mixed mesodermal
origins [23, 27] but detailed convincing lineage tracing
experiments of these muscles are not available. On theother hand, the satellite cells of the masseter muscle arederived from the first and second pharyngeal arches of
the cranial paraxial mesoderm with contributions fromthe splanchnic mesoderm [27]. At present, it is unclear
how the different origins of these satellite cells ultimately
affect the nature of the skeletal muscle from which theyarise. However, in the multiple types of myopathies,
including DMD, it has been suggested that differences in
the origin of satellite cells may contribute to the distinct
pathological susceptibility among different muscles [27].
Thus, the phenotypic difference between the masseterand tongue muscles observed in DMD rats may be due tothe difference in the nature of the satellite cells. However,
we cannot exclude the possibility that the differences indystrophic changes in each muscle are simply due to differences in the mechanical load on each muscle. In thisregard, studies that verify differences in muscle loading in
the rat are required.
The tongue muscle has a unique structural feature
among skeletal muscles in that it is covered by epithelium, and its development is indirectly influenced byepithelium [14, 18]. During development, the epithelium-derived sonic hedgehog (shh) acts on cranial neural crest cells (CNCC), which constitute the mesenchyme[14]. When the hedgehog signal in CNCC is defective,
migration of myoblasts from the occipital somite is perturbed, resulting in dysplasia of the tongue. Although
the involvement of the epithelium in the adult tongue
has not been reported, it is possible that factors derived
from the epithelium act directly or indirectly via mesenchyme on the tongue muscle, which may be protectiveYamanouchi et al. Skeletal Muscle(2022) 12:24

against degenerative changes of tongue myofibers causedby dystrophin deficiency. This point awaits further
investigation.
Macroglossia has been reported to occur not only inhuman DMD but also in canine X-linked muscular dystrophy (CXMD) [30]. In human DMD, it is known thatmacroglossia interferes with tongue function [36, 37],
but no detailed histological study has been performed.
On the other hand, it has been suggested that hypertrophy of myofibers is the cause of macroglossia in CXMD[30]. This is consistent with the results of the present
analysis in DMD rats. The mechanisms of skeletal muscle hypertrophy include increased protein synthesis,
decreased degradation, and increased number of myonuclei supplied to myofibers by satellite cells [12]. It has
been reported that some regenerated myofibers in mdx
mice and golden retriever muscular dystrophy (GRMD)
dogs show hypertrophy [4, 8, 15, 16, 24, 29], and one of
the mechanisms is the vigorous activation of the AktmTOR pathway during the period before the occurrenceof myofiber necrosis, which results in hypertrophy ofmyofibers leading to a subsequent reduced severity ofpathology [25]. Since the mechanism of myofiber hypertrophy in the tongue has not been analyzed in depth inany of the species reported [28, 30, 37], we attempted toclarify the mechanism of myofiber hypertrophy in the
tongue muscle of the rat DMD in the present study. As
a result, the signaling molecules involved in the regulation of protein synthesis and degradation were not significantly altered compared to WT rats, at least at theage of 6 months examined in this study. However, thisdoes not necessarily exclude the possibility that the AktmTOR pathway is involved in the hypertrophy of tongue
myofibers in DMD rats, and it still remains possible thatthe transient activation of this pathway at a younger age(before 6 months old) had led to the hypertrophy of muscle fibers. On the other hand, although not statisticallysignificant, the number of nuclei per myofiber showed anincreasing trend, suggesting that a supply of new myonuclei by satellite cells may have occurred. This is supported by the present result that myogenicity of satellite
cells derived from the tongue muscle of DMD rats was
relatively preserved in vitro, even at the age of 9 months,
unlike those from the masseter muscle.
The limitation of this study is that it is still unclear
whether the macroglossia in DMD rats influences tongue
function, and it is unknown whether the decreased foodintake seen in DMD rats is attributed to macroglossia.
Furthermore, it is also possible that the decreased foodintake seen in the DMD rats is caused by general fatigue
such as respiratory problems or cardiomyopathy. In fact,
the DMD rats show extensive fibrosis in their heart [22,33] and similar distributions and progression of heart

Page 11 of 13

involvement to those of patients with DMD [32]. Regarding these points, it is necessary to evaluate the tongue
function experimentally and to elucidate the exact causeof decreased food intake in DMD rats in the future.

Conclusions
We demonstrated that, unlike the masseter and hindlimb
muscles, degenerative/necrotic changes of myofibers,
decreased satellite cell proliferative activity, and increased
p16 and p19 expressions were seldomly observed in the
tongue muscle of DMD rats. Furthermore, the tongue
muscle of the rat showed higher Utrn transcription thanthe masseter muscle, suggesting that this compensatory
Utrn transcription might protect the tongue myofibers
from degenerative/necrotic changes under the lack ofdystrophin. Unraveling the mechanism involved in lessadvanced dystrophic changes in the tongue muscle in
DMD rats would benefit to establish an effective treatment for DMD.
Abbreviations
ANOVA: Analysis of variance; CNCC: Cranial neural crest cells; CSPG4: Chondroi‑
tin sulfate proteoglycan 4; CXMD: Canine X-linked muscular dystrophy; DMD:
Duchenne muscular dystrophy; DMD rat: A rat model of DMD; DSHB: Devel‑
opmental Studies Hybridoma Bank; EDTA: Ethylenediaminetetraacetic acid;
eMHC: Embryonic myosin heavy chain; HE: Hematoxylin–eosin; HPRT: Hypox‑
anthine‐guanine phosphoribosyltransferase; HRP: Horseradish peroxidase;
NICHD: National Institute of Child Health and Human Development; NDS:
Normal donkey serum; NGS: Normal goat serum; PBS: Phosphate-bufferedsaline; PFA: Paraformaldehyde; RIPA: Radioimmunoprecipitation assay; SDSPAGE: Sodium dodecyl sulfate–polyacrylamide gel electrophoresis; Shh: Sonic
hedgehog; TA: Tibialis anterior; WT: Wild-type; XY: Male WT; XDmdY: Male DMD.

Supplementary Information
The online version contains supplementary material available at https://​doi.​
org/​10.​1186/​s13395-​022-​00307-7.
Additional file 1: Supplementary Fig. 1. HE-stained sections of theextrinsic tongue muscle of DMD rats. At 3- and 8-month-old, regenera‑tive myofibers with centralized nuclei and basophilic cytoplasm wereobserved in some genioglossus muscles.
Additional file 2: Supplementary Fig. 2. Immunohistochemical analyses
of vascular endothelial (CD31) and mesenchymal progenitor (CSPG4)
cells. Representative immunohistochemistry of CD31 (red), and CSPG4(red) and laminin (green) in the tongue muscle of WT and DMD rats at
6-month-old. Nuclei were stained with Hoechst 33258 (blue). Grapheddata are quantitative analysis of CD31- and CSPG4-positive areas. The data
are expressed as mean+SE (CD31, WT (n=5) and DMD (n=6); CSPG4, WT(n=3) and DMD (n=3)).
Additional file 3: Supplementary Fig. 3. Western blot of molecules
involved downstream of Akt-mTOR pathway in the tongue muscle at
6-month-old. Two bands (doublet) were detected by the anti-4E-BP1
antibody. The upper (p-4E-BP1 (high)) and lower (p-4E-BP1 (low)) bands
correspond to highly phosphorylated 4E-BP1 and non- or lower-phos‑
phorylated 4E-BP1, respectively. α-tubulin was used as an internal control.
p-Akt (S473), Akt phosphorylated at ­Ser473. p-S6K (T389), S6 kinase phos‑
phorylated at ­Thr389. p-S6 (S240/244), S6 ribosomal protein phosphoryl‑
ated at ­Ser240 and ­Ser244. n, the number of samples.
Additional file 4: Supplementary Fig. 4. Representative immunocyto‑chemistry of satellite cells. The cultured cells were immunostained with

Yamanouchi et al. Skeletal Muscle(2022) 12:24

anti-Pax7 (A) and anti-MyoD (B) on day 4 of culture. Nuclei were stainedwith Hoechst 33258. Arrowheads (white) indicate positive cells.

Page 12 of 13

7.
8.

Authors’ contributions
KY conducted the experiments, analyzed the data, prepared the figures, andwrote the manuscript; YT, MI, RO, and HN conducted the experiments; SK pre‑pared tissue sections and conducted the experiments; JC, FH, SIT, TM, and KU
edited the manuscript. The authors read and approved the final manuscript.
Funding
The anti-Pax7 and anti-eMHC monoclonal antibodies were obtained from theDevelopmental Studies Hybridoma Bank developed under the NICHD andmaintained by the University of Iowa. This work was supported by Grant-inAid for Scientific Research (B) (20H03161, 19K22359, 16H05041 and 15K14883
to KY) from Japan Society for the Promotion of Science, Cross-ministerial
Moonshot Agriculture, Forestry, and Fisheries Research and Develop‑
ment Program, “Technologies for Smart Bio-industry and Agriculture” from
Bio-oriented Technology Research Advancement Institution, NARO (Grant
Number 20350956 to SIT), and Core-to-Core program from Japan Society for
the Promotion of Science (Grant Number JPJSCCA20210007 to SIT).

9.
10.

11.
12.
13.
14.
15.

Declarations
Ethics approval and consent to participate
All animal experiments performed in this study were in accordance with the
Guide for the Care and Use of Laboratory Animals of the University of Tokyoand were approved by the Institutional Animal Care and Use Committee ofthe University of Tokyo (P18-125). All studies were conducted in compliancewith the Helsinki Declaration.

16.
17.

Competing interests
The authors declare that they have no competing interests.

18.

Author details
1
Laboratory of Veterinary Physiology, Graduate School of Agricultural and Life
Sciences, The University of Tokyo, 1‑1‑1 Yayoi, Bunkyo‑ku, Tokyo 113‑8657,
Japan. 2 Laboratory of Veterinary Pathology, Graduate School of Agriculturaland Life Sciences, The University of Tokyo, 1‑1‑1 Yayoi, Bunkyo‑ku, Tokyo
113‑8657, Japan. 3 Laboratory of Animal Cell Regulation, Graduate Schoolof Agricultural and Life Sciences, The University of Tokyo, 1‑1‑1 Yayoi,
Bunkyo‑ku, Tokyo 113‑8657, Japan.

19.
20.
21.
22.

Received: 21 February 2022 Accepted: 6 October 2022
23.
24.
References
1. Ariga M, Nedachi T, Akahori M, Sakamoto H, Ito Y, Hakuno F, Takahashi S.
Signalling pathways of insulin-like growth factor-I that are augmented bycAMP in FRTL-5 cells. Biochem J. 2000;348(Pt 2):409–16.
2. Bodine SC, Latres E, Baumhueter S, Lai VK, Nunez L, Clarke BA,
Poueymirou WT, Panaro FJ, Na E, Dharmarajan K, Pan ZQ, Valenzuela
DM, DeChiara TM, Stitt TN, Yancopoulos GD, Glass DJ. Identificationof ubiquitin ligases required for skeletal muscle atrophy. Science.
2001;294:1704–8.
3. Botteron S, Verdebout CM, Jeannet PY, Kiliaridis S. Orofacial dysfunction in
Duchenne muscular dystrophy. Arch Oral Biol. 2009;54:26–31.
4. Coulton GR, Curtin NA, Morgan JE, et al. The mdx mouse skeletal muscle
myopathy: II. Contractile properties. Neuropathol Appl Neurobiol.
1988;14:299–314.
5. da Costa KVT, Ribeiro CMB, de Carvalho FD, Gonçalves LS, de Almeida
OP, de Carvalho Silva LT, Bastos YVP, Ferreira SMS. Dysphagia due to mac‑
roglossia in a patient with amyloidosis associated with multiple myeloma:
a case report. Spec Care Dentist. 2018;38:255–8.
6. Davalli P, Mitic T, Caporali A, Lauriola A, D’Arca D. ROS, cell senescence,and novel molecular mechanisms in aging and age-related diseases. Oxid
Med Cell Longev. 2016;2016:3565127.

25.
26.
27.
28.
29.
30.

31.

Deconinck AE, Rafael JA, Skinner JA, Brown SC, Potter AC, Metzinger L,
Watt DJ, Dickson JG, Tinsley JM, Davies KE. Utrophin-dystrophin-deficient
mice as a model for Duchenne muscular dystrophy. Cell. 1997;90:717–27.
DiMario JX, Uzman A, Strohman RC. Fiber regeneration is not persistent in
dystrophic (MDX) mouse skeletal muscle. Dev Biol. 1991;148:314–21.
Duan D, Goemans N, Takeda S, Mercuri E, Aartsma-Rus A. Duchenne
muscular dystrophy. Nat Rev Dis Primers. 2021;7:13.
Duddy W, Duguez S, Johnston H, Cohen TV, Phadke A, Gordish-Dressman
H, Nagaraju K, Gnocchi V, Low S, Partridge T. Muscular dystrophy in the
mdx mouse is a severe myopathy compounded by hypotrophy, hyper‑
trophy and hyperplasia. Skelet Muscle. 2015;5:16.
Dumont NA, Bentzinger CF, Sincennes MC, Rudnicki MA. Satellite cells
and skeletal muscle regeneration. Compr Physiol. 2015;5:1027–59.
Fukada SI, Ito N. Regulation of muscle hypertrophy: involvement of the
Akt-independent pathway and satellite cells in muscle hypertrophy. Exp
Cell Res. 2021;409:112907.
Grady RM, Teng H, Nichol MC, Cunningham JC, Wilkinson RS, Sanes JR.
Skeletal and cardiac myopathies in mice lacking utrophin and dystrophin:a model for Duchenne muscular dystrophy. Cell. 1997;90:729–38.
Jeong J, Mao J, Tenzen T, Kottmann AH, McMahon AP. Hedgehog signal‑
ing in the neural crest cells regulates the patterning and growth of facial
primordia. Genes Dev. 2004;18:937–51.
Kornegay JN, Childers MK, Bogan DJ, Bogan JR, Nghiem P, Wang J, Fan Z,
Howard JF Jr, Schatzberg SJ, Dow JL, Grange RW, Styner MA, Hoffman EP,
Wagner KR. The paradox of muscle hypertrophy in muscular dystrophy.
Phys Med Rehabil Clin N Am. 2012;23:149–72.
Kornegay JN, Cundiff DD, Bogan DJ, Bogan JR, Okamura CS. The cranial
sartorius muscle undergoes true hypertrophy in dogs with golden
retriever muscular dystrophy. Neuromuscul Disord. 2003;13:493–500.
Le Révérend BJ, Edelson LR, Loret C. Anatomical, functional, physiologi‑
cal and behavioural aspects of the development of mastication in earlychildhood. Br J Nutr. 2014;111:403–14.
Lin C, Fisher AV, Yin Y, Maruyama T, Veith GM, Dhandha M, Huang GJ, Hsu
W, Ma L. The inductive role of Wnt-β-Catenin signaling in the formation oforal apparatus. Dev Biol. 2011;356:40–50.
Massopust RT, Lee YI, Pritchard AL, Nguyen VM, McCreedy DA, Thompson
WJ. Lifetime analysis of mdx skeletal muscle reveals a progressive pathol‑ogy that leads to myofiber loss. Sci Rep. 2020;10:17248.
Matsumura K, Ervasti JM, Ohlendieck K, Kahl SD, Campbell KP. Associationof dystrophin-related protein with dystrophin-associated proteins in mdx
mouse muscle. Nature. 1992;360:588–91.
Motohashi N, Asakura A. Muscle satellite cell heterogeneity and selfrenewal. Front Cell Dev Biol. 2014;2:1.
Nakamura K, Fujii W, Tsuboi M, Tanihata J, Teramoto N, Takeuchi S, Naito
K, Yamanouchi K, Nishihara M. Generation of muscular dystrophy model
rats with a CRISPR/Cas system. Sci Rep. 2014;4:5635.
Parada C, Han D, Chai Y. Molecular and cellular regulatory mechanisms of
tongue myogenesis. J Dent Res. 2012;91:528–35.
Pastoret C, Sebille A. Mdx mice show progressive weakness and muscle
deterioration with age. J Neurol Sci. 1995;129:97–105.
Peter AK, Crosbie RH. Hypertrophic response of Duchenne and limbgirdle muscular dystrophies is associated with activation of Akt pathway.
Exp Cell Res. 2006;312:2580–91.
Rafael JA, Tinsley JM, Potter AC, Deconinck AE, Davies KE. Skeletal musclespecific expression of a utrophin transgene rescues utrophin-dystrophin
deficient mice. Nat Genet. 1998;19:79–82.
Randolph ME, Pavlath GK. A muscle stem cell for every muscle: variabilityof satellite cell biology among different muscle groups. Front Aging
Neurosci. 2015;7:190.
Renard D, Humbertclaude V, Labauge P. Macroglossia in adult Duchenne
muscular dystrophy. Acta Neurol Belg. 2010;110:288.
Sacco P, Jones DA, Dick JR, et al. Contractile properties and susceptibility
to exercise-induced damage of normal and mdx mouse tibialis anterior
muscle. Clin Sci. 1992;82:227–36.
Shimatsu Y, Yoshimura M, Yuasa K, Urasawa N, Tomohiro M, Nakura M,
Tanigawa M, Nakamura A, Takeda S. Major clinical and histopathologicalcharacteristics of canine X-linked muscular dystrophy in Japan. CXMDJActa Myol. 2005;24:145–54.
Spassov A, Gredes T, Gedrange T, Lucke S, Pavlovic D, Kunert-Keil C.
Histological changes in masticatory muscles of mdx mice. Arch Oral Biol.
2010;55:318–24.

Yamanouchi et al. Skeletal Muscle(2022) 12:24

Page 13 of 13

32. Sugihara H, Kimura K, Yamanouchi K, Teramoto N, Okano T, Daimon
M, Morita H, Takenaka K, Shiga T, Tanihata J, Aoki Y, Inoue-Nagamura
T, Yotsuyanagi H, Komuro I. Age-dependent echocardiographic andpathologic findings in a rat model with Duchenne muscular dystrophy
generated by CRISPR/Cas9 genome editing. Int Heart J. 2020;61:1279–84.
33. Sugihara H, Teramoto N, Nakamura K, Shiga T, Shirakawa T, Matsuo M,
Ogasawara M, Nishino I, Matsuwaki T, Nishihara M, Yamanouchi K. Cellularsenescence-mediated exacerbation of Duchenne muscular dystrophy.
Sci Rep. 2020;10:16385.
34. Takeuchi S, Nakano S, Nakamura K, Ozoe A, Chien P, Yoshihara H, Hakuno
F, Matsuwaki T, Saeki Y, Takahashi S, Yamanouchi K, Nishihara M. Roles ofchondroitin sulfate proteoglycan 4 in fibrogenic/adipogenic differentia‑
tion in skeletal muscle tissues. Exp Cell Res. 2016;347:367–77.
35 Torres LF, Duchen LW. The mutant mdx: inherited myopathy in themouse. Morphological studies of nerves, muscles and end-plates. Brain.
1987;110:269–99.
36. van den Engel-Hoek L, de Groot IJ, Sie LT, van Bruggen HW, de Groot SA,
Erasmus CE, van Alfen N. Dystrophic changes in masticatory muscles
related chewing problems and malocclusions in Duchenne muscular
dystrophy. Neuromuscul Disord. 2016;26:354–60.
37. van den Engel-Hoek L, Erasmus CE, Hendriks JC, Geurts AC, Klein WM,
Pillen S, Sie LT, de Swart BJ, de Groot IJ. Oral muscles are progressively
affected in Duchenne muscular dystrophy: implications for dysphagia
treatment. J Neurol. 2013;260:1295–303.
38. Waisman A, Norris AM, Elías Costa M, Kopinke D. Automatic and unbiased
segmentation and quantification of myofibers in skeletal muscle. Sci Rep.
2021;11:11793.
39. Yamanouchi K, Nakamura K, Takeuchi S, Hosoyama T, Matsuwaki T,
Nishihara M. Suppression of MyoD induces spontaneous adipogenesis inskeletal muscle progenitor cell culture. Anim Sci J. 2021;92:e13573.
40. Yoshioka K, Kitajima Y, Seko D, Tsuchiya Y, Ono Y. The body region specific‑
ity in murine models of muscle regeneration and atrophy. Acta Physiol(Oxf ). 2021;231:e13553.

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