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Protein arginine methyltransferases in murine skull development

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Protein arginine methyltransferases in murine skull development

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Protein Arginine Methyltransferases
in Murine Skull Development

By

Nicha Ungvijanpunya

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CRANIOFACIAL BIOLOGY)

May 2021

Copyright 2021 Nicha Ungvijanpunya
ii
Dedication
This dissertation is dedicated to my beloved parents, Mr. Suwit and Mrs. Somsri
Ungvijanpunya, who always believe in me.

iii
Acknowledgements
I would like to thank my PhD mentors: Dr. Jian Xu for teaching me scientific knowledge,
encouraging me to persevere, being my role model as a female researcher and a mother, and
kindly supporting me in every way; Dr. Stephen Yen for trusting in me, motivating me to do
research, providing me guidance on my project and career path, and being supportive throughout
this PhD journey. I want to thank all of my PhD dissertation committee members Dr. Michael
Stallcup, Dr. Amy Merrill-Brugger, and Dr. Zhipeng Lu for their expertise, guidance, and helpful
suggestions. I also would like to thank Dr. Baruch Frenkel for his advice on this project. I want to
acknowledge Dr. Michael Paine for always being supportive and answering all my questions
during my MS and PhD training. I would like to acknowledge Anandamahidol Foundation
(Thailand) along with CBY program for providing me a scholarship and support.
I want to thank my colleagues and friends, Dr. Jiang Qian (Julia) and Prerna Sehgal for
always having my back, creating happy work environment and giving me helpful suggestions;
previous lab members Dr. Yongchao Gou, Dr. Abhijit Shinde, and Dr. Olan Jackson-Weaver
whose work has inspired and brought me to this project; Dr. Jian Qin (Abby) for her detailed
answers to all the problems I came across; Dr. Xi Chen for her kind help.
I would like to thank Dr. Yang Chai for his advice and guidance. In addition, I am really
grateful for all of his lab members for sharing their knowledge and helping me throughout this
project especially Dr. Yuan Yuan, Dr. Jifan Feng, Dr. Xia Han (Summer), Dr. Tingwei Guo, Dr.
Jinzhi He, Dr. Jiahui Du, Dr. Eva Janeckova, and Dr. Shuo Chen. I would like to thank members
of Dr. Merrill’s lab including previous member Dr. Diana Rigueur for her guidance in skeletal
preparation and skull morphology analysis, and current member Lauren Bobzin for advice in PF
suture. I want to thank Dr. Rucha Arun Bapat for her advice in examination and dissertation
preparation.
iv
I would like to thank our collaborator, Dr. Weiqun Peng for helping me with IRI analysis. I
want to acknowledge the Flow Cytometry Facility at the USC Stem Cell Center for helping me
with the cell sorting and the Molecular Imaging Center for performing microCT scanning of my
samples. I want to thank Thach-Vu Ho for his helpful advice in microCT scanning analysis and for
resolving all the lab technical issues I encountered. I would like to thank Janice Bea, Magdalena
Morales, Elsa Miranda, Gina Nieto, Linda Hattemer, Zhounan Liu, Jian-Bao Xie, all the staff and
faculty members of the Center for Craniofacial Molecular Biology at USC who make CCMB feel
like my second home and for always being supportive.
I want to thank all of my friends in the US and in Thailand who keep me sane during tough
times, cheer me up and believe in me.
Lastly, I want to thank my family especially my parents, who always trust in my decision,
guide me when I am lost, encourage me to pursue my dreams, and inspire me to do good things
for other people. I would not be able to come this far without their support.

v
Table of Contents
Dedication………………………………………………………………………………..……………….ii
Acknowledgements…………………………………………………………………………………....iii
List of Tables…………………………………………………………………………………................x
List of Figures…………………………………………………………………………………..............xi
Abbreviations………..………………………………………………………………………..............xvi
Abstract………..………………………………………………………………………......................xvii
Chapter 1 Introduction………………………………………………………………………………….1
1.1 Craniofacial development………………………………………………………………………….1
1.2 Development of Craniofacial Bones……………………………………………………………..4
1.3 Craniofacial Disorders……………………………………………………………………………..5
1.4 Post-Translational Modification and Craniofacial Development……………………………6
Chapter 2 The Role of PRMT4 in Murine Posterior Frontal Suture Closure……………………8
2.1 Background…………………………………………………………………………………………..8
2.1.1 Protein Arginine Methyltransferase 4 (PRMT4)…………………………………………8
2.1.2 Frontal bone and posterior frontal (PF) suture development…………………………12
2.1.3 Runx2……………………………………………………………………………………...19
2.1.4 Previous findings of PRMT4 and Runx2.………………............................................23
2.2 Hypothesis………………………………………………………………………………………….24
2.3 Materials and Methods……………………………………………………………………………25
2.3.1 Animals………………………………………………………………………………...….25
2.3.2 MicroCT scanning………………………………………………..…………………...….25
2.3.3 Whole-mount Skeletal Staining……………………………………………………...….25
2.3.4 Tissue Processing and Immunofluorescence (IF) Staining…………………………..26
2.3.5 RNA In Situ Hybridization (ISH) using RNAScope……………………………..……..26
vi
2.3.6 Proximity Ligation Assay (PLA).……………………………..………………………….27
2.3.7 TUNEL assay…………………………..…………………………………………..…….27
2.3.8 Ex vivo Calvaria Culture and DiI labeling…………………………..………….……….28
2.3.9 Cell Culture, siRNA, and Plasmid Transfection……………………………….……….29
2.3.10 Migration assay…………………………..………….………………………………….29
2.3.11 SDS-PAGE and Immunoblotting…………………………..………………………….30
2.3.12 Bulk RNA-sequencing Analysis…………………………..………….………………..31
2.3.13 Chromatin Immunoprecipitation – Quantitative PCR (ChIP-qPCR)………………..32
2.3.14 Statistical analysis…………………………..………….……………………………....32
2.4 Results……………………………………………………………………………………………....33
2.4.1 NCC-specific Prmt4 deletion leads to PF suture patency…………………………….33
2.4.2 PRMT4 is enriched at the osteogenic fronts…………………………………………...37
2.4.3 PRMT4 co-express with Runx2…………………………………………………………39
2.4.4 Methyl-Runx2 is reduced upon NCC-specific Prmt4 deletion at the osteogenic
fronts………………………………………………………………………………………..……40
2.4.5 NCC-specific Prmt4 deletion does not significantly alter cell proliferation or
apoptosis………………………………………………………………………………………...41
2.4.6 NCC-specific Prmt4 deletion impairs osteogenic front cells migration………………43
2.4.7 PRMT4 depletion reduces migration of pre-osteoblast MC3T3-E1 cells…………..46
2.4.8 Runx2 methylation by PRMT4 is important for cell migration………………………...47
2.4.9 NCC-specific Prmt4 deletion downregulates a group of ECM-related genes in PF
suture…………..………………………………………………………………………………..48
2.4.10 Depletion of PRMT4 decreases Runx2 binding with its target gene promoter in
calvarial preosteoblasts………………………………………………………………………..50
2.4.11 NCC-specific Prmt4 deletion leads to reduction of H3R17me2 and H3R26me2 in
PF suture………………………………………………………………………………………...52
vii
2.4.12 NCC-specific Prmt4 deletion leads to reduction of bone matrix expression without
changing osteogenic fate commitment or frontal bone thickness…………………………..52
2.5 Discussion
PRMT is a major protein arginine methyltransferase for Runx2……………………………58
The source of migratory cells at the OF lining the PF suture………………………………..58
The force that drives cell migration at the OF in the PF suture closure…………………….59
Regulatory role of the surrounding structures………………………………………………..59
Postnatal role of PRMT4 in PF suture development…………………………………………60
NCC-specific Prmt4 deletion does not alter cell proliferation……………………………….60
The impact of NCC-specific Prmt4 deletion on osteogenic differentiation…………………60
Runx2 methylation by PRMT4 specifically affects PF suture closure among NCC-derived
structures………………………………………………………………………………..………61
CNCC-specific Prmt4 deletion leads to downregulation of ECM-related genes…………..61
Depletion of PRMT4 decreased Runx2 binding with its target genes in calvarial
preosteoblasts…………………………………………………………………………..………62
2.6 Conclusions and Future Directions………..……………………………………………….….63
Chapter 3 The Role of PRMT1 in Murine Mandibular Development……………………………65
3.1 Background…………………………………………………………………………………………65
3.1.1 Mandibular development……………………………………………………..………….65
3.1.2 Alternative splicing……………………………………………………………………….67
3.1.3 Intron retention during development …………………………………………………...69
3.1.4 The importance of IR and NMD pathway in craniofacial development………………72
3.1.5 PRMT1 and craniofacial development…………………………………………………73
3.1.6 PRMT1 and mRNA splicing……………………………………………………………..75
3.2 Hypothesis……………………………………………………………………………………….…76

viii
3.3 Materials and Methods……………………………………………………………………………77
3.3.1 Animals…………………………………………………………………………………....77
3.3.2 Bulk RNA-sequencing analysis………………………………………………………....77
3.3.3 Intron Retention Analysis………………………………………………………………..78
3.3.4 qRT-PCR to validate RNA-sequencing results………………………………………..78
3.3.5 Cell Culture and siRNA transfection…………………………………………………....79
3.3.6 Statistical analysis………………………………………………………………………..80
3.4 Results………………………………………………………………………………………………80
3.4.1 PRMT1 regulates alternative splicing events in mouse mandible……………………80
3.4.2 PRMT1 represses intron retention on a genomic scale……………………………….81
3.4.3 Intron retention leads to changes in mRNA level………………………………………82
3.4.4 Downregulated genes which shows increased IRI are involved in extracellular
matrix…………………………………………………………………………………………….83
3.4.5 NCC-specific Prmt1 deletion leads to an increase in intron retention of ECM-related
genes in mandible……………………………………………………………………………....84
3.4.6 Increased intron retention may play a major role in gene repression via Nonsense-
Mediated Decay (NMD) pathway…………………………………………………………...…89
3.4.7 Depletion of splicing factors regulated by PRMT1 leads to increased intron retention
of Tnn in vitro…………………………………………………………………………………….92
3.5 Discussion………………………………………………………………………………………….96
NCC-specific Prmt1 deletion leads to an increase in intron retention of ECM-related genes
in mandible……………………………………………………………………………………....96
Increased intron retention may play a major role in gene repression via Nonsense-
Mediated Decay (NMD) pathway….…………………………………………………………..96
Depletion of splicing factors regulated by PRMT1 leads to increased intron retention of
Tnn in vitro……………………………………………………………………………………….97
ix
3.6 Conclusions and Future Directions…………………………………………………………….98
Bibliography…………………………………………………………………………………………….99

x
List of Tables
Table1.1. Derivatives of pharyngeal arches, including neural crest derived structures……………3
Table 2.1. Primer sequences of each target gene for ChIP-qPCR……………………………..…..51
Table 3.1. Number of PTC found in retained intron of 18 genes which showed downregulation of
transcript and increased IRI…………………………………………………………………………….90

xi
List of Figures
Figure1.1. Neural crest migration and its contribution to craniofacial structures…………………...2
Figure1.2. Pharyngeal arches with their components and derivatives ……………..……………….3
Figure1.3. Cranial vault bones and their origins…………………………………………………….....4
Figure1.4. Craniofacial bones and their origins ……………………………………………...………..4
Figure 2.1. Three types of PRMTs and their catalytic products……………………………………....9
Figure 2.2. PRMT4 expression is the highest among all PRMTs in three osteogenic cell lines –
mouse calvarial preosteoblasts (MC3T3-E1), mouse stromal mesenchymal cells (ST2), and
neonatal mouse calvarial osteoblasts (NeMCO)…………………………………….…………….....11
Figure 2.3. The expression of PRMTs in embryonic fibroblasts and embryonic stem cells………11
Figure 2.4. Human skull consists of neurocranium and viscerocranium …………………………..12
Figure 2.5. Human calvaria and mouse calvaria are anatomically and developmentally similar...13
Figure 2.6. Four main components of cranial sutures……………………...……………………..…13
Figure 2.7. Cranial sutures are different in tissue derivatives….…………………………………...16
Figure 2.8. H&E staining of cranial sutures from 6-day old mice showed that each cranial suture
has different structure……………………..…………………………………………………………....16
Figure 2.9. The location of PF suture and the coronal section of PF suture………………….……17
Figure 2.10. Runx2 regulates osteoblast differentiation…………………...……………………….20
Figure 2.11 Runx2 regulates chondrocyte differentiation………………..………………………....20
Figure 2.12. Alternative splicing of Runx2 and Runx2 domains…………………………………….21
Figure 2.13. Major characteristics of cleidocranial dysplasia patients……………………………..22
Figure 2.14. Runx2
+/-
mice phenocopied cleidocranial dysplasia patients…………...…………....23
Figure 2.15. Runx2 is methylated by PRMT3 and PRMT4………………………………..………...24
Figure 2.16. Organ culture dish with metal grid and membrane filter for ex vivo calvaria
culture………………………………………………………………………………………………….....29
xii
Figure 2.17 Whole-mount skeletal staining of E16.5 mice showed wider gap between frontal
bones in mutant mice. Other bony structures appeared unchanged……………………..………...34
Figure 2.18 MicroCT scanning of P0 mouse skull showed wider gap between frontal bones in
mutant than in control……………………………………………………………………………………34
Figure 2.19. MicroCT scanning of P0 mouse skull showed no apparent alteration in size or shape
of other CNC-derived craniofacial bones in mutant mice…………………………………………….35
Figure 2.20. Prmt4 mutant mouse had smaller head and proportionally smaller body size than
control mouse…………………………………………………………………………………….…...…36
Figure 2.21. At 6-week of age, PF suture was patent in Prmt4 mutant mice while it was completely
fused in control and Prmt4 heterozygous deletion mice………………………….……………….…36
Figure 2.22. MicroCT analysis of 6 weeks old mouse skull revealed that Prmt4 mutant mice had
shorter skull with alteration of PF suture structure…………..………………………………………..37
Figure 2.23. PRMT4 is enriched at the osteogenic fronts……………………………..…………….38
Figure 2.24. PRMT4 co-expresses with Runx2………………………………….……………….….40
Figure 2.25. Methyl-Runx2 is detected at the OFs and is reduced upon NCC-specific Prmt4
deletion……………………………………………………………………….…………………………..40
Figure 2.26. Cell proliferation at the OFs was not significantly altered with NCC-specific Prmt4
deletion at P0 and E16.5………………………………………………………………………………..42
Figure 2.27. Cell apoptosis at the OFs was not significantly altered with NCC-specific Prmt4
deletion at P0 and E16.5…………….……………………………………………………………….…43
Figure 2.28. Cells at the OF migrate during PF suture closure…………………….…………….....45
Figure 2.29. DiI-labeled mesenchymal cells did not migrate towards the OF in either control or
mutant……………………………………………………………………………………….……………46
Figure 2.30. Cell migration was impaired upon PRMT4 depletion………………………..……..…47
Figure 2.31. Runx2 methylation deficiency impaired cell migration……………………..……..…..48
xiii
Figure 2.32. Schematic drawing showed workflow of RNA-seq analysis from CNCC isolated from
OF…………………………………………………………………………………………………………49
Figure 2.33. GO Analysis revealed that downregulated genes in Prmt4 mutant were mostly
related with ECM. In addition, IPA identified that cell migration and cell invasion pathways were
significantly downregulated in Prmt4 mutant……………………………………………………….…49
Figure 2.34. Examples of downregulated ECM genes which were identified as Runx2 targets…50
Figure 2.35. Depletion of PRMT4 decreased Runx2 binding with its target ECM-related migration
genes in calvarial preosteoblasts………………………...………………………………………….…51
Figure 2.36. NCC-specific Prmt4 deletion leads to reduction of H3R17me2 and H3R26me2 at the
OF of PF suture……………………………………………………………………………………….…52
Figure 2.37. Bone matrix genes and Sp7 were downregulated in osteogenic fronts of Prmt4
mutant while osteogenic genes expression such as Alpl was not significantly reduced……….….54
Figure 2.38. Depletion of PRMT4 decreased Runx2 binding with its target genes………………..54
Figure 2.39. The number of Runx2 positive cells at the osteogenic fronts was not significantly
altered in Prmt4 mutant mice when compared with control mice at P0 and E16.5...………………55
Figure 2.40. The number of Sp7 positive cells at the osteogenic fronts was not significantly altered
in Prmt4 mutant mice when compared with control mice at P0 and E16.5…………...………….…56
Figure 2.41. NCC-specific Prmt4 deletion does not lead to reduction of frontal bone thickness at
P0 and E16.5………………………………………………………………………………….…………57
Figure 2.42. Summary of findings………………………………………….……………………….…64
Figure 3.1. Scheme of mandibular morphogenesis………………………………………………….66
Figure 3.2. Two types of splicing events – constitutive and alternative splicing……………..…....68
Figure 3.3. Fates of Intron Retention………………………………………………………………….70
Figure 3.4. Examples of proteins involved in IR-NMD pathway mentioned previously…………...73
Figure 3.5. NCC-specific Prmt1 deletion leads to multiple craniofacial defects…………………...74
xiv
Figure 3.6. RNA-seq analysis of Prmt1 mutant palates showed a downregulation of osteogenic
genes and an upregulation of chondrogenic genes……………………………..…………………...75
Figure 3.7. Increase in alternative splicing event upon PRMT1 depletion in MEC1 cells…..…….76
Figure 3.8. PRMT1-specific targets identified by Mass Spectrometry analysis………….………..76
Figure 3.9. A scheme shows how primers are designed………………….…………………….…..79
Figure 3.10. Increased alternative splicing events in NCC-specific Prmt1 deletion mandibles….81
Figure 3.11. PRMT1 represses intron retention on a genomic scale…………………….…………82
Figure 3.12. Intron retention leads to changes in mRNA level……………..…………………….....82
Figure 3.13. Most of downregulated genes (96.2%) showed increased IRI………………..…...…83
Figure 3.14A. Genes which showed increased IR transcripts and decreased spliced transcript in
Prmt1 mutant…………………………………………………...……………….....…………………….85
Figure 3.14B. Genes which showed increased IR transcripts and decreased spliced transcript in
Prmt1 mutant…………..………………………………………………………………………..……….86
Figure 3.14C. Genes which showed increased IR transcripts and decreased spliced transcript in
Prmt1 mutant…………..…….………………………………………………………..…………………86
Figure 3.15A. Genes which showed slightly increased IR transcripts and decreased spliced
transcripts in Prmt1 mutant…...………………………………………………………...………………87
Figure 3.15B. Genes which showed slightly increased IR transcripts and decreased spliced
transcripts in Prmt1 mutant….………………………………………………………………………….87
Figure 3.16A. Genes which showed decreased IR transcripts and decreased spliced transcripts
in Prmt1 mutant……………………………..…………………………………………………………...88
Figure 3.16B. Genes which showed decreased IR transcripts and decreased spliced transcripts
in Prmt1 mutant………..………………………………………………………………………………...88
Figure 3.17. Genes which showed increased IR transcripts and increased spliced transcripts in
Prmt1 mutant………………………………………………………………………………………….....89
Figure 3.18. qRT-PCR to confirm knockdown efficiency of siRNA…………………………………92
xv
Figure 3.19A. Fold change of Tnn intron-retaining transcripts upon splicing factor depletion in
ST2 cells………………………………………………………………………………………………….93
Figure 3.19B. Fold change of Tnn spliced transcripts upon splicing factor depletion in ST2
cells……………………………………………………………………………………………………….93
Figure 3.20A. Fold change of Matn2 intron-retaining transcripts upon splicing factor depletion in
ST2 cells………………………………………………………………………………………………….94
Figure 3.20B. Fold change of Matn2 spliced transcripts upon splicing factor depletion in ST2
cells……………………………………………………………………………………………………….94
Figure 3.21A. Fold change of Cdkn2c intron-retaining transcripts upon splicing factor depletion
in ST2 cells……………………………………………………………………………………………….95
Figure 3.21B. Fold change of Cdkn2c spliced transcripts upon splicing factor depletion in ST2
cells……………………………………………………………………………………………………….95

xvi
Abbreviations
CKO = Conditional Knockout
CNCC = Cranial Neural Crest Cell
CNC = Cranial Neural Crest
ChIP = Chromatin Immunoprecipitation
CT = Computed Tomography
DM = Dura Mater
ECM = Extracellular Matrix
GO = Gene Ontology
H3 = Histone 3
IF = Immunofluorescence
IR = Intron Retention
IRI = Intron Retention Index
NCC = Neural Crest Cell
NMD = Nonsense-Mediated Decay
OF = Osteogenic Front
PF suture = Posterior Frontal Suture
PRMT = Protein Arginine Methyltransferase
PTC = Premature Termination Codon
PTM = Post-Translational Modification
qPCR = Quantitative Polymerase Chain Reaction
SAG = Sagittal Suture
SM = Suture Mesenchyme
TUNEL = Terminal Deoxynucleotidyl Transferase Biotin-dUTP Nick End Labeling

xvii
Abstract
Cranial sutures separate the skull bones and accommodate for brain and skull growth
during embryonic and postnatal development. Genetic mutation has been associated with
premature closure or patency of cranial sutures. In cleidocranial dysplasia patients with RUNX2
mutation, open metopic suture is one of the major characteristics. Runx2 is a master regulator of
osteogenic differentiation and bone formation. Its activity is regulated by multiple types of post-
translational modifications, including phosphorylation, acetylation and glycosylation. Previous
findings identified that Protein Arginine Methyltransferase 4 (PRMT4/CARM1) methylated Runx2
at four specific arginine residues. In this study, I have uncovered their co-expression in
osteoprogenitors at the osteogenic fronts of posterior frontal (PF) suture during embryonic
development. Neural crest-specific Prmt4 deletion in mice caused patent posterior frontal
(metopic) suture in adult and a wider gap between front bones during embryonic stages. I further
demonstrated that Prmt4 deficiency delayed osteoprogenitor migration at the osteogenic fronts
without disrupting cell proliferation or apoptosis. In addition, PRMT4 depletion and methylation-
deficient Runx2 both impaired migration ability of mouse calvarial preosteoblasts (MC3T3-E1)
cells, suggesting that PRMT4-catalyzed Runx2 methylation regulates migration in fate-committed
osteoblasts. I then determined the transcriptional landscape in osteogenic fronts and revealed a
significant downregulation of ECM-related genes with enrichment in the cell migration pathway.
Runx2 enrichment at migration gene promoters was significantly reduced upon PRMT4 depletion,
indicating that PRMT4-Runx2 signaling directly control the transcription of these migration genes.
We also noted downregulation in a fraction of bone matrix genes and reduced enrichment of
Runx2 at Sp7 promoter. However, the number of Runx2-positive or Sp7-positive cells at
osteogenic fronts or the frontal bone thickness did not decline. Altogether, these results
demonstrate that PRMT4 methylates Runx2 and this arginine methylation at osteogenic fronts is
essential for the migration of osteoprogenitors during posterior frontal suture closure.
1
Chapter 1 Introduction
Craniofacial development is a complicated biological process which involves a great
number of cell types, cellular processes and signaling pathways. Disruption in any component of
this sophisticated developmental process can lead to craniofacial abnormalities (Chai, 2015).
Visible complications usually involve irregularities of affected structures, alteration in appearance,
and inability to function properly. Moreover, financial burden, psychological issues, and treatment
risks are unavoidable difficulties for patients and their families (Strauss & Cassell, 2009; Visram
et al, 2019). Therefore, comprehensive knowledge of how these complex structures develop is
required to help us find alternative treatment options or eventually, a prevention.
Humans are born with a limited number of genes. However, with post-transcriptional
modifications such as alternative splicing, the complexity of transcripts is substantially increased
(Ben-Dov et al, 2008; Blencowe, 2006; Pan et al, 2008). Post-translational modifications (PTM)
add another layer of complexity which expands the proteome boundary (Mann & Jensen, 2003;
Venne et al, 2014).
Protein arginine methyltransferases (PRMTs) are a group of enzymes required for arginine
methylation which is one of the major methylation events and is essential for several
developmental processes (Bedford & Richard, 2005; Guccione & Richard, 2019). PRMTs were
reported to be therapeutic targets since their functions can be modified by inhibitors, making them
valuable molecules to study during craniofacial development (Cha & Jho, 2012). The aim of this
dissertation is to study the role of PRMTs during craniofacial bone development.

1.1 Craniofacial Development
The craniofacial region is relatively small when compared to the total human body size.
However, a complete understanding of this process has not been established yet due to the
complexity of its developmental process. The sculpting of craniofacial structures starts when
neural plate folds and neural crest cells derived from ectoderm at the dorsal aspect acquire motility
2
through epithelial-mesenchymal transition. Upon neural tube closure, neural crest cells migrate
ventrally to several destinations (Figure 1.1A). These neural crest cells are pluripotent, give rise
to several cell types and contribute to multiple structures in our body (Selleck & Bronner-Fraser,
1995). There are 5 pharyngeal arches and 4 pharyngeal clefts. Each arch consists of ectoderm,
mesoderm, endoderm, and neural crest – derived structures (Figure 1.1B and 1.2). Cranial neural
crest cells (CNCC) migrate to pharyngeal arches to contribute to the craniofacial region (Gilbert,
2000) (Table 1.1).

Figure1.1. Neural crest migration (A) and its contribution to craniofacial structures (B) (Dubey & Saint-
Jeannet, 2017). NP = neural plate; PM = paraxial mesoderm; NO = notochord; SO = somite; NE = non-
neural ectoderm; NT = neural tube; EP = epidermis; MB = midbrain; HB = hindbrain; PA1-3 = pharyngeal
arch 1-3; FR = frontal bone; PA = parietal bone; AS = alisphenoid bone; SQ = squamosal bone; NA = nasal
bone; ZY = zygomatic bone; MX = maxillary bone; MD = mandible; HY = hyoid bone

3
Figure1.2. Pharyngeal arches with their components and derivatives (Graham & Richardson, 2012).

Pharyngeal
arch
Skeletal elements (neural
crest plus mesoderm)
Arches, arteries
(mesoderm)
Muscles (mesoderm) Cranial nerves
(neural tube)
1 Incus and malleus (from
neural crest); mandible,
maxilla, and temporal bone
regions (from crest dermal
mesenchyme)
Maxillary branch of
the carotid artery (to
the ear, nose, and
jaw)
Jaw muscles; floor of
mouth; muscles of the
ear and soft palate
Maxillary and
mandibular divisions of
trigeminal nerve (V)
2 Stapes bone of the middle
ear; styloid process of
temporal bone; part of hyoid
bone of neck (all from neural
crest cartilage)
Arteries to the ear
region: cortico-
tympanic artery
(adult); stapedial
artery (embryo)
Muscles of facial
expression; jaw and
upper neck muscles
Facial nerve (VII)
3 Lower rim and greater horns
of hyoid bone (from neural
crest)
Common carotid
artery; root of internal
carotid
Stylopharyngeus (to
elevate the pharynx)
Glossopharyngeal
nerve (IX)
4 Laryngeal cartilages (from
lateral plate mesoderm)
Arch of aorta; right
subclavian artery;
original spouts of
pulmonary arteries
Constrictors of
pharynx and vocal
cords
Superior laryngeal
branch of vagus nerve
(X)
6 Laryngeal cartilages (from
lateral plate mesoderm)
Ductus arteriosus;
roots of definitive
pulmonary arteries
Intrinsic muscles of
larynx
Recurrent laryngeal
branch of vagus nerve
(X)

Table1.1. Derivatives of pharyngeal arches, including neural crest derived structures (Gilbert, 2000)

4
1.2 Development of Craniofacial Bones
The skull is formed by bones derived from mesoderm and cranial neural crest cells (Chai
et al, 2000; Jiang et al, 2002) (Figure 1.3 and 1.4). Cranial vault bones are further sandwiched
between the periosteum and the dura mater during developmental process and these surrounding
structures modulate bone integrity (Jin et al, 2016; Slater et al, 2008). Due to different origins,
locations, and surrounding structures, signaling pathways which regulate craniofacial bone
development are context-dependent (Lenton et al, 2005; Levine et al, 1998). Knowledge of how
one bone develops cannot be simply applied to others.

Figure1.3. Cranial vault bones and their origins. Blue structures including dura mater (not shown) are neural
crest – derived. Red structures are mesoderm – derived (Lenton et al, 2005).

Figure1.4. Craniofacial bones and their origins (Chai & Maxson, 2006).

5
1.3 Craniofacial Disorders
Craniofacial anomalies are one of the most common birth defects occurring in
approximately 2-3% of babies. These abnormalities are the main cause of early mortality (Mossey
& Catilla, 2001 ). The etiologies of most craniofacial disorders are not completely understood
despite extensive research efforts due to complexities involving genetic, epigenetic, and
environmental factors.
Craniosynostosis is the premature fusion of single or multiple cranial sutures. It is one of
the most common craniofacial anomalies with a prevalence of 1:2,500. This abnormality leads to
several complications such as increased intracranial pressure, visual impairment, respiratory
problem. Treatment is usually complicated and intensive requiring surgical procedures which can
be risky to patients (Wilkie, 1997). Previous studies have revealed a list of genes that are involved
in this anomaly such as FGFR1-3, TWIST, MSX2, FBN1, EFNB1, EFNA4 (Bellus et al, 1996;
Howard et al, 1997; Jabs et al, 1993; Merrill et al, 2006; Sood et al, 1996; Twigg et al, 2004). Each
suture seems to possess distinct mechanisms. Sagittal suture is the most affected suture
accounting for 40-55% of all cases. Coronal suture is the second most affected suture with
prevalence of 20-25%. Metopic suture affects about 5-15% of all cases while lambdoid suture is
the least affected at a prevalence of less than 5% (Cohen & MacLean, 2000; Morriss-Kay & Wilkie,
2005). However, a more recent study has shown an increased prevalence of craniosynostosis to
19% in metopic suture of all cases ranking it as the second most affected sutures (Boulet et al,
2008).
Mandibular hypoplasia refers to a small mandible, which could be congenital or acquired
(Singh & Bartlett, 2005). The severity varies which leads to a broad spectrum of consequences,
ranging from fatality caused by airway obstruction to functional difficulty caused by malocclusion.
Mandibular hypoplasia could be involved with syndromes such as Pierre Robin Sequence which
causes serious respiratory problem to the newborn (Gangopadhyay et al, 2012). Early surgery to
allow airway opening is usually required in severe cases. Compared with other craniofacial
6
structures, the molecular regulation of mandible morphogenesis is less studied. Therefore, it is
essential to investigate into this structure.

1.4 Post-Translational Modification and Craniofacial Development
Transcription is regulated by cis-regulatory factors which is DNA sequence itself and trans-
regulatory factors including transcription factors that recognize specific DNA sequences, and
histone proteins and epigenetic modifiers that coordinate transcriptional activity. Cis and trans
acting elements influence each other and cooperate to determine transcriptional outcome
(Griffiths et al, 2000; Margueron & Reinberg, 2010). Post-translational modifications on trans-
regulatory proteins add another level of complexity to their roles in regulating gene expression
(Murr, 2010).
Post-translational modifications are reversible or irreversible processes that alter the
capacity of protein function. Examples of these modifications include phosphorylation, acetylation,
methylation, glycosylation, disulfide bond formation, hydroxylation, ubiquitination, lipidation, and
SUMOylation (Walsh et al, 2005). These PTMs are found to be important during developmental
processes including craniofacial development. For example, several histone marks were found to
be differentially expressed and involved in regulating multiple DNA binding factors in craniofacial
tissues (Wilderman et al, 2018). The importance of studying roles of PTMs during development
is essential and interesting because some PTMs are reversible and could be potential therapeutic
targets.
Protein methylation includes histone methylation and non-histone protein methylation.
Methylation of histone proteins is usually involved in chromatin remodeling which then can either
upregulate or downregulate gene transcription. For example, H3K27me3 is a histone mark of
transcriptional repression while H3K4me3 is associated with active transcription (Zhang &
Reinberg, 2001). Histone methylation regulates multiple processes throughout the development
such as embryogenesis, body patterning, cell lineage determination, and organogenesis
7
(Jambhekar et al, 2019). Non-histone protein methylation is reported more recently. A great
number of RNA binding proteins, DNA binding proteins including transcription factors, and cell
surface receptors were reported to be methylated by PRMTs. These various substrates lead to
regulation in multiple processes (Wei et al, 2014).
There are several studies about essential roles of protein methylation or expression of
methyltransferases in palatal development. However, compared to orofacial cleft, how protein
methylation regulates craniofacial bone formation is less understood (Alvizi et al, 2017; Dong et
al, 2019; Seelan et al, 2013; Seelan et al, 2019).

8
Chapter 2
The Role of PRMT4 in Murine Posterior Frontal Suture Closure
2.1 Background
2.1.1 Protein Arginine Methyltransferase 4 (PRMT4)
Methylation is generated by a large group of enzymes called methyltransferases. One of
the most widely studied methyltransferase subgroups is the family of Protein Arginine
Methyltransferase (PRMTs). PRMTs add methyl group(s) from the methyl donor S-adenosyl-L-
methionine (SAM) to the nitrogen atom of arginine residues of substrates (Gary & Clarke, 1998;
Zhang & Reinberg, 2001). PRMTs can be divided to 3 groups which are type I, II, and III
depending on the products they generate. Type I PRMTs, which are PRMT1, PRMT2, PRMT3,
PRMT4, PRMT6, and PRMT8, generate monomethylarginine (MMA) and asymmetric
dimethylarginine (ADMA). Type II PRMTs, which are PRMT5 and PRMT9, generate MMA and
symmetric dimethylarginine (SDMA). PRMT7 is the only Type III PRMT which generates only
MMA (Figure 2.1). This modification is important for several cellular processes (Blanc & Richard,
2017; Morales et al, 2016). The roles of PRMTs are diverse and important during development
and physiological maintenance. Some examples of reported cellular roles of PRMTs are involved
in RNA processing, transcriptional regulation, mRNA splicing, protein synthesis, response to DNA
damage, and signal transduction via growth factors (Bedford & Richard, 2005; Guccione &
Richard, 2019). In addition, dysregulation of PRMTs was reported to be associated with diseases
such as cancer and inflammatory conditions (Bedford & Clarke, 2009). Several inhibitors were
discovered and proved effective mostly in cancer treatment (Guccione & Richard, 2019).
9

Figure 2.1. Three types of PRMTs and their catalytic products (Morales et al, 2016).

Protein arginine methyltransferase 4 (PRMT4) or coactivator-associated arginine
methyltransferase 1 (CARM1) was first discovered by Dr. Stallcup’s group as a PRMT that
regulates transcription by association with coactivators. Despite its homology with PRMT1,
PRMT4 exhibits different substrate preference and efficiently methylates Histone 3 (H3), whereas
PRMT1 methylates Histone 4 (H4) (Chen et al, 1999). This finding led to further exploration of
PRMT4 function in transcriptional regulation and identification of H3R17 and R26 as PRMT4-
targeted residues (Schurter et al, 2001). Methylation of H3 by PRMT4 was further validated in
vivo for transcriptional activation by recruiting transcription elongation - associated PAF1 complex
(PAF1c) and discharging corepressor from chromatin (Ma et al, 2001; Wu et al, 2012; Wu & Xu,
2012). More PRMT4 substrates have been identified in cellular processes including mRNA
splicing (Cheng et al, 2007; Ohkura et al, 2005), nonsense-mediated decay (Sanchez et al, 2016),
autophagy (Shin et al, 2016; Yu et al, 2020), and DNA damage response (Lee et al, 2011).
The role of PRMT4 in development was first demonstrated by Yadav N. et al, which
showed that global Prmt4 knock-out mice are smaller than wild-type and die soon after birth while
Prmt4 heterozygous deletion mice survive with some lethality (Yadav et al, 2003). Prmt4-null mice
fail to breathe and show reduced alveolar air spaces due to increased pulmonary cell proliferation
and impaired alveolar cell differentiation which lead to a fatality after birth (O'Brien et al, 2010). In
addition, numerous studies showed essential roles of PRMT4 during development in early
10
embryonic cell fate (Hupalowska et al, 2018; Panamarova et al, 2016; Parfitt & Zernicka-Goetz,
2010; Torres-Padilla et al, 2007), hematopoiesis (Li et al, 2013) and differentiation of thymocyte
(Kim et al, 2004), adipocyte (Yadav et al, 2008), skeletal muscle (Dacwag et al, 2009), and neuron
(Fujiwara et al, 2006; Selvi et al, 2015). Furthermore, PRMT4 enzymatic-deficient knock-in mice
show phenotypes similar to Prmt4-null mice suggesting a requirement of PRMT4 enzymatic
activity during development (Kim et al, 2010a). In addition to aforementioned roles during normal
development, overexpression of PRMT4 is involved in breast, ovarian, prostate, liver, colorectal,
hematopoietic and lung cancers (Cheng et al, 2013; El Messaoudi et al, 2006; Elakoum et al,
2014; Greenblatt et al, 2019; Karakashev et al, 2018; Kim et al, 2010b; Majumder et al, 2006;
Osada et al, 2013; Yang & Bedford, 2013). PRMT4-specific inhibitors and type I PRMT inhibitors
have been developed for cancer treatment (Guccione & Richard, 2019).
Another documented role of PRMT4 lies in the regulation of endochondral ossification.
Prmt4-null embryos show significantly delayed endochondral bone formation with a reduction in
chondrocyte proliferation. Moreover, this study revealed that PRMT4 methylates Sox9, which
dissociates beta-catenin from Sox9 and leads to upregulation of Tcf/Lef activity and Cyclin D1
expression to enhance chondrocyte proliferation (Ito et al, 2009). PRMT4 was also required in
regulating 25-hydroxyvitamin D3 24-hydroxylase for bone homeostasis (Christakos et al, 2007).
In addition, a recent study has shown that a single nucleotide polymorphism (SNP) of PRMT4 is
associated with osteoporosis (Panach et al, 2020). However, knowledge of how PRMT4 may
regulate osteogenesis during development is unknown.
Previous post-doctoral research fellow in Dr. Xu’s lab examined PRMTs expression in
osteogenic cell lines which are mouse calvarial preosteoblasts (MC3T3-E1), mouse stromal
mesenchymal cells (ST2), and newborn mouse calvarial osteoblasts (NeMCO). PRMT4 was
expressed the highest in these osteogenic cell systems among all PRMTs (Figure 2.2), in contrast
11
to other mesenchymal cell lines in which PRMT1 expression is usually the highest (Figure 2.3).
This finding suggests that PRMT4 may play important role in osteogenic cells.
Figure 2.2. PRMT4 expression is the highest among all PRMTs in three osteogenic cell lines – mouse
calvarial preosteoblasts (MC3T3-E1), mouse stromal mesenchymal cells (ST2), and neonatal mouse
calvarial osteoblasts (NeMCO). (Shinde A., unpublished data).

Figure 2.3. The expression of PRMTs in embryonic fibroblasts and embryonic stem cells. PRMT1
expression is the highest among all PRMTs. Bar graphs are generated with data from Expression Atlas of
EMBL-EBI with originally publication at (Lienert et al, 2011). Note that data of PRMT8 expression in
embryonic fibroblast is not available.

12
2.1.2 Frontal bone and posterior frontal (PF) suture development
Human skull is made up of multiple bones, which can be separated into two parts –
viscerocranium and neurocranium. Viscerocranium contributes to facial bones while
neurocranium covers and protects the brain. Neurocranium is comprised of cranial base and
cranial vault which is also known as the skull cap or calvaria. There are 5 major bones of calvaria:
a pair of frontal bones, a pair of parietal bones, and the interparietal part of occipital bone (Jin et
al, 2016; Tubbs et al, 2012) (Figure 2.4).

Figure 2.4. Human skull consists of neurocranium (highlighted in blue) and viscerocranium (highlighted in
orange)(modified from (Tubbs et al, 2012)).

Cranial sutures are located between two opposing cranial bones. Sutures are widely open
in early postnatal period and called the fontanelles. These fontanelles confer flexibility to the
newborn head during delivery and allow brain growth during development. There are 4 major
cranial sutures: metopic suture located between frontal bones, coronal suture between frontal and
parietal bone, sagittal suture between parietal bones, and lambdoid suture between parietal and
occipital bone (Figure 2.5). Each suture undergoes fusion process at different time. Metopic
suture fuses the earliest, around 1-2 years of age, while sagittal, coronal, and lambdoid sutures
remain patent and fuse at much later time, around 20-40 years old (Sperber, 2010; Weinzweig et
al, 2003). Mouse cranial sutures are quite similar to human sutures (Cohen, 1997; Teng et al,
2019). There are also 4 major cranial sutures located in the same positions in which posterior
13
frontal suture is equivalent to metopic suture (Figure 2.5). This similarity makes mouse a suitable
study model for the study of cranial suture development.

Figure 2.5. Human calvaria (A) and mouse calvaria (B) are anatomically and developmentally similar (Teng
et al, 2019).

Each cranial suture consists of 4 main components which are the periosteum, dura mater
(DM), suture mesenchyme (SM), and osteogenic fronts (OF) of two opposing cranial bones
(Figure 2.6).

Figure 2.6. Four main components of cranial sutures (Zhao et al, 2015).

Role of dura mater during suture closure was studied by Opperman’s group which showed
that dura mater is essential for maintaining coronal suture patency via multiple soluble heparin-
binding factors (Opperman et al, 1995; Opperman et al, 1996; Opperman et al, 1993). In addition,
A B
14
a direct contact with dura mater was found to be important for PF suture closure (Roth et al, 1996).
However, this role is limited to prenatal stage since postnatal ex-vivo suture culture stayed patent
even without dura mater (Kim et al, 1998). This finding also suggested the possibility of different
mechanisms in regulating suture closure prenatally and postnatally. Furthermore, the role of dura
mater in maintaining suture patency was found to be location specific. When dura mater was
surgically displaced, the dura mater of PF suture led to sagittal suture closure while the dura mater
of sagittal suture maintained PF suture patency (Levine et al, 1998). This finding led to another
discovery about PF suture-derived dural cells showing higher osteogenic induction but lower cell
proliferation than sagittal suture-derived dural cells (Mehrara et al, 1999; Warren et al, 2003b).
Periosteum was reported to be a source for progenitor cells during bone repair (Lin et al,
2014; Roberts et al, 2015). However, there was not enough data to support this role of periosteum
during suture closure. In addition, a study by Opperman et al showed that periosteum did not play
role in maintaining coronal suture patency unlike dura mater (Opperman et al, 1994).
Suture mesenchyme is a stem cell niche for cranial bones in the postnatal period, which
is important for suture patency maintenance and bone injury repair (Zhao et al, 2015). Gli1+ cells
derived from suture mesenchyme contributed to calvarial bone, periosteum and dura mater. In
the sagittal (SAG) suture, during embryonic period, suture mesenchyme close to osteogenic
fronts (OF) were found to differentiate into osteoblasts (Lana-Elola et al, 2007). SAG suture
mesenchyme also expressed Twist1 (Rice et al, 2003), which inhibits differentiation of osteoblasts,
and Msx1, Msx2, and Runx2, which promote differentiation, although Runx2 expressed in SAG
suture mesenchyme was not as high as that in the OF (Kim et al, 1998; Rice et al, 2003). FGF2
was found to induce Twist1 expression in suture mesenchyme (Rice et al, 2003). However, in
contrast to SAG, the stem cell population marked by Gli1 were not detected in PF suture (Zhao
et al, 2015).
OFs are the front ends of two opposing calvarial bones where proliferating and
differentiating preosteoblasts locate. Therefore, expression of osteogenic differentiation markers
15
such as Runx2 and Bsp are detected in the OF (Rice et al, 2003). This part of calvarial suture,
together with aforementioned suture mesenchyme were studied extensively, in which involvement
of a large group of transcription factors and signaling pathways have been uncovered. Growth of
parietal bones depends largely on proliferation of osteoprogenitors around the osteogenic fronts
regulated by multiple signaling pathways including BMP signaling (via Msx2, Alx4), FGF signaling,
and Foxc1 (Eswarakumar et al, 2002; Rice et al, 2005). Fgfr1 and Fgfr2 were expressed in cranial
bones in which Fgfr1 is involved in osteoblast differentiation while Fgfr2 is involved in OF cell
proliferation in coronal suture (Iseki et al, 1999). Bmp2 and Bmp4 expression was observed in
embryonic SAG suture (Kim et al, 1998) in which BMP2 and BMP4 induce Msx2 expression in
OF (Rice et al, 2003). In addition, Shh and Ptc expression was observed in embryonic PF and
SAG suture but not in coronal suture (Kim et al, 1998), which suggests diversity in regulatory
mechanisms among all calvarial sutures.
Apoptosis was also found to be involved during normal suture closure in several locations
of calvarial sutures including the mid-suture mesenchyme, osteogenic fronts, and dura mater
(Rice et al, 1999). Furthermore, similar apoptotic activity but different apoptotic pathways in the
fusing suture versus patent suture were observed (Fong et al, 2004).
Despite the similarity in main components, calvarial sutures do not behave the same,
possibly due to their location and developmental origin. Among major calvarial bones, frontal bone
is the only one derived from neural crest cells. Its underlying dura mater is also neural crest
derived. Therefore, sagittal and coronal sutures are formed with a mixed origin of neural crest and
mesoderm-derived cells, while PF suture is solely derived from neural crest cells (Chai et al, 2000;
Jiang et al, 2002) (Figure 2.7). In addition, most sutures are closed by intramembranous
ossification of two opposing cranial bones. However, posterior frontal suture closure undergoes
endochondral ossification (Bradley et al, 1996; Sahar et al, 2005). The PF suture is the only suture
that fuses in mice by 6-week of age (Bradley et al, 1996), while the other sutures remain patent
throughout life.
16
When looking at the suture appearance in adult mice, two opposing bones in PF suture
and sagittal suture meet in the middle of the suture while in the coronal suture, they overlap
(Figure 2.8).

Figure 2.7. Cranial sutures are different in tissue derivatives (Slater et al, 2008). Blue color represents
neural crest derived structures while pink color represents mesoderm derived structures. OF = Osteogenic
fronts; SM = Suture mesenchyme; FB = Frontal bone; PB = Parietal bone; DM = Dura mater, PC =
Pericranium (Periosteum).

Figure 2.8. H&E staining of cranial sutures from 6-day old mice showed that each cranial suture has
different structure (Lenton et al, 2005). OBT = outer bone table; IBT = inner bone table; CT = connective
tissue; PC = pericranium (periosteum); SM = suture mesenchyme; OF = osteogenic front; DM = dura mater;
SS = sagittal sinus; PB = parietal bone; FB = frontal bone.

As displayed in the Figure 2.8, PF suture is composed of ectocranial (OBT) and
endocranial layers (IBT), in which only the endocranial layer undergoes fusion while the
ectocranial layer remains unfused. This fusion is mediated by endochondral ossification, another
unique feature of the PF suture. The PF suture closes from the anterior to posterior direction and
progresses from inner (endocranial) to outer (ectocranial) layer (Bradley et al, 1996).
Posterior frontal suture
Coronal suture
Sagittal suture
Posterior frontal suture
Coronal suture
Sagittal suture
17
Comprehensive study in PF suture development during postnatal period (Sahar et al, 2005)
showed distinct morphology of this suture and the role of Sox9 in regulating chondrogenesis at
the beginning of endochondral ossification. The suture mesenchyme condensation in
chondrogenesis along with upregulation in Sox9 started around P7 in a discontinuous pattern
along the PF suture preceding the bony bridge formation starting in P9-P10 and Osteocalcin
upregulation. This study also confirmed previous finding that PF suture fuses from anterior to
posterior direction and ectocranial layer remains open while endocranial layer fuses which makes
PF suture unique (Figure 2.9).

Figure 2.9. The location of PF suture (A) and the coronal section of PF suture (B) (Sahar et al, 2005).
Jugum limitans (JL) separates frontal suture to 2 parts: anterior frontal (AF) suture and posterior frontal (PF)
suture. F = frontal bone; P = parietal bone; SO = supraoccipital bone; COR = coronal suture; SAG = sagittal
suture; LAM = lambdoid suture

Studies in PF suture have shown the involvement of multiple factors and signaling
pathways. Msx1 and Msx2 functioned synergistically to regulate osteogenic differentiation in the
frontal bone (Han et al, 2007; Satokata et al, 2000; Satokata & Maas, 1994). In addition,
overexpression of Ameloblastin, which is an enamel matrix protein, suppressed Msx2 expression
and decreased cell proliferation which then led to PF suture patency (Atsawasuwan et al, 2013).
Furthermore, Msx1 was also found to coorperate with Dlx5 in regulating cell proliferation and
osteogenic differentiation in the frontal bone (Chung et al, 2010). Canonical Wnt signaling is also
important for PF suture closure during endochondral ossification regulated by Twist1 (Behr et al,
2010). In addition, deletion of Axin2, a component in Wnt signaling, led to ectopic cartilage but
18
absence of endochondral ossification which delayed PF suture fusion though the closure finally
occurred (Behr et al, 2013). Twist1 is an inhibitor of chondrogenesis (Reinhold et al, 2006).
Though expression of Msx2 and Twist1 was more restricted to suture mesenchyme of SAG suture
in later embryonic stages and were found to inhibit osteogenic differentiation, both of them were
reported to function together in regulating proliferation and cell differentiation in frontal bone
rudiments in early embryonic stages (Ishii et al, 2003). Disruption in Hedgehog signaling also led
to ectopic bone formation between frontal bones and wider suture (Veistinen et al, 2012; Veistinen
et al, 2017). Noggin, which is an antagonist of BMPs was found to be low expressed in fused PF
suture unlike patent sutures (Warren et al, 2003a). Prickle1 homozygous mutant exhibited wider
PF suture with significant reduction of both Wnt and Hedgehog signaling in frontal bone
primordium (Wan et al, 2018). Furthermore, TGF-beta signaling acts in frontal bone primordium
by regulating cell proliferation via Fgfr2 and regulating osteogenic differentiation via Dlx5 and
Twist1 (Sasaki et al, 2006). Engrailed 1 deletion mice led to patent PF suture due to a reduction
in cell proliferation and osteogenic differentiation (Deckelbaum et al, 2006). In addition, Ephrin-
B1 mutation delayed PF suture fusion by impairing osteogenic differentiation via gap junction
communication (Davy et al, 2006). A deletion of Lmx1b, an anti-osteogenic gene which controls
the differentiation of early migrating mesenchyme (EMM) from supra-orbital mesenchyme (SOM)
in early calvaria patterning, led to premature fusion of multiple sutures including PF suture
(Cesario et al, 2018; Chen et al, 1998). Apart from common signaling pathways regulating
craniofacial development, histone methyltransferase G9a was also found to regulate cell
proliferation and osteogenic differentiation in the frontal bone by interacting with Runx2 (Ideno et
al, 2020). In addition, G9a deposited H3K9me2 at Twist genes loci and deletion of G9a from
CNCC led to PF suture patency (Higashihori et al, 2017). These altogether suggested a nexus of
signaling pathways and transcription factors network in concert to regulate PF suture
development.
19
Most studies of PF suture closure in postnatal period focused on endochondral ossification,
while most studies during embryonic stages focused on cell proliferation, osteogenic
differentiation, and cell survival. Here, my dissertation on embryonic stages shows a new
molecular mechanism of how PF suture closure is regulated and provides another missing piece
of PF suture development.

2.1.3 Runx2
Transcription factor Runx2 (also known as Cbfa1, Aml3, Pebp2aA) is a member of the
Runx family in which all members, Runx1 (Cbfa2), Runx2, and Runx3 (Cbfa3) share the Runt
domain. The Runt domain is the DNA-binding domain and also essential for heterodimerization
with Cbfb, which enhances DNA binding (Ogawa et al, 1993) and stabilizes Runx2 by inhibiting
ubiquitination (Lim et al, 2015; Qin et al, 2015). Runx1 was reported to be important in
hematopoiesis (Wang et al, 1996). Runx3 was found to be involved in metastasis of several
cancer types (Chen et al, 2016), neuronal development (Inoue et al, 2008), lymphocyte
development (Ebihara et al, 2015), and regulate chondrocyte differentiation in cooperation with
Runx2 (Yoshida et al, 2004).
Runx2 is a master regulator for bone formation, as loss of Runx2 abolished bone formation
in vivo (Komori et al, 1997; Otto et al, 1997). Runx2 regulates osteoprogenitor proliferation in vivo
via Fgfr2 and Fgfr3 (Kawane et al, 2018) and osteoblast differentiation (Komori et al, 1997) (Figure
2.10). It is also essential in chondrocyte proliferation (Yoshida et al, 2004) and differentiation
(Inada et al, 1999) (Figure 2.11). Runx2 is required for mesenchymal cell differentiation to form
osteoprogenitors. Subsequently, Runx2 induces Sp7 expression and both of them with Wnt
signaling are required for preosteoblast differentiation to immature osteoblasts. However, Runx2
expression level is decreased at later stages of differentiation. Runx2 also dictates cartilage
phenotype to be either temporary (i.e. a cartilage during endochondral ossification) or permanent
(i.e. articular cartilage) (Ueta et al, 2001). In addition to proliferation and differentiation, Runx2
20
also regulates other cellular processes including cell apoptosis via Bcl2 (Eliseev et al, 2008), TNF-
alpha (Olfa et al, 2010), and p53 (Ozaki et al, 2013).

Figure 2.10. Runx2 regulates osteoblast differentiation (Komori, 2018).

Figure 2.11 Runx2 regulates chondrocyte differentiation (Komori, 2018).

Cell migration is another cellular process regulated by Runx2. Runx2 cooperates with
PI3K-Akt signaling to promote the migration of osteoblasts and chondrocytes (Fujita et al, 2004).
Runx2 also enhances migration/invasion and bone metastasis of multiple cancer types including
21
breast cancer cell (Ferrari et al, 2013), prostate cancer cell (Akech et al, 2010; Ge et al, 2016),
thyroid cancer cell (Sancisi et al, 2012), and hepatic cancer cell (Wang et al, 2016).
Expression of Runx2 is regulated by alternative splicing. Runx2 can be transcribed from
a distal P1 promoter or a proximal P2 promoters, leading to type II Runx2 transcript and type I
Runx2 transcript respectively (Figure 2.12A). Type II Runx2 is more specific to osteogenic cells
while type I Runx2 is expressed more widely also in other cell types and tissues such as skeletal
muscle (Xiao et al, 2003).
Runx2 activity is regulated by multiple domains. There are 3 activation domains (AD1-3)
in Runx2, in which AD2 and AD3 are involved in Runx2 transactivation function
(Thirunavukkarasu et al, 1998), while VWRPY motif and repression domain (RD) is involved with
repression (Westendorf, 2006). The nuclear localization signal domain (NLS), common among
Runx proteins, regulates Runx2 nuclear localization (Thirunavukkarasu et al, 1998) while the
nuclear matrix target signal domain (NMTS) regulates Runx2 intranuclear localization (Zeng et al,
1998) (Figure 2.12B).

Figure 2.12. Alternative splicing of Runx2 (A) and Runx2 domains (B) (Ziros et al, 2008): AD = Activation
Domain; QA = Glutamine-Alanine rich domain; RUNT = Runt Homology Domain (DNA binding domain);
NLS = Nuclear Localization Signal; PST = Proline-Serine-Threonine rich region; NMTS = Nuclear Matrix
Targeting Signal; RD = Repression Domain

A
B
22
Heterozygous mutation of RUNX2 in human causes cleidocranial dysplasia (Mundlos et
al, 1997) with reported prevalence of 0.12 per 10,000 (Stevenson et al, 2012). These patients
present with major characteristics which are short statue, large anterior fontanelle, frontal bossing,
absent or small clavicles, supernumerary teeth, incomplete eruption of permanent dentition,
maxillary hypoplasia, and Class III malocclusion (Mundlos, 1999) (Figure 2.13). Mice with Runx2
heterozygous deletion phenocopied cleidocranial dysplasia patients by showing PF suture
patency and hypoplasia of clavicles (Figure 2.14) along with other skeletal defects such as
delayed intramembranous ossification, small cranial bones, small hyoid bone, and abnormal
xiphoid process (Komori et al, 1997; Otto et al, 1997). Recent study showed that more than 50%
dosage of Runx2 is required to induce osteogenic differentiation from calvarial mesenchymal cells
to osteoprogenitors via hedgehog, Fgf, Wnt, and Pthlh signaling pathways. Failure in doing so led
to cleidocranial dysplasia phenotypes in mice (Qin et al, 2019).

Figure 2.13. Major characteristics of cleidocranial dysplasia patients – patent metopic suture (A), absent
clavicles (B), and unerupted/supernumerary teeth (C) (Tokuc et al, 2006).

A
B
C
23

Figure 2.14. Runx2
+/-
mice showed hypoplasia of clavicles (b) and open metopic suture (d) which
phenocopied cleidocranial dysplasia patients (Otto et al, 1997).

Previous studies have shown multiple post-translational modification of Runx2 including
phosphorylation, acetylation, and ubiquitination which affected its functions (reviewed by (Bae &
Lee, 2006; Vimalraj et al, 2015)). However, studies in arginine methylation of Runx2 is still lacking.

2.1.4 Previous findings of PRMT4 and Runx2:
Dr. Yongchao Gou in Dr. Xu’s lab performed in vitro and in vivo methylation assays and
discovered that PRMT3 and PRMT4 methylated Runx2 (Figure 2.15A and B). In addition, he
identified 4 arginine methylation sites of Runx2 which are R244, R253, R372, and R377. No
methylation was detected when these arginines (R) were mutated to alanine (A) (Figure 2.15C).
These 4 arginines are located in the transcriptional activation/repression domain of Runx2. While
PRMT3 is expressed mainly in the cytosol (Tang et al, 1998), PRMT4 is expressed in both
cytoplasm and nucleus. Runx2 is a transcription factor localized predominantly in the nucleus
(Zeng et al, 1997). Therefore, PRMT4 seems to be a major Runx2 arginine methyltransferase.

Runx2
+/+
Runx2
+/+
Runx2
+/-
Runx2
+/-

24

Figure 2.15. Runx2 is methylated by PRMT3 and PRMT4. In vitro methylation assay shows that Runx2
can be methylated by only PRMT3 and PRMT4 (A). In vivo methylation assay also shows the strongest
methyl-Runx2 bands when PRMT3 and PRMT4 are present (B). This Runx2 methylation is undetectable
when four arginines (R244, R253, R372, and R377) are mutated to alanine (C), confirming that these four
arginine are Runx2 methylation sites for PRMT3 and PRMT4 (Gou Y., unpublished data).

2.2 Hypothesis
Based on the literature and preliminary findings, I hypothesize that PRMT4 is essential
during craniofacial bone development via Runx2 methylation. To test the hypothesis, I will
characterize the craniofacial bone phenotype following neural crest-specific Prmt4 deletion,
determine the functional significance of PRMT4-mediated Runx2 methylation in craniofacial
development, and dissect the mechanistic roles of Runx2 methylation in craniofacial development.

A
25
2.3 Materials and Methods
2.3.1 Animals
Wnt1-Cre and R26R
tdTomato
mice were purchased from Jax (#009107 and #007914).
Prmt4
fl/fl
mice was generously provided by Dr. Mark Bedford at MD Anderson (Yadav et al, 2003).
All animal care and experiments were carried out in accordance with protocols approved by the
Institutional Animal Care and Use Committee (IACUC) at the University of Southern California.

2.3.2 MicroCT scanning
Mouse skulls were collected at P0 and 6-week old of age. The samples were fixed in 4%
paraformaldehyde in PBS overnight, washed several times in PBS to remove 4% PFA, and then
placed in PBS. The skulls were scanned at the USC Molecular Imaging Center, followed by dicom
files reconstruction and analysis using Avizo Lite software (Thermo Fisher Scientific) as previously
demonstrated (Gou et al, 2018b).

2.3.3 Whole-Mount Skeletal Staining
Whole mount skeletal staining was performed following the protocol described previously
(Rigueur & Lyons, 2014) with some modifications. Mouse skulls were harvested at E16.5 and 6-
week of age. Skin, eyes and adipose tissue were removed. Skulls were immersed in 95% EtOH
overnight for fixation, then 100% Acetone for 1-2 days to remove remaining fat. Then, the skulls
were placed in 0.03% w/v Alcian Blue solution for 1-2 days before washing with 70% and 95%
EtOH overnight. The next day, 6-week old skulls were placed in 1% KOH for clearing the tissue
4 hours before replacing with ½ dilution of 0.005% w/v Alizarin red solution for 2 days and 1/3
dilution of 0.005% w/v Alizarin red solution for another 2 days. For E16.5, skulls were placed in
0.005% w/v Alizarin red solution for 3-4 hours. Then, the skulls were placed in 1%KOH for clearing
and gradually changed to 25% glycerol in 1%KOH, 50% glycerol in 1%KOH, 75% glycerol in
1%KOH and finally to 100% glycerol for storage. Skull pictures were taken under Leica M125
microscope.
26
2.3.4 Tissue Processing and Immunofluorescence (IF) Staining
Mouse skulls were collected at E16.5 and P0. The tissues were fixed in 4%
paraformaldehyde in PBS overnight, except the cultured calvaria tissues which were fixed for 4
hours before proceeding to the next step. P0 heads were decalcified in 10% EDTA overnight. For
cryoprotection, tissues were immersed in 15% sucrose in PBS, 30% sucrose in PBS, 30%
sucrose in Optimal Cutting Temperature medium (OCT, Sakura), and finally embedded in OCT.
Tissue blocks were sectioned at 8 µm thickness in coronal orientation on positively charged glass
slides. The tissue samples were washed in PBS 20 minutes to remove OCT and in 0.5% Triton
X-100 in PBS 20 minutes for permeabilization. Next, tissue sections were washed briefly with
PBS, blocked with 10% goat serum for 1 hour, and incubated with primary antibody diluted in 10%
goat serum at 4°C overnight in moisture chamber. Primary antibodies used in this study were:
PRMT4 (1:100, Bethyl Laboratory Inc. Cat# A300-421A-M), RUNX2 (1:100, Cell Signaling
Technology Cat# 12556), SP7 (1:200, Abcam Cat# ab22552), Ki67 (1:100, Abcam Cat# ab16667),
H3R17me2a (1:200, Active Motif Cat# 39709), and H3R26me2 (1:100, Millipore Cat# 07-215).
Finally, the samples were blocked with secondary antibody (1:300, goat anti-rabbit IgG (H+L)
Alexa Fluor 488 Cat# A-11008) diluted in 10% goat serum for 1 hour, DAPI diluted 1:1000 in PBS
for 10 minutes and then mounted with mounting media (Electron Microscopy Science Cat#
1798510) and covered with cover glass. Images were visualized under Keyence BZ-X800 and
Leica DMI 3000B microscopes. Quantification was performed using ImageJ software.

2.3.5 RNA In Situ Hybridization (ISH) using RNAScope
Tissue samples were prepared under RNase-free condition before sectioning. RNA ISH
was carried out by using RNAScope 2.5HD Duplex Detection kit (Advanced Cell Diagnostics
Cat#322430). Following manufacturer’s protocol, target retrieval for 10 minutes and protease
treatment for 15 minutes were performed. Then, PRMT4 probe (Advanced Cell Diagnostics, Mm-
Carm1 C1, Cat# 528841) or PRMT3 probe (Advanced Cell Diagnostics, Mm-Prmt3 C1, Cat#
27
528831) and Runx2 probe (Advanced Cell Diagnostics, Mm-Runx2-C2, Cat# 414021-C2) were
mixed in 50:1 ratio respectively. Tissue sections were incubated with mixed probes for 2 hours at
40°C in the HybEZ oven (Advanced Cell Diagnostics). Hybridization and signal detection steps
were followed as manufacturer’s instruction. Nuclei were counterstained with 20% Hematoxylin.
Tissue sections were mounted with VectaMount Mounting Medium (Vector Laboratories Cat# H-
5000). Images were visualized under Keyence BZ-X800.

2.3.6 Proximity Ligation Assay (PLA)
PLA was performed using Duolink In Situ kit (Millipore Sigma Cat# DUO92101-1KT),
Runx2 antibody (Cell Signaling Technology Cat# 12556) and pan-methyl arginine antibody,
Asymmetric Di-Methyl Arginine (ADMA) (Cell Signaling Technology Cat# 13522) to detect methyl-
Runx2. Since the antibodies for both proteins were anti-rabbit-IgG, Duolink In Situ Probemaker
MINUS (Millipore Sigma Cat# DUO92010-1KT) and PLUS (Millipore Sigma Cat# DUO92009-1KT)
were applied to generate PLA conjugated primary antibodies for Runx2 and ADMA respectively.
The frozen tissue sections were permeabilized with 0.5% Triton X-100 for 15 minutes before
following the manufacturer’s instruction. Tissue sections were blocked with the Duolink blocking
solution in humidity chamber for 1 hour at 37°C before incubating with PLA conjugated primary
antibodies (diluted 1:50 in Duolink PLA probe diluent) overnight at 4°C. The next day, tissue
sections were incubated with ligase solution for 30 minutes at 37°C and amplification solution for
100 minutes at 37°C. After final washes, tissue sections were mounted with Duolink In Situ
mounting media with DAPI to counterstain nuclei. Images were visualized under Leica DMI 300B
microscopes. The number of PLA signals which appear as red dots in the nuclei were quantified.

2.3.7 TUNEL Assay
TUNEL Assay was performed using Click-iT Plus TUNEL Assay kit (Thermo Fisher
Scientific Cat# C10617) to detect in situ apoptosis. Briefly, tissue sections were incubated with
28
Proteinase K solution for 15 minutes. For positive control, DNase (Qiagen Cat# 79254) was
applied for 10 minutes to induce DNA strand breaks. Next, the enzyme terminal deoxynucleotidyl
transferase (TdT) solution was incubated with the sections for 1 hour. Then, the Click-iT Plus
cocktail was applied for 30 minutes before nuclear staining with DAPI and mounting. Images were
visualized under Leica DMI 300B microscopes.

2.3.8 Ex vivo Calvaria Culture and DiI Labeling
Calvaria were dissected from mouse embryos at E15.5 and cultured as previously
described (Rice et al, 2003) with slight modifications. Calvaria (above the eyes) were dissected
in Hanks’ Balanced Salt Solution (HBSS) (Thermo Fisher Scientific Cat# 14175079) under the
dissecting microscope. The skin and brain tissue were removed while dura mater was carefully
maintained. CellTracker CM-DiI dye (Thermo Fisher Scientific Cat# C7000) was chosen for dye
labeling. Micropipettes (Drummond Scientific Company Cat# 2-000-001) were used to push the
dye to label at either osteogenic fronts or suture mesenchyme. After labelling, the calvaria were
placed on the 0.8 µm membrane filter (Millipore Sigma Cat# AABP04700) on the metal grid, which
rested on the inner edge of the organ culture dish (Corning Cat# 353037) (Figure 2.16).
Dulbecco’s Modified Eagle’s Medium (DMEM) (Genesee Scientific Cat# 25-501) which is
supplemented with 10% fetal bovine serum (FBS), 0.1mg/ml ascorbic acid, and 1%
penicillin/streptomycin was added to the inner well of the organ culture dish to cover the calvaria
base. The calvaria photos were taken every day from Day 0 (when the dye was injected) to Day
3 under Leica MZ10F microscope. The media was replaced every day. After 3-day observation,
the calvaria tissues were processed for cryosection and immunostaining. Photos were used to
measure the PF suture closure percentage and compare between control and mutant groups.
29
Figure 2.16. Organ culture dish with metal grid and membrane filter for ex vivo calvaria culture

2.3.9 Cell Culture, siRNA, and Plasmid Transfection
Mouse calvarial preosteoblasts (MC3T3 – E1) were cultured in Minimum Essential
Medium a (MEM-a) without ascorbic acid supplemented with 10% FBS in incubator until they
reached confluence. Then, cells were split to three groups in a cell density which would reach
about 90% confluence the next day for reverse transfection with scrambled siRNA (siControl:
Qiagen Cat# 1027310) and two independent siRNA targeting Prmt4 (siPrmt4 #3: Qiagen Cat#
SI00941437 and siPrmt4 #5: Qiagen Cat# SI04422614) at 40 nM using Lipofectamine RNAiMax
transfection reagent (Invitrogen) for siRNA delivery. For plasmid transfection: GFP, flag-tagged
WT Runx2, and flag-tagged 4RK Runx2 were transfected (forward) to 80% confluent cells with
Lipofectamine 2000 (Thermo Fisher Scientific Cat# 11668019) at ratio plasmid 1.25ug to
lipofectamine 2.5ul.

2.3.10 Migration assay
After 24 hours of reverse transfection with siRNA or forward transfection with plasmids,
cells were re-plating to ensure confluency after overnight culture. Then, a 200ul-pipette tip was
used to generate the cell-free area in the middle of each well. Media was changed afterwards,
and T0 pictures were taken 30 minutes after the scratch was made under Leica DMI 3000B
microscope. T1 pictures were taken at 12 hours and 16 hours after scratching for siRNA and
plasmid groups respectively. In each well, pictures were taken at 4 different areas along the
30
wound edges. To analyze the wound, straight lines were drawn at the edges of the wound. Wound
size was measured as a width of the gap between these two straight lines of the wound at each
timepoint. Percentage of wound closure of each sample were calculated using the formula
described in the previous study (Yue et al, 2010) as shown below.

2.3.11 SDS-PAGE and Immunoblotting
MC3T3-E1 cells were scraped and lysed in lysis buffer containing 50 mM Tris-HCl pH 7.5,
250 mM NaCl, 2mM EDTA, 0.1% NP-40, 10% glycerol, and protease inhibitor cocktail.
Quantification of protein concentration was performed using Bio-Rad protein assay (Bio-Rad
Laboratories). Each protein sample (40 ug) was separated using SDS-PAGE at 80-110V and
separated proteins were transferred to 0.45 μm PVDF membrane at 100V for 3 hours. The
membrane was blocked with 5% powdered milk in TBST (TBS, 0.1% Tween 20) for 1 hour before
incubated with primary antibody (PRMT4 (1:5,000 Bethyl Laboratory Inc. Cat# A300-421A-M), b-
actin (1:1,000 Santa Cruz Cat# sc-47778), Runx2 (1:1,000 MBL Cat# D130-3)) diluted in 3% BSA
in TBST with sodium azide at 4°C overnight. Then, membrane was washed in TBST briefly and
blocked with HRP-conjugated secondary antibody diluted 1:10,000 in 5% powdered milk in TBST
for 1 hour. Finally, membrane was washed in TBST and incubated in ECL (GE Healthcare) for 1
minutes to detect protein bands using HyBlot CL film (Denville Scientific).

Wound Closure (%) = 0
W
!"#$
−W
!"∆$
W
!"#$
2 × 100%

W
!"#$
is the gap distance of the wound measured 30 minutes after scratching
W
!"∆$
is the gap distance of the wound measured h hours after scratching

31
2.3.12 Bulk RNA-sequencing Analysis
Calvaria from mouse embryos from control (Wnt1-cre; R26R
tdTomato
) and Prmt4 mutant
(Wnt1-cre; R26R
tdTomato
; Prmt4
fl/fl
) groups at E15.5 were dissected in HBSS under dissecting
microscope and the osteogenic fronts (including small part of adjacent suture mesenchyme and
frontal bone) were cut out for further steps. Tissues were digested in TrypLE solution (Thermo
Fisher Scientific Cat# 12605010) with rotation in 37°C chamber for 30 minutes. Then, 10%FBS
was added to stop the digestion. The supernatant was transferred to collecting tube with 40-µm
cell strainer. Cells were collected by centrifuging at 500 rcf for 5 minutes and were resuspended
in PBS. DAPI was added to each sample to help exclude dead cells during fluorescence activated
cell sorting (FACS). After cell sorting, only tdTomato-positive neural crest cells were collected for
RNA extraction with RNeasy Micro Kit (Qiagen Cat# 74004). NCC from nine control OFs and four
mutant OFs were sequenced with the Illumina HiSeq 4000 at 20 million sequencing depth and
100 bp paired end sequencing. Partek Flow (Partek Inc.) was utilized for analysis. Reads were
trimmed from both ends based on a quality score with minimum read length of 25. Then, reads
were aligned with STAR to Mus musculus (mouse) – mm10 assembly and GENCODE Genes –
release M25 aligner index. Next, reads were quantified to annotation model (Partek E/M). Noise
reduction filter and normalization were applied. GSA was chosen to generate differential analysis
comparing between Prmt4 mutant and control group. Upregulated and downregulated genes were
analyzed for pathways involved and Gene Ontology (GO) with Ingenuity Pathway Analysis
(Qiagen Inc.) and Metascape (metascape.org) (Zhou et al, 2019) respectively. Altogether, six
RNA samples from a pool of nine control OFs and four mutant OFs were sequenced and analyzed.

32
2.3.13 Chromatin Immunoprecipitation – Quantitative PCR (ChIP - qPCR)
MC3T3-E1 cells were transfected with siControl, siPrmt4 #3, or siPrmt4 #5 and cultured
to reach 100% confluency in 6-cm dish before proceeding with Chromatrap Enzymatic ChIP-seq
kit (Chromatrap Cat#500191). Briefly, cells were crosslinked with 1% formaldehyde, quenched
with 0.65M glycine, scraped in cold PBS, and centrifuged. Next, cells were lysed in hypotonic
buffers and enzymatic digestion was performed to shear chromatin. Shearing efficiency was
validated by running an aliquot of the samples on agarose gel to confirm that chromatin fragment
size ranges between 100-600 bp. Then, 20% of each sample was saved as ‘input’. The remaining
samples were incubated with anti-Runx2 antibody (Cell Signaling Cat# 12556S) or rabbit IgG (Cell
signaling Cat# 2729S) as negative control overnight at 4°C. After that, chromatin was eluted using
columns and proceeded with reverse cross-linking procedures together with ‘input’ samples and
incubated at 65°C overnight. Finally, DNA was eluted and used for qPCR with PowerUP SYBR
Green Master Mix (Thermo Fisher Scientific Cat# A25743). The quantification of enrichment from
Ct value was normalized with percentage of input as described in previous study (Lin et al, 2012).
DCt (normalized ChIP) = Ct(sample) – [Ct(input) – log2(input dilution factor)]. Then, the percentage
of each sample per input was calculated from 100 x [2
-DCt (normalized ChIP)
]. Finally, the enrichment
fold of Runx2 to its target genes was normalized over IgG group (negative control).

2.3.14 Statistical analysis
Presented data were mean ± SE from at least 3 independent experiments. Comparison
analysis was conducted using t-test (for 2 groups) or 1-way ANOVA (for ≥ 3 groups) with Tukey
HSD post hoc test. Statistical significance was considered at P ≤ 0.05.

33
2.4 Results
2.4.1 NCC-specific Prmt4 deletion leads to PF suture patency
Since it was reported previously that global Prmt4 knock-out mice die soon after birth
(Yadav et al, 2003), I decided to generate conditional Prmt4 knock-out mice. To study the role of
PRMT4 in CNCC behavior and craniofacial morphogenesis, I deleted Prmt4 in the neural crest
cell lineage using Wnt1-Cre and characterized the skull morphology of control (Prmt4
fl/fl
), and
Prmt4 mutant (Wnt1-Cre; Prmt4
fl/fl
) mice. At E16.5, I noted a wider gap between frontal bones in
the mutant mice (Figure 2.17A), while other bony structures appeared unchanged (Figure 2.17B).
I further performed morphometric analysis using microCT scanning at the newborn stage (P0),
which revealed a significantly larger gap between frontal bones in mutant mice when compared
with control mice (Figure 2.18A and B). Other cranial neural crest (CNC)-derived craniofacial
bones, including premaxillae, maxillae, palatine bones, and mandibles showed no apparent
alteration in size or shape (Figure 2.19A-D). I further examined Prmt4 mutant mice in adulthood.
The Prmt4 mutant mice had smaller heads and proportionally smaller body size (Figure 2.20).
Posterior frontal (PF) suture undergoes complete fusion around 5 to 6 weeks after birth (Bradley
et al, 1996). Therefore, I performed whole-mount skeletal preparation and morphometric analysis
of 6 week-old mouse heads and found that PF suture was patent in all the Prmt4 homozygous
deletion mice, while PF suture fused completely in control and Prmt4 heterozygous deletion
(Wnt1-Cre; Prmt4
fl/Wt
) mice (Figure 2.21). In addition, coronal sections of PF suture from microCT
scanning confirmed an alteration of PF structure in Prmt4 homozygous deletion mice (Figure 2.22).
These data demonstrated that Prmt4 deletion in CNCs led to posterior frontal suture patency.

34

Figure 2.17 Whole-mount skeletal staining of E16.5 mice showed wider gap between frontal bones
(highlighted in yellow) in mutant mice (A). Other bony structures appeared unchanged (B). Scale bar = 1mm

Figure 2.18 MicroCT scanning of P0 mouse skull showed wider gap between frontal bones in mutant (A-b)
than in control (A-a). Gap width, which is the distance between point1 on the left (L1) and right side (R1)
was significantly larger in mutant mice (B). Scale bar = 1mm, *p-value ≤ 0.05

B
35

Figure 2.19. MicroCT scanning of P0 mouse skull showed no apparent alteration in size or shape of other
CNC-derived craniofacial bones in mutant mice: premaxilla (A), maxilla (B), palatine bone (C), mandible
(D). Scale bar = 1mm

36

Figure 2.20. Prmt4 mutant mouse (left) had smaller head and proportionally smaller body size than control
mouse (right). These mice were 8 months old. Scale bar = 1cm

Figure 2.21. At 6-week of age, PF suture was patent in Prmt4 mutant mice (A-c’) while it was completely
fused in control (A-a’) and Prmt4 heterozygous deletion mice (A-b’). Scale bar = 2mm

Wnt1-cre; Prmt4
fl/fl
Prmt4
fl/fl

37

Figure 2.22. MicroCT analysis of 6 weeks old mouse skull revealed that Prmt4 mutant mice had shorter
skull (c) with alteration of PF suture structure (f). Scale bar = 2mm (a-c) and 1mm (d-f)

2.4.2 PRMT4 is enriched at the osteogenic fronts
I assessed PRMT4 expression in PF suture components with immunofluorescent staining
using calvaria from control (Prmt4
fl/fl
; R26R
tdTomato
) and mutant mice (Wnt1-cre; Prmt4
fl/fl
;
R26R
tdTomato
). TdTomato reporter was incorporated in these mice to label neural-crest derived
tissue (Figure 2.23A-g). At P0, PRMT4 was expressed in the developing frontal bone primordium,
suture mesenchyme and dura mater, and significantly enriched in osteogenic fronts (Figure
2.23A-a’). Similar PRMT4 expression pattern was observed at E16.5 (Figure 2.23B). In Prmt4
mutant mice, Prmt4 was successfully deleted in CNC-derived tissues including the frontal bone
primordium, suture mesenchyme and dura mater (Figure 2.23A-d’). In addition to Prmt4, Runx2
was highly expressed at the osteogenic fronts (Figure 2.23C).

38

Figure 2.23. PRMT4 is enriched at the osteogenic fronts. At P0, PRMT4 was expressed in the frontal bone,
suture mesenchyme (SM), and dura mater (DM), and significantly enriched in the osteogenic fronts (OF)
(A-a’). tdTomato labeled neural crest-derived tissue (A-g). In Prmt4 mutant mice, Prmt4 was successfully
deleted (A-d’). Similar PRMT4 expression pattern was observed at E16.5 (B). In addition to Prmt4, Runx2
was highly expressed at the OF (C). The red line on the skull marked the area where tissue sections were
obtained from (A-h). Scale bar = 100µm

39
2.4.3 PRMT4 co-expresses with Runx2
PRMT4 is highly expressed in cells at the osteogenic front, where Runx2 positive
osteoprogenitors are enriched. To examine whether Runx2 and PRMT4 are co-expressed in
osteoprogenitors, I performed RNA in situ hybridization with RNAScope to assess the mRNA
expression of Runx2 and Prmt4. PRMT4 was co-expressed with Runx2 in both osteogenic fronts
and suture mesenchyme (Figure 2.24A-a and A-b). In addition, Prmt4 transcripts were decreased
in the Prmt4 mutant while Runx2 transcripts remained unaltered (Figure 2.24A-c and A-d).
Moreover, PRMT3 expression pattern was similar to PRMT4 but at a much lower level, as
demonstrated by weak signals for Prmt3 transcripts (Figure 2.24B).

40
Figure 2.24. PRMT4 co-expresses with Runx2. RNA in situ hybridization of PF suture at P0 showed that
PRMT4 co-expressed with Runx2 at OF (A-a) and SM (A-b) in control while Prmt4 transcripts were
decreased in mutant (A-c and A-d). In addition, PRMT3 expression pattern was similar to PRMT4 but at
much lower level (B). RNA-seq data from OF showed significant reduction of Prmt4 transcripts in mutant
(D) while Runx2 and Prmt3 transcripts were not significantly altered in mutant (C and E). Scale bar = 50µm,
*p-value ≤ 0.05.

2.4.4 Methyl-Runx2 is reduced upon NCC-specific Prmt4 deletion at the osteogenic fronts
To further assess whether Runx2 is methylated in cells at the osteogenic fronts, I
performed a Proximity Ligation Assay (PLA), which is a method to detect protein-protein
interaction and protein modification in situ. In this case, the colocalization of arginine methylation
and Runx2 was visualized in red dots (Figure 2.25A-a’). There was a significant reduction of red
signals in the Prmt4 mutant when compared with control (Figure 2.25A-b’). This finding suggested
a significant reduction of Runx2 methylation at the osteogenic fronts when Prmt4 is deleted.

Figure 2.25. Methyl-Runx2 is detected at the OFs as red dots, pointed out by yellow arrows (A-a’) and is
reduced upon NCC-specific Prmt4 deletion at the OFs (A-b’). Quantification of number of reactions (red
dots) was shown as a bar graph (B). Scale bar = 50 µm (a and b) and 25 µm (a’ and b’). ** p-value ≤ 0.01

41
2.4.5 NCC-specific Prmt4 deletion does not significantly alter cell proliferation or apoptosis
Since I found loss of Runx2 methylation in NCC-specific Prmt4 deletion mice at the
osteogenic fronts of the patent PF suture, I questioned if this phenotype resulted from disturbance
of Runx2 functions. Runx2 was found to regulate several cellular processes including cell
proliferation, cell differentiation, cell apoptosis, and cell migration. Therefore, I decided to
investigate in these processes.
I examined proliferation of osteogenic cells by staining PF suture with Ki67, which is a
protein marker expressed in dividing cells and peaks in mitosis. Ki67 was highly expressed in
osteogenic fronts where proliferation of pre-osteoblasts takes place, but Ki67 expression was not
altered in PF suture of mutant mice for all timepoints observed (Figure 2.26).
To detect cell apoptosis, I performed TUNEL assay and found that the number of apoptotic
cells was not changed significantly in Prmt4 mutant (Figure 2.27). Altogether, these findings
indicated that CNC-specific deletion of Prmt4 did not affect the proliferation or apoptosis of cells
at the osteogenic fronts in PF suture development.

42

Figure 2.26. Cell proliferation at the OFs was not significantly altered with NCC-specific Prmt4 deletion at
P0 (A) and E16.5 (C). Quantification of Ki67 positive cells at the OF was shown as a bar graph for each
time point (B and D). Scale bar = 100µm, p-value > 0.05 (not statistically significant)

43

Figure 2.27. Cell apoptosis at the OFs was not significantly altered with NCC-specific Prmt4 deletion at P0
(A) and E16.5 (C). Quantification of positive cells at the OF was shown as a bar graph for each time point
(B and D). Scale bar = 100µm, p-value > 0.05 (not statistically significant)

2.4.6 NCC-specific Prmt4 deletion impairs osteogenic front cells migration
Migration of pre-osteoblast/osteoblast is one of important processes during bone
formation, bone remodeling, and bone repair (Thiel et al, 2018). It was found to be regulated by
several factors such as TGF-beta1, BMP-2, dynamin GTPase, Nck, and transcription factors like
Runx2 (Aryal A C et al, 2015; Eleniste et al, 2014; Fiedler et al, 2002; Fujita et al, 2004; Tang et
al, 2009). Previous studies have shown apical migration of frontal bone primordium drives frontal
bone development in early embryonic stage (Machida et al, 2014; Yoshida et al, 2008), and that
suture mesenchyme progenitors are recruited to the osteogenic fronts during the postnatal period
(Zhao et al, 2015). However, mechanism of cell migration at the developmental stage of my focus
is unclear. Therefore, I decided to investigate for cell migration at both osteogenic fronts and
suture mesenchyme.
44
I hypothesized that cell migration at OF is reduced by Prmt4 deletion, which leads to PF
suture patency. I proposed 2 hypotheses here. The first hypothesis is that cells at the osteogenic
fronts migrate towards the midline and this migration is impaired in Prmt4 mutant. The second
one is that cells at the suture mesenchyme are recruited to the OF and this recruitment is impaired
in Prmt4 mutant.
To analyze cell migration in live calvaria, I used a chloromethylbenzamido derivative of DiI
(1,1’-Dioctadecyl-3,3,3’,3’-Tetramethylindocarbocyanine Perchlorate; DiLC18(3)) called
CellTracker CM-DiI. This dye has been used for labeling and tracking cells in several studies
(Lana-Elola et al, 2007; Machida et al, 2014; Ting et al, 2009; Yoshida et al, 2008). It is harmless
to cells and does not diffuse to adjacent cells. Once it passes through cell membrane, it is well-
retained and stable in fixative reagents.
To test the first hypothesis, I injected DiI at the osteogenic front on both sides. I expected
that the majority of the dye is incorporated into the OF cells and moves towards the midline. Some
dye may retain in the matrix of the injection area (Figure 2.28A). Calvaria photos from
fluorescence microscope showed that the dye-labeled cells at the OF on both sides moved
towards the midline from Day 0 to Day 3 in control (Figure 2.28B). However, this movement was
impaired in Prmt4 mutant which was in accordance with a reduction in percentage of PF suture
closure when compared with control (Figure 2.28C). I further co-stained coronal sections of these
calvaria tissue with Runx2 to locate the osteoprogenitors and the osteogenic fronts. The majority
of the DiI was found at the tip of osteogenic front and traces of DiI could be visualized in the distal
part of the frontal bones (Figure 2.28D). These findings supported the first hypothesis and
demonstrated that osteogenic cells at the osteogenic fronts migrate during PF suture closure and
this migration was impaired in Prmt4 mutant.
To test the second hypothesis, I injected DiI at the suture mesenchyme. I expected that
dye-labeled progenitors migrate from the suture mesenchyme to the osteogenic fronts. Some dye
may retain in the matrix at the injection site (Figure 2.29A). However, calvaria photos from
45
fluorescence microscope revealed that DiI-labeled suture mesenchymal cells did not migrate to
the OF in either control or Prmt4 mutant (Figure 2.29B). I further performed coronal sectioning
and co-staining with Runx2 and examined consecutive sections carefully to confirm this
observation (Figure 2.29C). These findings invalidated the second hypothesis.
Taken together, the data indicate that osteogenic cells at the OF are the predominate drive
for migration during OF suture closure at E15~18 and this migration requires PRMT4.
Figure 2.28. Cells at the OF migrate during PF suture closure. A schematic drawing of the first hypothesis
(A). Fluorescence microscope photos showed that DiI-labeled cells at the OF moved towards the midline
from Day0 to Day3 in control and this movement was impaired in mutant (B) (Scale bar = 1mm). Percentage
of PF suture closure was shown as a bar graph (C) (*p-value ≤ 0.05). IF staining showed a majority of the
DiI the tip of OF which supported the first hypothesis (D) (Scale bar = 100µm (a,b) and 50µm (a’, b’)
46

Figure 2.29. A schematic drawing of the second hypothesis was shown (A). Fluorescence microscope
photos showed that DiI-labeled mesenchymal cells did not migrate towards the OF in either control or
mutant (B) (Scale bar = 1mm). IF staining further invalidated the second hypothesis (C) (Scale bar = 100µm
(a,b) and 50µm (a’, b’)

2.4.7 PRMT4 depletion reduces migration of pre-osteoblast MC3T3-E1 cells
To directly assess the role of PRMT4 in cell migration, I used MC3T3-E1 cells which is a
pre-osteoblast cell line derived from mouse calvaria. Then, PRMT4 was depleted using 2
independent siRNAs and migration assay was performed (Figure 2.30A). After 12 hours,
percentage of wound closure was lower in both PRMT4 depletion groups when compared with
siControl group (Figure 2.30B and C). This finding indicated that cell migration was impaired upon
PRMT4 depletion.

47

Figure 2.30. Cell migration was impaired upon PRMT4 depletion. PRMT4 was depleted from MC3T3-E1
cells (A) and scratch assay was performed. After 12 hours, wound gap was larger in siPrmt4 groups when
compared with siControl group (B). Percentage of wound closure was shown as a bar graph. *p-value ≤
0.05, **p-value ≤ 0.01

2.4.8 Runx2 methylation by PRMT4 is important for cell migration
To investigate whether PRMT4-regulated arginine methylation of Runx2 regulates its pro-
motility function in osteoblasts, wild-type Runx2 (WT Runx2) and methyl-deficient Runx2 (4RK
Runx2) were transfected to MC3T3-E1 cells to compare wound closure percentage in scratch
assay (Figure 2.31A). At 16 hours after scratching, 4RK Runx2 group showed a significantly lower
wound closure percentage when compared with control (GFP) and WT Runx2 groups (Figure
2.31B and C). This finding suggested that Runx2 methylation deficiency impaired cell migration.

B
C
48

Figure 2.31. Runx2 methylation deficiency impaired cell migration. WT Runx2 and 4RK Runx2 were
transfected to MC3T3-E1 cells (A) and scratch assay was performed. After 16 hours, wound gap was larger
in 4RK Runx2 group when compared with GFP and WT Runx2 group (B). Percentage of wound closure
was shown as a bar graph. **p-value ≤ 0.01 when compared with GFP, ## p-value ≤ 0.01 when compared
with WT Runx2

2.4.9 NCC-specific Prmt4 deletion downregulates a group of ECM-related genes in PF
suture
To determine the molecular mechanism by which PRMT4-Runx2 regulates migration, I
isolated CNCC from osteogenic fronts of PF suture at E15.5, purified by cell sorting with tdTomato,
and analyzed with RNA-sequencing (Figure 2.32).
The list of downregulated genes in osteogenic fronts of Prmt4 mutant (≥1.3-fold reduction,
P≤0.05) was analyzed for Gene Ontology (GO). In terms of cellular components, collagen-
containing extracellular matrix (GO:0062023) was mostly associated with the downregulated
genes. In addition, downregulated genes were highly related to invadopodium (GO:0071437) and
actin cytoskeleton (GO:0015629) which are involved in extracellular matrix as well (Figure 2.33A).
With Ingenuity Pathway Analysis (IPA) (Qiagen), cell migration and cell invasion pathways were
identified to be downregulated significantly in Prmt4 mutant (Figure 2.33B). Using IPA, many of
the downregulated cell migration/invasion and matrix genes were further identified to be Runx2
targets (Figure 2.34).
49

Figure 2.32. Schematic drawing showed workflow of RNA-seq analysis from CNCC isolated from OF.

Figure 2.33. GO Analysis revealed that downregulated genes in Prmt4 mutant were mostly related with
ECM (A). In addition, IPA identified that cell migration and cell invasion pathways were significantly
downregulated in Prmt4 mutant (B).

50

Figure 2.34. Examples of downregulated ECM genes which were identified as Runx2 targets.
*p-value ≤ 0.05, **p-value ≤ 0.01

2.4.10 Depletion of PRMT4 decreases Runx2 binding with its target gene promoters in
calvarial preosteoblasts.
To determine whether downregulation of Runx2 target genes in Prmt4-deletion osteogenic
fronts was due to a reduction in Runx2 binding to the promoter region, I analyzed ChIP-seq data
generated from Dr. Pike’s group (Meyer et al, 2014) using UCSC genome browser and selected
ECM-related migration genes (Timp1, Serpine1, and Pik3cb) which show robust Runx2 binding
peak. Next, I performed ChIP-qPCR to check for enrichment of Runx2 on these gene promoters
in MC3T3-E1 cells treated with control siRNA and two independent Prmt4 siRNAs. I observed a
reduction of Runx2 enrichment at the promoter region of selected genes in Prmt4 siRNA groups
when compared with control siRNA group (Figure 2.35). These findings suggested that PRMT4
depletion leads to a reduction of Runx2 binding to ECM-related genes.

51

Figure 2.35. Depletion of PRMT4 decreased Runx2 binding with its target ECM-related migration genes in
calvarial preosteoblasts. For each gene, Runx2 binding peak (Meyer et al, 2014) was shown at the top with
red arrows specifying the location which is targeted by primers (A,C,E) and ChIP-qPCR result shown in bar
graphs below (B,D,F). *p-value ≤ 0.05 compared with IgG, **p-value ≤ 0.01 compared with IgG
# p-value ≤0.05 compared with Runx2 IP of siControl, ## p-value ≤0.01 compared with Runx2 IP of siControl

Target
gene
Forward primer sequence Reverse primer sequence Coordinate
Sp7 TTAGCTGCTGCCCCCTCC AAAATAAACCGACCGCAGGGA chr15:102193208-
102193314
Alpl CCTGGTGTGCCAATGTCGAT AATGCTGACTGAGCTTCGGA chr4:137,349,646-
137,349,770
Timp1 TGACTCACTAACTTCTTTTACCATC AGGCAGGACACAATCTTGACA chrX:20446559-
20446667
Serpine1 TGGAGACGGGACGTTTTTCT CAGCCAACAAGAGCCAATCAC chr5:137547917-
137548067
Pik3cb ATTCCCTGCGATGTGAACGA GTGGTGCAGAGACGAACACA chr9:99040930-
99041058

Table 2.1. Primer sequence of each target gene for ChIP-qPCR
A
B
C
D
E
F
52
2.4.11 NCC-specific Prmt4 deletion leads to reduction of H3R17me2 and H3R26me2 in PF
suture
H3R17 and H3R26 were discovered to be methylated by PRMT4 (Schurter et al, 2001)
and involved in transcriptional activation (Ma et al, 2001). Since several gene sets were
downregulated upon Prmt4 deletion, I questioned if methylation of H3R17 and H3R26 is affected.
Immunostaining of PF suture at P0 showed a significant reduction of both H3R17me2 and
H3R26me2 in PF suture (Figure 2.36A and B).

Figure 2.36. NCC-specific Prmt4 deletion leads to reduction of H3R17me2 (A) and H3R26me2 (B) at the
OF of PF suture. Scale bar = 100 µm

2.4.12 NCC-specific Prmt4 deletion leads to reduction of bone matrix expression without
changing osteogenic fate commitment or frontal bone thickness.
Besides downregulation of migration-related ECM genes, I also observed reduction in a
fraction of bone matrix genes Bglap, Bglap2, Spp1, Ibsp, Dmp1, and transcription factor Sp7 in
osteogenic fronts of Prmt4 mutant (Figure 2.37A to F). To determine whether downregulation of
Sp7 in Prmt4-deletion osteogenic fronts is due to a reduction in Runx2 binding at the promoter
region, I performed ChIP-qPCR and revealed that enrichment of Runx2 at the Sp7 promoter
region was significantly reduced by PRMT4 depletion in MC3T3-E1 cells (Figure 2.38A and B). I
53
also noticed a set of osteogenic genes that are known to be Runx2 transcriptional targets were
not reduced in expression, such as Alpl (Figure 2.37G). When ChIP-qPCR was performed, I found
that Runx2 enrichment at the promoter region was not altered by PRMT4 depletion either (Figure
2.38C and D).
I next assessed whether osteogenic differentiation is disturbed by Prmt4 deletion in vivo,
by analyzing Runx2 and Sp7, which are important for early and late phase of osteogenic
differentiation. Runx2 is known as an early pre-osteoblast fate commitment marker, and Sp7
functions downstream of Runx2 as a marker for mature osteoblasts (Komori et al, 1997;
Nakashima et al, 2002). These two osteoblast differentiation markers showed similar expression
pattern in both groups, which are mainly at the osteogenic fronts and along the surfaces of frontal
bones. At E16.5 and P0 stages, the number of Runx2 and Sp7 positive cells at the osteogenic
fronts was not significantly altered in mutant mice when compared with control mice (Figure 2.39
and 2.40), suggesting that the number of Runx2+ and Sp7+ cells was not reduced. I further
measured bone thickness and showed that Prmt4 deletion caused no reduction of skull thickness
(Figure 2.41).
Altogether, these findings indicated that CNC-specific deletion of Prmt4 does not impair
osteogenic fate commitment, nor reduction of Sp7+ osteoblast number or frontal bone thickness,
but Prmt4 deficiency leads to a moderate reduction in the expression of Sp7 and a subset of bone
matrix genes.

54

Figure 2.37. Bone matrix genes (A-E) and Sp7 (F) were downregulated in osteogenic fronts of Prmt4
mutant while osteogenic genes expression such as Alpl was not significantly reduced (G).
*p-value ≤ 0.05, **p-value ≤ 0.01

Figure 2.38. Depletion of PRMT4 decreased Runx2 binding with its target genes. For each gene, Runx2
binding peak (Meyer et al, 2014) was shown at the top with red arrows specifying the location which is
targeted by primers (A,C) and ChIP-qPCR result shown in bar graphs below (B,D).
*p-value ≤ 0.05 compared with IgG, **p-value ≤ 0.01 compared with IgG
# p-value ≤0.05 compared with Runx2 IP of siControl

C
B
D
55

Figure 2.39. The number of Runx2 positive cells at the osteogenic fronts was not significantly altered in
Prmt4 mutant mice when compared with control mice at P0(A) and E16.5(C) (Scale bar = 100 µm).
Quantification of Runx2 positive cells at the OF was shown as a bar graph for each time point (B and D).
p-value > 0.05 (not statistically significant)

56

Figure 2.40. The number of Sp7 positive cells at the osteogenic fronts was not significantly altered in Prmt4
mutant mice when compared with control mice at P0(A) and E16.5(C) (Scale bar = 100 µm). Quantification
of Sp7 positive cells at the OF was shown as a bar graph for each time point (B and D). p-value > 0.05 (not
statistically significant)
57

Figure 2.41. NCC-specific Prmt4 deletion does not lead to reduction of frontal bone thickness at P0(A) and
E16.5(C) (Scale bar = 100 µm). Quantification of frontal bone thickness was shown as a bar graph for each
time point (B and D). p-value > 0.05 (not statistically significant)

58
2.5 Discussion
PRMT4 is a major protein arginine methyltransferase for Runx2
Previous findings from Dr. Xu’s lab showed that both PRMT3 and PRMT4 methylates
Runx2. When I generated CNCC-specific Prmt4 deletion mice, I questioned if the reduction of
Runx2 methylation from Prmt4 deletion would be compensated by upregulation of Prmt3 to
ameliorate the phenotype. I examined Prmt3 expression in PF suture with RNA in situ
hybridization and observed that Prmt3 expression level was similar between control and Prmt4
mutant. Prmt3 expression in PF suture was also significantly lower than Prmt4 expression as
demonstrated by both in situ and RNA-seq, which suggested PRMT4 as the dominant PRMT to
methylate Runx2. However, I observed about 50% reduction of methyl-Runx2 in OF by Prmt4
deletion, suggesting that endogenous PRMT3 may methylate Runx2 in the absence of PRMT4.
Further evaluation is needed to explore this possibility.
For these Prmt4 mutant mice, Exon2 and 3 were deleted (amino acid 117-187 or 349-
561bp as explained in (Yadav et al, 2003)). The Prmt4 probe used in RNAScope spanned across
316-1308 bp, so this probe can bind to the residual Prmt4 exons which can be transcribed.
Therefore, I observed a reduction instead of complete depletion of Prmt4 expression in Prmt4
mutant.
The source of migratory cells at the OF lining the PF suture
Using DiI labeling, I showed that there was a significant cell migration at the OF towards
the midline, but cell migration from suture mesenchyme towards OF was undetectable. My
findings here showed that frontal bone primordium migration acts as the dominant drive for PF
suture closure, which supports earlier studies by Yoshida et al. and Ting et al. showing that
migration of frontal bone primordium contributed frontal bone growth during early development
(E13.5) (Ting et al, 2009; Yoshida et al, 2008). My finding in the suture mesenchyme area is also
in line with the study of Lana-Elola et al., in which the labeled mid-suture mesenchymal cells
showed little movement and did not integrate into OF except a small population which located
59
adjacent to the OF (Lana-Elola et al, 2007). The possibility that SM cells adjacent to the OF are
recruited during suture formation cannot be ruled out and needs to be further investigated. To
study this possibility rigorously, DiI labeling of the OF combined with DiO labeling of the adjacent
SM and genetic lineage tracing studies are required. Holmes et al. previously reported Lrig1 and
Prxx1 as potential markers of the SM which can be utilized for lineage tracing studies of SM-
derived cell contribution to the OFs (Holmes et al, 2020).
The force that drives cell migration at the OF in PF suture closure
In the study of Lana-Elola et al., the movement of OF cells was interpreted as a
consequence of pushing force from OF cell proliferation rather than an active migration force
(Lana-Elola et al, 2007). However, in my study, Prmt4 deletion impaired > 50% of the OF cell
movement while OF cell proliferation remained unaltered, suggesting additional forces at work.
My RNA-seq analysis revealed that Prmt4 deletion downregulated genes involved in migration,
including genes encoding proteins responsible for matrix degradation and cytoskeleton
reorganization. I further demonstrated the direct role of PRMT4 and Runx2 methylation in cell
migration using MCT3T-E1 cells. Taken together, these findings suggested that PRMT4-Runx2
pathway promotes active migration as a driving force for PF suture closure and frontal bone
expansion at this stage of development. Consistent with my findings, a recent study using RNA-
seq analysis of PF suture also showed cell migration as one of the top GO involved in embryonic
PF suture closure (Holmes et al, 2020).
Regulatory role of the surrounding structures
Dura mater interacts with overlying suture and dictates its patency during embryonic stage
(Opperman et al, 1995; Opperman et al, 1996; Opperman et al, 1993; Roth et al, 1996). My
immunostaining and RNA in situ hybridization showed that PRMT4 is expressed in the dura mater.
Therefore, there is a possibility that the Prmt4-deficient dura mater may be involved in delayed
PF suture closure. However, this may not be associated with Runx2 since Runx2 expression in
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the dura mater is only slightly higher than the background. The potential role of PRMT4 in dura
mater awaits further investigation.
Postnatal role of PRMT4 in PF suture development
It was reported that Prmt4 null mice exhibit delayed endochondral ossification and PRMT4
methylates Sox9, a major regulator in this process (Ito et al, 2009). Since PF suture closure
requires endochondral ossification during postnatal period, Prmt4 deletion may impair this
process and lead to PF suture patency. Endochondral ossification occurs during postnatal period
starting around P7 (Sahar et al, 2005). Investigation at later time period is required to test this
hypothesis.
NCC-specific Prmt4 deletion does not alter cell proliferation.
Cell proliferation at the OF is one of the major drives for calvarial suture closure (Lana-
Elola et al, 2007) and Fgfr2 regulates osteogenic cell proliferation (Iseki et al, 1999). Although
PRMT4 has been shown to be required for cell proliferation in many cancer cells (Al-Dhaheri et
al, 2011; Wu et al, 2020; Zhang et al, 2017), in the osteogenic cells at this developmental stage,
loss of Prmt4 did not reduce cell proliferation as assessed by Ki67 staining. I also examined Fgfr2
expression in the RNA-seq data and found that Fgfr2 expression in the Prmt4 mutant OF was not
reduced (1.02 folds, p-value = 0.913). This is consistent with my findings that cell proliferation
was not reduced.
The impact of NCC-specific Prmt4 deletion on osteogenic differentiation
In Prmt4 mutant PF suture, the number of Runx2+ osteoprogenitors and Sp7+ osteoblasts
did not decrease, indicating that osteogenic cell fate commitment was not impaired. In addition,
frontal bone thickness was not altered either, indicating that mineralization and bone formation
was not blocked. However, RNA-seq analysis showed a moderate reduction of Sp7 expression
(1.36 folds, p-value = 0.04), suggesting that Sp7 expression among Sp7+ cells may be decreased,
as 20-30% decrease in protein expression may be not accurately reflected by immunostaining
approach. I also observed downregulation of a subset of bone matrix gene transcripts, including
61
Bglap, Bglap2, Spp1, Ibsp, and Dmp1, while other osteogenic differentiation gene expression was
not significantly altered in the osteogenic fronts of Prmt4 mutant such as Alpl (1.37 folds, p-value
= 0.07) and Runx2 (1.19 folds, p-value = 0.28). Based on these findings, I conclude that
osteogenic fate commitment was not impaired, but bone matrix deposition is slightly reduced in
the osteogenic fronts of Prmt4 mutant mice.
Osteogenic differentiation and bone matrix deposition are essential steps of
intramembranous ossification in flat bone growth (Opperman, 2000). Therefore, the reduction in
bone matrix deposition may cooperate with the impairment of cell migration and contribute to PF
suture patency.
Runx2 methylation by PRMT4 specifically affects PF suture closure among NCC-derived
structures
Shirai et al (Shirai et al, 2019) generated NCC-specific Runx2 deletion and observed that
in heterozygous deletion, PF suture and palate were the only affected NCC-derived structures,
suggesting importance and unique roles of Runx2 regulation in these two craniofacial structures.
In the Prmt4 mutant model, I performed thorough histological analysis of palate morphogenesis
at E13.5~15.5 and noted no obvious alterations. However, PF suture patency phenotype was
manifested in all Prmt4 mutant mice examined. In the mandibles, although PRMT4 and Runx2
showed high levels of expression (data not shown), there was no significant alteration in
mandibular morphology. Therefore, Prmt4 deletion specifically affected PF suture and one
question that begs explanation is the reason for specificity. Since impaired cell migration was
identified as a major factor for PF suture patency, it suggested that cell migration may not be a
major factor for mandible morphogenesis.

CNCC-specific Prmt4 deletion leads to downregulation of ECM-related genes
My study showed a downregulation of ECM-related genes in Prmt4 mutant. A recent study
which performed RNA-seq in embryonic PF suture also suggested the importance of ECM-related
62
genes during PF suture development (Holmes et al, 2020). In addition, they observed a difference
in alternative splicing of ECM-genes and splicing factor genes between SM and OF. Alternative
splicing mechanisms may also contribute to the downregulation of ECM genes in Prmt4 mutant
since PRMT4 was reported in regulating alternative splicing events especially exon skipping
(Cheng et al, 2007).
Another possible cause of downregulation is a reduction of H3R17me2 and H3R26me2
upon Prmt4 deletion since these histone marks were involved in transcriptional activation (Ma et
al, 2001; Schurter et al, 2001). These possibilities need to be further investigated.

Depletion of PRMT4 decreased Runx2 binding with its target genes in calvarial
preosteoblasts.
PRMT4-targeted Runx2 arginine methylation sites locate in the transcriptional
activation/repression domain, but not the DNA-binding domain. I observed a significant reduction
of Runx2 binding with its target gene promoters. This finding suggested two possibilities. One is
that Runx2 methylation induces conformational changes that reduces the DNA binding affinity.
The second possibility is that arginine methylation controls Runx2 cofactor interaction, especially
cofactors that bind to the methylated region. Loss of PRMT4, which leads to the loss of Runx2
methylation, may decrease Runx2 interaction with co-activators such as Smad (Xu et al, 2015),
or increase Runx2 interaction with co-repressors such as HDAC (Schroeder et al, 2004), to alter
the genomic localization of Runx2. These possibilities need to be further investigated.
For ECM-related migration genes such as Timp1, Serpine1 and Pik3cb, I observed a
downregulation of these genes and significant reduction of Runx2 binding to these gene
promoters, which suggested that for these migration genes, Runx2 methylation is required for
Runx2 binding with other co-activators which recruit Runx2 to these migration gene promoters
and enhancers. However, for osteogenic genes such as Alpl, there was only a slight reduction in
63
Runx2 binding which suggested that methylation of Runx2 is not required for DNA binding but
may be required for recruiting additional co-activators to efficiently induce gene transcription.

2.6 Conclusions and Future Directions
This study shows that neural crest-specific Prmt4 deletion leads to posterior frontal suture
patency. This deletion caused a significant reduction of PRMT4-mediated Runx2 methylation and
H3R17/R26 methylation in the osteogenic fronts of posterior frontal suture, which resulted in
impaired cell migration without a significant change in cell differentiation, proliferation, or
apoptosis. In addition, I demonstrated that PRMT4 depletion or methylation-deficient Runx2
reduced cell migration in vitro suggesting that PRMT4-Runx2 pathway directly regulated
osteoblast migration. Further transcriptomic analysis showed a downregulation of genes in cell
migration/invasion pathways. This downregulation could be a result from alteration in binding of
Runx2 with its target promoters and enhancers as well as cofactors upon loss of arginine
methylation by PRMT4. Moreover, changes in transcriptional regulation from depletion of H3R17
and H3R26 methylation is another possibility. Further experiments should be performed to
validate possibilities mentioned above using ChIP-seq and biochemical approaches. In addition,
this study focused on the role of PRMT4 in PF suture during embryonic development. Its role in
postnatal period may be as significant as embryonic period and further investigation is required.

64

Figure 2.42. Summary of findings. PRMT4 methylated Runx2. Methylation of Runx2 was required for
osteoblast migration and regulated ECM-related gene transcription and these components are essential for
PF suture closure. In addition, Prmt4 deletion led to a reduction of H3R17me2 and H3R26me2 which may
downregulate ECM-related genes in PF suture.

65
Chapter 3
The Role of PRMT1 in Murine Mandibular Development
3.1 Background
3.1.1 Mandibular development
Mandible is a movable bone, which is important for speech and eating. The position of
mandible also has a high impact on airway space since the tongue, several muscles and tissues
are attached to the mandible. Retrusive mandible, which is a result of underdeveloped mandible,
is associated with malocclusion and narrow airway in infants and adults, which could lead to a
serious sleep disorder - obstructive sleep apnea (OSA) (Joshi et al, 2014; Knappe & Sonnesen,
2018).
Mandible is one of the CNCC-derived structures arising from the 1
st
pharyngeal arch. In
mice, the mandibular process is separated from the maxillary process around E10.5 and grows
outwards. Later around E12.5, Meckel’s cartilage starts to form. At E13.5, osteoprogenitors form
and differentiate adjacent to the Meckel’s cartilage. As Meckel’s cartilage grows along the
mandibular primordium on both sides, it finally reaches the mandibular midline forming a
symphysis while the other end contributes to malleus and incus bones (Parada & Chai, 2015)
(Figure 3.1). Meckel’s cartilage in the developing mandible is important in controlling the size
(Mori-Akiyama et al, 2003).

66

Figure 3.1. Scheme of mandibular morphogenesis (Parada & Chai, 2015). E = eye; T = tongue; PS =
palatal shelves; M = malleus; I = incus; MC = Meckel’s cartilage; S = symphysis

Multiple factors that regulate chondrogenic or osteogenic differentiation play roles during
mandibular morphogenesis. First, Sox9, which is the master regulator of chondrogenesis is
required for Meckel’s cartilage formation. Deletion of Sox9 leads to absent Meckel’s cartilage
(Akiyama et al, 2005). Dlx genes are also involved in mandibular patterning in which different
combinations of Dlx deletion (among Dlx1, 2, 3, 5, and 6) lead to alterations of various severities
(Depew et al, 2005). Mandible undergoes intramembranous ossification (Parada & Chai, 2015);
therefore, Runx2 and Sp7 are also involved. Runx2 deletion mice showed absent condylar or
angular cartilage although Meckel’s cartilage was formed, indicating impaired secondary cartilage
formation (Shibata et al, 2004). NCC-specific Sp7 deletion resulted in small mandibles with normal
Meckel’s cartilage (Baek et al, 2013). TGF-beta signaling is required for cell proliferation via
connective tissue growth factor (CTGF) and osteogenesis via Msx1 (Oka et al, 2007). Mice with
gain-of-function Fgfr2 mutation showed enlarged mandible and Meckel’s cartilage due to
increased proliferation of osteocytes and chondroblasts respectively (Motch Perrine et al, 2019).
Inhibition of BMP signaling is also important for normal mandibular growth since gain-of-function
67
BMP-signaling mutation mice show overgrowth and endochondral ossification of Meckel’s
cartilage instead of physiological degeneration (Wang et al, 2013). A recent study also showed
that transcription factor Meis2 is important for mandibular development via Sonic hedgehog (Shh)
pathway since NCC-specific Meis2 deletion caused mandibular defects with ectopic ossification
(Fabik et al, 2020). In addition to aforementioned differentiation genes and signaling pathways,
extracellular matrix (ECM) components are involved in mandibular development. Has2 gene,
which encodes hyaluronic acid synthase (HAS) that is responsible for hyaluronic acid (HA)
production and required for palatogenesis, was also highly expressed in the mandibular
mesenchyme including Meckel’s cartilage (Lan et al, 2019). NCC-specific Has2 deletion resulted
in short Meckel’s cartilage with reduced glycosaminoglycans (GAGs) formation (Yonemitsu et al,
2020). However, more investigations are needed to elucidate roles of ECM components in
mandibular morphogenesis.

3.1.2 Alternative splicing
Human genome contains genetic information stored in the form of DNA, which serves as
template for the production of proteins that regulate numerous functions in the human body.
Genetic information is first copied into messenger RNA (mRNA) via a process called transcription.
The mRNA molecules contain intron and exon sequences which undergo another process called
“splicing” that generates mature mRNA as a template for the translation process in order to make
proteins. There are two types of splicing which are constitutive splicing and alternative splicing
(Figure 3.2). Constitutive splicing is when introns are spliced out and exons are ligated together
in their original sequence. With this type of splicing, our body can generate only limited number
of mRNAs with limited number of genes that humans were born with. This number of mRNAs
would not be enough to generate all proteins our body needs to function. On the other hand,
alternative splicing, in which some exons are skipped during the splicing process while others are
68
included, can produce a variety of mature mRNAs. Alternative splicing was found in almost 95%
of human genes (Wang et al, 2008) and further categorized into 5 main types (Figure 3.2).
1. Mutually exclusive exons: Either of the two exons is included after the splicing.
2. Cassette alternative exon (Exon skipping): In this event, a particular exon is spliced out or
retained. This is the most frequent type of alternative splicing in mammals.
3. Alternative 3’ splice site: mRNA isoforms are generated from alternative 3’ splice site
which changes the exon junction at the 3’ end.
4. Alternative 5’ splice site: mRNA isoforms are generated from alternative 5’ splice site
which changes the exon junction at the 5’ end.
5. Intron retention (IR): Single or multiple introns are retained in mature mRNA after splicing.

Figure 3.2. Two types of splicing events – constitutive and alternative splicing (Wang et al, 2015).
Alternative splicing is further categorized into 5 major types. Grey blocks represent exons separated by
introns represented in black solid lines. Dashed lines indicate splicing events.

Alternative splicing events are controlled by cis- and trans-acting elements. Cis-acting
elements, including splicing enhancers and silencers, are areas located on pre-mRNA sequences
recognized by trans-acting elements, which are splicing factors, to activate or suppress splicing.
69
Various combinations of cis- and trans-elements affect spliceosome assembly and the outcome
of splicing event (Wang et al, 2015; Wang & Burge, 2008). Epigenetic factors such as DNA
methylation and histone methylation also affect the recruitment and activity of splicing factors
(reviewed by (Monteuuis et al, 2019)).

3.1.3 Intron retention during development
Intron retention is more common and widely studied in fungi and plants than in mammals.
However, improved technology in sequencing and bioinformatic analysis have brought attention
to the role of intron retention in mammals (Braunschweig et al, 2014; Grabski et al, 2021). A study
by Middleton R. et al. emphasizing IR in mammals showed that retained introns are involved in
more than 80% of genes regulating cell differentiation and cell cycle (Middleton et al, 2017).
Recently, more studies have been published revealing the importance of IR in neuronal
development (Mauger et al, 2016; Yap et al, 2012) and differentiation process in the
hematopoietic lineage (Ni et al, 2016; Ullrich & Guigó, 2020; Wong et al, 2013). In addition, intron
retention was identified in cancers (Dvinge & Bradley, 2015) and cellular response to
environmental stimuli (Shalgi et al, 2014). Intron retention was found to control gene expression
in a cell-type and developmental stage-specific manner. It contributes to the tuning of mammalian
transcriptome, with higher prevalence in low expression genes that are associated with
developmental processes (Braunschweig et al, 2014).
Retained intron can be stored in the nucleus or exported to the cytoplasm. Both events
then lead to multiple consequences depending on several factors as reviewed by Jacob and Smith
below (Figure 3.3).
- IR causes mRNA storage within the nucleus and stabilizes mRNA for a delayed signal-
induced splicing. These are called detained introns (Boutz et al, 2015) (Figure 3.3A).
- IR causes mRNA retention within the nucleus and degradation (Figure 3.3B).
70
- The mRNA with retained intron is exported to cytoplasm. However, the retained intron
contains premature termination codon (PTC) which triggers nonsense-mediated decay
(NMD) to degrade mRNA (Braunschweig et al, 2014) (Figure 3.3C).
- The retained intron is located at 3’ untranslated region (UTR). When the mRNA is exported
to the cytoplasm, retained intron at the 3’UTR will affect mRNA stability and translational
efficiency (Thiele et al, 2006) (Figure 3.3D).
- When mRNA with retained intron at the 5’UTR is exported to the cytoplasm, retained intron
may activate or repress mRNA translation (Weatheritt et al, 2016) (Figure 3.3E).
- Some mRNA with retained intron is exported to cytoplasm and translated to generate
different protein isoforms (Braunschweig et al, 2014). In this case, it is difficult to define
whether retained intron is an intron or alternative exon (Figure 3.3F).

Figure 3.3. Fates of Intron Retention (Jacob & Smith, 2017).
A
B
C
D
E
F
71
In brief, intron retention controls gene expression and is well-conserved among species
(Schmitz et al, 2017). The most frequent consequence of intron retention identified so far is
associated with a decreased level of protein production via Nonsense-Mediated Decay
(NMD)(Braunschweig et al, 2014; Middleton et al, 2017).
Protein synthesis involves a series of processes from transcription to translation in which
earlier steps influence later events. Any error occurring along the way will be passed on and may
aggravate to severe problems. Therefore, a quality control is needed in every single step to
promptly detect and remove the errors (Isken & Maquat, 2007). Premature termination codon
(PTC) is an abnormal stop codon located within the mRNA. The majority of PTC leads to
translational stall and degradation of the mRNA in a process called “Nonsense-mediated decay
(NMD)”, although a small fraction of PTC can induce abnormal splicing called “Nonsense-
associated altered splicing (NAS)” (Dahlberg & Lund, 2004; Hentze & Kulozik, 1999).
NMD is the process to detect spliced mRNAs which contain an error – a premature stop
codon (PTC) and degrade these mRNAs, thus preventing the translation of truncated or
nonfunctional protein which may cause harm once accumulated (Cheng & Maquat, 1993; Hentze
& Kulozik, 1999). In mammals, NMD machinery can either start as early as when mRNAs are
newly-spliced in the nucleus, or it can act on spliced mRNAs that are exported to the cytoplasm.
Both functions are supported by multiple studies (Dahlberg & Lund, 2004; Iborra et al, 2004; Schell
et al, 2002). The NMD machinery are formed by multiple components including, UPF proteins
(UPF1, UPF2, UPF3, UPF3X), SMG proteins (SMG1, SMG5, SMG6, SMG7, SMG8, SMG9,
SMG10) and the newly identified RUVBL1, RUVBL2, and RPB5 (Conti & Izaurralde, 2005; Izumi
et al, 2010; Maquat, 2004; Yamashita et al, 2009).

72
3.1.4 The importance of IR and NMD pathway in craniofacial development
Human and mouse genetic mutations in several important regulators of IR and critical
components of NMD pathway have been associated with craniofacial deformities. For example:
1. ZMYND11 (or BS69), bound to the chromatin via H3.3K36me3, interacts with spliceosome
protein EFTUD2 to destabilize the spliceosome complex (Guo et al, 2014) and represses
RNA Pol II elongation. Mutation of ZMYND11 in human leads to developmental delay and
several craniofacial abnormalities such as prominent forehead and high palate (Tumiene
et al, 2017).
2. Eftud2 is a spliceosome protein. Mutation of Eftud2 leads to intron retention and exon
skipping which triggers NMD (Lei et al, 2017). Human with EFTUD2 mutation present with
Mandibulofacial Dysostosis with Microcephaly (MFDM) (Lines et al, 2012).
3. Smg1 is a key component of the NMD complex, as the kinase activity of Smg1 is required
for the catalytic activation of multiple components in the NMD machinery. Mutation of
Smg1 therefore contributes to an accumulation of PTC-containing transcripts as a result
of impaired NMD (McIlwain et al, 2010). Mice with loss-of-function Smg1 mutation show
prognathic mandible and overgrown incisors (Roberts et al, 2013).
4. Smg9 is a subunit of the SMG1 complex critical for NMD machinery (Yamashita et al,
2009). SMG9 mutation in human leads to several craniofacial defects such as
micropthalmia, cleft lip, cleft palate, hypertelorism. Mice with Smg9 mutation also show
malformed head and eyes (Shaheen et al, 2016).
These findings suggest important physiological functions of IR and NMD pathway in
normal craniofacial development and genetic perturbation of IR or/and NMD pathway may cause
craniofaical defects.

73

Figure 3.4. Examples of proteins involved in IR-NMD pathway mentioned in the text above: Zmynd11(A),
Eftud2(B), Smg1(C), and Smg9(D).

3.1.5 PRMT1 and craniofacial development
PRMT1 is a type I PRMT which is responsible for the majority of protein arginine
methylation in mammals (Tang et al, 2000). PRMT1 is important in multiple cellular processes
through methylation of histone and non-histone proteins, including transcriptional regulation via
methylating H4R3 (Strahl et al, 2001; Wang et al, 2001), DNA damage response via methylating
MRE11(Boisvert et al, 2005b) and 53BP1 (Boisvert et al, 2005c), cell survival via methylating
FOXO which is a key regulator for cell survival downstream of PI3K-AKT pathway (Yamagata et
al, 2008). PRMT1 methylates hnRNPs and other RNA binding proteins to affect their subcellular
localization and activity in mRNA splicing (Boisvert et al, 2005a; Côté et al, 2003; Hartel et al,
2019; Ostareck-Lederer et al, 2006). PRMT1 also regulates signaling pathways including BMP
signaling (Xu et al, 2013) and Wnt signaling (Bikkavilli & Malbon, 2012; Cha et al, 2011). In
addition, PRMT1 is involved in several types of cancers including breast cancer, prostate cancer,
lung cancer, colon cancer, and leukemia (reviewed by (Yang & Bedford, 2013)).
74
Global Prmt1 null mice die early during embryogenesis before E7.5 (Pawlak et al, 2000;
Yu et al, 2009) which suggests its significance during development. Tissue-specific deletion of
Prmt1 demonstrated roles in heart development (Jackson-Weaver et al, 2020), neuronal
development (reviewed by (Hashimoto et al, 2021)), and inflammatory response of the periodontal
tissue (Zhang et al, 2018).
The role of PRMT1 in craniofacial development was assessed by Dr. Yongchao Gou in
Dr. Xu’s lab. He showed that NCC-specific Prmt1 deletion leads to multiple craniofacial
deformities including a small head, cleft palate, alveolar bone and incisor defects, and small
mandibles (Gou et al, 2018a; Gou et al, 2018b) (Figure 3.5). RNA-seq analysis from palatal
samples of control and Prmt1 conditional knockout (CKO) mice showed a downregulation of
osteogenic genes and an upregulation of chondrogenic genes which suggests that PRMT1 may
regulate cell fate switch in CNCC (Gou Y, unpublished data) (Figure 3.6). However, molecular
and cellular mechanisms by which PRMT1 regulate mandibular morphogenesis was not clear.
Therefore, my study will focus on the mandibles.

Figure 3.5. NCC-specific Prmt1 deletion leads to multiple craniofacial defects including a small head (B,
H,J), cleft palate (D,F), and small mandibles (L) (Gou et al, 2018a; Gou et al, 2018b).

75

Figure 3.6. RNA-seq analysis of Prmt1 mutant palates showed a downregulation of osteogenic genes and
an upregulation of chondrogenic genes (Gou Y, unpublished data).

3.1.6 PRMT1 and mRNA splicing
Many RNA binding proteins and splicing factors harbor RGG or RG motifs in their
sequence, which are recognized by PRMT1 for arginine methylation, which affects their
subcellular localization and activity in mRNA splicing (Boisvert et al, 2005a; Côté et al, 2003;
Hartel et al, 2019; Ostareck-Lederer et al, 2006). A recent study by Olan et al. also showed that
Prmt1 depletion in embryonic ventricular epicardial cells (MEC1) led to an increase in alternative
splicing events (Jackson-Weaver et al, 2020) (Figure 3.7). In addition, a study by Dr. Graham’s
group explored PRMT1 targets using PRMT1-depleted 293T cells and mass spectrometry
analysis. There are multiple splicing proteins which were found to be PRMT1-specific targets
since they showed decrease of dimethylation when PRMT1 is depleted (Hartel et al, 2019). These
proteins included EWSR1, G3BP1, HNRNPA1, SFPQ, TAF15, TRA2B, and WDR70 (Figure 3.8).
These findings identified splicing factors that are arginine methylated in a PRMT1-dependent
manner.
Upregulation of chondrogenic genes Downregulation of osteogenic genes
76
Figure 3.7. Increase in alternative splicing event upon PRMT1 depletion in MEC1 cells (Jackson-Weaver
et al, 2020). Each bar graph represents the number of alternative splicing events which increase in PRMT1
siRNA (with or without TGF-beta) when compared with control group. SE = skipped exon; RI = Retained
intron; MXE = Mutually exclusive exon; A5SS = Alternative 5’ splice site; A3SS = Alternative 3’ splice site

Figure 3.8. PRMT1-specific targets identified by Mass Spectrometry analysis (Hartel et al, 2019).
Combination of chemical and immunoaffinity approaches were performed. Methylation level of PRMT1
shRNA group compared with control shRNA was shown as Log2 fold change. Bar graph with black outline
indicates FDR q-value < 0.1. (SCX = strong cation exchange; IAP = immunoaffinity purification; DMA =
dimethylarginine; SDMA = symmetric dimethylargine; ADMA = asymmetric dimethylarginine; MMA =
monomethylarginine)

3.2 Hypothesis
I hypothesize that neural crest-specific Prmt1 deletion increases intron retention which
affects mandibular development.
77
3.3 Materials and Methods

3.3.1 Animals
Wnt1-Cre and R26R
tdTomato
mice were purchased from Jax (#009107 and #007914).
Prmt1
fl/fl
mice were kindly provided by Stephanie Richard (McGill University). All animal care and
experiments were carried out in accordance with protocols approved by the Institutional Animal
Care and Use Committee (IACUC) at the University of Southern California.

3.3.2 Bulk RNA-sequencing analysis
Mandibles from mouse embryos from control (Wnt1-cre; Td/Td) and Prmt1 mutant (Wnt1-cre;
Prmt1
fl/fl
; Td/Td) groups at E13.5 were dissected in HBSS under dissecting microscope. Tongue
tissues were removed. Tissues were chopped and digested in TrypLE solution (Thermo Fisher
Scientific Cat# 12605010) with rotation in 37°C chamber for 30 minutes. After that, 10%FBS was
added to stop the digestion. The supernatant was transferred to collecting tube with 40-µm cell
strainer. Cells were collected by centrifuging at 500 rcf for 5 minutes and were resuspended in
PBS. DAPI was added to each sample to help exclude dead cells during fluorescence activated
cell sorting (FACS). After cell sorting, only tdTomato positive - neural crest cells were collected.
RNA was extracted with RNeasy Mini Kit (Qiagen Cat# 74104). NCC from three control and two
mutant mandibles were sequenced with the Illumina HiSeq 4000 at 40 million sequencing depth
and 100 bp paired end sequencing. Partek Flow (Partek Inc.) was utilized for analysis. Reads
were trimmed from both ends based on a quality score with minimum read length of 25. Then,
reads were aligned with STAR to Mus musculus (mouse) – mm10 assembly and GENCODE
Genes – release M23 aligner index. Next, reads were quantified to annotation model (Partek E/M).
Noise reduction filter and normalization were applied. GSA was chosen to generate differential
analysis comparing between Prmt1 mutant and control group. Upregulated and downregulated
genes were analyzed for pathways involved and Gene Ontology (GO) with Ingenuity Pathway
78
Analysis (Qiagen Inc.) and Metascape (metascape.org) (Zhou et al, 2019) respectively. For
alternative splicing analysis, reads were re-aligned with STAR, mapped with GENCODE M23
annotation, and analyzed with rMATS. The parameters used for splicing analysis are FDR < 0.05
and p-value < 0.05. Altogether, two RNA samples from a pool of three control OFs and two mutant
OFs were sequenced and analyzed.

3.3.3 Intron Retention Analysis
Intron Retention events were analyzed by our collaborator, Dr. Weiqun Peng at George
Washington University using IRTools to calculate the Intron Retention Index (IRI). IRI is a read
density of shared intronic region divided by a read density of shared exonic region (Ni et al, 2016).
Genes with low expression (RPKM < 1) were discarded from the analysis.

3.3.4 qRT-PCR to validate RNA-sequencing results
Downregulated ECM-related genes with a fold change ≥ 1.5, p-value ≤ 0.05, and
increased intron retention were selected for qRT-PCR validation. For each gene, the retained
intron was chosen based on the following criteria.
1) The intron which show high/the highest IRI ratio between mutant and control
2) The intron with IRI at and above 0.004 in mutant group
3) The intron which is shared among isoforms.
Once the candidate intron was chosen, 2 sets of primers were designed as explained in
previous publication (Wong et al, 2013) (Figure 3.9). The first set of primers was designed to
amplify the region that showed the highest intron reads in the specific intron and aimed to detect
the retained intron. The second set of primer was designed to span the adjacent exons on either
side of the chosen intron and aimed to detect the spliced transcript. Primers were designed using
Primer BLAST (NCBI) for a product size range of 50-120 bp, with the least unintended targets,
the least self-complementarity, and the least Tm difference between forward and reverse primers.
79
RNA was extracted from samples harvested as described in 3.3.2. Complementary DNA
(cDNA) was constructed from RNA samples with Maxima H Minus cDNA Synthesis Master Mix
with dsDNase (Thermo Scientific Fisher Cat#M1681). The quantification of mRNA was performed
using PowerUP SYBR Green Master Mix (Thermo Fisher Scientific Cat# A25743). The mRNA
expression was normalized with GAPDH. Relative quantification was calculated using DDCt
method (Pfaffl, 2001).

Figure 3.9. A scheme shows how primers are designed (modified from (Wong et al, 2013)). After splicing,
a majority of transcripts in control group were predicted to be correctly spliced while a majority of transcripts
in Prmt1 mutant group were predicted to show retained intron. Two sets of primer are designed to detect
sequences located in retained intron (1) and sequences located at the junction of two adjacent exons from
correctly spliced mRNA (2). Set (1) primer will be called intron primer and set (2) primer will be called exon
primer.

3.3.5 Cell Culture and siRNA transfection
Mouse stromal mesenchymal cells (ST2) were cultured in RPMI1640 medium (Genesee
Scientific Cat# 25-506) supplemented with 10% FBS in incubator until they reached confluence.
Then, cells were split in a cell density which would reach about 70-80% confluence the next day
for reverse transfection with control siRNA (Qiagen Cat# 1027310), hnRnpa1 siRNA #1 (Qiagen
Cat# SI01068207), hnRnpa1 siRNA #3 (Qiagen Cat# SI01068221), Srsf1 siRNA #2 (Qiagen Cat#
SI01415890), Srsf1 siRNA #3 (Qiagen Cat# SI01415897), Sfpq siRNA #5 (Qiagen Cat#
80
SI05783848), Sfpq siRNA #9 (Qiagen Cat# SI05783876), Ewsr1 siRNA #1 (Qiagen Cat#
SI00997115), Ewsr1 siRNA #2 (Qiagen Cat# SI00997122), G3bp1 siRNA #3 (Qiagen Cat#
SI00207830), G3bp1 siRNA #5 (Qiagen Cat# SI02692858), Taf15 siRNA #1 (Qiagen Cat#
SI01440775), Taf15 siRNA #3 (Qiagen Cat# SI01440789), Tra2b siRNA #1 (Qiagen Cat#
SI01415911), Tra2b siRNA #4 (Qiagen Cat# SI01415932), Wdr70 siRNA (Qiagen Cat#
SI01211147), and Wdr70 siRNA #1 (Qiagen Cat# SI02854775) at 40 nM using Lipofectamine
RNAiMax transfection reagent (Invitrogen) for siRNA delivery (For each gene, 2 siRNAs were
combined). Cells were cultured for 48 hours before being harvested for RNA extraction with
TRIzol (Thermo Fisher Scientific Cat#15596018).

3.3.6 Statistical analysis
Presented data were mean ± SE from at least 3 independent experiments. Comparison was
analyzed using t-test. Statistical significance was considered at P ≤ 0.05.

3.4 Results

3.4.1 PRMT1 regulates alternative splicing events in mouse mandible
For alternative splicing analysis, rMATS was used to quantify the five types of splicing events:
skipped exon (SE), alternative 5’ splice site (A5SS), alternative 3’ splice site (A3SS), mutually
exclusive exons (MXE), and retained intron (RI). The results, plotted in a bar graph, showed an
overall increase in splicing events in Prmt1 mutant mandible by a total of 2410 events (Figure
3.10). Among these, skipped exon exhibited the highest number which were 1,443 events. The
second highest was retained introns which were 545 events, accounting for 22.6% of all increased
alternative splicing events. Alternative 3’ and 5’ splice site events were almost similar in number,
183 and 164 respectively. Mutually exclusive exons increased the least, by 75 events.

81

Figure 3.10. Increased alternative splicing events in NCC-specific Prmt1 deletion mandibles. Bar graph
represents the number of alternative splicing events which increase in Prmt1 CKO when compared with
control group. SE = skipped exon; A5SS = Alternative 5’ splice site; A3SS = Alternative 3’ splice site; MXE
= Mutually exclusive exon; RI = Retained intron

3.4.2 PRMT1 represses intron retention on a genomic scale
When analyzed further, only IR showed asymmetric response (Braunschweig et al, 2014),
because retained introns in Prmt1 mutant mandibles predominantly increased, while the other
four types of alternative splicing events exhibited roughly half increase and half decrease.
Asymmetry is a typical feature of coordinated IR program (Braunschweig et al, 2014), indicating
PRMT1 as an endogenous regulator of IR.
IR is then analyzed by IRTools through collaboration with Dr. Weiqun Peng’s group.
During their analysis, Intron Retention Index (IRI) is used as a read density of shared intronic
region divided by a read density of shared exonic region. Therefore, higher IRI refers to higher
intron retention. The bar graph showed IRI distribution for the whole genome for control and Prmt1
mutant mandible. As shown in the graph, IRI distribution of Prmt1 mutant mandible shifted to the
right side (Figure 3.11). This result suggested an increase of median IRI in the whole genome
when Prmt1 was deleted.

82

Figure 3.11. PRMT1 represses intron retention on a genomic scale. Genes of the whole genome were
categorized based on IRI and were plotted in this graph. Density of relative frequency indicates frequency
of genes that display the corresponding Log2 IRI value. IRI distribution for the whole genome of Prmt1
mutant mandible (Red) shifted to the right side of bar graph when compared with control (Blue), suggesting
an increase of median IRI in the whole genome when Prmt1 was deleted.

3.4.3 Intron retention leads to changes in mRNA level
When intron retention analysis data was overlapped with differential gene expression, IRI
was increased (IRI > 0.001, which represents the right-sided half of the mutant plot in Figure 3.11)
in 10,403 genes. Within these increased IRI genes, there were 2,853 downregulated genes
(27.4%) and 1,546 upregulated genes (14.9%) while the remaining genes showed no change
(Figure 3.12). These findings suggested that intron retention leads to changes in transcripts.
Figure 3.12. Intron retention leads to changes in mRNA level. Among increased IRI genes, 27.5% showed
decreased transcripts, 15% showed increased transcripts, while the remaining remained unchanged.

83
3.4.4 Downregulated genes which show increased IRI are involved in extracellular matrix
With 1,011 genes which displayed more than 1.5-fold downregulation of expression in
Prmt1 mutant, there were 973 genes (96.2%) which showed increased IRI (Figure 3.13A). When
I analyzed this set of genes which were downregulated and showed increased IRI for Gene
Ontology (GO), I found that extracellular matrix organization and cell adhesion were the most
enriched categories of downregulated genes (Figure 3.13B).

Figure 3.13. Most of downregulated genes (96.2%) showed increased IRI (A). GO Analysis revealed that
these downregulated genes with increased IRI in Prmt1 mutant were mainly involved in ECM and cell
adhesion (B).

A
B
84
3.4.5 NCC-specific Prmt1 deletion leads to an increase in intron retention of ECM-related
genes in mandible
A set of 32 ECM-related genes were chosen to be validated based on IRI analysis. The
majority of selected genes, which were 23 genes, showed increased intron-retaining transcripts
(IR transcripts) and decreased spliced transcripts (which was indicated by decreased flanking
exon junction) in Prmt1 mutant when compared with control (Figure 3.14 and 3.15). There were
14 genes which showed significant increased IR transcripts (>1.2 fold) and decreased spliced
transcripts: Tnn, Col14a1, Matn2, Scara5, Dcn, Kitl, Loxl1, Eln, Adamts12, Adam12, Lox,
Adamts2, Nfix, and Cd44 (Note that only intron primer was tested with Cd44 since I could not
successfully amplify the exon specific to the sequence) (Figure 3.14A to C). Within this group of
genes, fold differences were statistically significant (p-value ≤ 0.05) in 6 genes which were Tnn,
Matn2, Scara5, Loxl1, Adamts12, and Lox. There were 9 genes which showed slightly increased
IR transcripts and decreased spliced transcripts in Prmt1 mutant which were Ibsp, Ltbp4, Fosl2,
Bgn, Cd34, Alpl, Mmp14, Adamts17, and Fbln2 (Figure 3.15A and B).
However, there were 8 genes which showed decreased intron reads and transcript on the
selected loci: Dpt, Htra1, Col16a1, Thbs2, Mmp2, Col6a1, Emilin3, and Ltbp3 (Figure 3.16A and
B). On the other hand, Col6a2 is the only gene which showed an increase in both IR transcripts
and spliced transcripts (Figure 3.17).
I also included Cdkn2c, which is a cell cycle regulator since it also showed increased IRI
and might be involved in cell proliferation during mandible development. This gene showed
increased IR transcripts and decreased spliced transcripts similar to the bioinformatics analysis
result (Figure 3.14A).
Smg1 showed increased IRI but upregulated transcripts. However, I included this protein
to the validation since it is involved in nonsense-mediated decay (NMD) pathway and might
represent IR-induced delayed splicing, another important function of IR during development.
85
Results from the qRT-PCR validation showed increased IR transcripts and spliced transcripts in
Prmt1 mutant which confirmed findings from the bioinformatics analysis (Figure 3.17).
Genes that were validated to show significantly increased IR transcripts and decreased
spliced transcripts in Prmt1 mutant, such as Tnn, Matn2, and Cdkn2c were used as IR indicators
for further study to uncover downstream mediators of PRMT1 that control intron splicing.

Figure 3.14A. Genes which showed increased IR transcripts and decreased spliced transcripts in Prmt1
mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control

**
**
*
*
*
**
**
**
A
86

Figure 3.14B. Genes which showed increased IR transcripts and decreased spliced transcripts in Prmt1
mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control
Figure 3.14C. Genes which showed increased IR transcripts and decreased spliced transcripts in Prmt1
mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control

**
**
*
**
*
**
B
*
*
** **
**
**
C
87

Figure 3.15A. Genes which showed slightly increased IR transcripts and decreased spliced transcripts in
Prmt1 mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control

Figure 3.15B. Genes which showed slightly increased IR transcripts and decreased spliced transcripts in
Prmt1 mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control
**
*
**
**
A
**
*
**
**
B
88

Figure 3.16A. Genes which showed decreased IR transcripts and decreased spliced transcripts in Prmt1
mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control

Figure 3.16B. Genes which showed decreased IR transcripts and decreased spliced transcripts in Prmt1
mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control
**
*
* **
**
A
**
**
**
**
**
B
89

Figure 3.17. Genes which showed increased IR transcripts and increased spliced transcripts in Prmt1
mutant. *p-value ≤ 0.05 when compared with control; **p-value ≤ 0.01 when compared with control

3.4.6 Increased intron retention may play a major role in gene repression via Nonsense-
Mediated Decay (NMD) pathway
Among the top 20 downregulated genes, there were 18 genes which showed increased
IRI. I analyzed the sequence of all 18 transcripts and observed a great number of premature
terminal codons (PTC) in retained introns of all these genes (Table 3.1). PTC can trigger
nonsense-mediated decay process and eventually lead to mRNA degradation. Therefore, this
finding suggested that intron retention may play a major role in gene repression by degrading
mRNA via NMD pathway. Further experiments need to be performed in order to support this
hypothesis.

*
*
90
Table 3.1. Number of PTC found in retained intron of 18 genes which showed downregulation of transcript
and increased IRI

No. Gene Isoform Retained Intron No. Number of PTC
1 Dpt NP_062733.1 (201aa) 3 66
2 Ibsp NP_032344.2 (324aa) 4 71
3 Abca8a NP_694785.3 (1619aa) 25 40

26 130

30 26

31 79
4 Tnn NP_808507.2 (1560aa) 12 27

13 26

14 62

21 39
5 Col14a1 NP_001355351.1 (1797aa) 14 286

40 269

42 69

44 46

45 269
6 Kitl NP_038626.1 (273aa) 1 568

2 222

6 112

9 (3'UTR) 123
7 Htra1 NP_062510.2 (480aa) 1 341

3 246
8 Aspn NP_079987.2 (373aa) 1 (5'UTR) 139

2 36

5 87
9 Scara5 NP_083179.2 (491aa) 2 286

3 600

4 105

5 26

7 256

8 28
10 Ccdc3 NP_083080.1 (273aa) 2 1460
11 Atp1b1 NP_033851.1 (304aa) 2 130

4 30

91
Table 3.1(Continued). Number of PTC found in retained intron of 18 genes which showed downregulation
of transcript and increased IRI

No. Gene Isoform Retained Intron No. Number of PTC
12 Matn2 NP_001345709.1 (956aa) 1 (5'UTR) 156

2 437

3 179

4 370

5 164

6 148

8 139

10 204

11 20

12 39

13 40

14 45

16 5

18 44
13 Acta2 NP_031418.1 (377aa) 1 (5'UTR) 49

3 61

7 16

8 18
14 Fndc1 NP_001074885.2 (1781 aa) 1 331

3 66

12 95

16 58

17 60

18 26

19 9
15 Emc10 NP_932108.3 (258aa) 2 25

3 4
16 Mgmt NP_032624.1 (211aa) 1 (5'UTR) 870

2 2055

3 603

4 90
17 Nppc NP_035063.1 (126aa) 2 (3'UTR) 40
18 Loxl1 NP_034859.2 (607aa) 1 186

2 37

6 28

92
3.4.7 Depletion of splicing factors regulated by PRMT1 leads to increased intron retention
of Tnn in vitro
Deficiency in splicing factor activity has been documented to be a cause of intron retention.
To determine whether splicing factors regulated by PRMT1 are involved in intron retention of
ECM-related genes, I started with in vitro experiments using ST2 mesenchymal stromal cell line.
First, I chose two independent siRNAs for each of the eight splicing proteins, which are hnRNPA1,
Srsf1, Sfpq, Ewsr1, Taf15, G3bp1, Tra2b, and Wdr70, tested these siRNAs in ST2 cells and
confirmed that they induced efficient knockdown (Figure 3.18). Then, to determine whether any
of these splicing factors is responsible for intron splicing, I depleted splicing factors individually
and examined whether loss of any splicing factor(s) increases intron retention using three genes
Tnn, Matn2, and Cdkn2c as indicators.

Figure 3.18. qRT-PCR to confirm knockdown efficiency of siRNA. *p-value ≤ 0.05 when compared with
siControl group.

93
I observed increased IR transcripts and decreased spliced transcripts in Tnn when
hnRnpa1, Sfpq, Taf15, and G3bp1 is depleted (Figure 3.19A and B), but none of the splicing
factor depletion leads to increased IR transcripts with decreased spliced transcripts (Figure 3.20A
and B) and Cdkn2c (Figure 3.21A and B).

Figure 3.19A. Fold change of Tnn intron-retaining transcripts upon splicing factor depletion in ST2 cells.
*p-value ≤ 0.05 when compared with siControl group, **p-value ≤ 0.01 when compared with siControl group.

Figure 3.19B. Fold change of Tnn spliced transcripts upon splicing factor depletion in ST2 cells.
*p-value ≤ 0.05 when compared with siControl group, **p-value ≤ 0.01 when compared with siControl group.
A
B
94

Figure 3.20A. Fold change of Matn2 intron-retaining transcripts upon splicing factor depletion in ST2 cells.
*p-value ≤ 0.05 when compared with siControl group

Figure 3.20B. Fold change of Matn2 spliced transcripts upon splicing factor depletion in ST2 cells.
*p-value ≤ 0.05 when compared with siControl group

A
B
95

Figure 3.21A. Fold change of Cdkn2c intron-retaining transcripts upon splicing factor depletion in ST2 cells.
*p-value ≤ 0.05 when compared with siControl group, **p-value ≤ 0.01 when compared with siControl group

Figure 3.21B. Fold change of Cdkn2c spliced transcripts upon splicing factor depletion in ST2 cells.
*p-value ≤ 0.05 when compared with siControl group, **p-value ≤ 0.01 when compared with siControl group

A
B
96
3.5 Discussion
NCC-specific Prmt1 deletion leads to an increase in intron retention of ECM-related genes
in mandible
Bioinformatics analysis demonstrated increased intron reads and decreased mRNA
transcript in large set of ECM-related genes in Prmt1 mutant when compared with control. I further
validated these findings in the majority of selected ECM genes, which conformed to my hypothesis
that NCC-specific Prmt1 deletion leads to IR which then leads to downregulation of transcript.
However, there were some genes which showed not only increased IR but also upregulation of
mRNA transcript. This may suggest another possibility of IR which can lead to mRNA stabilization
either in the nucleus (storage and delayed splicing) or cytoplasm (3’UTR-regulated stabilization).
In addition, some genes showed decreased retained intron and downregulation of mRNA
transcript. This may suggest that a downregulation of these genes was not a result of IR but
repressed transcription.

Increased intron retention may play a major role in gene repression via Nonsense-
Mediated Decay (NMD) pathway
I observed PTC in retained intron of all top 18 downregulated genes. A previous study
showed that retained intron with PTC mostly lead to mRNA degradation via NMD and only small
amount does not undergo degradation (Braunschweig et al, 2014). I then suggest that these
downregulated genes may be caused by NMD. Further experiments need to be conducted to
confirm that NMD is responsible for the decreased transcripts.

97
Depletion of splicing factors regulated by PRMT1 leads to increased intron retention of
Tnn in vitro
I found that a depletion of 1 out of 3 PRMT1-targeted splicing factors in ST2 cells led to
increased retained intron and decreased spliced transcript of Tnn. Not all splicing factor depletion
led to similar result suggesting several possibilities: (1) only a subset of splicing factors are
disrupted upon Prmt1 deletion, which means methylation from PRMT1 may be required for the
activity, subcellular localization, or recruitment of these splicing factors, since PRMTs were found
to regulate splicing proteins by assisting assembly of spliceosome, subcellular localization (Shen
et al, 1998), and cotranscriptional recruitment (Yu, 2011); (2) only a subset of splicing factors are
required for intron splicing. Additionally, histone modification by PRMTs can also affect alternative
splicing events by associating with splicing factors recruitment. These could be possible
mechanisms. However, there were some variations in the experiments which suggests that IR
events we observed from NCC-specific Prmt1 deletion mandibles may be specific to NCC and
confirms that repeated experiment in NCC is required to validate the hypothesis.
Another experiment to confirm if these splicing factors are responsible for IR is to generate
mutant mice with deletion of interested splicing factor and compared phenotypes and IR events
with the NCC-Prmt1 deletion mice. According to my literature review, I have not come across
studies with mutation of these splicing factors yet. This investigation will be challenging but may
be required.

98
3.6 Conclusions and Future Directions
Findings from this study show that Prmt1 deletion in murine neural crest cells leads to
increased intron retention in genomic level and a downregulation of ECM-related genes. This
gene repression could be a result of IR-mediated NMD process. However, there are several
experiments required to validate the hypothesis. These future experiments are listed as follows.
- Depletion of splicing factors in primary in murine CNCC to determine their role in intron
splicing of CNCC
- Validation of in vivo interaction and methylation of splicing factors in the developing
mandibles using PLA
- Confirmation of a reduction in extracellular matrix protein deposition in NCC-specific Prmt1
deletion mandibles with mass spectrometry, Western Blot, and immunostaining
In addition, studying IR in several developmental stages would uncover how IR regulates
mandibular morphogenesis during normal development. Furthermore, this intron retention may
not be specific to mandibular development. Further studies in the role of intron retention in other
craniofacial tissues could lead us to another mechanism for developmental control in addition to
cellular and molecular processes which we have been focused on.

99
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Abstract (if available)

Abstract Cranial sutures separate the skull bones and accommodate for brain and skull growth during embryonic and postnatal development. Genetic mutation has been associated with premature closure or patency of cranial sutures. In cleidocranial dysplasia patients with RUNX2 mutation, open metopic suture is one of the major characteristics. Runx2 is a master regulator of osteogenic differentiation and bone formation. Its activity is regulated by multiple types of post-translational modifications, including phosphorylation, acetylation and glycosylation. Previous findings identified that Protein Arginine Methyltransferase 4 (PRMT4/CARM1) methylated Runx2 at four specific arginine residues. In this study, I have uncovered their co-expression in osteoprogenitors at the osteogenic fronts of posterior frontal (PF) suture during embryonic development. Neural crest-specific Prmt4 deletion in mice caused patent posterior frontal (metopic) suture in adult and a wider gap between front bones during embryonic stages. I further demonstrated that Prmt4 deficiency delayed osteoprogenitor migration at the osteogenic fronts without disrupting cell proliferation or apoptosis. In addition, PRMT4 depletion and methylation-deficient Runx2 both impaired migration ability of mouse calvarial preosteoblasts (MC3T3-E1) cells, suggesting that PRMT4-catalyzed Runx2 methylation regulates migration in fate-committed osteoblasts. I then determined the transcriptional landscape in osteogenic fronts and revealed a significant downregulation of ECM-related genes with enrichment in the cell migration pathway. Runx2 enrichment at migration gene promoters was significantly reduced upon PRMT4 depletion, indicating that PRMT4-Runx2 signaling directly control the transcription of these migration genes. We also noted downregulation in a fraction of bone matrix genes and reduced enrichment of Runx2 at Sp7 promoter. However, the number of Runx2-positive or Sp7-positive cells at osteogenic fronts or the frontal bone thickness did not decline. Altogether, these results demonstrate that PRMT4 methylates Runx2 and this arginine methylation at osteogenic fronts is essential for the migration of osteoprogenitors during posterior frontal suture closure.

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Creator Ungvijanpunya, Nicha (author)

Core Title Protein arginine methyltransferases in murine skull development

School School of Dentistry

Degree Doctor of Philosophy

Degree Program Craniofacial Biology

Publication Date 03/25/2021

Defense Date 03/08/2021

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag calvaria,CARM1,ECM,extracellular matrix,frontal bone,intron retention,mandible,migration,OAI-PMH Harvest,osteogenic front,osteoprogenitor,posterior frontal suture,PRMT,PRMT1,PRMT4,protein arginine methyltransferase,RUNX2

Language English

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Advisor Xu, Jian (committee chair), Lu, Zhipeng (committee member), Merrill-Brugger, Amy (committee member), Stallcup, Michael (committee member), Yen, Stephen (committee member)

Creator Email dr.nichaung@gmail.com,ungvijan@usc.edu

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