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RESEARCH ARTICLE
Feedback regulation of Notch signaling and
myogenesis connected by MyoD–Dll1 axis
Haifeng ZhangID
1
, Renjie ShangID
1,2
, Pengpeng BiID
1,2
*
1Center for Molecular Medicine, University of Georgia, Athens, Georgia, United States of America,
2Department of Genetics, University of Georgia, Athens, Georgia, United States of America
*pbi@uga.edu
Abstract
Muscle precursor cells known as myoblasts are essential for muscle development and
regeneration. Notch signaling is an ancient intercellular communication mechanism that
plays prominent roles in controlling the myogenic program of myoblasts. Currently whether
and how the myogenic cues feedback to refine Notch activities in these cells are largely
unknown. Here, by mouse and human gene gain/loss-of-function studies, we report that
MyoD directly turns on the expression of Notch-ligand gene Dll1 which activates Notch path-
way to prevent precautious differentiation in neighboring myoblasts, while autonomously
inhibits Notch to facilitate a myogenic program in Dll1 expressing cells. Mechanistically, we
studied cis-regulatory DNA motifs underlying the MyoD–Dll1–Notch axis in vivo by charac-
terizing myogenesis of a novel E-box deficient mouse model, as well as in human cells
through CRISPR-mediated interference. These results uncovered the crucial transcriptional
mechanism that mediates the reciprocal controls of Notch and myogenesis.
Author summary
The formation of skeletal muscle tissue during development and regeneration is orches-
trated by controlling the expression levels of muscle-specific transcriptional factors
including MyoD gene. Previous studies have identified the key function of Notch signal-
ing, an evolutionarily conserved cell-cell communication pathway, in blocking the expres-
sion of MyoD and the generation of functional muscle cells. Therefore, at the beginning
of myogenesis, the activity of Notch pathway needs to be downregulated in order to pro-
mote MyoD expression thus the induction for expression of muscle structural genes.
Here, we identified a key regulatory mechanism by which MyoD directly induces the tran-
scription of Dll1 gene which encodes a classical Notch ligand. Interestingly, Dll1 can
dampen Notch cell-autonomously to promote muscle cell differentiation while activate
Notch in neighboring cells to block their myogenic differentiation. The discovery of the
MyoD–Dll1–Notch gene control axis fills the knowledge gap for our understanding of the
molecular regulation of myogenesis.
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OPEN ACCESS
Citation: Zhang H, Shang R, Bi P (2021) Feedback
regulation of Notch signaling and myogenesis
connected by MyoD–Dll1 axis. PLoS Genet 17(8):
e1009729. https://doi.org/10.1371/journal.
pgen.1009729
Editor: Anthony B. Firulli, Indiana University Purdue
University at Indianapolis, UNITED STATES
Received: March 16, 2021
Accepted: July 20, 2021
Published: August 9, 2021
Copyright: ©2021 Zhang et al. This is an open
access article distributed under the terms of the
Creative Commons Attribution License, which
permits unrestricted use, distribution, and
reproduction in any medium, provided the original
author and source are credited.
Data Availability Statement: All relevant data are
within the manuscript and its Supporting
Information files.
Funding: This work was supported by the starting
up fund from the University of Georgia to P.B. The
funders had no role in study design, data collection
and analysis, decision to publish, or preparation of
the manuscript.
Competing interests: The authors have declared
that no competing interests exist.
Introduction
Skeletal muscle accounts for around 40% of adult human body weight. Myogenesis involves a
series of events that begins with the specification of muscle lineage by master transcriptional
regulators, including Pax7 and Muscular Regulatory Factor (MRF), followed by the expression
of a vast number of genes that establish muscle structure and function [1–6]. In adult tissue,
muscle stem cells normally remain in a quiescent state, but can be promptly activated by
injury, following which the stem cells enter cell-cycle to generate a pool of precursors which
either differentiate to repair myofiber or self-renew to replenish stem cell pool [7,8]. The pre-
cise control of these diverse states of muscle stem cell is key for proper tissue homeostasis [9–
11].
Notch signaling is an ancient intercellular communication mechanism that determines the
cell-fate for various tissue types in metazoan [12–15]. Alterations of this pathway underline a
spectrum of disease and cancer [16–20]. Transduction of Notch signaling is initiated upon
binding of a Notch receptor to a ligand located on a neighboring cell [14,21]. Notch ligands
are members of the DSL (Delta/Serrate/LAG-2) family protein that include Delta-like (Dll1,
Dll3, Dll4) and Jagged (Jag1, Jag2) in mammals. Endocytosis of Notch-bound ligand generates
a mechanical pulling force that drives conformational change of the Notch receptor [22]. This
facilitates subsequent proteolytic cleavage of Notch receptor and produces Notch intracellular
domain (NICD) [23]. As a transcriptional activator, NICD then translocates to nucleus where
it binds with Rbpj and recruits a transcriptional complex to activate the expression of down-
stream target including Hairy/enhancer of split (Hes) and Hes-related with YRPW motif
(Hey) family genes [24]. Despite the simplicity in design, the biological function of Notch sig-
naling is highly context-dependent [12,25]. This is in part due to versatile ligand-utilizations
for signaling in various biological processes [25,26].
In skeletal muscle, Notch is popularly known as a potent inhibitor of myogenic differentia-
tion [27–31]. Genetic studies also unveiled a key paradigm of this signaling pathway in enforc-
ing the quiescent state of muscle stem cells [30,32–35]. Of note, deletion of Rbpj or Notch-
ligand Dll1 in mouse resulted in depletion of muscle progenitor cells accompanied by severe
muscle hypotrophy and failure of muscle regeneration [30,36–38]. On the other hand, consti-
tutive activation of Notch in myocytes is sufficient to induce cellular dedifferentiation that re-
expression of stem-cell markers [39]. Notch signaling is also essential to build a specialized
microenvironment or niche that controls muscle stem cell function [40–42]. In the proximity
to stem cell, endothelial cells from microvasculature can supply Notch-ligand Dll4 that acti-
vates Notch and maintains a quiescent state of muscle stem cells [43]. Similarly, differentiating
muscle cell can also convey a self-renewing signal by providing Dll1 [44,45]. Although Dll1
and Dll4 appeared to exert similar function for skeletal muscle cell in mouse, opposing func-
tion of these ligands in regulation of myogenesis were identified in chick somites [46].
Currently, it remains largely unknown whether and how the core myogenic program feed-
back to restrain Notch activity; and how does this reciprocal regulation determine the differen-
tiation dynamics of myoblast. Here, by gene gain/loss-of-function studies in human and
mouse cells, we unraveled the crucial role of MyoD in controlling Dll1 expression and Notch
activity. Strikingly, deletion of MyoD in mouse or human myoblast abolished Dll1 expression,
whereas forced expression of MyoD robustly induced Dll1 transcription in fibroblast. Utilizing
a novel line of mutant mouse, we also probed the cis-regulatory element whereby MyoD con-
trols Dll1 expression. Employing heterologous cell-mixing experiment, we report the cis-inhib-
itory and trans-activation roles of Dll1 on Notch transduction whereby divergent myogenic
programs can be established among the subpopulations of myoblast. These results provide an
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insight to the evolutionarily conserved control mechanism of myogenesis for mouse and
human.
Results
Strong induction of Notch ligand Dll1 expression during human and
mouse myogenesis
Notch ligands play either redundant or divergent functions depending on the biological con-
text [26,47]. To understand which Notch ligand(s) may participate the human myogenic pro-
gram, we performed gene expression analyses in low-passage human myoblasts that were
derived from a healthy donor as described previously [48,49]. Of note, these cells display
robust myogenic and fusogenic potentials (Fig 1A). Shortly after three days’ induction, huge
myosin+ syncytia that contained hundreds nuclei can be observed (Fig 1A), accompanied by
prompt inductions of myogenin (MyoG) and myosin expression (Fig 1B). Therefore, these
human myoblasts can serve as an ideal model to dissect the genetic mechanism of human
myogenesis.
Among all five Notch ligands, expression of DLL1 was the highest induced and peaked at 36
hours post human myoblast differentiation (Fig 1C). Its expression was downregulated there-
after (Fig 1C). By comparison, expression of JAG1 was induced toward later stages of myoblast
differentiation. In parallel, we also examined the expression patterns of Notch ligand genes
during mouse myogenic differentiation in vitro and muscle regeneration in vivo by querying
RNA-seq datasets generated from previous studies [50,51]. Similar with human data, expres-
sion of Dll1 and Jag1 were promptly induced upon differentiation of mouse myoblasts (Fig
1D), though only Dll1 showed a dramatic increase during muscle regeneration (Fig 1E).
Fig 1. Robust inductions of Dll1 expression by myogenic cues in both mouse and human cells. (A) Human myoblasts
display high myogenic and fusogenic potentials in culture. Cells were labelled by GFP to visualize the syncytium at early
stages of differentiation. Scale bar, 100 μm. (B) Western blotting results of myosin (MF20 antibody) and Myogenin
(MyoG) in human myoblasts at various stages of differentiation. (C) qPCR results of Notch ligand genes during human
myoblast differentiation. The Ct values for each gene at growth-culture condition (time 0) are Dll1 (25.6), Dll3 (26.8),
Dll4 (32.4) and Jag1 (28.6). Jag2 expression was not detected. (D,E) RNA sequencing results of Notch ligand genes and
Notch target Heyl during mouse myoblast differentiation (D, GSE20846) and muscle regeneration (E, GSE97764). CTX
(cardiotoxin) samples were from day 3 post injury. FC, expression fold change.
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Expression of canonical Notch target gene Heyl was also significantly upregulated in these
myogenic conditions (Fig 1D and 1E). These results indicate that Dll1 may play an evolution-
arily conserved role in controlling myogenesis for the various mammal species.
DLL1 transactivates Notch signaling which inhibits human myoblast
differentiation
Depending on the model of selection, activation of Notch by Dll1 was reported to either pro-
mote myogenesis in chicken [46] or inhibit myogenesis in mice [37,44]. To probe the role of
Dll1 in regulations of Notch signaling and human myogenesis, we set up a unique cell-mixing
and gene expression assay (Fig 2A). First, we overexpress Dll1 in mouse fibroblasts which were
later co-cultured with human myoblasts. RNA from the mixing culture was collected and used
Fig 2. Dll1 transactivates Notch which inhibits human myogenic differentiation. (A) Schematics of heterologous
cell-mixing assay and Notch signaling pathway. (B) DNA electrophoresis results that validated the specificity of human
qPCR primers. (C) qPCR results of human HEY1 and HEYL using primers tested in B. Cells were differentiated for 24
hours after mixing. EV: empty control vector. The presence or absence of indicated cell types is indicated by plus or
minus sign, respectively. n= 3. (D) Fluorescence images of mouse fibroblasts and human myoblasts in separate or
mixing-culture conditions. Human myoblasts were labelled by red fluorescence protein Cherry; mouse fibroblasts
(10T1/2) were infected by control (EV) or Dll1 expressing retroviruses. Cells were differentiated for 24 hours. Scale
bar, 25 μm. (E) Quantification results of MyoG+ myoblasts as shown in D. n= 3. (F) Myosin immunostaining of
human myoblasts that were differentiated for 96 hr. Scale bar, 100 μm. (G) Measurement of myoblast fusion as in F.
n= 3. (H,I) qPCR results for genes in primary mouse myoblasts after mixing culture with fibroblasts. Data are
means ±SEM. P<0.05, P<0.01, P<0.001. ns, not significant.
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to measure Notch-target gene expression with human sequence-specific primers. This allows
us to gauge Notch activity in human myoblasts without force separating them from fibroblasts.
Given Notch activation strictly requires physical contact between adjacent cells, this experi-
ment design avoids potential disruption of Notch signaling from scenarios where cells have to
be first separated before gene expression analysis.
We selected Notch target genes HEY1 and HEYL to gauge Notch activity because they are
abundantly expressed in human myoblasts while the species-specificity of qPCR primers were
confirmed (Fig 2B). Through these experiments, we detected dramatic inductions of HEY1
and HEYL expression in human myoblasts when co-cultured with mouse fibroblasts express-
ing Dll1 (Fig 2C), suggesting that Dll1 is capable of transactivating Notch pathway in human
muscle cells.
We continued to dissect the biological function of Dll1 in human myoblasts by examining
its impact on the myogenic differentiation. Human myoblasts were pre-labelled with red fluo-
rescence protein Cherry and induced to differentiate after mixing with mouse Dll1+ fibroblasts.
Notably, this co-culture scheme strongly inhibited myogenic differentiation evidenced by
drastic reductions of MyoG and myosin expressions (Fig 2D–2F). Accordingly, myoblast
fusion, a hallmark of skeletal myogenesis, was completely abolished when the cells were co-cul-
tured with Dll1+ fibroblasts (Fig 2F and 2G).
Beyond the inhibition of myogenic differentiation, Notch signaling is also essential for
maintaining the quiescence of muscle stem cells and the expression of Pax7, a faithful marker
and master regulator of muscle stem cells [52]. Because the immortalized human myoblasts do
not express Pax7, we chose primary mouse myoblasts to examine a role of Dll1-transduced
Notch signaling on the expression of muscle stem cell markers. As expected, when these cells
were mixed with Dll1+ fibroblasts, Pax7 expression was significantly increased, along with
other markers that are either exclusively (Calcr,Tenm4) or abundantly (Cdh15) expressed by
quiescent muscle stem cells (Fig 2H and 2I).
Dll1 cis-inhibits Notch and promotes human myogenic differentiation
Dll1 expression can be promptly induced in myocytes by myogenic stimulation. Given the
potent anti-myogenic activity of Notch, permissive Notch activations by signaling within Dll1
+ myocytes would cause the termination myogenic differentiation. Therefore, we hypothesized
that Dll1+ myocytes may employ an intrinsic mechanism to escape Notch activation in order
to complete the myogenic program.
Intriguingly, forced expression of Dll1 in human myoblasts significantly reduced the
expression of Notch target genes HEY1,HEYL and HES1 by 75%, 45% and 70% respectively
(Fig 3A). This result hints at a cis-inhibitory action of Dll1, i.e. ectopic ligand expression in sig-
nal-receiver cells blocked the signaling from senders, a paradigm that was previously reported
in other developmental processes [53]. To prove this, we again employed heterologous cell-
mixing experiment that can distinguish the cis- and trans-effects of Dll1 (Fig 3B). Consistently,
overexpression of Dll1 in mouse fibroblasts can faithfully activate Notch in human myoblasts
shown by inductions of HEY1 and HEYL expression (Fig 3C), accompanied by drastic reduc-
tions of MyoD expression (Fig 3D–3F) and myoblast fusion (Fig 3E and 3G). As expected,
such trans-acting effects of Dll1 were abolished by γ-secretase inhibitor DAPT (Fig 3E–3G,
group 5 vs 4). Interestingly, similar to DAPT, forced expression of Dll1 in human myoblasts
(signal receivers) also blocked Notch activation and restored MyoD expression, myogenic dif-
ferentiation and myoblast fusion (Fig 3C–3G, group 6 vs 4). Together, these results reveal the
cis-inhibitory role of Dll1 that blocks Notch activation while promotes myogenic
differentiation.
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Deletion of DLL1 from human myoblasts abolishes its trans-activating and
cis-inhibitory effects on Notch signaling
To examine the role of endogenous DLL1 in human myoblasts, we performed DLL1 loss-of-
function study by CRISPR/Cas9 mediated mutagenesis (Fig 4A). First, human myoblasts were
transduced with lentiviruses that deliver the expression of Cas9 and one pair of guide RNA
(gRNA) targeting the coding exon 4 of DLL1 gene (Fig 4B). Later, we obtained myoblast clones
that were expanded from single-cell isolations after CRISPR treatment (Fig 4A). Genotyping
analysis by PCR (Fig 4C) and sequencing (Fig 4D) revealed biallelic frameshift mutations in
one DLL1 knockout clone (DLL1
KO
). Possibly due to nonsense-mediated decay of resultant
messenger RNA, expression of DLL1 in DLL1
KO
myoblasts was reduced by 87%, compared to
control clones transduced with only Cas9 (Fig 4E). As expected, the signaling of Notch from
human to mouse cells in the mixing culture was compromised when DLL1 was deleted in
human myoblasts (Fig 4F). The dampened Notch activity in mouse myoblasts is also accompa-
nied by a lower expression level of Pax7 (Fig 4G and 4H) and higher expression levels of myo-
genic differentiation markers, e.g. MyoG and Myh3 (Fig 4I). Thus, these results confirmed the
trans-acting effects of DLL1 on Notch and myogenic gene expression.
Fig 3. Dll1 cis-inhibits Notch signaling and promotes human myoblast differentiation. (A) qPCR results of Notch
target genes in human myoblasts in response to Dll1 overexpression. Cells are cultured in growth medium. (B)
Schematic of Notch signaling in heterologous cell-mixing assays for panels C–G. (C,D) Relative mRNA levels of Notch
target genes HEY1/HEYL (C) and MyoD (D) in human myoblasts. Cells were differentiated for 24 hours. n= 3. (E)
Fluorescence images of mouse fibroblasts (10T1/2) and human myoblasts in separate or mixing-culture conditions.
Human myoblasts were labelled by red fluorescence protein Cherry; mouse fibroblasts or human myoblasts were
infected by control (EV) or Dll1 expressing retroviruses. Cells were mixed at day-2 post infection followed by
myogenic differentiation for 24 hours (top) or 96 hours (bottom). Scale bar, 100 μm. (F) Quantification results of
MyoD+ cells as in E (top). n= 3. (G) Measurements of fusion index and nucleus number per syncytium of human
myotubes as in E (bottom). Cells were differentiated for 96 hours. Treatments in groups 1–6 are same for panels E–G.
n= 3. Data are means ±SEM. P<0.05, P<0.01, P<0.001.
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Studies of cis-inhibition of Notch by their ligands have historically based on overexpression
strategy [53]. We wish to examine the cis-inhibitory effect of DLL1 at its endogenous expres-
sion level by comparing WT or DLL1
KO
myoblasts. To achieve this goal, we co-cultured
Fig 4. DLL1 loss-of-function study in human myoblasts. (A) Schematic of CRISPR/Cas9 approach to generate gene-
knockout clones from human myoblasts. (B) Schematic of human DLL1 gene structure and the sequences of gRNAs in
exon 4. (C) DNA electrophoresis results of DLL1 genotyping PCR. (D) Sanger sequencing analysis of human DLL1
genotyping PCR products. The premature stop-codon was detected in exon 5 and exon 7 for the –95 bp and –19 bp
allele, respectively. (E) qPCR results of DLL1 measured using primers located in exon 3 (forward) and exon 4 (reverse).
DLL1 has total 11 exons. (F,G) qPCR results for genes in mouse myoblasts that were co-cultured with human
myoblasts of indicated genotypes. (H) Western blotting analysis of protein lysates from mixing culture of mouse/
human myoblasts. Note that the immortalized human myoblasts do not express Pax7. (I) qPCR results for genes in
mouse myoblasts that were co-cultured with human myoblasts of indicated genotypes. (J) qPCR results that measured
HEY1 and HEYL expression in WT or DLL1
KO
human myoblasts after co-culturing with Dll1+ fibroblasts. (K)
Myosin immunostaining of human myoblasts. (L) Quantification of myosin+ cells as in panel K. n= 3. Data are
means ±SEM. P<0.05, P<0.01, P<0.001.
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human WT or DLL1
KO
myoblasts with Dll1+ fibroblasts which serve as signal senders. Indeed,
the expression levels of Notch target genes HEY1 and HEYL in DLL1
KO
myoblasts were signifi-
cantly higher than those in WT myoblasts after mixing with fibroblasts (Fig 4J). The higher
level of Notch is associated with drastic reduction of myogenic potentials of DLL1
KO
myoblasts
(Fig 4K and 4L). These results validate the cell-autonomous inhibition of Notch by DLL1
which is essential for the proper induction of myogenic differentiation.
Our results thus far suggest that only the signaling between Dll1– receiver and Dll1+ sender
cell is effective, whereas the signaling among the Dll1+ sender cells is less fruitful. Reinforcing
the signaling directionality, we show that Notch activation, recapitulated by direct expression
of NICD, down-regulated DLL1 expression by 87% in human myocytes, together with drastic
inhibitions of myogenic differentiation and markers’ expression (S1 Fig). Therefore, the Notch
effector NICD and ligand Dll1 can reciprocally inhibit the other’s abundance, thus establishing
Notch polarity in human myoblasts.
Cis-inhibitory role of Dll1 is mediated by its extracellular domain
Following ligand-receptor interaction, Notch activity can be refined at multiple steps along the
signal transduction pathway [14]. To probe the stage where Dll1 exerts cis-inhibitory effect on
Notch, we expressed NICD alone or together with Dll1 in human myoblasts. Recapitulating
the effects of Notch activation, expression of NICD significantly induced the expression of
Notch targets and blocked the expression of MyoD/MyoG and myogenic differentiation (Fig
5A and 5B). Of note, co-expression of Dll1 failed to mitigate these effects from NICD (Fig 5A
and 5B). Therefore, Dll1 may cis-inhibit Notch at upstream of the generation of NICD.
Previous structural studies of Notch ligand-receptor interactions suggest that both the
trans- and cis-effects of ligand are carried by its extracellular domain through binding with
Notch receptor [53,54]. To validate this model, we employed DLL1
KO
myoblasts to examine
the cis-inhibition function of mutated Dll1 proteins where either the extracellular or intracellu-
lar domain was removed (Fig 5C). As a validation of our experiment design, we show that
overexpression of Dll1 in fibroblasts (Fig 5D–5G, groups 2 vs 1) can consistently upregulate
expression of Notch target genes in myoblasts and blocks their myogenic differentiation and
fusion; such changes can be abolished when full-length Dll1 is re-expressed in DLL1
KO
myo-
blasts (Fig 5D–5G, groups 3 vs 2). In this context, deletion of the extracellular domain but not
intracellular domain of Dll1 protein abolished the cis-regulatory function of Dll1 (Fig 5D–5G,
groups 4/5 vs 3). Therefore, consistent with previous biochemical studies in non-muscle cells,
Dll1 in myoblasts also exerts the cis-regulatory function through the action of its extracellular
domain.
MyoD is the key regulator of Dll1 expression in human and mouse
myoblasts
Given the significant roles of Dll1 in human myoblasts, we continued to dissect the transcrip-
tional mechanism that governs its expression. The robust inductions of Dll1 during myogenic
differentiation suggest that myogenic regulators, e.g., MyoD and MyoG, may directly govern
Dll1 transcription. Supporting this notion, co-expression of MyoD or MyoG can restore Dll1
expression (S1C Fig) together with the myogenic differentiation of human myoblasts (S1D
Fig) that were blocked by NICD expression (S1D Fig).
To directly examine the regulatory relationship, we generated both mouse and human
MyoD
KO
myoblasts following a CRISPR–Cas9 workflow (Fig 6A). For mouse MyoD gene, one
gRNA from the first coding exon was selected for the targeting. Following clonal expansion,
genotyping analysis by sequencing revealed disruptions of MyoD ORFs (Fig 6B). As a result,
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MyoD protein was depleted from MyoD
KO
myoblasts shown by immunostaining (Fig 6C).
Notably, inactivation of MyoD significantly downregulated Dll1 expression (Fig 6D), accom-
panied by drastic reductions of Hey1 and Heyl expression (Fig 6E). In comparison, expression
of Hes1 showed a trend of reduction yet was not statistically significant compared with control
group (Fig 6E).
When induced for myogenic differentiation, mouse MyoD
KO
group did not to show any
myosin+ cells (Fig 6F). Consistently, expression of myogenin (MyoG), myoblast-fusion factors
myomixer (Mymx) and myomaker (Mymk) [55,56] was undetectable in MyoD
KO
cells (Fig
6G). To ascertain that such phenotypes were attributed to the exact loss of MyoD but not rare
CRISPR off–target effect (if any), we performed rescue experiments. Indeed, myogenic differ-
entiation (Fig 6F and 6G), expression of Dll1 (Fig 6H) and Notch target genes (Fig 6I) were all
restored when MyoD was re-introduced into MyoD
KO
cells. Interestingly, forced expression of
MyoG also achieved similar rescue effects (Figs 6F–6I).
In parallel, we examined the conservation of this regulatory mechanism in human cells. As
such, we generated human MyoD
KO
myoblasts by CRISPR mutagenesis using a pair of gRNA
validated in our previous study [49]. Sequencing confirmed the biallelic frame-shift mutations
in CRISPR treated cells (S2A Fig). Absence of MyoD protein was again validated by immunos-
taining (S2B Fig). Similar with the mouse data, deletion of MyoD from human myoblasts also
abolished myogenic differentiation (S2B and S2C Fig) and significantly inhibited the expres-
sion of DLL1 (S2D Fig) and HEY1 (S2E Fig), which were collectively normalized upon re-
expression of MyoD or MyoG protein in human MyoD
KO
cells (S2B–S2E Fig).
Fig 5. The extracellular domain of Dll1 is required for the cis-inhibition of Notch. (A) Immunostaining results of
human myoblasts with retroviral expression of NICD alone or together with Dll1. Scale bar, 100 μm. (B) qPCR results
of human myoblasts with retroviral expression of NICD alone or together with Dll1. Cells were differentiated for 24
hours. n= 3. (C) Schematic of Notch signaling and cell mixing experiment. (D–F) Myosin immunostaining (D) and
qPCR (E, F) results of the co-cultured myoblast-fibroblast cells with expression of full length or mutated Dll1 proteins.
EV, empty vector control. Note that deletion of extracellular domain abolishes the cis-inhibitory function of Dll1. (G)
Measurement of myoblast fusion as shown in panel D. Treatments in groups1–5 are same for panels D–G. Data are
means ±SEM. P<0.05, P<0.01, P<0.001. ns, not significant.
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To model the regulation of Notch activity by MyoD-Dll1 axis, we performed heterolo-
gous cell-mixing experiment. In this assay, WT or MyoD
KO
human myoblasts were mixed
with mouse myoblasts (Fig 6J). As such, the trans-acting effect upon disruption of MyoD
can be examined by comparing Notch target gene expression between various cell-mixing
groups. We first validated the mouse sequence-specificity of qPCR primers (Fig 6K). Of
note, after mixing with WT-human myoblasts, Heyl expression was significantly induced
in mouse myoblasts (Fig 6L). However, this effect was abolished when human MyoD
KO
myoblasts were used for the mixing experiment (Fig 6L). Such deficiency of Heyl expres-
sion was rescued when Dll1 or MyoD was provided back to human MyoD
KO
myoblasts
(Fig 6L). These results suggest that MyoD, by upregulation of Dll1, can transactivate Notch
signaling in adjacent myoblasts.
Fig 6. MyoD is essential for Dll1 expression in mouse myoblasts. (A) Schematic of CRISPR/Cas9 approach to
generate gene-knockout clones from mouse C2C12 myoblasts. (B) Sanger sequencing results of mouse MyoD
genotyping PCR products. (C) Myod immunostaining confirmed depletions of MyoD protein in MyoD
KO
clones.
Scale bar, 50 μm. (D,E) qPCR results of Dll1 (D) and Notch target genes (E) in WT and MyoD
KO
myoblasts. Cells were
differentiated for 48 hours. n= 3. (F) Immunostaining results of MyoD (top) or MyoG (bottom) together with myosin.
Note that MyoD
KO
myoblasts failed to differentiate and such a defect was rescued by retroviral expression of MyoD or
MyoG. Cells were differentiated for 72 hours. Scale bar, 100 μm. (G) Western blotting results to show the expression
levels of muscle proteins in WT or MyoD
KO
myoblasts. Cells were differentiated for 72 hours. (H,I) qPCR results of
Dll1 (H) and Notch target genes (I) in mouse MyoD
KO
cells. Myoblasts were differentiated for 72 hours. n= 3. (J)
Schematic of Notch signaling that depicted the rationale of human and mouse myoblast-mixing assays. (K) Gel
electrophoresis result that validated the specificity of mouse qPCR primers. (L) qPCR results that measured mRNA
level of mouse Heyl using primers validated in K. Before mixing with mouse myoblasts, WT or MyoD
KO
Human
myoblasts were infected by retroviruses expressing Dll1 or MyoD. Cells were differentiated for 24 hours. n= 3. Data
are means ±SEM. P<0.05, P<0.01, P<0.001. ns, not significant.
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Disparities of Dll1 expression changes upon genetic deletions of MyoG
from human and mouse myoblasts
MyoG is a direct target gene of MyoD during myogenic differentiation [57,58]. Of note, simi-
lar with Dll1, expression of MyoG was also abolished in both mouse (Fig 6F and 6G) and
human (S2C Fig)MyoD
KO
cells. Intriguingly, the biological deficiency of MyoD
KO
cells can be
rescued by forced expression of MyoG, indicating that MyoD may control Dll1 expression
through MyoG.
To directly test a role of MyoG in MyoD–Dll1 regulatory axis, we inactivated MyoG gene
through CRISPR mutagenesis in mouse (Fig 7A) and human myoblasts (S3A Fig) [49]. Suc-
cessful depletions of MyoG proteins were confirmed by Western blotting (Fig 7B) and immu-
nostaining (Figs 7C and S3B) using an antibody that detects the carboxyl terminus of
myogenin. Given the high efficiency of CRISPR editing in C2C12 myoblasts (Fig 7B and 7C),
we first characterized the bulk CRISPR-treated cells before isolating single-clone mutants. Of
note, the majority of MyoG
KO
C2C12 myoblasts did not express myosin after full-term myo-
genic differentiation (Fig 7D). In alignment with the fusogenic defects, expression of Mymx
and Mymk genes was abolished in mouse MyoG
KO
cells (Fig 7B). For Dll1 expression however,
we only observed a mild though significant reduction (33% lower) in mouse MyoG
KO
cells
(Fig 7E). In comparison, the reductions for expression of the Notch target gene Heyl were
more pronounced (83% lower) (Fig 7E). Validating the specific effect of CRISPR-knockout,
myoblast differentiation (Fig 7F, top row) as well as gene expression changes (Figs 7G and 7H)
were rescued upon re-expression of MyoG. More interestingly, forced expression of MyoD
Fig 7. Responses of Dll1 expression upon deletion of MyoG from mouse myoblasts. (A) Schematic of mouse MyoG gene
structure and an example of Sanger sequencing result after CRISPR/Cas9 mediated gene editing in C2C12 mouse myoblasts. (B)
Western blotting results to show the expression levels of myogenic markers in WT and MyoG
KO
myoblasts. Star highlights a non-
specific band. (C,D) Immunostaining results of mouse WT and MyoG
KO
myoblasts at day 2 (C) and day 7 (D) post differentiation.
(E) qPCR results of Dll1 and Heyl in WT and MyoG
KO
myoblasts. Cells were differentiated for 48 hours. n= 3. (F) Immunostaining
result of MyoG and myosin for bulk CRISPR-treated myoblasts (top) and two isolated KO clones (bottom). Note that although
MyoG were invariably depleted, the two KO clones showed a large variation of myogenic capacity. Cells were induced for
differentiated for 72 hours. (G) Western blotting results to show the expression levels of myogenic differentiation markers in and
bulk CRISPR-treated MyoG
KO
myoblasts. Cells were differentiated for 72 hours. (H,I) qPCR results for gene expression in WT and
bulk CRISPR-treated MyoG
KO
myoblasts (H) or MyoG
KO
single-knockout clones (I). Cells were differentiated for 48 hours. n= 3.
Data are means ±SEM. P<0.05, P<0.01, P<0.001. ns, not significant. Scale bars, 100 μm.
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Human muscle differentiation is controlled by MyoD-Dll1 axis
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can also rescue the phenotypes of MyoG
KO
cells, suggesting that MyoD can regulate Notch and
myogenic gene expression independent of MyoG.
In contrast to the complete myogenic failure of MyoD
KO
myoblasts, myosin+ cells were
spotted from the bulk CRISPR-treated MyoG
KO
myoblasts (Fig 7F). We postulated that this
may reflect a heterogeneity of C2C12 myoblasts [59] which showed various responses upon
MyoG deletion, or simply due to the lack of uniformity of gene knockout. Thus, we set out to
isolate and characterize the MyoG
KO
single-clones. Indeed, while the majority of MyoG
KO
clones showed the complete failure of differentiation (e.g. A1 clone), myosin+ cells were read-
ily detected among a few clones after myogenic induction (e.g. A4 clone) (Fig 7F). Interest-
ingly, the changes of Dll1 expression corelated well with myogenic potentials of MyoG
KO
clones (Fig 7I). Similarly, human MyoG
KO
myoblasts, which retain myogenic potential (myo-
sin+) despite of a fusion defect (S3B and S3C Fig), showed a relatively normal level of expres-
sion for DLL1, compared with control group (S3D Fig).
To rule out an effect of cell immortalization on the gene expression changes above, we also
repeated MyoD/MyoG CRISPR experiments in primary mouse myoblasts. Because these cells
are not amendable for clonal expansion and characterization, we opted to directly examine the
expression of Dll1 gene in the bulk CRISPR-treated cells (S4A Fig). Reflecting high efficiency
of knockout, we observed a drastic reduction of MyoD or MyoG expression in the CRISPR
treated groups (S4B Fig). Of note, inactivation of MyoD also reduced the expression of MyoG,
while deletion of MyoG has no obvious effect on the expression level of MyoD (S4B Fig), con-
sistent with the regulation relationship between these factors. Nonetheless, both MyoD and
MyoG CRISPR treatments caused major defects of myogenic differentiation (S4B Fig), accom-
panied by significant reductions of Dll1 expression (S4C Fig). Together, our analyses of mouse
and human gene knockout myoblasts suggest that the induction of Dll1 expression is an inte-
gral part of myogenic program for muscle cells under the control by MyoD and MyoG.
MyoD induces Dll1 transcription in non-muscle cells
Our gain- and loss-of-function experiments unveiled the crucial roles of MyoD in controlling
Dll1 expression in both human and mouse myoblasts. We continued to study the mechanistic
basis underlying this regulation and focused on two questions. First, is the transactivator
MyoD sufficient to induce Dll1 expression? Second, how MyoD transactivates Dll1
expression?
We investigated the first question by performing sufficiency test in fibroblasts which do not
express any myogenic factor. We transduced fibroblasts with MyoD or (and) MyoG. Consis-
tent with our previous report that MyoD is sufficient to induce the expression of Mymx and
Mymk [49], syncytia of fibroblasts are produced (Fig 8A). Strikingly, expression of MyoD in
fibroblast robustly activated Dll1 expression by 66 folds (Fig 8B and 8C). MyoG also induced
Dll1 expression though at much weaker levels compared with MyoD (Fig 8C). Again, expres-
sion of MyoG was also strongly upregulated by MyoD (Fig 8B, right) whereas MyoG does not
significantly induce expression of MyoD (Fig 8B, left). Consistent with the dominant effect of
MyoD, the inductions of Dll1 expression by MyoD was not significantly boosted by adding
MyoG (Fig 8C).
During the fate conversion of fibroblasts, MyoD induces a pan-myogenic program exempli-
fied by the upregulation of MyoG expression (Fig 8B, right) [60–63]. We continued to examine
whether MyoD requires other myogenic factors at its downstream to induce Dll1 expression.
To test this, we concomitantly inhibited protein translation by cycloheximide (CHX) with the
control of MyoD transcriptional activities (Fig 8D). The later was achieved through command-
ing nuclear importing of a MyoD-estrogen receptor fusion protein (MyoD
ER
) [64] with
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Human muscle differentiation is controlled by MyoD-Dll1 axis
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treatment of 4-hydroxytamoxifen (TMX). We reasoned that if MyoD protein is self-sufficient
to activate Dll1 expression, such an induction should not be negated by CHX treatment which
blocks translations of other myogenic factors, e.g. MyoG; by contrary, if CHX compromises
the action of MyoD, it would suggest that MyoD requires function from other factor(s) to
cooperatively activate Dll1 expression.
This experiment design was validated previously [49] and also confirmed here by showing
the nuclear import of MyoD
ER
protein in response to TMX treatment (Fig 8E). Notably, acti-
vation of MyoD
ER
robustly induced Dll1 transcription (Fig 8F). Such an effect was completely
abolished when CHX was administered together with TMX (Fig 8F). Therefore, additional fac-
tor(s), which can be induced by MyoD, is required by MyoD to activate Dll1 transcription.
This result is in contrast with the report that MyoD is self-sufficient to induce Dll1 expression
during Xenopus embryogenesis [31]. As such, mammals may have evolved an extra layer of
regulatory mechanism to fine-tune Dll1 expression during myogenesis.
Fig 8. Sufficiency test of Dll1 inductions by MyoD in fibroblasts. (A) Fluorescence images of cell cytosol dye CMFDA (green) to
highlight fibroblast syncytia induced by MyoD or MyoG. Scale bar, 100 μm. (B,C) qPCR results of MyoD,MyoG (B), and Dll1 in
mouse fibroblasts with forced expression of MyoD, MyoG or both. n= 3. (D) Schematic of experimental design to test the
sufficiency of MyoD transcriptional activity in activating Dll1 expression in fibroblasts. (E) MyoD immunostaining of fibroblasts.
TMX: tamoxifen. CHX: cycloheximide. Scale bar, 25 μm. (F) qPCR results of Dll1 expression in fibroblasts after treatment
illustrated in D. Note that induction of Dll1 expression by MyoD was abolished upon CHX treatment (5 hours). n= 3. Data are
means ±SEM. P<0.05, P<0.01, P<0.001. ns, not significant.
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Perturbations of cis-regulatory motifs on Dll1 promoter affect its
expression in vivo and in vitro
As a bHLH transcriptional factor, MyoD activates the expression of its target genes through
binding to E-box motifs (CANNTG) [62]. Using FIMO, a motif discovery tool that empirically
predicts transcriptional factor binding sites [65], we discovered three highly conserved MyoD-
binding motifs in the intron 4 of Dll1 gene (Fig 9A). Of note, analysis of an ENCODE ChIP-
seq dataset [66] also discovered a MyoD-binding peak in this region during C2C12 myoblast
differentiation (Fig 9B).
We then utilized a CRISPR interreference tool to interrogate gene regulatory mechanism as
previously reported [49,67–69]. For this experiment, a catalytically dead Cas9 (dCas9) was
guided to these E-box motifs to dissect their function underlying the MyoD–Dll1 regulatory
axis (Fig 9C). We hypothesized that MyoD can bind to the intronic E-box motifs to induce
Dll1 expression during myogenic differentiation (Fig 9C, state a); when recruited by gRNA,
Fig 9. Regulation of Dll1 expression by MyoD through intronic E-box elements in human and mouse cells. (A)
Bioinformatic predictions of MyoD binding motifs in the Dll1 intronic regions from distantly related mammalian
species. (B) ENCODE ChIP-seq results of MyoD from mouse myoblasts. Green box highlights the mouse sequence
displayed in A. (C) Schematic of experiment design and rationale to probe cis-regulatory elements by dCas9 mediated
interference. CRISPRi: CRIPSR interference; (D) qPCR results of human WT myoblasts with expression of dCas9 and
gRNA indicated. n= 3. (E) Sequencing result of mouse Dll1
Δ232
mutant allele generated by CRISPR/Cas9 mediated
gene targeting. (F) H&E staining results of cross-sections from control and injured tibialis anterior muscles. Arrows
point to regenerated myofibers with centronuclei. Scale bar, 200 μm. (G) qPCR results that measured Dll1 expression
in mouse fibroblasts that infected by control or MyoD-expressing retroviruses. n= 4. (H) Schematic summary of
transcriptional regulations and ligand-receptor interactions between Notch signal sending or receiving cells. Data are
means ±SEM. P<0.05, P<0.01, P<0.001.
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the positioning of dCas9 to these E-box motifs should block MyoD binding thus repressing the
expression of Dll1 (Fig 9C, state b). Consistent with our hypothesis, expression of gRNA that
recruits dCas9 to each of these three E-box motifs significantly inhibited Dll1 expression by an
average of 65% in human myoblasts (Fig 9D). No additive effect was observed when these E-
boxes were simultaneously blocked (Fig 9D). Therefore, MyoD can activate Dll1 expression in
human myoblasts at least partially through binding to the evolutionarily conserved E-
box motifs located in the intron 4 of Dll1 gene.
The CRISPR interference results prompted us to test the function of these cis-regulatory ele-
ments during myogenesis in vivo. Using CRISPR genome editing, we generated a novel mouse
model that deleted these E box motifs from intron 4 of Dll1 gene. Sequencing confirmed the
removal of a 232 bp region centered on these conserved E-box elements (Figs 9E and S5A).
We named this allele as Dll1
Δ232
. Germline transmission of Dll1
Δ232
allele was confirmed after
breeding the founder with WT mice. Intercrosses of Dll1
Δ232/+
mice produced homozygous
mutants at expected Mendelian ratios (S5B Fig). Dll1
Δ232/Δ232
mutants were overly normal and
fertile, which are in stark contrast to the premature lethality that was observed for Dll1 null
mutants [37].
Given Dll1 expression was strongly induced by muscle injury (Fig 1E), we first tested
whether deletion of the intronic E-box elements can affect Dll1 expression and muscle regen-
eration. Surprisingly, in contrast to major defect of muscle regeneration in Dll1 gene knockout
model [70], normal progressions of muscle regeneration at day–4, –7 and –28 post injury were
observed for Dll1
Δ232/Δ232
mutant mice (Figs 9F and S5C and S5D). To better interpret this
result, we examined expression of Dll1. Fibroblasts were isolated from control and mutant
mice followed by overexpression of MyoD. Interestingly, the Dll1 expression in mutant fibro-
blasts showed a 71% decrease compared with control group (Fig 9G). This effect is similar
with that achieved by CRISPR interference (65% lower) in human myoblasts (Fig 9D). In
reviewing the ChIP-seq data [66], we did not observe additional MyoD binding peak in other
introns or intergenic regions. Lastly, a recent study using luciferase assay also identified this
intronic region as MyoD binding site [70]. Therefore, we reasoned that Dll1
Δ232/Δ232
mutant
cells may have used alternative E-box motifs from distal enhancer of an unknown region to
compensate the knockout or CRISPR-interference effects in vivo.
Discussion
In summary, our study uncovered the crucial regulations of Dll1 expression and Notch activa-
tion by myogenic factor MyoD. This intercellular regulatory loop (Fig 9H) involves three
major parts that could be initiated by asymmetric divisions of muscle precursor cells [71–73]
that allows one daughter cell to inherit a higher level of MyoD protein than the other. First, the
MyoD
high
myoblast can promptly turn on the expression of Dll1 which transactivates Notch in
adjacent MyoD
low
cell. Second, activation of Notch in MyoD
low
cell will further downregulate
transcriptions of MyoD and Dll1 genes. Third, even in presence of any residual expression of
Dll1 from MyoD
low
cell, binding of these ligand with Notch receptor in MyoD
high
cells will
unlikely activate Notch in the latter due to cis-inhibition from the abundantly expressed Dll1.
Similarly, Notch signaling between MyoD
high
cells will also be inhibited. Collectively, the
polarities of Notch can be established that allow daughter cells to adapt opposite cell-fates, i.e.
differentiation or self-renewal. Specifically, prompt upregulation of Dll1 in differentiating cells
can refrain cell-autonomous Notch activity. Given the strong inhibitory effects of Notch on
myogenesis, such cis-inhibitory mechanism is a prerequisite for completion of myogenic dif-
ferentiation. On the other side, transactivation of Notch pathway may prevent precocious dif-
ferentiation and promote a quiescence state and the self-renewal of muscle stem cells for long-
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Human muscle differentiation is controlled by MyoD-Dll1 axis
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term muscle regeneration (Fig 9H). Together with previous studies [27,35], these results
enabled a more complete understanding of the feedback mechanism whereby Notch polarity
and the divergent cell-fates can be established during myogenesis.
Notch activity is sensitive to the molar ratios of ligand and receptor. The cis-inhibitory
effect of ligand on Notch was initially observed in the developmental processes of Drosophila
[53,74]. Reflecting the complexity of mammalian system, cis-interaction of receptor with
ligand was shown to either inhibit or activate Notch in different biological processes [26,75,
76]. Nonetheless, our study provides the direct evidence that Dll1 can attenuate Notch signal-
ing cell-autonomously to facilitate myogenic differentiation of myoblasts. Our structure and
function analysis of Dll1 revealed the key roles of its extracellular domain for the cis-inhibition.
This result is consistent with previous structural determination of the Notch ligand/receptor
protein complexes [54]. The cis interaction of ligand with receptor may either titrate receptor
and limit it from trans-interacting with ligand from sender cells [53] or the ligand in cis can
stabilize Notch receptor such that the proteolytic processing and activation of receptors are
interfered [53].
Another interesting question that can be explored is the role of Jag1 in this model. During
myogenic differentiation of mouse and human myoblasts, expression of Jag1 was also induced,
though not as strongly as Dll1. Of note, previous study showed that Jag1 is the least-efficient
ligand in activating Notch in myoblast [77]. Therefore, the upregulation of Jag1 expression
during myoblast differentiation may have smaller impact on the Notch activity in adjacent
cells, compared with that from Dll1.
In addition to the gene gain- and loss-of-function experiments, we also examined the cis-
regulatory elements underlying the control of Dll1 transcription through two complementary
approaches: CRISPR-interference in human and CRISPR-knockout in mouse. Results from
these experiments consistently revealed the crucial role of the intronic E-box motifs in driving
Dll1 expression. As such, the interference and knockout of these elements produced 65% and
71% reductions of Dll1 expression, respectively. However, this magnitude of change failed to
elicit any obvious phenotype in vivo during muscle development or regeneration. A few possi-
bilities may explain this result. First, downregulation of Dll1 may triggered upregulation of
other Notch ligands in muscle cells that compensated for the reduction of Dll1. Second, a
residual 29% of Dll1 expression in mutant myoblasts could be sufficient to elicit cis-inhibitory
and trans-activating effects such that the Notch activity and cell fate are not impacted. Never-
theless, the genetic mouse model that we generated could be applied to understand whether
these cis-regulatory sites can be utilized by other bHLH transcriptional factors, e.g., NeuroD
[78], during other developmental processes or diseases. In addition, future efforts are war-
ranted to comprehensively dissect the unknown cis-regulatory sequence whereby MyoD drives
Dll1 expression during myogenic differentiation. Lastly, the mysterious factor(s) at down-
stream of MyoD that cooperatively induces Dll1 expression also needs to be identified.
Although MyoG appeared as a tempting candidate, the observation that MyoD can robustly
induce Dll1 expression from MyoG
KO
cells indicates that other factor(s) must exist in regula-
tion of this process.
Materials and methods
Generation of Dll1
Δ232
mutant mice
All animal procedures were approved by the Institutional Animal Care and Use Committee
(IACUC) at the University of Georgia. Dll1
Δ232
mouse model was generated by oviduct elec-
troporation as previously reported [79]. Briefly, the copulated female mice were used for sur-
gery to exposes oviduct. CRISPR gene editing cocktails were freshly assembled and contained
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6μM Cas9 protein (IDT, 1081058, Lot # 0000405530), 30 μM gRNA (crRNA annealed with
tracrRNA, IDT, 1072534, Lot # 0000403961). This cocktail was delivered into oviduct through
microcapillary injection. Oviduct electroporation was performed using CUY21EDIT II elec-
troporator with the following protocol: Pd A: 100 mA, Pd on: 5 ms, Pd off: 50 ms, three cycles,
decay 10%. Muscle injury was induced by injecting 1.2% barium chloride (50 μl) into tibialis
anterior muscle.
Mouse genotyping analysis
Dll1 genotyping PCR was performed using genomic DNA extracted from the toe clipping with
the primers, forward: AGAACCTCTGTTCGTGCCTG and reverse: GCGTCTAGGACAAA
AGGGCT. For Sanger sequencing, PCR products were first gel purified and cloned into pCRII
Topo vector (Thermo Fisher Scientific, K460001) and sequenced with T7 or SP6 primers.
Germline transmission of the Dll1
Δ232
allele was confirmed by genotyping F1 generation mice.
Cell cultures
Human myoblasts (ID: hSkMC-AB1190) were immortalized as previously described [48].
Human myoblasts were cultured in Skeletal Muscle Cell Growth Medium (PromoCell, C-
23060). Mouse 10T1/2 fibroblasts (ATCC, CCL-226) and C2C12 myoblasts (ATCC, CRL-
1772) were maintained in 10% FBS with 1% penicillin/streptomycin (Gibco, 15140122) in
DMEM (Dulbecco’s Modified Eagle’s Medium-high glucose, D5796). Primarily myoblasts
were isolated by enzyme digestion following the protocol [80]. Human and mouse myoblast
differentiation medium contains 2% horse serum supplemented in DMEM with 1% antibiotics
penicillin/streptomycin. Cells were tested mycoplasma negative using Universal Mycoplasma
Detection Kit (ATCC, 30-1012K).
Lentivirus preparation and CRISPR-Cas9 knockout experiments in vitro
Lenti-CRISPR v2 vector [81] used for gene knockout experiments in vitro was a gift from Feng
Zhang (Addgene plasmid # 52961). The guide RNAs that target the coding regions of human
and mouse MyoD and MyoG genes were individually cloned into the Lenti-CRISPR v2 vector
and sequenced to verify the correct insert. The sequences for gRNA used in this study are
provided.
Mouse Dll1 intron 4 gRNA1: GCACGTAGCCTTCTACCAAT
Mouse Dll1 intron 4 gRNA2: GGGGCTTTGTGCATAGAATT
Mouse MyoD gene: GTCAAGTCTATGTCCCGGAG
Mouse MyoG gene gRNA1: ACACCTTACATGCCCACGGC
Mouse MyoG gene gRNA2: CCACACTGAGGGAGAAGCGC
Human MyoD gene: CGTCGAGCAATCCAAACCAG
Human MyoG gene gRNA1: ACCACCAGGCTACGAGCGGA
Human MyoG gene gRNA2: CCACACTGAGGGAGAAGCGC
Human DLL1 gene gRNA1: ACCCAGAGGCACCTGACGGT
Human DLL1 gene gRNA2: TGACGAACACTACTACGGAG
Lentivirus was produced by transfecting Lenti-X 293T cells (Clontech, 632180) using
FuGENE6 transfection reagent (Promega, #E2692) with pLenti-V2, psPAX2 and pMD2.G
plasmids. 48 hours post transfection, lentivirus was collected to infect human or myoblast
myoblasts. psPAX2 vector was a gift from Didier Trono (Addgene plasmid # 12260). pMD2.G
vector was a gift from Didier Trono (Addgene plasmid # 12259).
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Retroviral vector preparations and expression
Retroviral expression vector pMXs-Puro (Cell Biolabs, # RTV-012) was used for gene cloning
and expression in human and mouse myoblasts. The open reading frames for red fluorescent
protein Cherry and mouse Dll1 were cloned into pMXs-Puro vector by In-Fusion cloning.
The identities of the DNA inserts in the plasmids were Sanger sequencing verified. MyoD-
pCLBabe plasmid [82] was a gift from Stephen Tapscott (Addgene plasmid # 20917). Myo-
blasts were labelled by retroviral infection that delivers the pMXs-Cherry expression vector.
MyoDER expression and chemical treatments were performed as previously reported [49].
Differentiation index and fusion index measurements
Differentiation index was calculated as the percentage of the nuclei number within muscle
cells (MF20+) divided by the total nuclei number in the imaging area. Fusion index was calcu-
lated as the nuclei number in myotubes (3 nuclei) as a percentage of the total number of
nuclei inside muscle cells.
RNA extraction and gene expression analysis
RNA was extracted by using TRIzol Reagent (Thermo Fisher Scientific). Before used for
reverse transcription, RNA quality and concentration were assessed by using a spectropho-
tometer (Nanodrop One, Thermo Fisher Scientific). cDNA was synthesized by reverse tran-
scription using random oligos with M-MLV reverse transcriptase (Invitrogen, 28025013). To
analyze gene expression changes, the Real-time PCR was performed using QuantStudio 3
Real-Time PCR System (Thermo Fisher Scientific) with SYBR Green Master Mix (Roche) and
qPCR primers. The 2
ΔΔCt
method was used to analyze gene expression after normalization to
expression of Gapdh or 18S rRNA. Primer sequences used in this study are provided.
Primers for DLL1 qPCR-F: GATTCTCCTGATGACCTCGCA
Primers for DLL1 qPCR-R: TCCGTAGTAGTGTTCGTCACA
Primers for JAG1 qPCR-F: GTCCATGCAGAACGTGAACG
Primers for JAG1 qPCR-R: GCGGGACTGATACTCCTTGA
Primers for DLL3 qPCR-F: CACTCCCGGATGCACTCAAC
Primers for DLL3 qPCR-R: GATTCCAATCTACGGACGAGC
Primers for DLL4 qPCR-F: GTCTCCACGCCGGTATTGG
Primers for DLL4 qPCR-R: CAGGTGAAATTGAAGGGCAGT
Primers for JAG2 qPCR-F: TGGGCGGCAACTCCTTCTA
Primers for JAG2 qPCR-R: GCCTCCACGATGAGGGTAAA
Primers for HEY1 qPCR-F: AACTGTTGGTGGCCCTGAAT
Primers for HEY1 qPCR-R: CAATTGACCACTCGCACACC
Primers for HEYL qPCR-F: ATGAGTCCTGGGAGAGACCC
Primers for HEYL qPCR-R: GCCAGTCAGTCATTGCTCCT
Primers for HES1 qPCR-F: TTTTTGGCGGCTTCCAAGTG
Primers for HES1 qPCR-F: GGTGGGCTAGGGACTTTACG
Primers for MYOD1 qPCR-F: CGACGGCATGATGGACTACA
Primers for MYOD1 qPCR-R: TATATCGGGTTGGGGTTCGC
Primers for MYOG qPCR-F1: GCCAACCCAGGGGATCAT
Primers for MYOG qPCR-R1: CCCGGCTTGGAAGACAATCT
Primers for MYH3 qPCR-F: ATTGCTTCGTGGTGGACTCAA
Primers for MYH3 qPCR-R: GGCCATGTCTTCGATCCTGTC
Primers for GAPDH qPCR-F: CACCAGGTGGTCTCCTCTGA
Primers for GAPDH qPCR-R: CAAGGGGTCTACATGGCAACT
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Primers for 18S qPCR-F: GTAACCCGTTGAACCCCATT
Primers for 18S qPCR-R: CCATCCAATCGGTAGTAGCG
Primers for Dll1 qPCR-F: CAGGACCTTCTTTCGCGTATG
Primers for Dll1 qPCR-R: AAGGGGAATCGGATGGGGTT
Primers for Hey1 qPCR-F: GCACGCCACTATGCTCAATG
Primers for Hey1 qPCR-R: GGGGACCTAGACTACCAGCA
Primers for Heyl qPCR-F: CCACTGGCGCAGATGAGTTA
Primers for Heyl qPCR-R: ATCCTGTTGGCTTGGGATGG
Primers for Hes1 qPCR-F: TTTTTGGCGGCTTCCAAGTG
Primers for Hes1 qPCR-R: GGTGGGCTAGGGACTTTACG
Primers for Gapdh qPCR-F: TCTCCTGCGACTTCAACAGC
Primers for Gapdh qPCR-R: AGTTGGGATAGGGCCTCTCTT
Primers for 18s qPCR-F: ACCGCAGCTAGGAATAATGGA
Primers for 18s qPCR-R: GCCTCAGTTCCGAAAACCA
Western blotting
Protein lysate was prepared by using RIPA buffer (Sigma, R0278) supplemented with complete
protease inhibitor (Sigma). Lysates were centrifuged at 16,000 x g for 15 minutes. The resultant
supernatant was mixed with 4x Laemmli sample buffer (Bio-Rad, #161–0747). Total 20 μg pro-
tein lysates were separated by SDS-PAGE gel electrophoresis. The proteins were transferred to
a polyvinylidene fluoride membrane and blocked in 5% milk at room temperature, and incu-
bated with following primary antibodies diluted in 5% milk overnight at 4˚C. Gapdh (Santa
Cruz Biotechnology, sc-32233), α-Tubulin (Santa Cruz Biotechnology, sc-8035), myosin
(DSHB, MF20), MyoD (Santa Cruz Biotechnology, SC-304), MyoG (DSHB, F5D). The HRP-
conjugated secondary antibodies: Donkey anti-sheep IgG-HRP (Santa Cruz Biotechnology, sc-
2473), Goat Anti-Mouse IgG (H+L)-HRP Conjugate (Bio-Rad, 170–6516) and Goat Anti-Rab-
bit IgG (H + L)-HRP Conjugate (Bio-Rad, 170–6515) were diluted at 1:5,000. Signal detection
was performed using Western Blotting Luminol Reagent (Santa Cruz Biotechnology, sc2048).
CRISPR interreference assays
Lenti-SAM v2 was a gift from Adam Karpf (Addgene plasmid # 92062). Lenti-SAM v2 plasmid
was used for gRNA cloning after the removal of VP64 expression cassette. The gRNA
sequences that target the control and E-box motif regions of human DLL1 intronic region
were provide below.
Human DLL1 intron 4 gRNA1: CTGCCCCAGCGCAACAATGC
Human DLL1 intron 4 gRNA2: ATGGAGCAGCTGTCCTGCCC
Human DLL1 intron 4 gRNA3: AGAGTGGACAGCTGGTATGC
Immunostaining
Immunostaining was performed as previously reported [49]. Briefly, cells were fixed in 4%
PFA for 10 minutes, membrane was permeabilized using 0.2% Triton X-100. Cells are blocked
with 3% BSA at room temperature. Primary antibody incubation was performed overnight at
4˚C. Immunostaining signal was detected by incubating with fluorescence conjugated second-
ary antibodies and Hoechst 33342 to visualize nucleus. Fluorescence images were collected
using the BioTek Microscope System or Olympus FV1200 Confocal Laser Scanning
Microscope.
PLOS GENETICS
Human muscle differentiation is controlled by MyoD-Dll1 axis
PLOS Genetics | https://doi.org/10.1371/journal.pgen.1009729 August 9, 2021 19 / 25
Quantification and statistical analysis
Experiments were repeated at least three times. All quantitative results were analyzed with stu-
dent’s ttest with a two-tail distribution. Comparisons with Pvalues <0.05 were considered
significant.
Supporting information
S1 Fig. Notch activation inhibits DLL1 expression in human cells through MyoD/MyoG.
(A–C) qPCR results of myogenic markers (A), DLL1 (B, C) in human myoblasts with retroviral
expression of NICD. Note that inhibitory effect of NICD on DLL1 expression was rescued by
MyoD or MyoG expression (C). Cells were differentiated for 72 hours. n= 3. Data are
means ±SEM. P<0.01, P<0.001. (D) MyoG and myosin immunostaining results of
human myoblasts after differentiation for 72 hours. Note that myogenic differentiation and
fusion defects of NICD can be rescued by co-expression of MyoD or MyoG. Scale bar, 100 μm.
(TIF)
S2 Fig. MyoD is essential for DLL1 expression in human myoblasts. (A) Human MyoD gene
structure and sequencing results that confirmed biallelic frame-shifts of MyoD ORFs in one
isolated MyoD
KO
clone similar to our previous report [49]. Arrow points to a 1bp insertion;
arrowhead points to a 1bp deletion. (B) Immunostaining results of MyoD (top) and myosin
(bottom) of human WT and MyoD
KO
myoblasts. MyoD staining confirmed the depletion of
MyoD proteins in MyoD
KO
cells. Scale bar, 100 μm. (C) Western blotting results that showed
the absence of MyoG and myosin expression in human MyoD
KO
myoblasts at 48 hours post
differentiation. (D,E) qPCR results of human WT and MyoD
KO
myoblasts with retroviral
expression of MyoD or MyoG. Cells were differentiated for 48 hours. n= 3. Data are
means ±SEM. P<0.05, P<0.01, P<0.001. ns, not significant.
(TIF)
S3 Fig. MyoG deletions from human myoblasts do not affect DLL1 expression. (A) Human
MyoG gene structure and an example of sequencing results similar to our previous report [49].
(B) Immunostaining result of MyoG and myosin to show the complete depletion of MyoG
proteins and relatively mild defect of differentiation for one clonally derived human MyoG
KO
myoblasts. Scale bar, 100 μm. (C) Quantifications of myoblast fusion. (D) qPCR result to show
that DLL1 expression was not significantly affected upon deletion of MyoG from human myo-
blasts. Data are means ±SEM. P<0.01, P<0.001. ns, not significant.
(TIF)
S4 Fig. Deletion of MyoD or MyoG affects Dll1 expression in mouse primary myoblasts.
(A) Schematic of experiment design. (B) Immunostaining results of mouse primary myoblasts
to show the expression levels of MyoD/MyoG before and after CRISPR treatments. Arrows
point to cells that show cytoplasmic staining signals which are likely non-specific signals. (C)
qPCR results of Dll1 in CRISPR treated mouse primary myoblasts. n = 3. Data are
means ±SEM. P<0.001.
(TIF)
S5 Fig. Characterizations of Dll1
Δ232 /Δ232
mouse. (A) Representative gel electrophoresis
result of genotyping PCR of WT and Dll1
Δ232/Δ232
mice. (B) Summary of Dll1
Δ232/Δ232
genotyp-
ing results. (C) Schematic of experiment design and the timeline of treatments and tissue col-
lections. (D) H&E staining results of cross-sections from control and injured tibialis anterior
muscles. Arrows point to small regenerating myocytes. Scale bar, 200 μm.
(TIF)
PLOS GENETICS
Human muscle differentiation is controlled by MyoD-Dll1 axis
PLOS Genetics | https://doi.org/10.1371/journal.pgen.1009729 August 9, 2021 20 / 25
Acknowledgments
We thank Gaurav Gopu, Alina Baiju, William Maley, Hector Romero-Soto, Emily Marie
Hicks and other trainees in the lab for technical assistance. We thank Myoline platform of the
Myology Institute for immortalized human cell lines and Dr. Shihuan Kuang from Purdue
University and Dr. Xiaochun Li from UTSW for sharing reagents.
Author Contributions
Conceptualization: Pengpeng Bi.
Formal analysis: Haifeng Zhang, Pengpeng Bi.
Funding acquisition: Pengpeng Bi.
Investigation: Haifeng Zhang, Renjie Shang.
Methodology: Haifeng Zhang, Renjie Shang, Pengpeng Bi.
Project administration: Haifeng Zhang.
Resources: Pengpeng Bi.
Supervision: Pengpeng Bi.
Validation: Haifeng Zhang, Renjie Shang.
Writing – original draft: Pengpeng Bi.
Writing – review & editing: Pengpeng Bi.
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