University of Massachusetts Medical School eScholarship@UMMS
GSBS Dissertations and Theses Graduate School of Biomedical Sciences
2019-03-07
Understanding the Role of Prdm12b in Zebrafish Development
Ozge Yildiz University of Massachusetts Medical School
Let us know how access to this document benefits ou.y Follow this and additional works at: https://escholarship.umassmed.edu/gsbs_diss
Part of the Developmental Biology Commons, and the Developmental Neuroscience Commons
Repository Citation Yildiz O. (2019). Understanding the Role of Prdm12b in Zebrafish Development. GSBS Dissertations and Theses. https://doi.org/10.13028/f500-5006. Retrieved from https://escholarship.umassmed.edu/ gsbs_diss/1013
Creative Commons License
This work is licensed under a Creative Commons Attribution 4.0 License. This material is brought to you by eScholarship@UMMS. It has been accepted for inclusion in GSBS Dissertations and Theses by an authorized administrator of eScholarship@UMMS. For more information, please contact [email protected].
UNDERSTANDING THE ROLE OF PRDM12B IN ZEBRAFISH DEVELOPMENT
A Dissertation Presented
By
ÖZGE YILDIZ
Submitted to the Faculty of the
University of Massachusetts Graduate School of Biomedical Sciences, Worcester
in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
March 7, 2019
Interdisciplinary Graduate Program
UNDERSTANDING THE ROLE OF PRDM12B IN ZEBRAFISH DEVELOPMENT
A Dissertation Presented By
ÖZGE YILDIZ
This work was undertaken in the Graduate School of Biomedical Sciences Interdisciplinary Graduate Program Under the mentorship of
______Charles Sagerström, Ph.D., Thesis Advisor
______Claudio Punzo, Ph.D., Member of Committee
______Jaime Rivera, Ph.D., Member of Committee
______Ingolf Bach, Ph.D., Member of Committee
______Rolf Karlstrom, Ph.D., External Member of Committee
______Anthony Imbalzano, Ph.D., Chair of Committee
______Mary Ellen Lane, Ph.D., Dean of the Graduate School of Biomedical Sciences
March 7, 2019
III
ACKNOWLEDGEMENT
First, I would like to thank Dr. Charles Sagerström for his support and mentorship for all these years. I have learnt a lot from him both professionally and personally. I am very thankful to him for believing in me and giving me a chance to prove myself when I first joined his lab as a rotation student. I have learnt much from him in being critical, focused and calm at all times.
I would also like to thank my TRAC/DEC committee members for their constructive feedback and support for all these years.
Graduate school was challenging as well as being fun at times. The reason I enjoyed many aspects of it is due to the support of my lab mates and friends. I owe a special thanks to Ami, Jillian, Hulya, Numana and Elif Kamber. I could only get relieved with your friendship outside of the lab. Meanwhile, I could only go through this incredible journey of my life because I had very special lab mates- Will, Jen, Priya, Frank and Denise. Sharing highs and lows of graduate school with you was priceless due to your constant support. I should also thank graduate school to give me lifelong friends. I would have never been able to complete this work without Kaya’s helpful troubleshooting and discussions.
Finally, I would like to thank my family for their encouragement and love.
My parents and sister have always seen the worst part of my struggle and helped me to be stronger in each time. My special thanks are for my husband and my son for their unconditional love. IV
ABSTRACT
Function of the adult nervous system relies on the appropriate
establishment of neural circuits during embryogenesis. In vertebrates, the
neurons that make up motor circuits form in distinct domains along the
dorsoventral (DV) axis of the neural tube. Each domain is characterized by a
unique combination of transcription factors (TFs) that promote a specific fate,
while repressing the fates of adjacent domains. The prdm12 TF is required for
the expression of eng1b and the generation of V1 interneurons in the p1 domain,
but the details of its function remain unclear.
We used CRISPR/Cas9 genome editing technology to generate the first
germline mutants for the prdm12 gene and used this resource, together with
classical luciferase reporter assays and co-immunoprecipitation experiments, to
study prdm12b function in zebrafish. We also generated germline mutants for
bhlhe22 and nkx6.1 to examine how these TFs act with prdm12b to control p1
formation.
We find that prdm12b mutants lack eng1b expression in the p1 domain
and also possess an abnormal Mauthner cell-dependent escape response. Using cell culture-based luciferase reporter assays, we demonstrate that Prdm12b acts as transcriptional repressor, most likely by recruiting EHMT2/G9a. We also show that the Bhlhe22 TF binds to the Prdm12b zinc finger domain to form a
Bhlhe22:Prdm12b complex. However, bhlhe22 mutants display normal eng1b V expression in the p1 domain. While prdm12 has been proposed to promote p1 fates by repressing expression of the nkx6.1 TF, we do not observe an expansion of the nkx6.1 domain upon loss of prdm12b function, nor is eng1b expression restored upon simultaneous loss of prdm12b and nkx6.1.
We conclude that prdm12b germline mutations produce a phenotype that is indistinguishable from that of morpholino-mediated loss of prdm12 function. In terms of prdm12b function, our results indicate that Prdm12b acts as transcriptional repressor and interacts with both EHMT2/G9a and Bhlhe22.
However, bhlhe22 function is not required for eng1b expression in vivo, perhaps indicating that other bhlh genes can compensate for its loss during embryogenesis. Lastly, we do not find evidence for nkx6.1 and prdm12b acting as a repressive pair in the formation of the p1 domain – suggesting that prdm12b is not solely required to repress non-p1 fates, but is also needed to promote p1 fates.
VI
TABLE OF CONTENTS
ACKNOWLEDGEMENT ...... III
ABSTRACT ...... IV
LIST OF FIGURES ...... X
LIST OF TABLES ...... XII
LIST OF VIDEOS ...... XIII
CHAPTER I: INTRODUCTION ...... 1
VERTEBRATE CENTRAL NERVOUS SYSTEM DEVELOPMENT ...... 2
The neural plate formation ...... 2
Transformation of neural plate into neural tube ...... 5
Neural tube patterning...... 6
PRDM FAMILY MEMBERS ...... 15
PRDMs as gene expression regulators ...... 16
Role of PRDMs in tissue specification and differentiation ...... 17
PRDMs in nervous system ...... 20
ZEBRAFISH LOCOMOTION CIRCUITRY ...... 24
Spinal cord neurons ...... 24
Motor behaviors in zebrafish ...... 27
CONTRIBUTION OF THIS WORK TO THE FIELD ...... 31 VII
CHAPTER II: ZEBRAFISH PRDM12B ACTS INDEPENDENTLY OF NKX6.1
REPRESSION TO PROMOTE ENG1B EXPRESSION IN THE NEURAL TUBE
P1 DOMAIN ...... 33
BACKGROUND ...... 34
METHODS ...... 37
Zebrafish care ...... 37
Generation of CRISPR/Cas9 mutant zebrafish lines ...... 37
Antisense morpholino oligonucleotide injections ...... 40
In situ RNA hybridization ...... 40
Luciferase reporter assays ...... 41
Co-immunoprecipitation and Western blotting ...... 42
Immunocytochemistry ...... 42
Behavioral analysis ...... 43
Genotyping ...... 43
RESULTS ...... 44
Germ line disruption of prdm12b blocks eng1b expression in the p1 domain
...... 44
The defective escape response in prdm12b mutants is caused by disruption
of the Mauthner cell pathway ...... 58
Prdm12b acts as a repressor in vitro...... 66
Prdm12b interacts with the Bhlhe22 transcription factor and the EHMT2
methyltransferase...... 72 VIII
bhlhe22 is not required for eng1b expression in the zebrafish p1 domain ...76
prdm12b does not maintain the p1 domain by repressing nkx6.1 ...... 81
DISCUSSION ...... 91
prdm12b germ line mutants recapitulate the phenotype observed using
antisense-based approaches ...... 92
prdm12b is a bona fide transcriptional repressor ...... 93
An undefined domain persists at the p1 position in prdm12b mutants ...... 95
CONCLUSION...... 97
AUTHORS' CONTRIBUTIONS ...... 97
ACKNOWLEDGEMENTS ...... 98
CHAPTER III: DISCUSSION ...... 99
CORRELATION BETWEEN MORPHOLINO-INDUCED AND MUTANT
PHENOTYPES IN PRDM12B LOCUS ...... 101
THE FATE OF THE REMAINING P1 DOMAIN IN PRDM12B MUTANTS IS
UNDEFINED ...... 102
THE ROLE OF PRDM12B IN DORSOVENTRAL PATTERNING IS COMPLEX
...... 105
Further in vivo analysis of Prdm12b interaction partners namely Bhlhe22 and
EHMT2/G9a is required ...... 105
What are the regulatory relationships of a subset of genes in the p0-p2?.. 109 IX
THE RELATIONSHIP OF GENE EXPRESSION THROUGH THE P0-P2
DOMAIN MARKERS DOES NOT INVOLVE SIMPLE DIRECT
INTERACTIONS ...... 112
REMAINING QUESTIONS AND FUTURE DIRECTIONS OF THIS WORK .. 115
What are the prdm12b direct targets? ...... 115
Do p1-derived neurons restrict Mauthner cells to fire only once? ...... 117
CONCLUSION...... 120
APPENDIX: THE EFFECT OF PRDM12B AND BHLHE22 MISEXPRESSION
...... 122
INTRODUCTION ...... 123
METHODS ...... 123
Zebrafish care ...... 123
Zebrafish were handled as discussed in Chapter II...... 123
Zebrafish embryonic injections ...... 123
In situ RNA hybridization ...... 124
RESULTS ...... 125
Gain of function prdm12b and/or bhlhe22 does not affect eng1b expression
...... 125
DISCUSSION ...... 128
REFERENCES...... 129
X
LIST OF FIGURES
Figure 1.1. Formation of zebrafish neural tube...... 3
Figure 1.2. Regional patterning of neural tube...... 7
Figure 1.3. Schematic diagram of a neural tube cross section...... 9
Figure 1.4. Summary of gene expression along dorsoventral axis...... 14
Figure 1.5. Mechanism of touch response...... 29
Figure 2.1. Generation of germ line prdm12b mutants...... 48
Figure 2.2. Sequence of mutant prdm12b, bhlhe22 and nkx6.1 alleles...... 51
Figure 2.3. Characterization of the prdm12bsa9887 mutant...... 53
Figure 2.4. prdm12b germ line mutants lack eng1b expression in the p1 domain.
...... 56
Figure 2.5. prdm12b mutant fish display an abnormal touch evoked response. .61
Figure 2.6. Detailed analysis of the touch-evoked escape response in prdm12b mutant and wild type animals...... 63
Figure 2.7. The zinc finger domain is necessary for Prdm12b-mediated repression...... 68
Figure 2.8. Expression of GAL4DBD-Prdm12b constructs used in transfection experiments...... 70
Figure 2.9. Prdm12b interacts with Bhlhe22 and EHMT2/G9a...... 75
Figure 2.10. Generation of bhlhe22 germ line mutant...... 78
Figure 2.11. Analysis of bhlhe22 mutant zebrafish...... 80 XI
Figure 2.12. Generation of germ line nkx6.1 mutants...... 84
Figure 2.13. prdm12b does not maintain the p1 domain by repressing nkx6.1. ..87
Figure 2.14. prdm12b controls the size of the p1 domain...... 90
Figure 3.1. Regulatory relationships of a subset of genes expressed in the p0-p2 domains...... 111
Figure 3.2. Schematic diagram depicting core elements of locomotion circuitry of distinct ventral interneuron classes with transcriptional identity of p1 derived V1 interneurons ...... 119
Figure A.1. prdm12b and bhlhe22 overexpression has no overt effect on eng1 expression in zebrafish...... 127
XII
LIST OF TABLES
Table 2.1. The characteristics of CRISPRs targeting prdm12b, bhlhe22 and nkx6.1 ...... 39
Table 2.2. Sequences of oligos to generate single guide RNAs...... 39
Table 2.3. Primers used for genotyping of prdm12b, bhlhe22 and nkx6.1 mutant fish ...... 44
XIII
LIST OF VIDEOS
Video 1. Movie of wild type touch-evoked response...... 64
Video 2. Movie of prdm12b mutant touch-evoked escape response ...... 64
Video 3. Movie of prdm12b mutant touch-evoked escape response ...... 65
Video 4. Movie of prdm12b mutant touch-evoked escape response ...... 65
CHAPTER I: INTRODUCTION
2
VERTEBRATE CENTRAL NERVOUS SYSTEM DEVELOPMENT
Function of the central nervous system relies on finetuning of appropriate
neuronal connections at the right time and place. A very intricate and highly-
regulated process controls every step of neurogenesis ranging from neural plate
induction to synapse formation. Generation of regional identity within the neural
tube and specification of neurons are dependent on an elaborate genetic and
molecular program. Correct interpretation of messages relayed by the molecular program, by neural progenitors, results in differentiation of neurons, subsequent motor coordination and the formation of memories and complex behaviors. Thus,
proper timing and management of genetic and molecular cues is highly critical for
neurogenesis and the establishment of the neural circuit. Any disturbances
occurred during neural development are thought to be linked to neuropsychiatric
disorders such as autism and schizophrenia [1].
The neural plate formation
Neural induction and formation of the central nervous system starts with
the transformation of a uniform layer of the neural plate into the neural tube
(Figure 1.1). The neural plate is composed of undifferentiated ectodermal cells
formed during gastrulation. Neural features are induced in neural plate by the 3
Figure 1.1. Formation of zebrafish neural tube.
The major steps of the transformation of neural plate into neural tube.
4
effect of specialized mesodermal cells beneath the ectoderm derived from the organizer. The importance of neural inducing capacity of organizer comes from the study of Spemann and Mangold on 1924. Transplantation of amphibian dorsal tip of blastopore- organizer region- into ventral side of another embryo
caused to form second neural tube [2]. This experiment shows that organizer has the capacity to influence the fate of its neighboring tissue. Neural inducing signals coming from the organizer and its derivatives, prechordal mesoderm, include noggin, chordin, follistatin and other bone morphogenic protein (BMP) inhibitors. Fibroblast growth factors (FGFs) have also been implicated as signals required for neural induction along with repression of BMP signaling (reviewed in
[3]). The coordinated play between BMP and FGF signaling accounts for neurulation while wingless-integrated (Wnt), retinoic acid (RA) and hedgehog
pathways also contribute to the complexity of vertebrate neural plate formation.
Not only do the extrinsic factors mentioned above regulate neural
induction, intrinsic transcription factors also take part in the interplay of key
regulators in early neurogenesis. Members of SRY-box containing genes B1
(SoxB1) family constitute the basis of neural default program. In zebrafish,
sox1(a/b), sox2, sox3 and sox19(a/b) are all implicated in neural ectodermal fate
determination while Pou2/Oct4 and Sox2 are direct repressors of neural
differentiation (reviewed in [4]).
The influence of intrinsic factors cannot be considered separately from the
extrinsic ones. For example, sox3 expression is dependent on FGF signaling in 5
zebrafish [5]; however, it later directs the expression of early BMP factors, namely bmp2b/7/4 [6].
Transformation of neural plate into neural tube
In chick, mice, and amphibians shortly after neural plate has formed, the
neural groove is formed beneath the overlying of ectoderm via uplifting of neural
plate edges and eventually fusion of the two halves of the neural plate (reviewed
in [7]). The zebrafish neural tube formation differs in terms of lumen formation.
Cavitation of neural tube first forms a neural keel. Through so-called ‘secondary
neurulation’, the fish neural keel expands and transforms into the typical neural
tube (Figure 1.1). Despite the disparities in lumen formation between zebrafish
and mammals, formation of all types of neurons and their relative arrangement in
the neural tube are highly conserved (reviewed in [4]).
Formation of neural lineages follows neural tube formation. Neural progenitors divide asymmetrically or symmetrically, and eventually become committed to their particular fate during development [8]. There are both extrinsic and intrinsic factor playing important roles for neural tube regionalization and neurogenesis. Of the extrinsic factors, Sonic hedgehog (Shh) secreted from the notochord and the floor plate, and BMP signals secreted from roof plate
demarcate the ventral and dorsal regions of the neural tube, respectively.
Regulation of neurogenesis is further amplified by Notch signaling and 6
expression of basic Helix-Loop-Helix (bHLH) and homeodomain transcription factors. An imbalance in Notch signaling between cells results in pro-neural fate decisions. Activation of Notch leads to non-neural fates; however, inhibition of the
Notch signaling endorses pro-neural fates (reviewed in [3]). Neurogenic bHLH transcription machinery such as neurogenin1 and asc1, drives the primary neural network along with Notch signaling (reviewed in [4]).
Neural tube patterning
Regional patterning of the neural tube is divided into two axes: anterior- posterior (A-P) and dorsoventral (D-V). All mature neural and glial cell types arise from multipotent progenitor cell domains along both axes. Anterior-posterior axis of the neural tube is compartmentalized into four regions: forebrain, midbrain, hindbrain and spinal cord. Regionalization of the neural tube is all dependent on secreted signals from localized organizing centers such as the zona limitans intrathalamica and the isthmus secreting Shh; FGF and Wnt signals, respectively. Retinoids also direct spinal cord regionalization (reviewed in [3])
(Figure 1.2). This very complex network of signals along with different
transcriptional profiling establishes neural diversity along the neural tube.
Signals emitted from three distinct regions, pattern the D-V axis of the
neural tube. These regions are the roof plate, floor plate and notochord. The roof 7
Figure 1.2. Regional patterning of neural tube.
Dorsoventral axis is structured under the influence of BMP and Shh. Anterior- posterior axis is structed by Shh, FGF, Wnt and RA.
[3]
8 plate is a thin, single layer of cells at the dorsal tip of the neural tube. The floor plate is located at the ventral tip but is wider in size compared to the roof plate.
The region between these two tips is packed with proliferative neural progenitors
(neuroblasts are post-mitotic) which eventually differentiate into sensory neurons, interneurons, and motor neurons. The notochord, which is a mesodermal structure beneath the floor plate, is a very important patterning factor for the neural tube. The mesoderm also gives rise to somites. The effect of mesoderm on floor plate is confirmed via grafting of an extra notochord into chick embryos, which results in extra floor plate at high levels and extra motor neurons at low levels [9].
The interplay between dorsalizing and ventralizing signals coming from roof and floor plate establishes domain identity along neural tube. Shh, FGF, Wnt and BMP proteins all take part neural tube patterning (Figure 1.2).
Sonic hedgehog (Shh)
Inductive signals emerging from notochord and the floor plate generate all the ventral neurons of both the brain and the spinal cord. Secreted glycoprotein Shh is the key mediator of ventralizing signals, which acts as a concentration gradient. Shh induces V0-V3 interneuron classes and motor neurons (Figure 1.3)
(reviewed in [10]). Shh induced ventral neuron formation was studied in notochord deficient chick embryos [11], zebrafish [12] and Shh null mice [13]. 9
Figure 1.3. Schematic diagram of a neural tube cross section.
Dorsoventral domains are established by opposing concentrations of BMP and
Shh which affect progenitor gene expression and in turn establish domain identities.
10
Shh performs its ventralizing effect via regulating both Class I and Class II homeodomain transcription factors. Class II genes are activated upon high levels of Shh, whereas Class I genes which are activated by RA are negatively regulated by high levels of Shh (reviewed in [10]). The establishment of cross repression activities between Class I (Pax7, Pax3, Dbx1, Dbx2, Irx3 and Pax6) and Class II (Nkx6.1, Nkx6.2, Olig2, Nkx2.2 and Nkx2.9) transcription factors ensures the formation of discrete ventral domains along the D-V axis [14,15].
Specific domain restricted gene expression, and thus specific identities, is due to such cross repressive interactions at the domain boundaries. Three such examples are the p1/p2 domain boundary where Dbx2/Nkx6.1, the p2/pMN boundary where Olig2/Irx3 and the pMN/p3 boundary where Pax6/Nkx2.2 pairs repress the expression of each other [16-18].
Bone Morphogenic Proteins (BMPs)
Dorsalizing signals coming from epidermal ectoderm and roof plate have the ability to induce neural crest cells which are required for the initiation of neural tube formation. The key mediators of dorsal neural cell types include BMP proteins, namely Bmp4 and Bmp7 and their antagonists noggin, chordin and follistatin [19-21].
Dorsal interneuron fate, specifically dl1- dl3, is promoted by dorsal signals including Bmp2, Bmp4, Bmp5, Bmp7, Gdf6/7, Dsl1 and Activin-b. These dorsal 11
signals affect progenitor domains that express helix loop helix family transcription
factors neurogenin1/2, Mash and Math1. Exogenous treatment in chick neural plate with dorsalizing signals leads to generation of dl1 and dl3 interneurons, whereas treatment with their antagonists leads to loss of dl1-dl3 interneurons [20] and (reviewed in [7]).
Fibroblast Growth Factors (FGFs) and Retinoic Acid (RA)
Neural induction by the organizer is the very first and most crucial step in
the initiation of nervous system development. It has been recently discovered
that the organizer is a center which controls many secreted signals including
FGFs. Particularly, expression of Fgf8 during gastrulation is linked to the neural
induction along with the fact that its downregulation is required for neural
differentiation [22,23].
Somite derived RA signaling is also equally significant for the generation
of neural precursors through antagonist effect between RA/Fgf8 and regulation of
Pax6 gene in mice [24,25].
Wnts
Other extracellular signals present in roof plate are Wnt proteins which
take part in a variety of embryonic cell signaling events specifically Wnt1 and
Wnt3a (reviewed in [21]). The evidence for the involvement of Wnt signaling in 12
dorsoventral patterning came from a study showing that Wnt1 and Wnt3a null
mouse has reduced dl1 and dl2, and increased dl3 neurons [26]. Downregulation of Wnt1 and Wnt3a had been observed in BMP receptor knock-out mice, which further proves the involvement of Wnt proteins in maintaining dorsal domain
identity along the dorsoventral axis [27].
Dorsoventral domain identity
The morphogens and the transcription factors form the gene regulatory
network. This gene regulatory network will define the domain identities along the
D-V axis. Each domain along D-V axis has a unique transcription profile resulting
in a specific domain identity, giving rise to specific classes of neurons from each
region.
Induction of gene expression through Shh and BMP subdivides the neural
tube into distinct progenitor domains (Figure 1.3). Under the graded influence of
Shh, five ventral domains are formed (p0-p3). The concentration gradient of Shh manifests itself in the activation of Nkx6.1, Nkx6.2, Nkx2.2, Olig2 and repression
of Pax3, Pax6, Pax7, Dbx1, Dbx2, Irx3 [13-15,17,18,28-31]. Besides, domain restricted expression of progenitor genes is further controlled by cross repression
between neighboring domain markers in order to prevent hybrid identities. To
illustrate, olig2 and irx3 repress each other (Figure 1.4) to limit their expression at
dorsal and ventral domain boundaries, respectively [18,32]. 13
Dorsal domains (dP1-dP6) under the influence of BMP are characterized
by their expression of bHLH (Math, Mash, Ngn) and LIM (Lbx, Lmx) transcription
family of proteins [33]. Experimentation in increasing and decreasing BMP signaling has led to either decrease or increase of dorsal domain markers, respectively [27,34]. This further proves key roles of morphogens in establishing
of domain identities.
This combinatorial code for gene expression establishes eleven progenitor
domains that give rise to six dorsal sensory neurons originating from dP1-dP6,
four interneurons originating from p0-p3 and motor neurons originating from
pMN. Progenitor domains commit to their specific neuronal type later in
development with the help of gene regulatory network and ultimately giving rise
to functional neural circuits.
14
Figure 1.4. Summary of gene expression along dorsoventral axis.
Schematic representation of expression domains of progenitor and post-mitotic genes along dorsoventral axis of the neural tube.
15
Gene regulatory network for neural circuit formation is composed of many different types of transcription factor families. As mentioned above, bHLH, Class I and Class II homeodomain transcription families take part in establishment of domain identities along D-V axis of neural tube [7,10,35]. Emerging data indicates that PRDM family of transcription factors are also expressed in neural progenitor domains (Figure 1.4). Recent studies that question the roles of PRDM genes in multiple aspects of neural development revealed that early patterning of neural tube and neural differentiation are partly due to function of PRDM family members.
PRDM FAMILY MEMBERS
The name “PRDM” is derived from one of its members, called positive
regulatory domain l-binding factor one (PRD1-BF1) [36]. The PRDM family,
totaling sixteen family members, is characterized by the presence of an N
terminus PR domain, which is similar to the catalytically active SET histone
methyltransferases (HMTs), and variable number of Zinc fingers, except Prdm11,
which has no Zinc fingers. Prdm factors have been implicated in tissue specific
gene regulation and many developmental processes while a dysfunction of Prdm
family is observed in many types of cancer and diseases (reviewed in [37]).
16
PRDMs as gene expression regulators
The functional mechanism for several Prdm proteins is still under
question. In contrast to SET domains, most PR domain members lack the
conserved H/RxxNHxC motif that is essential for HMT activity [38]; however,
intrinsic HMT activity has been found in some members of Prdm proteins, namely, Prdm2, Prdm3, Prdm6, Prdm8, Prdm9, Prdm13 and Prdm16 [39-43].
Even among Prdm factors with HMT activity, mechanism of action differs. Prdm2
and Prdm8 were both shown to associate with repressive mark of H3K9me2,
whereas Prdm9 catalyzes H3K4me3 [39,40,44].
Some Prdm members overcome the necessity of intrinsic HMT activity by
recruiting histone modifying enzymes to mediate transcriptional repression or
activation depending on both time and context. Examples of such histone
modifying enzymes include G9a, SuV39H, Polycomb repressive complex 2
(PRC2), Prmt5 and Lsd1as histone methyltransferases and p300/CBP and
P/CAF as histone deacetylases [45-52] (reviewed in [53]).
Aside from interactions with chromatin modifying enzymes, some Prdm
family members require co-repressors. Binding to Groucho by Prdm1 and
recruitment of CtBP (C terminal binding protein) by Prdm1, Prdm3 and Prdm16
are examples of such interactions between PRDMs and repressors [54-61]
(reviewed in [53]). 17
Prdms as gene expression modifiers need to have access to DNA at
target gene regulatory regions. Direct DNA binding has been found only in
Prdm1, Prdm3, Prdm5, Prdm9, Prdm13, Prdm14 and Prdm16 via ChIP-seq
experiments [62-70]. Prdm members that cannot bind to DNA themselves, recruit
DNA binding proteins to gain access to the target genomic locations. For example, Prdm8 binds Bhlhb5 (also known as Bhlhe22) [71] and Prdm16 forms a complex with C/EBP-b enabling access to the regulatory regions and control of gene expression [72]. So far, only a subset of Prdm members has been fully
characterized in terms of both direct DNA binding and interaction partners.
Role of PRDMs in tissue specification and differentiation
Prdm1
Prdm1, also known as Blimp1, is the most studied member of Prdm family.
Many roles have been attributed to Prdm1 in various cell differentiation events ranging from lymphoid cell differentiation to photoreceptor formation. As the name B-lymphocyte-induced maturation protein 1 (Blimp1) implies, Prdm1 drives differentiation and maturation of B cell lineage. Bi-directional transcriptional regulation has been observed for Prdm1 when determining plasma cell differentiation by repressing anti B cell fates contrary to activating interleukin 10 in regulatory T cell lineage (reviewed in [53]). 18
Prdm1 action in the retina ensures maintenance of photoreceptor identity over bipolar cells. Chx10 (also known as Vsx2) expressing precursor cells give rise to both photoreceptors and bipolar cells depending on Chx10 up or down regulation [73,74]. Prdm1 acts as the switch for Chx10 downregulation, stabilizing photoreceptor cell fate.
Prdm5
PRDM5 locus on human genome 4q26 has been associated with several types of cancer ranging from ovarian to lung [75]. Prdm5 acts as dual gene expression regulator which functions as both repressor and activator [62]. This complicates the mechanistic role of Prdm5 in carcinogenesis. However, recent work suggested that Prdm5 is a tumor suppressor (reviewed in [37]). Studies performed in zebrafish have shown that Prdm5 negatively regulates Wnt/PCP signaling pathway [76]. Constitutively active Wnt signaling due to the lack of a negative regulator, such as Prdm5, may serve as the basis for carcinogenesis.
This may partially explain why Prdm5 inactivation is associated with human cancers.
Prdm6
Prdm6 was first identified as a gene involved in smooth muscle proliferation and named PR domain In Smooth Muscle (PRISM) [45]. Two 19 different isoforms of Prdm6 used in two independent studies in mice have been shown to take part in vascular development. One claimed that Prdm6 promotes cardiac smooth muscle proliferation by suppressing differentiation signals [45] while the other argued that a longer Prdm6 isoform active in hematovascular cell precursors, inhibits endothelial cell differentiation with the help of its intrinsic
HMTase activity and interaction with chromatin remodeling enzymes such as
H4K20 [42].
Prdm14
Prdm14 upregulation has been observed in several types of cancers namely lymphoma, leukemia and breast cancers [77-79]. Deregulation of Prdm14 mRNA in many cancer types stems from the fact that Prdm14 helps to establish embryonic stem cell pluripotency under the regulation of Oct4, Sox2 and Nanog
[80].
Prdm16
Prdm16 is one of the Prdm family member that can act as both repressor and activator depending on the context. Prdm16 is a key regulator of the switch between brown adipose tissue formation versus white adipose tissue.
Understanding the mechanism of brown versus white adipose tissue formation is important in understanding the underlying cause of obesity and weight gain. Mice 20
overexpressing Prdm16 showed resistance to obesity due to increased formation
of brown fat [81]. The underlying mechanism of brown fat formation via Prdm16
is that Prdm16 directs the activation of brown preadipocyte genes such as Pparg
and Ppargc1a after the recruitment of DNA binding protein C/EBPb [70,72,82].
Along with brown adipose tissue activation, Prdm16 and co-repressor CtBP are
recruited to the promoters of white adipose tissue genes, namely myocyte genes,
to repress white adipose tissue formation over brown fat [57]. Myocytes and
brown adipose tissue derive from the same precursors. Prdm16 acts as switch
between white versus brown adipose tissue depending on recruitment of either
CtBP or C/EBPb, respectively.
PRDMs in nervous system
Prdm13
Prdm13 is a member of transcriptional regulatory complex that specifies glutamatergic (excitatory) versus GABAergic (inhibitory) neural fate. Prdm13 is
directly regulated by Ptf1, bHLH transcription factor defining inhibitory neurons, in
order to repress excitatory neural fate by silencing Tlx1 and Tlx3 [83]. Tlx3
expression is under tight regulation by Prdm13. Not only does Prdm13 form a
complex with another transcription factor Ascl1 and inhibit Tlx3, whose
expression is driven by Ascl1 alone, but it also reduces the activity of Neurog2 on
Tlx3 activation [68,83]. 21
Prdm8
A screen for target genes of Bhlhb5 (Bhlhe22), a transcriptional repressor functioning in neural circuit development, revealed Prdm8 as a direct target gene.
Prdm8 is expressed at the p1-pMN domains of hindbrain and spinal cord. Mice lacking either interactors have shown similar neural abnormality such as mistargeting of neurons at dorsal telencephalon, unusual movement and increased scratching [71]. Further analysis of Bhlhb5 and Prdm8 uncovered that they form a repressive complex to bind at Cdh11 gene to regulate the formation of corticospinal motor neurons. The interaction between Bhlhb5 and Prdm8 is twofold. Bhlhb5 ensures DNA binding via its canonical E-box motif, whereas recruitment of Prdm8 by Bhlhb5 mediates repression [71] (reviewed in [84]).
Prdm14
A zebrafish mutant named short lighting (slg) showing shortening of CaP motor neurons was identified after Tol2 transposon-mediated gene trap screen.
Sequencing indicated that prdm14 locus was affected in slg mutants. Loss of prdm14 in zebrafish resulted in CaP axon shortening and embryonic movement defects while other motor neuron types seemed unaffected. This phenotype was largely due to decreased islet2 expression upon prdm14 downregulation. Further analysis indicated that islet2 is a Prdm14 direct target which regulates late stage 22
differentiation of CaP neurons but no other types of motor neurons [85]. Prdm14
that was already shown to bind DNA via its zinc finger domain [69], has been
shown to bind to upstream regulatory site at islet2 locus [85]. islet2 is a target of
prdm14 at CaP motor neurons, whereas islet1 has also been found to act
downstream of prdm1a in Rohon-Beard cells, mechanosensory neurons [86].
This shows that Prdm family members can direct neural cell fate at different
locations/times via the same mediators.
Prdm12
The role of Prdm12 was first addressed in the context of chronic myeloid
leukemia caused by a large deletion on human chromosome 9 including the
PRDM12 region [87,88]. In P19 carcinoma cells, retinoic acid induced Prdm12
expression demonstrates anti-proliferative effect by regulating G1 cell cycle [89].
After the reports of Prdm12 expression in nervous systems of both mouse and
zebrafish, research on Prdm12 has opened the way to understand the functional
role of Prdm12 in neurogenesis. Prdm12 was expressed in ventricular zone- p1 ventral spinal cord progenitor region, dorsal root ganglia and cranial placodes in mouse [90]. Similarly, in zebrafish, expression was seen in tegmentum,
cerebellum, hindbrain and olfactory placodes [91]. More detailed analysis of
prdm12b in zebrafish has revealed that prdm12b expression is confined to the p1 domain of hindbrain and spinal cord (Figure 1.4). Knock-down of prdm12b in 23 zebrafish via MO (anti-sense morpholinos) leads to the loss of eng1b expression in p1 domain in both hindbrain and spinal cord and the loss of eng1b+ V1 interneurons. Fish lacking prdm12b, and in turn V1 interneurons, show abnormal escape response [92]. Xenopus Prdm12 also showed identical expression pattern as in zebrafish and drives V1 cell fate by regulating en1 (eng1b) through direct repression of dbx1, nkx6.1 and nkx6.2 [93]. Regulation of eng1b by
Prdm12 was found to be G9a dependent [93] since vertebrate Prdm12 was shown to recruit G9a methyltransferase to execute H3K9me2 [89,94]. The interaction between G9a and Prdm12 can be partially explained by the lack of intrinsic histone methyl transferase activity of Prdm12. The transcriptional regulatory network governing V1 cell fate suggested that Prdm12 directly represses dbx1 and nkx6 genes in RA treated Xenopus ectodermal explants derived either from wild-type or prdm12 overexpressing embryos [93]. This idea would support that Prdm12 acts as repressor in establishing the p1 domain boundary with neighboring progenitor domains; however, such direct targets still await validation from in vivo experiments in the endogenous state.
Prdm12 has been also implicated in nociceptor neurogenesis in humans
[94,95]. Ten different homozygous mutations in PRDM12 were found in individuals with chronic insensitivity to pain (CIP). The observed point mutations were scattered along the PRDM12 gene, not confined to a particular domain.
Carriers of these mutations show insensitivity to pain, cold, heat from birth and severe injuries later in life [94]. 24
ZEBRAFISH LOCOMOTION CIRCUITRY
Vertebrate central nervous development ensures precise neural circuit formation that drives particular behaviors ranging from feeding to swimming. The complexity of molecular basis of behavior stems from the neuronal network governed by intricate genetic composition and subsequent circuit formation.
Spinal cord neurons
Anterior-posterior and dorsoventral patterning of the spinal cord is
governed by the morphogen gradients such as- caudalizing Fgf, rostralizing RA,
ventralizing Shh and dorsalizing BMP and Wnt signaling. These signaling
pathways regulate transcriptional programs specify progenitor genes within a
well-defined progenitor domain. When neural progenitor cells stop proliferating,
they exit the cell cycle, become post-mitotic and express a variety of transcription
factors determine the type of neurons they give rise to (Figure 1.3 and Figure
1.4). To date, more than 20 classes of neurons have been described in the
developing spinal cord (reviewed in [96]).
The most studied spinal cord neurons belong to the family of motor
neurons derived from the pMN domain of the spinal cord. The pMN domain is
localized ventral to the p2 domain marked by Irx3 which represses Olig2, dorsal
to the p3 domain expressing Nkx6.1 and Nkx6.2 (Figure 1.4). Cross repressive 25
actions on Olig2 result in the motor neuron specific expression of Hb9, Ngn2 and
Isl1 [15-17]. The pMN progenitor domain, marked by Olig2 expression, is
subdivided into motor columns which give rise to clusters of motor neurons
innervating a single muscle. Motor columns are subdivided into medial (MMC)
and hypaxial (HMC) motor columns depending on their innervation of either the
back or the trunk, respectively. Motor column divisions are called lateral motor
columns (LMC) at dorsal and ventral portions of the limb. The last class of motor
columns, which are not somatic, is called visceral preganglionic motor neurons
(PGC) (reviewed in [25]).
Contrary to motor neurons, interneurons are much more diverse and stay
within spinal cord which make them hard to distinguish. However,
electrophysiology, cell morphology, axonal trajectory and gene expression
profiling have all helped to describe spinal cord interneurons in recent years
(reviewed in [97]). Basically, interneurons can be classified into two main groups
called ventral interneurons “V” and dorsal interneurons “dI”. V0-V3 ventral interneurons mainly function together with motor neurons, whereas dorsal interneurons are associated with sensory function.
V0 interneurons arise from dorsal most ventral progenitor pool, p0, and are characterized by their expression of Dbx1, Dbx2 and Evx1 transcription factors. They are primarily contralateral with some ipsilateral projections, either acting as inhibitory or excitatory neurons. They send their axons 2-4 spinal segments rostrally [98,99]. To date, four different classes of V0 interneuron 26 subtypes were identified: V0V, V0D, V0C, V0G depending on the cells derived from
Evx1, Dbx1, Lhx3, Lhx5 expressed lineages (reviewed in [96]).
V1 interneurons are born at p0 progenitor domain with the expression of
Pax6, Dbx2, Nkx6.2 (Figure 1.3 and Figure 1.4). After post-mitotic differentiation,
V1 interneurons are marked by their expression of Engrailed-1 (En1) [100-104].
Classification based on electrophysiology led to the identification of two V1 interneuron populations: Renshaw cells and V1a inhibitory interneurons constitute a total of 25% of all V1 interneurons [105]. In the zebrafish spinal cord, eng1b is expressed by a class of ipsilateral ascending inhibitory interneurons called CiA neurons which are the homologs of V1 interneurons in mammals
([101] and reviewed in [97]). The function of V1 interneurons in motor circuitry was examined in three independent studies all aiming to ablate V1 interneurons either genetically or using neurotoxins such as diphtheria A and allatostatin.
Upon inactivation/loss of V1 interneurons, the locomotor step cycle has extended
([106], reviewed in [107]). These studies suggest that loss of inhibitory connections in left-right motor coordination causes out of phase locomotion.
V2 interneurons are comprised of two different subpopulations: excitatory
V2a and inhibitory V2b. This segregation was observed in zebrafish embryos and revealed that same V2 progenitor cells give rise to both subtypes depending on
Chx10 expression in V2a and Gata2/3 expression in V2b [108]. Segregation of
V2 interneurons was proposed to be mediated by Notch/Delta pathway (reviewed in [96]). Ablation of V2a interneurons via expressing diphtheria toxin in Chx10 27
expressing neurons resulted in disruption of left-right coordination governed by spinal cord neural network capable of generating organized motor activity [109].
V3 interneurons that arise from ventral most p3 domain are marked by
their expression of Nkx2.2, Nkx2.9 and Sim1 [14,110]. Two subdivisions of V3 interneurons, V3V and V3D, constitute the major class of excitatory commissural neurons. V3 neurons are also a component of motor network, specifically important for balanced locomotor rhythm. Electrophysiological and morphological
properties of V3 interneurons are mostly studied in mice. The fish homolog of V3
interneurons has not been identified yet; however, a class of excitatory
commissural neurons in zebrafish hindbrain, called spiral fiber neurons, shares
similarities with the mouse V3 interneurons and they are capable of regulating
the Mauthner neuron escape pathway (reviewed in [107]).
Motor behaviors in zebrafish
The central pattern generator (CPG) neural network that governs motor
output is evolutionarily conserved between terrestrial and aquatic animals. As a
result, the zebrafish is a powerful model organism for studying the spinal cord
development and performing behavioral studies (reviewed in [107]). The
complexity of mammalian nervous system and the time and the cost needed to
genetically analyze spinal cord circuits have enabled zebrafish usage in
functional analysis of spinal cord development. 28
It is crucial to understand how neural circuits dictate complex behaviors
such as locomotion. Zebrafish embryos are well-matched for behavioral studies
due to their large numbers progeny and development. Developmental
characteristics of locomotion in zebrafish are well described [111]. In recent
years, several mutations have been found to have an effect on locomotion in
zebrafish [112].
Zebrafish motor behaviors can be divided into three categories during development. Spontaneous contractions, touch response and swimming are
three sequential motor behaviors observed in zebrafish embryos starting at 17
hours post fertilization (hpf) [113]. The first movement is spontaneous
contraction, which can be described as slow side to side contraction of tail.
Spontaneous contraction can be observed between 17 hpf to 26 hpf. Starting at
21 hpf, embryos start to respond to touch on either head or tail. Touch evoked
response can be described as a rapid coil followed by slower undulations. The
third movement is swimming which starts by 26 hpf. Swimming can be defined as
alternating forward body movement which needs to be at least one body length
[113].
Sensory information, such as touch to head and acoustic stimulus, is
relayed to the brainstem for processing (Figure 1.5). To interpret and respond to
stimulus, the Mauthner neuron escape pathway is activated. Mauthner neurons are a pair of easily identifiable neurons born in rhombomere 4 of the hindbrain in
zebrafish and amphibians. They send their axons contralaterally along the body. 29
Figure 1.5. Mechanism of touch response.
Sensory organs relay stimulus to Mauthner neurons through anterior lateral line nerve and midbrain/hindbrain nuclei. Output of touch stimulus is executed by spinal cord interneurons and motor neurons.
30
The activation of Mauthner neurons triggers the first large amplitude body bend, called C bend, followed by lower amplitude body bends resulting in moving away from threatening stimulus [114]. This escape behavior is thought to have evolved to escape from potential predators. Mauthner neurons fire only once and elicit escape response via spinal cord neurons. Mauthner neurons are the core component of CPG for locomotor output; however, the input and feedback mechanisms transmitted from spinal interneurons are also crucial for successful operation of motor circuitry. For example, electrophysiological studies in fish and frog revealed that En1 positive V1 interneurons provide inhibitory synaptic input received by motor neurons [101]. Functional assays, testing the involvement of any motor circuit component like Mauthner neurons, interneurons and motor neurons, provide valuable information. Touch evoked response analysis is a classical method of testing of locomotion in zebrafish [114].
All embryonic zebrafish behaviors are analyzed using video recordings.
For quantitative analysis, head versus tail angles along fish movements are measured via a computer program. Quantification can include the number of body bends, duration of the response, distance traveled, bend intervals and so on.
31
CONTRIBUTION OF THIS WORK TO THE FIELD
Despite extensive studies of prdm12b and its role in zebrafish neurogenesis, mechanistic insight into prdm12b function in zebrafish embryonic development has not been achieved in germ line mutants. The severity of germ line prdm12b loss proves that prdm12b is vital not only for neural tube patterning during zebrafish embryogenesis but also for its survival.
This work shows that the phenotype of loss of function prdm12b allele, which lacks eng1b expressing V1 interneurons and presents subsequent defective Mauthner neuron dependent locomotion, is indistinguishable from prdm12b morphants. Consistency and severity of the phenotype are further confirmed by two different germ line CRISPR/Cas9 prdm12b mutants, morpholino knockdown, and ENU-derived prdm12b allele. Further characterization of Prdm12b revealed that Zinc finger domain of Prdm12b is essential for its repression and mediates the interaction with Bhlhe22. I have also found that Prdm12 interaction with EHMT2/G9a is not through any particular domain.
Since the phenotype resulting from prdm12b loss is confined to the p1 domain, I have suggested that loss of prdm12b is actually affecting size of the p1. I have further demonstrated the decrease in the p1 domain size upon prdm12b loss. 32
Taken together, the results presented in this study have deepened and expanded the state of the art of prdm12b’s role in zebrafish embryogenesis.
CHAPTER II: ZEBRAFISH PRDM12B ACTS INDEPENDENTLY OF NKX6.1 REPRESSION TO PROMOTE ENG1B EXPRESSION IN THE NEURAL TUBE P1 DOMAIN
Ozge Yildiz, Gerald B. Downes and Charles G. Sagerström
This chapter submitted as a manuscript to Neural Development and accepted on February 13, 2019.
34
BACKGROUND
Appropriate function of the adult nervous system requires the
establishment of neural circuits during embryonic development. For such circuits
to form properly, neurogenesis has to occur at the right time and place, neurons must migrate to the correct site and they must make appropriate connections.
Disruptions to any step in this process result in improper neural circuit formation and such disruptions are thought to underlie many neurodevelopmental disorders
– including schizophrenia and autism [1].
The embryonic vertebrate neural tube represents a well-studied system of neural circuit formation where various progenitor types form in distinct domains arrayed along the dorsoventral (DV) axis. These progenitor domains form in response to morphogen gradients – particularly dorsally derived Bone morphogenic protein (BMP) and ventrally derived Sonic hedgehog (Shh;
reviewed in [3,21]). In response to these morphogens, each progenitor domain
acquires a unique gene expression profile that initially consists primarily of
transcription factors (TFs). Strikingly, TFs unique to one progenitor domain
frequently cross-repress the expression of TFs associated with adjacent
domains, thereby establishing distinct boundaries that delineate individual
progenitor domains along the DV axis. The graded morphogen signal, and the
resulting distinct transcriptional programs, leads to the development of sensory
neurons in the dorsal domains (pd1-pd5) and interneurons and motor neurons in 35
the ventral domains (pd6-p0, p1, p2, pMN, p3) of the neural tube. Neurons from
each of these domains then make connections to establish motor circuits that
control the activity of limb and trunk musculature [10].
Many TFs that control establishment of progenitor domains along the DV
axis belong to the homeodomain (HD) and basic Helix-Loop-Helix (bHLH) families. For instance, work in mouse and chick indicate that Shh activates genes
such as Nkx6.1, Nkx6.2, Nkx2.2, and Olig2, while it represses Pax3, Pax6, Pax7,
Dbx1, Dbx2 and Irx3 [13-15,17,18,28-31]. These TFs then repress each other’s
expression to establish distinct progenitor domains. For instance, Irx3 and Olig2 are mutually repressive at the p2/pMN boundary [18,32] such that loss of Olig2 leads to a ventral expansion of Irx3 expression, causing the pMN domain to give
rise to V2 interneurons and astrocytes in place of motor neurons and
oligodendrocytes [32]. More recently, members of the Prdm TF family have also
been implicated in the formation of progenitor domains and the establishment of
functional motor circuits (reviewed in [84]). The Prdm family consists of many
members (Prdm1-16) that harbor an N-terminal PR domain, as well as a variable
number of zinc fingers [37,53], and that appear to preferentially act in complexes
with bHLH TFs [84]. Hence, Prdm13 acts together with Ascl1 to promote
formation of GABAergic neurons [68,83], while Prdm8 interacts with the Bhlhe22
(a.k.a. Bhlhb5) TF to regulate axon outgrowth [71]. Of particular interest, Prdm12
is expressed in the developing CNS of mouse, frog, chick and zebrafish
[90,92,93] – specifically in the p1 domain, which gives rise to V1 interneurons. 36
Prdm12 deficiency in zebrafish and frog results in loss of eng1 expression from the p1 domain and animals lacking prdm12 function demonstrate a defective touch-evoked escape response [92,93], suggesting that the V1 interneurons are absent. However, key aspects of Prdm12 function remain unclear. First, Prdm12 activity has only been assessed via overexpression and transient knock-down approaches – particularly antisense morpholino oligonucleotides (MOs) – that have recently come under scrutiny as prone to non-specific off-target effects.
Furthermore, Prdm12 is suggested to act as a transcriptional repressor, but this is based on overexpression in fish and frog embryos [93,115] and has not been stringently tested. Here, we generate and characterize the first germ line prdm12 mutants using CRISPR/Cas9 to inactivate zebrafish prdm12b. prdm12b mutants display embryonic lethality and, in accordance with previous prdm12b MO analyses, we find that prdm12b mutants exhibit loss of eng1b expression in the p1 domain together with an abnormal touch-evoked escape response that we demonstrate is due to defects in a Mauthner cell-dependent pathway. We also employ luciferase reporter assays to reveal that Prdm12b acts as a bona fide repressor. This repression requires a conserved zinc finger domain that interacts with the Bhlhe22 TF, but, when we generate a bhlhe22 germ line zebrafish mutant, it displays a normal p1 territory – indicating that bhlhe22 does not need to act with prdm12b for p1 progenitor formation in vivo. Lastly, while Nkx6.1 is known to repress p1 fates in other systems, we find that prdm12b and nkx6.1 does not form a reciprocally repressive TF pair in the zebrafish. Therefore, 37 instead of the p1 domain taking on a p2 fate, a residual domain with unknown properties persists at the p1 position in prdm12b zebrafish mutants.
METHODS
Zebrafish care
Wild type and mutant zebrafish were raised in the University of
Massachusetts Medical School Aquatics Facility. All embryos were staged according to previously described morphological standards [116].
Generation of CRISPR/Cas9 mutant zebrafish lines
We designed single guide RNAs (sgRNA) for the zebrafish prdm12b, bhlhe22 and nkx6.1 genes (Table 2.1) using the CHOPCHOP web tool [117].
Each sgRNA was assembled by annealing two single stranded oligonucleotides containing the T7 promoter and the target sequence (Table 2.2) followed by PCR amplification, purification and in vitro transcription using T7 RNA polymerase
(Promega) as described previously [118]. A linearized plasmid encoding cas9 was used for in vitro transcription using the SP6 mMessage mMachine Kit
(Ambion) according to the manufacturer’s instructions [119]. cas9 mRNA and sgRNA was co-injected into 1-cell stage zebrafish embryos at the following concentrations: 150ng/µL sgRNA plus 200 ng/µL cas9 mRNA for prdm12b, 38
100ng/µL sgRNA plus 200 ng/µL cas9 mRNA for bhlhe22 and 150ng/µL sgRNA plus 200 ng/µL cas9 mRNA for nkx6.1. The next day, injected embryos were assayed for sgRNA activity by DNA extraction, PCR amplification, restriction digestion and DNA sequencing (Table 2.1). Detection of F0 founders was done by crossing sgRNA/cas9-injected animals with wildtype zebrafish and screening their offspring for mutagenic events using the diagnostic restriction enzymes listed in Table 2.1. Confirmed founders were crossed to wildtype animals to raise
F1 carriers for each mutant. 39
Table 2.1. The characteristics of CRISPRs targeting prdm12b, bhlhe22 and nkx6.1
Target Start Target sequence Enzyme Strand Mutagenesis Transmission gene Coordinate Ratea Rateb prdm12b Chr5:66656496 GCTGGGGGAACACCTGTTCG Taq1a + 1/4 71/92 um318 43/79 um319 bhlhe22 Chr24:25069884 TTCACACACAAAGATCCGGT BstYI - 6/14 24/37 um320 nkx6.1 Chr21:17886500 AGTGGAGGATGCTGGTCCAG AvaII - 8/12 18/21 um321 a The fraction of screened F0 animals that carried a mutagenic event. b The fraction of screened F1 animals that were heterozygous for a mutagenic event.
Table 2.2. Sequences of oligos to generate single guide RNAs
Target Gene Target sequence a PAM b First oligo c Second oligo d prdm12b GCTGGGGGAACACCTGTTCG AGG TTAATACGACTCACTATAGGGC AAAAAAGCACCGACTCGGTGCCAC TGGGGGAACACCTGTTCGGTT TTTTTCAAGTTGATAACGGACTAGC TTAGAGCTAGAAATAGCAAG CTTATTTTAACTTGCTATTTCTAGC TCTAAAAC bhlhe22 TTCACACACAAAGATCCGGT CGG TTAATACGACTCACTATAGGTT AAAAAAGCACCGACTCGGTGCCAC CACACACAAAGATCCGGTGTTT TTTTTCAAGTTGATAACGGACTAGC TAGAGCTAGAAATAGCAAG CTTATTTTAACTTGCTATTTCTAGC TCTAAAAC nkx6.1 AGTGGAGGATGCTGGTCCAG TGG TTAATACGACTCACTATAGAGT AAAAAAGCACCGACTCGGTGCCAC GGAGGATGCTGGTCCAGGTTT TTTTTCAAGTTGATAACGGACTAGC TAGAGCTAGAAATAGCAAG CTTATTTTAACTTGCTATTTCTAGC TCTAAAAC a Genomic sequence targeted by the single guide RNA b Protospacer adjacent motif (PAM) sequence in the genomic DNA recognized by Cas9 c Sequence of the first oligo used to assemble the guide template containing the T7 promoter sequence d Sequence of the second oligo used to assemble the guide template containing the constant region 40
Antisense morpholino oligonucleotide injections
Antisense morpholino oligonucleotides (MOs) were obtained from Gene
Tools LLC. MO injections were performed into the yolk of 1-cell stage embryos
using 1-2ng of solution containing dilutions of 3mM morpholino stock, distilled
water and phenyl red. An MO with the sequence 5’-
GCAGGCAACACTGAACCCATGATGA-3’ was used to target the prdm12b
translation start site. This MO was reported previously [92] and our analyses in
this manuscript demonstrate that the effects of MO-mediated prdm12b knockdown are indistinguishable from the effects of prdm12b germ line mutations.
In situ RNA hybridization
Embryos were fixed in 4% paraformaldehyde (PFA) and stored in 100% methanol at -20°C. In situ RNA hybridization was performed as described [120]
followed by a color reaction using NBT/BCIP in 10% polyvinyl alcohol. RNA
probes for the genes eng1b, evx1, vsx2, pax3, nkx6.1, dbx1 and prdm12b were
synthesized as previously described [118]. Embryos were dissected from the yolk
and flat mounted in 80% glycerol for imaging on bridged coverslips or sectioned
as described [121]. Images were captured using a Nikon Eclipse E600
microscope equipped with spot RT color camera (model 2.1.1). Images were
imported into Adobe Photoshop and adjustments were made to contrast, levels, 41 color matching settings and cropping only. All adjustments were made to the entire image.
Luciferase reporter assays
0.5x106 HEK293T cells were seeded in 6-wells plate and cultured in antibiotic free Dulbecco’s Modified Eagle Medium (DMEM; Gibco) supplemented with 10% fetal bovine serum (Hyloclone) overnight. Transient transfections were performed using Lipofectamine 2000 reagent (Invitrogen) according to the manufacturer’s instructions. For each transfection, 200ng of the pGL4.31[luc2P/GAL4UAS/Hydro] reporter plasmid and 50ng pRL-SV40 control plasmid was combined with varying concentrations of GAL4DBD fusion plasmids
(exact concentrations are given in figure legends). Empty vector DNA was included to keep the total amount of DNA constant for all transfections.
Luciferase activity was measured 24hours post transfection and firefly luciferase levels were normalized to renilla luciferase levels using the Dual Luciferase
Reporter Assay System (Promega) following the manufacturer’s instructions in a
Perkin Elmer Envision 2104 Multiplate reader. For Trichostatin A (TSA) treatment, transfected cells were exposed to either DMSO, 50nM or 250nM TSA for 12 hours starting 24 hours after transfection and then harvested for luciferase assays.
42
Co-immunoprecipitation and Western blotting
3x106 HEK293T were seeded in 10cm dishes and transfected as above.
Transfected cells were lysed in 4mL of ice-cold co-IP buffer (50mM Tris-HCl
pH7.5, 150mM NaCl, 0.2mM EDTA, 1mM DTT, 0.5% Triton X100, 1X Complete
Protease Inhibitor (Roche)) followed by incubation on ice for 30min. Cell lysates
were centrifuged at 2,000g for 10 min at 4°C to eliminate cell debris. For
immunoprecipitation, 8μg of the mouse anti-Flag antibody (Sigma-Aldrich,
F3165) was used in each sample and incubated at 4°C overnight. 40μL of
Dynabeads was added in each sample and incubation was done for 4hr at 4°C.
Four washes of 1mL co-IP buffer was used to eliminate non-specific binding.
Lastly, immune complexes were eluted in 80μL of 1X Laëmmli buffer (Biorad) containing 2.5% beta-mercaptoethanol. Samples were agitated at 95°C for five minutes prior to Western blotting. Western Blotting was performed using rabbit
HA antibody (Abcam, ab9110) as described previously [122].
Immunocytochemistry
Primary antibodies: mouse 3A10 (1:100; Developmental Studies
Hybridoma Bank (DSHB) [123]), mouse F310 (1:100; DSHB [124]), mouse anti-
Isl (39.4D5, 1:100; DSHB [125]), mouse 81.5C10 (Hb9; 1:400; DSHB [126]).
Alexa Fluro secondary antibodies: 488, 568 goat anti-mouse (both at 1:200;
Molecular probes). Embryos were fixed in 4% AB fix (4% paraformaldehyde, 8% 43
sucrose, 1x PBS) overnight at 4°C. Whole-mount fluorescent labeling was
performed as described [127]. Images were captured on either Nikon Eclipse
E600 (3A10, Isl1 and Hb9 staining) or a Zeiss LSM700 confocal microscope
(F310 staining). Images were imported into Adobe Photoshop and adjustments
were made to contrast, levels, color matching settings and cropping only. All
adjustments were made to the entire image.
Behavioral analysis
Escape responses were elicited by a light tap to the head or tail of an embryo with a 3.22/0.16 g of force Von Frey filament. A high-speed digital camera (Fastec Imaging, San Diego, CA) mounted to a 35mm lens (Nikon,
Melville, NY), recorded each response at 1000 frames/s. Computer software generated in the Downes laboratory [128] quantified the head-tail angle for each
frame, which was then plotted in Prism. The calculated escape response began
in the frame preceding the first movement until movement was no longer
observed.
Genotyping
CRISPR-generated mutant alleles of prdm12b, bhlhe22 and nkx6.1 were genotyped by Taq1a, BstYI or AvaII restriction digest, respectively, of PCR products amplified from genomic DNA using primers listed in Table 2.3. 44
prdm12bsa9887 mutants were genotyped by sequencing of PCR products amplified
from genomic DNA using primers listed in Table 2.3. Total RNA from 24hpf WT and bhlhe22 zebrafish whole embryos was extracted with the RNeasy kit
(Qiagen) following manufacturer’s instructions. Total RNA was then used in cDNA kit (ThermoFisher Scientific). Wildtype and bhlhe22 mutant transcripts were identified by sequencing of PCR products amplified from cDNA using primers listed in Table 2.3.
Table 2.3. Primers used for genotyping of prdm12b, bhlhe22 and nkx6.1 mutant fish
Primer name Sequence Purpose prdm12b-1 GGTTCGGCTCATCATGGGTTC Forward primer used to genotype um318, um319 prdm12b-2 GCAAACTGACCTCCAGAGAA Reverse primer used with primer prdm12b-1 prdm12b-3 TTCCAGCTTAGTTCTGCCAAGTG Forward primer used to genotype sa9887 prdm12b-4 CGACCTCCAAGTTCTGTTCTT Reverse primer used with primer prdm12b-3 bhlhe22-1 AGAATAAACTTGGGCGGAGAC Forward primer used to genotype um320 bhlhe22-2 CATTGCTTACACAGGCTGGA Reverse primer used with primer bhlhe22-1 bhlhe22-3 GCATCCGACTTTCTGGAGAC Forward primer used to genotype um320 bhlhe22-4 GCTGGAGGTGACATTGTTGAA Reverse primer used with primer bhlhe22-3 nkx6.1-1 GGTCACTGTCCTGCTTCTTG Forward primer used to genotype um321 nkx6.1-2 CCACACCTTGACTTGACTCTC Reverse primer used with primer nkx6.1-1
RESULTS
Germ line disruption of prdm12b blocks eng1b expression in the p1 domain
The prdm12 TF is known to be expressed in the developing CNS of mouse,
chick, Xenopus and zebrafish [90,92,93] – particularly in sensory ganglia and in
the p1 domain of the neural tube. The p1 domain gives rise to eng1b-expressing 45
V1 interneurons that regulate motor circuits in several vertebrate species
[101,103,106]. Disruption of prdm12 function using antisense morpholino oligonucleotides (MOs) leads to the loss of eng1b expression in the p1 domain, but not in other eng1b expressing tissues – such as the midbrain-hindbrain boundary (MHB) and the somites – in zebrafish and Xenopus [84,93], but there have been no germ line mutations for prdm12 produced in any organism.
Importantly, recent work has demonstrated several cases where apparently specific MO-derived phenotypes do not match the phenotypes of germ line mutants for the same gene [129]. The underlying causes of such discrepancies are varied, but include off-target effects, as well as compensatory changes in the expression of genes with similar functions to the targeted gene [130]. Hence, it is essential to confirm MO-derived phenotypes by comparisons to the phenotypes of germ line mutant animals. To this end, we used the CRISPR/Cas9 genome editing system [131,132] to generate prdm12b germ line mutant zebrafish. We tested five sgRNAs targeting the first exon of the prdm12b gene and identified one that efficiently disrupts a diagnostic Taqa1 site at position 129 of prdm12b exon 1 in 24hpf zebrafish embryos (Figure 2.1A, B). Injected embryos were raised to adulthood and screened to identify founders that carry mutations in the prdm12b gene (Figure 2.1C). In this manner, we identified one mutant F0 founder out of four tested (Table 2.1). Since zebrafish F0 founders are usually mosaic, this founder was crossed to wildtype fish and the resulting F1 generation raised to adulthood (Figure 2.1D). Genotyping revealed that the F0 founder 46
transmitted mutations to 77% (114/171) of its F1 offspring (Table 2.1).
Subsequent sequencing of genomic DNA from individual F1 fish identified two different alleles (prdm12bum318 and prdm12bum319; Figure 2.1E, F; Figure 2.2). In
both alleles, the mutant sequence leads to a frameshift and premature
termination of translation upstream of the conserved PR domain and the zinc
finger domains. In addition, while we were in the process of generating prdm12b
mutants, a mutant prdm12b allele became available from the zebrafish
information resource center (ZIRC) as a product of the zebrafish mutation project
(ZMP). This mutant allele (prdm12bsa9887) is ENU-derived and carries a T>C
change in an essential splice site at the beginning of intron 2, within the PR
domain and upstream of the zinc finger domains (Figure 2.3A). We obtained this
line from ZIRC and confirmed the presence of the expected mutation by
sequencing (Figure 2.3B, C).
Since the effects of MOs wear off as development progresses (largely due
to MO degradation) they are not a reliable tool to assess genetic effects on
embryo viability. However, having generated prdm12b germ line mutants, we
were able to examine the effect of prdm12b on viability by crossing heterozygous
carriers and genotyping the resulting offspring at different stages of
embryogenesis. prdm12b mRNA does not appear to be maternally deposited
(Figure 2.4A, B) and is not detected until the end of gastrulation [84], suggesting
a relatively late role in development. Accordingly, we observe the expected ~25%
homozygous prdm12b mutants (26/139 for um318 and 29/116 for um319) at 4dpf 47
48
Figure 2.1. Generation of germ line prdm12b mutants.
A. Schematic showing genomic sequence of prdm12b. Exons are indicated as boxes and black lines represent introns. The PR domain and three zinc fingers
(ZnF) are highlighted in dark red and blue, respectively. The CRISPR target sequence is shown in red with the Taqa1 restriction site bracketed and the black arrow indicating the Taqa1 cut site. B. Identification of functional guide RNAs. sgRNA and cas9 mRNA was injected into 1-cell stage embryos. Injected embryos were raised to 24hpf and Taqa1 digestion of PCR amplicons from pools of embryos was used to identify CRISPR-induced mutations (black arrow). C.
Identification of individual F0 founders. sgRNA/cas9 injected embryos were raised to adulthood and crossed to wildtype fish. Taqa1 digests of PCR amplicons from pools of embryos was used to identify F0 mosaic founders (black arrow). D. Identification of F1 animals. Adult F0 mosaic founders were out- crossed to wildtype fish and the F1 offspring raised to adulthood. Taqa1 digests of PCR amplicons from individual fin clip genomic DNA was used to identify heterozygous F1 animals. E. Sequencing of F1 genomic DNA revealed the transmission of two different mutant alleles (um318, um319). um318 carries a 42 base pair deletion (black dashes) and a 16 base pair insertion (blue), while um319 carries a 17 base pair deletion (black dashes). The CRISPR target sequence is shown in red. F. Predicted amino acid sequence of mutant alleles.
The um318 peptide shares its first 41 amino acids, and the um319 peptide its first 43 amino acids, with wildtype Prdm12b. The two mutant peptides then utilize 49
a different reading frame that terminates at a premature stop codon N-terminal to
the conserved PR domain. inj = sgRNA/Cas9-injected embryos, uninj = uninjected control embryos.
50
51
Figure 2.2. Sequence of mutant prdm12b, bhlhe22 and nkx6.1 alleles.
The predicted amino acid sequence for each mutant allele was aligned to the corresponding wildtype sequence using Clustal Omega.
52
53
Figure 2.3. Characterization of the prdm12bsa9887 mutant.
A. Schematic showing genomic sequence of prdm12b. Exons are indicated as
boxes and black lines represent introns. The PR domain and three zinc fingers
(ZnF) are highlighted in dark red and blue, respectively. The black arrow
indicates a single base pair change in the second intron of prdm12bsa9887. B, C.
Sequence traces confirming the expected single nucleotide change in wildtype
(B) versus prdm12b+/sa9887 (C) animals. D-G. eng1b expression in 24hpf embryos from a cross of prdm12b+/sa9887 animals. Embryos are shown in dorsal (D, E) or
lateral (F, G) view with anterior to the left. eng1b expression is lost in 27% of
embryos compared to 73% of embryos showing wildtype eng1b staining.
54
55
56
Figure 2.4. prdm12b germ line mutants lack eng1b expression in the p1
domain.
A, B. prdm12b is not maternally deposited. In situ hybridization detects prdm12b
expression at 24hpf (B), but not at 2.5hpf (A), in wildtype embryos. C. Bar chart
depicting the frequency of each genotype at various time points in broods from
crosses of prdm12b heterozygous animals. Error bars indicate ±S. E. (n=3). dpf =
days post fertilization, mo = months. D, E. Morphology of 15dpf prdm12b+/+ (D)
and prdm12bum319/um319 (E) fish. F-S. eng1b expression in 24hpf embryos from
crosses of prdm12b+/um318 heterozygotes (F-K), or prdm12b+/um319 heterozygotes
(L-S). Numbers in each panel indicate the fraction of animals with the specified
phenotype. T, U. evx1 expression in 24hpf embryos from a cross of
prdm12b+/um319 heterozygotes. V, W. vsx2 expression in 24hpf embryos from a cross of prdm12b+/um319 heterozygotes. Embryos are shown in dorsal (F-H, L-N,
T-Y) or lateral (I-K, O-Q) view with anterior to the left, or in cross section (R, S)
with dorsal to the top. Brackets indicate r4, arrows mark V1 interneurons and
arrowheads mark somites. MHB = midbrain–hindbrain boundary, HB = hindbrain
and SC = spinal cord.
57
(Figure 2.4C), but by 15dpf only ~13% of embryos are homozygous mutant
(22/172 for um319) and by 21dpf we no longer detect any homozygous mutants
(0/129 for um319). We also do not observe homozygous mutants when genotyping adult offspring (2 months of age; 0/92 for um318 and 0/145 for um319) from these crosses. Since prdm12b mutants start dying between 4dpf and 15dpf, we monitored developing embryos more closely during this time interval and noticed that a fraction of embryos grew at a slower rate (Figure 2.4D,
E). When the smaller embryos were genotyped, 82% (18/22) turned out to represent homozygous prdm12b mutants. This slower rate of growth suggests that the mutants may be unable to feed properly (perhaps due to the motility defects described below). However, when fed brine shrimp, even the mutant embryos show evidence of food in their digestive tract (orange/yellow color in
Figure 2.4D, E). Hence, the mutants are capable of feeding, although we cannot exclude the possibility that they do so sub-optimally.
Since loss of eng1b expression in the p1 domain is the key feature of the zebrafish prdm12b morphant phenotype, we next assayed eng1b expression in all three prdm12b mutant alleles by in situ hybridization at 24hpf. For both
CRISP/Cas9-generated alleles, ~25% of embryos from crosses of heterozygous carriers lack eng1b expression in hindbrain and spinal cord (Figure 2.4F-Q). In the affected embryos, eng1b expression is lost from the p1 domain, but persists at the MHB and in the somites (whole mount in Figure 2.4H, K, N, Q and section in Figure 2.4S). Genotyping revealed that all embryos lacking eng1b expression 58
in the p1 domain represent homozygous prdm12b mutants (45/45 for
prdm12bum318 and 13/13 for prdm12bum319). Similarly, eng1b expression is lost in
both hindbrain and spinal cord in 27% of embryos from a cross of prdm12bsa9887/+
heterozygous fish, while the remaining embryos show unaffected eng1b
expression (Figure 2.3D-G). We conclude that germ line mutants for prdm12b display the same loss of eng1b expression as previously reported for prdm12b
morphants.
The defective escape response in prdm12b mutants is caused by disruption of
the Mauthner cell pathway
V1 inhibitory interneurons are responsible for the modulation of motor
circuits in many species, including zebrafish, Xenopus and mouse ([116,123],
reviewed in [107]). Accordingly, we previously demonstrated that prdm12b morphants display abnormal movements in response to touch [84]. The touch-
evoked escape response is a classical method of assessing functionality of motor
output in aquatic species [114] and it has been applied to zebrafish [133,134]. In
this test, a touch stimulus causes the fish to undergo a large amplitude body
bend (C bend), which reorients the animal away from the stimulus. The initial
large amplitude body bend is followed by lower amplitude counter bends,
allowing the fish to propel itself away. Strikingly, the escape response of
prdm12b morphants is exaggerated, such that morphants perform not just one, 59
but several repetitive C-bends and, compared to a wild type response – which
lasts ~100ms – the response of prdm12b morphants is prolonged and may
continue for several hundred milliseconds [84]. To determine if this defect is
observed also in germ line mutants, we assessed the escape response in 4dpf
old prdm12b mutant fish to a head tap, followed by genotyping. We find that all
prdm12b mutants (9/9 for um318 and 8/8 for um319), respond by carrying out
repetitive C-bends (up to seven C-bends) for extended periods of time (Figure
2.5A, B; Figure 2.6, Video 1, 2). We extended this analysis to also score the
response of prdm12bum319 homozygous mutant animals when tapped on the tail.
We observed no differences between responses to head versus tail stimulation – in all 11 cases were the responses exaggerated to both stimuli (Figure 2.5C, D;
Figure 2.6, Video 1, 3, 4). The touch-evoked escape response is mediated via
reticulospinal neurons – most notably the Mauthner cells, but also MiD2 and
MiD3 cells – and our results therefore indicate that this pathway is abnormal in
prdm12b mutants. Notably, there is no known circuit connecting V1 interneurons
to the reticulospinal cells, suggesting that the abnormal escape response
observed in prdm12b mutants may be independent of the loss of V1
interneurons. Indeed, the behavior of the mutants is consistent with enhanced or
excessive activity of this pathway, perhaps due to impaired synapse function or
circuit regulation. Accordingly, we do not detect structural defects in either the
morphology of Mauthner cells (Figure 2.5E), or the structure of trunk/tail
musculature (Figure 2.5F). We conclude that prdm12b germ line mutant animals 60
61
Figure 2.5. prdm12b mutant fish display an abnormal touch evoked
response.
A-D. Representative kinematic traces for 10 wildtype (A) and 11 prdm12b mutant
(B) fish stimulated with a head touch, as well as for 11 prdm12b mutants first
assayed with a head touch (C) and subsequently with a tail touch (D). Zero
degrees on the y-axis indicate a straight body while positive and negative angles
represent body bends in opposite directions. All fish were at 4dpf. E.
Neurofilament staining via anti-3A10 labeling of Mauthner neurons in a cross of prdm12b+/um318 heterozygotes (n=117). F. Staining of myosin light chain via anti-
F310 in a cross of prdm12b+/um398 heterozygotes (n=16).
62
63
Figure 2.6. Detailed analysis of the touch-evoked escape response in prdm12b mutant and wild type animals.
A, B. Representative kinematic traces of individual wild type (A) and prdm12b mutant (B) animals stimulated with a head tap (from Figure 2.5A, B). C.
Quantification of number of body bends with an amplitude similar to the C-bend
(defined as exceeding 100°; from data collected in Figure 2.5A, B). D.
Quantification of C bend duration (from data collected in Figure 2.5A, B). E, F.
Representative kinematic traces of individual prdm12b mutant animals stimulated with a head (left panels) or a tail (right panels) tap (from Figure 2.5C, D)
64
Video 1. Movie of wild type touch-evoked response.
Movie of representative wild type animal tapped on the head (from Figure 3A; recorded at 1000 frames/second)
Video 2. Movie of prdm12b mutant touch-evoked escape response
Movie of representative prdm12b mutant animal tapped on the head (from Figure
3B; recorded at 1000 frames/second)
65
Video 3. Movie of prdm12b mutant touch-evoked escape response
Movie of representative prdm12b mutant animal tapped on the head (from Figure
2.5C; recorded at 1000 frames/second)
Video 4. Movie of prdm12b mutant touch-evoked escape response
The same prdm12b mutant animal as in Video 3 was instead tapped on its tail
(from Figure 2.5D; recorded at 1000 frames/second).
66
display a defective escape behavior that is qualitatively and quantitatively
indistinguishable from that of prdm12b morphants.
Prdm12b acts as a repressor in vitro
The fact that prdm12b belongs to a family of transcription factors, together with the finding that loss of prdm12b function abolishes eng1b expression, suggests that this factor may function as a transcriptional activator. Accordingly, transfection of prdm12 into P19 cells upregulates p27 mRNA and protein levels
[89]. However, recent reports instead suggest that prdm12 acts as a repressor
[93], but this conclusion was based on overexpression experiments in vivo and
has not been tested directly. To more directly determine whether prdm12b acts
as an activator or repressor, we made use of classical reporter assays. While prdm12b possesses three putative zinc-fingers (ZnFs), it is not clear if these are
sufficient for DNA binding and there is no well-defined genomic motif for
Prdm12b binding. We therefore fused the well-characterized DNA binding domain (DBD) from the GAL4 transcription factor in-frame to the N-terminus of zebrafish Prdm12b (Figure 2.7A; Figure 2.8). Transcriptional activity was
measured using the pGL4.31 reporter vector that contains multiple GAL4 binding
sites (upstream activation sequence; UAS) in front of the firefly luciferase gene.
Co-transfection of the reporter plasmid together with the GAL4-DBD alone led to a modest increase in Luciferase activity (Figure 2.7B). Strikingly, when the
GAL4DBD-Prdm12b fusion protein was instead co-transfected with the reporter 67
68
Figure 2.7. The zinc finger domain is necessary for Prdm12b-mediated
repression.
A. Diagram of GAL4DBD-Prdm12b constructs. FL = full-length, PR = PR domain,
ZnF = zinc finger domain. B-E. Reporter assays in HEK293 cells testing activity of GAL4DBD-Prdm12b constructs. For each experiment, the pRL-SV40 renilla- luciferase control plasmid and the pGL4.31 UAS:Firefly-luciferase reporter plasmid were co-transfected with the indicated GAL4DBD-Prdm12b construct or with a plasmid containing the GAL4DBD alone. Each construct was tested in triplicate and luciferase activity is expressed as mean fold induction ± SE over pGL4.31 reporter alone. Transfection efficiency was corrected by normalizing to renilla luciferase activity.
69
70
Figure 2.8. Expression of GAL4DBD-Prdm12b constructs used in
transfection experiments.
A. Immunoblot showing expression of GAL4DBD-Prdm12b constructs in transfected HEK293T cells. All constructs are stable except Myc-GAL4-∆PR- prdm12b. B. Immunoblot showing expression of Myc-Flag-G9a and Myc-Flag-
Bhlhe22 constructs in transfected HEK 293T cells.
71 plasmid, a dose-dependent reduction in Luciferase activity was observed (Figure
2.7B), indicating that the Prdm12b protein functions as a repressor.
Prdm12b contains two types of conserved domains – the PR domain and the zinc fingers. The PR domain is related to SET domains that function as histone lysine methyl transferases (HMTs). Most PR domain proteins lack the
H/RxxNHxC motif that is essential for HMT activity [38]; however, Prdm2, Prdm3,
Prdm6, Prdm8, Prdm9 and Prdm13 were recently shown to exhibit intrinsic methyltransferase activity [39-43]. Accordingly, the PR domain of Prdm12b has been postulated to act as a H3K9 methyltransferase – to deposit methyl groups onto lysine 9 of histone 3 –thereby repressing gene expression [115]. A recent study of Prdm9 demonstrated that cysteine 321 (Cys321) is highly conserved among Prdm family members that have intrinsic histone methyl transferase activity and that substituting Cys321 with a proline decreases Prdm9 activity
~1000 fold [135]. Our sequence comparison of Prdm1, 9, 10 and 12b revealed that Prdm12b carries a cysteine residue (Cys164) at the analogous position to
Cys321 in Prdm9, while Prdm1 and Prdm10 (that lack methyltransferase activity) contain a proline at this position. To determine the functional contribution of
Cys164, we tested the activity of several substitution mutants using the luciferase assay, but neither a cysteine -> proline, nor a cysteine -> alanine, substitution at position 164 affected the repressive activity of Prdm12b (Figure 2.7C). Deletion of the entire PR domain proved to be uninformative as this protein was unstable in HEK293 cells (Figure 2.8). Previous work also demonstrated that some Prdm 72 proteins act as repressors by recruiting histone deacetylases (HDACs) via the
PR domain [45,48,50], but we find that Trichostatin A (TSA; a HDAC inhibitor) does not affect the repressive activity of Prdm12b (Figure 2.7D). Lastly, we deleted the conserved zinc fingers in Prdm12b in order to determine if they might be required for its repressive function. Strikingly, deletion of the ZnFs completely abolished the repressive activity of Prdm12b and instead appears to produce a protein with slight activator activity (Figure 2.7E). Taken together, our results indicate that Prdm12b functions as a repressor and that this activity requires intact zinc finger domains, at least in the context of a GAL4DBD fusion protein.
Prdm12b interacts with the Bhlhe22 transcription factor and the EHMT2 methyltransferase
As discussed, it is unclear if Prdm12b binds DNA directly and it may instead be recruited to genomic binding sites by forming complexes with a DNA- binding factor. Since prdm12b is expressed only in the p1 domain, we focused our search for DNA-binding Prdm12b-interactors to ones that are co-expressed with prdm12b in the p1 domain. Based on this criterion, the Bhlhe22 transcription factor (also known as Bhlhb5) represents a potential binding partner for
Prdm12b. In particular, bhlhe22 is expressed in the pdl6, p1, p2 and p3 domains and has been implicated in the specification of V1 and V2 interneurons [136].
Furthermore, Bhlhe22 has been shown to form complexes with Prdm8, 73
suggesting that it may act broadly as a partner for Prdm proteins [71]. Using co- immunoprecipitation, we confirmed the interaction between Bhlhe22 and Prdm8
(Figure 2.9A, lane 9) and further demonstrated robust binding between Bhlhe22 and Prdm12b (Figure 2.9A, lane 6). More detailed analyses using Prdm12b deletion constructs indicated that the ZnF domain – that we already identified as
necessary for Prdm12b-mediated repression (see Figure 2.7D) – is required for
Bhlhe22 binding (Figure 2.9A, lane 7). In contrast, the PR domain does not
appear to be absolutely required for the Prdm12b-Bhlhe22 interaction (Figure
2.9A, lane 8).
Moreover, since Prdm12b appears to lack intrinsic methyltransferase
activity, it must function by recruiting factors to mediate its repressive effects.
Accordingly, Prdm family members recruit various transcriptional repressors
([54,57-59] and reviewed in [53]). In particular, Prdm1, 5 and 6, as well as
Prdm12, have been shown to bind EHMT2/G9a – a H3K9 methyltransferase
[45,48,62,137]. In the case of Prdm12, binding to EHMT2/G9a is reportedly
mediated by the ZnF domains [89]. Since this is the same domain that we find to
be required for binding to Bhlhe22, we examined this in further detail. We
confirmed that Prdm12b interacts with EHMT2/G9a (Figure 2.9B, lane 2), but find
that neither the ZnF, nor the PR domain, is required for this binding (Figure 2.9B,
lanes 5 and 8).
74
75
Figure 2.9. Prdm12b interacts with Bhlhe22 and EHMT2/G9a.
A, B. Co-immunoprecipitation experiments assaying interactions between
Prdm12b and Bhlhe22 or EHMT2/G9a. The indicated constructs were co- transfected into HEK293T cells followed by immunoprecipitation with anti-Flag and Western blotting with anti-HA. Arrows at right indicate the expected sizes of each protein. Figure 2.8B demonstrates that Flag-G9a and Flag-Bhlhe22 are stable upon transfection into HEK293 cells.
76
We conclude that Prdm12b binds to both Bhlhe22 and EHMT2/G9a.
Additionally, the Prdm12b ZnF domain – that is essential for Prdm12b-mediated
repression – is required for binding to Bhlhe22, but not to EHMT2.
bhlhe22 is not required for eng1b expression in the zebrafish p1 domain
Previous work reported that siRNA-mediated knock-down of bhlhe22 in
the chick spinal cord leads to a reduction in eng1 expression in the p1 domain
[136], akin to the effect we observe in prdm12b mutants. The similarity of the
bhlhe22 and prdm12b loss-of-function phenotypes, taken together with our finding that these two proteins form complexes, suggests that bhlhe22 and prdm12b may cooperate to control eng1b expression. To test this possibility, we generated germ line mutants for zebrafish bhlhe22 using the CRISPR/cas9 system. Specifically, a sgRNA targeting the 5’ end of the bhlhe22 coding sequence (that is contained on a single exon) was used to generate six founders carrying mutations in the bhlhe22 gene (Table 2.1; Figure 2.10A-D). One founder was characterized further and found to transmit a small deletion that introduces a frameshift, which is predicted to cause premature termination of Bhlhe22 protein synthesis upstream of the bHLH domain (Figure 2.2B, Figure 2.10E). We find that animals homozygous for this mutant allele (bhlhe22um320) are viable to
adulthood (Figure 2.11A). As expected, sequencing of bhlhe22
77
78
Figure 2.10. Generation of bhlhe22 germ line mutant.
A. Schematic showing genomic sequence of bhlhe22 with the bHLH domain indicated in blue. Note that bhlhe22 is contained on a single exon. The CRISPR target sequence is shown in red with the BstYI restriction site bracketed and the black arrow indicating the BstYI cut site. B. Identification of functional guide
RNAs. sgRNA and cas9 mRNA was injected into 1-cell stage embryos. Injected
embryos were raised to 24hpf and BstYI digestion of PCR amplicons from pools
of embryos was used to identify CRISPR-induced mutations (black arrow). C.
Identification of individual F0 founders. sgRNA/cas9 injected embryos were
raised to adulthood and crossed to wildtype fish. BstYI digests of PCR amplicons
from pools of embryos was used to identify F0 mosaic founders (black arrow). D.
Identification of F1 animals. Adult F0 mosaic founders were out-crossed to
wildtype fish and the F1 offspring raised to adulthood. BstYI digests of PCR
amplicons from fin clip genomic DNA was used to identify heterozygous F1
animals. E. Sequencing of F1 genomic DNA revealed the transmission of one
mutant allele (um320) carrying a 5 base pair deletion (black dashes). The
CRISPR target sequence is shown in red. F. Predicted amino acid sequence of
mutant allele. The um320 peptide shares its first 67 amino acids with the wildtype
protein before going out of frame and terminating at a premature stop codon N-
terminal to the bHLH domain.
79
80
Figure 2.11. Analysis of bhlhe22 mutant zebrafish.
A. Chart depicting the frequency of each genotype at various timepoints in
broods from crosses of bhlhe22+/um320 heterozygous fish. mo = month, y/o = year
old. B. Sequencing traces of transcripts from wild type versus bhlhe22um320/um320 animals showing the expected 5bp deletion. C-E. Expression of eng1b (C), evx1
(D) and vsx2 (E) in 24hpf wildtype and bhlhe22um320/um320 mutant embryos.
81
transcripts from such homozygous animals detected only the mutant sequence confirming presence of the mutant allele (Figure 2.11B). To test if bhlhe22 might
function with prdm12b in p1 formation, we examined eng1b expression in
bhlhe22um320 animals by in situ hybridization. We find that expression of eng1b is
unaffected in homozygous bhlhe22 mutants (Figure 2.11C). Since siRNA-
mediated knock-down of bhlhe22 reportedly disrupts gene expression in p0-p2 of
chick embryos [136], we also examined expression of vsx2 in the p2 domain
(Figure 2.11D) and evx1 in the p0 domain (Figure 2.11E), but do not observe any
disruptions. We conclude that, in contrast to the situation in chick, zebrafish
bhlhe22 is not required for p1 domain formation.
prdm12b does not maintain the p1 domain by repressing nkx6.1
Repressive interactions are common during formation of the neural tube,
whereby mutually repressive pairs of TFs are involved in the establishment of
individual progenitor domains (reviewed in [3,84]). Since prdm12b appears to act
as a repressor, it is plausible that it forms a repressive pair with nkx6.1 to
establish the p1 domain and permit eng1b expression. Accordingly, nkx6.1
mutant mice display a ventral expansion of the p1 domain at the expense of the
p2, pMN and p3 domains [16]. Furthermore, dorsal expansion of nkx6.1 has
been reported in prdm12 MO-injected fish and frog embryos [92,93] and
overexpression of prdm12 inhibits nkx6.1 expression in frog embryos. To test this 82
model further, we generated nkx6.1 mutant zebrafish by targeting a sgRNA to the
5’ end of exon 1. This produced eight founders carrying mutations in the nkx6.1
gene (Table 2.1; Figure 2.2C; Figure 2.12). Five of these were characterized
further and found to transmit two different mutant alleles. The nkx6.1um321 allele
contains a 23bp deletion while the nkx6.1um322 allele carries a 1bp insertion (as well as three single base pair substitutions). In both alleles, this leads to frameshifts that terminate at a premature stop codon upstream of the HOX domain. Accordingly, immunostaining with an anti-Nkx6.1 antibody revealed loss of Nkx6.1 protein in homozygous nkx6.1um321/ um321 mutants (Figure 2.13A).
Similar to the situation with prdm12b mutants, we find that homozygous
nkx6.1um321 mutant animals are observed at the expected ratio during early
development, but we detect only a few homozygous nkx6.1um321 animals at
adulthood (Figure 2.13B). While nkx6.1 mutant mice display a profound loss of
motor neurons [16], nkx6.1 MO-injected zebrafish show defective formation in only a subset of motor neurons and only at later stages of development
[138,139]. In general agreement with these MO-based zebrafish studies, we do not detect overt changes in expression of the hb9 motor neuron marker in nkx6.1 mutant zebrafish (Figure 2.13C), but we do observe subtle defects in the formation of branchiomotor neurons in the hindbrain (Figure 2.13D) 83
84
Figure 2.12. Generation of germ line nkx6.1 mutants.
A. Schematic showing genomic sequence of nkx6.1 with the homeodomain indicated in green. The CRISPR target sequence is shown in red with the AvaII restriction site bracketed and the black arrow indicating the AvaII cut site. B.
Identification of functional guide RNAs. sgRNA and cas9 mRNA was injected into
1-cell stage embryos. Injected embryos were raised to 24hpf and AvaII digestion of PCR amplicons from pooled embryos was used to identify CRISPR-induced mutations (black arrow). C. Identification of individual F0 founders. sgRNA/cas9 injected embryos were raised to adulthood and crossed to wildtype fish. AvaII digests of PCR amplicons from pools of embryos was used to identify F0 mosaic founders (black arrow). D. Identification of F1 animals. Adult F0 mosaic founders were out-crossed to wildtype fish and the F1 offspring raised to adulthood. AvaII digests of PCR amplicons from fin clip genomic DNA was used to identify heterozygous F1 animals. E. Sequencing of F1 genomic DNA revealed the transmission of two mutant alleles (um321, um322). um321 carries a 23 base pair deletion (black dashes) while um322 carries a 1 base pair insertion (green) and 3 base pair substitutions (blue). The CRISPR target sequence is shown in red. F. Predicted amino acid sequence of mutant alleles. The um320 and um321 peptides share their first 44 amino acids with the wildtype sequence before going out of frame and terminating at a premature stop codon N-terminal to the conserved homeodomain. G. Quantification of the size (along the dorsoventral 85
axis) of the nkx6.1 expression domain in prdm12b MO-injected embryos (data from Figure 2.14).
86
87
Figure 2.13. prdm12b does not maintain the p1 domain by repressing
nkx6.1.
A. Anti-Nkx6.1 immunostaining of nkx6.1um321/um321 mutant (left) and wildtype
(right) embryos at 30hpf. B. Chart indicating the frequency of each genotype at
various time points in broods from crosses of nkx6.1+/um321 heterozygous
mutants. C. Hb9 immunostaining in wildtype (left) versus a cross of nkx6.1+/um321 heterozygous embryos (right) at 33hpf. D. Islet-1/2 immunostaining of 50hpf embryos from a cross of nkx6.1+/um321 heterozygotes. E. Expression of eng1b in
24hpf embryos from a cross of nkx6.1+/um321 heterozygotes. F. Expression of
eng1b in 24hpf uninjected wildtype embryos (left panels), 24hpf prdm12b MO-
injected wildtype embryos (middle panels) and 24hpf prdm12b MO-injected
embryos from a cross of nkx6.1+/um321 heterozygotes (right panels). G.
Expression of prdm12b in a representative wildtype embryo (left panel) and a
representative embryo from a cross of nkx6.1+/um321 heterozygotes (middle panel)
at 24hpf. Right panel shows quantification of the size of the prdm12b expression
domain in 11 wildtype embryos and 20 embryos from a cross of nkx6.1+/um321 heterozygotes. Numbers in panels indicate the fraction of embryos displaying the phenotype shown.
88
We next used the nkx6.1 mutant fish to test if nkx6.1 and prdm12b act as
a repressive pair to establish the p1 domain and enable eng1b expression.
However, we do not find evidence for expansion of the eng1b (Figure 2.13E) or
prdm12b (Figure 2.13G) expression domains in nkx6.1 mutants. In accordance
with previous reports, we observe a slight expansion of the nkx6.1 domain in
prdm12b loss of function animals, but this effect falls below the level of statistical
significance (Figure 2.12G). Furthermore, if nkx6.1 and prdm12b act as a
repressive pair, nkx6.1 would expand into the p1 domain in prdm12b mutant
animals, thereby expanding the p2 domain at the expense of the p1 domain and leading to loss of eng1b expression. Therefore, we would expect eng1b
transcripts to be present in the p1 domain of animals lacking both nkx6.1 and
prdm12b function. To test this, we microinjected the prdm12b MO (that we know
phenocopies the prdm12b germ line mutant; see Fig. 1-3 [92]) into embryos from
a cross of heterozygous nkx6.1um321 carriers. We find that eng1b expression is
absent in all MO-injected embryos, regardless of nkx6.1 status (Figure 2.13F),
indicating that loss of eng1b expression is not the result of nkx6.1-mediated
expansion of the p2 domain. Lastly, if the loss of eng1b expression in prdm12b
mutants is due to expansion of adjacent domains, we would expect the p1
domain to be absent in prdm12b loss of function animals. Using five different
combinations of domain-specific genes as markers, we find that the p1 domain is
significantly smaller, but still present, in the absence of prdm12b function (Figure
2.14A-O). Since we measured the ratio of neural tube to p1 domain, the smaller 89
90
Figure 2.14. prdm12b controls the size of the p1 domain.
Expression of pax3/nkx6.1 (A, B), dbx1/nkx6.1 (D, E), pax3/vsx2 (G, H), dbx1/vsx2 (J, K) and evx1/nkx6.1 (M, N) in 24hpf wildtype (A, D, G, J, M) or prdm12b MO-injected (B, E, H, K, N) embryos. Panels show cross sections through the spinal cord with dorsal to the top. C, F, I, L, O show quantification of the size (along the dorsoventral axis) of the p0/p1 domain (C, I) or the p1 domain
(F, L, O) relative to the neural tube. At least 10 representative sections were used for each gene pair.
91
size of prdm12b mutant fish does not contribute to the reduced p1 domain size.
We conclude that prdm12b is required for establishing an appropriately sized p1
domain, not for preventing nkx6.1-mediated dorsal expansion of adjacent
domains.
DISCUSSION
We report the first germ line mutants disrupting function of the prdm12 TF.
In particular, we find that three distinct zebrafish prdm12b mutant alleles produce
an identical phenotype. We use these lines to extend previous characterization of
prdm12 loss of function animals to demonstrate that prdm12b is essential for
embryonic development, specifically for the formation of a Mauthner cell-
dependent neural circuit controlling a classical escape response. Using in vitro approaches, we further demonstrate that Prdm12b functions as a bona fide transcriptional repressor – most likely by recruiting EHMT2/G9a. Although
Prdm12b binds via its essential zinc-finger domain to the Bhlhe22 TF, generating and analyzing a bhlhe22 germ line zebrafish mutant revealed no effects on eng1
expression in the p1 domain – indicating that prdm12b and bhlhe22 do not need
to act together for p1 formation in vivo. Lastly, it has been suggested that prdm12b and nkx6.1 form a cross-repressive TF pair essential for the establishment of p1 domain fates. We tested this hypothesis by generating a nkx6.1 germ line zebrafish mutant and analyzing it along with our prdm12b 92
mutant, but do not find support for such a cross-repressive arrangement. In fact,
instead of the p1 domain taking on a p2 fate in prdm12b mutants, a domain
persists at the p1 position, but it does not express genes indicative of a specific
progenitor class.
prdm12b germ line mutants recapitulate the phenotype observed using antisense-based approaches
Prdm12 function has been addressed previously, but only by transient loss
of function approaches. In particular, antisense morpholino oligos (MOs) were
first used in zebrafish [92] and subsequently in frog [93,115] to disrupt prdm12
function. The resulting animals lack expression of eng1 in the p1 domain of the
neural tube, but gene expression appears relatively normal in adjacent domains.
eng1-expressing progenitors in the p1 domain are known to give rise to V1
interneurons that act in motor circuits (reviewed in [107]). Accordingly, fish and
frogs lacking prdm12 function display abnormal escape responses [92,93], but
the nature of this effect (excessive C-bends) suggests a defect in a reticulospinal
cell-controlled circuit that is likely independent of the loss of V1 interneurons.
Importantly, recent work has highlighted significant concerns with MO-based
approaches. In particular, there are many instances where germ line mutations
do not confirm previous reported MO-based phenotypes [129]. While some of
these cases may be explained by underappreciated compensatory mechanisms 93
[140], there are striking examples of MO phenotypes that turn out to be due to
non-specific or off-target effects [118]. Against this background, it is essential to
determine the phenotype of prdm12 germ line mutants. To address this, we used
CRISPR/Cas9 to generate two lines carrying frameshift mutations in the
zebrafish prdm12b gene and also obtained an ENU-induced splice-site mutation from the zebrafish resource center. All three lines display a phenotype that is in good agreement with MO-derived data. In particular, germ line mutants lack eng1b expression and display escape response defects indistinguishable from
those in MO injected embryos. Hence, our findings indicate that, in this case, the
various MOs act specifically. Since there is currently no available prdm12
knockout line in mouse, it remains possible that there will be species-specific
differences in prdm12 function, as was recently observed when comparing MO-
injected, zebrafish germ line mutants and mouse germ line mutants of the PG1
hox genes [141].
prdm12b is a bona fide transcriptional repressor
The Prdm12 TF has been suggested to act as a repressor based on
overexpression studies in vivo and in dissected frog embryos [93,115], but as an
activator based on transfection experiments in P19 cells [89]. To address this
discrepancy, we made use of classical reporter assays and find that zebrafish
Prdm12b efficiently represses expression from a luciferase reporter gene. Other 94
members of the Prdm family have been reported to act as repressors, but appear
to use distinct mechanisms to do so. For instance, several Prdm TFs recruit
histone deacetylases (HDACs) to repress transcription, but we find that an HDAC
inhibitor does not affect the repressive properties of Prdm12b, indicating that it
functions independently of HDACs. Overexpression of Prdm12 also promotes the
deposition of repressive methyl marks on H3K9 [89,93,115]. Accordingly, the PR
domain of some Prdm proteins exhibits methyltransferase activity and this
domain is required for Prdm12 function in Xenopus [93]. However, we find that mutating a key conserved PR domain residue does not affect the repressive activity of prdm12b. Accordingly, in vitro analyses using core histone substrates failed to detect intrinsic methyltransferase activity for Prdm12 [89]. Notably,
murine Prdm12 binds EHMT2/G9a (an H3K9 methyltransferase; [89]) and
EHMT2/G9a is required for Prdm12 function in Xenopus [93], suggesting that
Prdm12 may act as a repressor by recruiting EHMT2/G9a. We show that zebrafish Prdm12b also binds EHMT2/G9a, but in contrast to the situation in the mouse, the Prdm12b zinc finger domains are not required for this interaction.
In spite of the presence of several zinc finger domains, many Prdm
proteins require interactions with other TFs for targeting to genomic binding sites.
In particular, several Prdm proteins form complexes with bHLH TFs [84]. For
instance, Bhlhe22 is known to interact with Prdm TFs [71] and is required for
expression of eng1 in the chick neural tube [136], making it a candidate
interaction partner for Prdm12b. Indeed, we show by co-immunoprecipitation that 95
Prdm12b and Bhlhe22 can form a complex. Furthermore, this interaction requires
the Prdm12b zinc finger domain that we find is required for Prdm12b repressor
activity. To test the role for bhlhe22 in vivo, we used CRISPR/Cas9 to generate a germ line mutant in zebrafish, but we do not find any evidence that bhlhe22 is required for formation of the p1 domain in zebrafish embryos. It is not clear why loss of bhlhe22 function produces different effects in zebrafish versus chick, but this may stem from the different approaches used – germ line mutation in zebrafish versus transient siRNA-mediated knock-down in chick. The lack of a phenotype may also be the effect of compensatory mechanisms, either by other bHLH TFs - which are broadly expressed in the neural tube [35] – or by more general mechanisms operating to suppress the effects of genetic lesions [142].
We conclude that Prdm12b acts as a repressor of transcription – most likely by recruiting EHMT2/G9a – and that the Prdm12-mediated induction of genes such as p27 is most likely the result of indirect events.
An undefined domain persists at the p1 position in prdm12b mutants
The mechanism whereby prdm12 promotes formation of the p1 domain remains unclear. Mutual repression between TFs expressed in adjacent domains is the predominant mechanism for the creation of distinct domains along the dorsoventral axis of the vertebrate neural tube. Since prdm12 functions as a repressor, it is possible that it acts to repress the formation of adjacent domains. 96
Indeed, overexpression and MO-based approaches in the frog have led to the suggestion that prdm12 and nkx6.1 (that is expressed in the p2, p3 and pMN domains) forms such a cross-repressive pair [93]. In this model, loss of prdm12 would lead to loss of eng1 expression due to nkx6.1 expression (and p2 fates) expanding into the p1 domain. However, our initial analyses of nkx6.1 mutant zebrafish do not support this model. First, if prdm12b is required for eng1 expression in the p1 domain due to its repression of nkx6.1, eng1b should be restored to the p1 domain in embryos lacking both nkx6.1 and prdm12b, but this is not what we observe. Second, if prdm12b and nkx6.1 cross-repress each other’s expression, prdm12b expression should expand ventrally in nkx6.1 mutants and vice versa, but this also does not occur. Lastly, when one member of a cross-repressive pair is mutated, the corresponding progenitor fate is usually replaced by the adjacent fate, but this is not the case in prdm12b mutants – where a domain persists at the p1 position, albeit in a narrower form. Since this domain does not express any of the genes diagnostic for various fates along the
DV axis, its exact state is not clear. We note that prdm12 is reported to have anti- proliferative activity [89] and that p1 progenitor cells must exit the cell cycle prior to differentiating into V1 interneurons. It is therefore possible that prdm12 is required for this transition and that loss of prdm12 leaves cells in a proliferative progenitor state.
97
CONCLUSION
Our results demonstrate an essential role for prdm12b in zebrafish
neurogenesis. By generating germ line mutations, we show that a loss of function
prdm12b allele results in lack of eng1b expressing V1 interneurons and
subsequent defective Mauthner cell dependent locomotion, which is
indistinguishable from prdm12b morphants, and ultimately embryonic lethality.
Further analyses revealed that the Prdm12b zinc finger domain, which is
essential for repression, is also necessary for binding to the Bhlhe22 TF, but not
to EHMT2/G9a. We generated a bhlhe22 mutant zebrafish line, but find no
evidence for bhlhe22 function in the formation of the p1 domain in zebrafish
embryos. Lastly, upon examination of cross-repressive interaction between
prdm12b and nkx6.1, we do not find evidence for nkx6.1 and prdm12b acting as
a repressive pair in the formation of the p1/p2 boundary. Our results suggest that
prdm12b does not only regulate eng1b expression in the p1 domain, but also
takes part in regulating the size of this domain.
AUTHORS' CONTRIBUTIONS
OY participated in the design of the study, generated the CRISPR-based
mutant lines, performed all in situ and immunochemistry analysis of the mutant lines, carried out all reporter assays and co-immunoprecipitation experiments, 98
performed the computational analyses of escape responses and drafted the
manuscript. GBD carried out and recorded the escape response, assisted in data interpretation, reviewed, edited and approved the manuscript. CGS conceived
the study, secured funding, participated in study design, and finalized the
manuscript.
ACKNOWLEDGEMENTS
We are grateful to Dr. Scot Wolfe for advice regarding CRISPR/Cas9
mutagenesis. We acknowledge ZIRC for providing the prdm12bsa9887 mutant. The
following monoclonal antibodies were obtained from the Developmental Studies
Hybridoma Bank, created by the NICHD of the NIH and maintained at The
University of Iowa, Department of Biology, Iowa City, IA 52242: Isl1/2 (39.4D5)
antibody and HB9 (81.5C10) developed by T. M. Jessell and S. Brenner-Morton;
3A10 antibody developed by T. M. Jessell, J. Dodd, and S. Brenner-Morton;
F310 antibody developed by F.E. Stockdale. This work was supported by NIH
grant NS038183 to CGS and NSF grant IOS1456866 to GBD. CHAPTER III: DISCUSSION
100
Prdm12b function was addressed previously via transient loss of function
approaches. The use of antisense morpholino oligos (MOs) both in zebrafish and
frogs revealed that loss of prdm12b is associated with lack of expression of eng1b in the p1 domain. Since eng1b is an indicative marker for V1 interneuron formation, prdm12b knock-down animals display abnormal Mauthner neuron- dependent locomotion [92,93]. However, recent developments regarding the non- specific and off-target effects of MOs have raised much concern for MO-induced
phenotypes [129]. Several MO-induced phenotypes cannot be verified by germ line mutant studies [118]. Therefore, the generation of a prdm12b germline mutant became essential to determine the role of prdm12b in the context of
complete loss of function in vivo.
In this work, I present utilization of the CRIPSR/Cas9 genome editing tool
to create two mutant lines carrying frameshift mutations in the zebrafish prdm12b
gene. The first report of germline mutants disrupting the function of prdm12b is in
good agreement with MO-induced phenotype along with ENU-induced splice site
mutation from the Zebrafish Resource Center, which lacks of eng1b expression
in the p1 domain and results in abnormal touch-evoked response. Further
analysis of Prdm12b revealed that Prdm12b acts as a transcriptional repressor
most likely via recruiting histone methyltransferase EHMT2/G9a. Prdm12b also
binds the Bhlhe22 transcription factor via its zinc finger domain most likely to
access genomic sites in chromatin to execute transcriptional repression.
However, analysis of bhlhe22 germline mutants did not reveal a requirement of 101
bhlhe22 in the formation of the p1 domain in zebrafish. Lastly, our additional
analysis on the possibility of cross-repressive state between prdm12b and nkx6.1 to form the p1/p2 domain boundary showed that the requirement of prdm12b to drive eng1b in p1 is not through the virtue of repressing nkx6.1. Embryos doubly deficient for prdm12b and nkx6.1 failed to restore eng1b expression.
The mutant lines generated here provide insight into the role of prdm12b in nervous system development and that is critical for embryonic survival.
Despite the absence of a clear genetic hierarchy to fit prdm12b into neural tube patterning, this work shows key evidence for the role of prdm12b in transcriptional regulation within the p1 domain and p1-derived neurons.
CORRELATION BETWEEN MORPHOLINO-INDUCED AND MUTANT
PHENOTYPES IN PRDM12B LOCUS
MOs have a long history in zebrafish research to knock-down a targeted
gene. Recent advances revealed that many germ line mutants do not phenocopy
the morphant phenotype [130,142,143]. Recent research has even showed that
around 80% of published morphant phenotypes from Sanger Zebrafish Mutation
Project failed to confirm genetic mutations in the same genes [143]. Therefore, it is important to validate previously published morphant phenotypes in germ line mutants. In our project we aimed to test prdm12b germ line mutants not only for 102 validation of the relevant MO-induced phenotype but also in order to fully understand the role of prdm12b in zebrafish nervous system development.
The discrepancies between phenotypes with the two different approaches, i.e. MO vs. mutants, can be attributed to potential off-target effects of MOs, difficulty in establishing a standard MO dose, and the unknowns of level of homology required by each MO sequence for efficient binding [130]. In this project, I have analyzed prdm12b MO and three distinct zebrafish prdm12b mutant alleles (two from CRISPR/Cas9 genome editing and one from the
Zebrafish Mutation Project). They all yielded identical phenotype enabling us to use the prdm12b mutants as reliable tools for characterizing the function of prdm12b. For this reason, I have utilized both prdm12b morphants and mutants throughout this project to address this particular aspect of zebrafish neural development.
THE FATE OF THE REMAINING P1 DOMAIN IN PRDM12B MUTANTS IS
UNDEFINED
The characterization of prdm12b mutants revealed that the loss of eng1b expression in prdm12b mutants is the most striking phenotype. The change in eng1b expression and the subsequent loss of V1 interneurons led us to consider the remaining composition of cell types in the p1 domain after prdm12b knockout.
Since both prdm12b germ line mutants and morphants have exactly the same 103 phenotype and due to the trouble caused by continuous dependence on heterozygous mutant in-crosses for any assay using prdm12b germ line mutants because they are not homozygous viable, I have moved forward with prdm12b morphants to analyze the size of the p1 domain in prdm12b morphants with double in situs followed by sectioning.
The cross-repressive nature of dorsoventral patterning during neurogenesis has also led to a shift of domain identities into neighboring domain characteristics upon loss of specific domain marker genes [15-17,144]. In particular, if loss of eng1b expression in prdm12b germ line mutants is due to expansion of nkx6.1-dependent repression, I would expect the p1 domain to be replaced by an expanded p2 domain. In this case, the p1 domain would be shifted to have p2 or p0 domain characteristics. To answer this, I have used five different combinations of domain-specific genes as markers namely pax3/nkx6.1, dbx1/nkx6.1, pax3/vsx2, dbx1/vsx2 and evx1/nkx6.1. Our results revealed that p1 domain size is smaller upon prdm12b knockdown and there is no significant dorsal expansion of nkx6.1, contrary to the previous publications where nkx6.1 dorsal expansion was observed upon prdm12b knockdown (Figure 2.14) [92,93].
Since the p1 domain does not express any of the genes tested above specifically upon prdm12b loss, the exact composition of the p1 domain is still under question. Determining the composition of the remaining p1 domain in prdm12 morphants would require analyzing the gene markers at later stages of embryonic development. We know that prdm12b mutants are smaller in body 104 size and do not reach adulthood (Figure 2.4C). Expression analysis after 24 hpf may reveal the trajectory of the p1 domain in terms of its composition. In particular, expression of additional post-mitotic genes such as pax2, lhx1/5, lhx3, foxd3 and foxp2 would be the first to test whether the remaining p1 retains its “p1 domain” identity [25]. Not only expression analysis but also quantification of in situ and antibody staining would allow subtle changes upon prdm12b loss. Since the effect of prdm12b loss on eng1 expression is highly restricted-only V1 interneurons- the importance of quantification of any gene markers would reveal a specific patterning defect.
It is worth considering whether the reduction in size of the p1 domain is caused by cell death. Apoptotic cells can be detected via various methods such as acridine orange (AO) staining, TUNEL assay and activated Cas3 assay [145].
All three methods are straightforward and well-established, but standard techniques for detecting apoptotic cells are technically challenging in whole mounts due to the need for co-labeling in order to identify the specific cell types undergoing apoptosis. Besides, we do not necessarily know the exact genetic composition for the remaining p1 domain in prdm12 morphants and the choice of a co-labeling marker for apoptotic cells in the p1 domain has many limitations.
However, recent advances in genome editing can also be applied to visualization of apoptotic cells. Successful CRISPR/Cas9 mediated knock-in zebrafish could be one option to adapt protocols for apoptotic cells [146]. For example, live imaging of apoptotic cells in GFP knock-in prdm12b fish would be more indicative 105
of whether prdm12b+ cells are dying in the course of neural development. To
better illustrate, the coding sequence of GFP or any other fluorescent reporter
can be knocked-in upstream of the ATG site of the prdm12b locus, thus, placing it under the control of the endogenous promoter/enhancer elements with designed disruption of prdm12b gene expression. This approach would enable to
knockout prdm12b and knockin GFP reporter line at the same time with the use
of a single CRISPR/Cas9 targeting event.
Despite the evidence supporting the role of prdm12b in regulating eng1b
expression during embryogenesis, the exact genetic pathway driving V1
interneuron differentiation in p1 domain remains unclear. How this pathway can
be understood, is described below.
THE ROLE OF PRDM12B IN DORSOVENTRAL PATTERNING IS COMPLEX
Further in vivo analysis of Prdm12b interaction partners namely Bhlhe22 and
EHMT2/G9a is required
Bhlhe22 interaction
As presented in Chapter II, I have used a candidate approach to identify
Prdm12b interactor proteins. I had narrowed down my search into two strong
candidates, Bhlhe22 and EHMT2/G9a. 106
According to the Bhlhe22 binding assay, I speculate that Prdm12b does not contain a DNA binding domain since three zinc fingers (ZnF) may not be sufficient for DNA binding. Indeed, there are several examples of Prdm proteins acting in complexes with DNA-binding transcription factors [68,71]. I have limited my search for potential Prdm12b binding partners to those expressed in the p1 domain like bhlhe22. Surprisingly, Bhlhe22 is not only expressed in the pdl6, p1, p2 and p3 domains, but also implicated in the formation of spinal interneurons
[136]. Based on these criteria, the Bhlhe22 transcription factor represents a potential binding partner for Prdm12b. Using co-immunoprecipitation, we demonstrated robust binding between Bhlhe22 and Prdm12b (Figure 2.9A, lane
6), more specifically that the ZnF domain is essential for this interaction while the
PR domain is not.
Previous work reported that siRNA-mediated knock-down of bhlhe22 in the chick spinal cord leads to a decrease in eng1 expression in the p1 domain and subsequent reduction in V1 interneurons [136]. In line with our findings regarding the Prdm12b-Bhlhe22 interaction, I have hypothesized that bhlhe22 and prdm12b may cooperate in vivo to regulate eng1b expression and V1 interneuron formation in the p1 domain. I generated a mutant line lacking bhlhe22 using the CRISPR/Cas9 system. In contrast to studies done in chicks, bhlhe22 germ line mutants did not show any change in eng1b expression in the p1 domain [136]. This led us to conclude that zebrafish bhlhe22 is not required 107
for V1 interneuron formation despite their robust interaction with Prdm12b in
vitro.
This discrepancy can be explained by compensatory mechanisms
suggested by a recent study in zebrafish. The activation of compensatory
mechanisms is observed in gene knockouts but not in morphants [142].
Compensatory mechanisms may have mitigated any deleterious effects of the
bhlhe22 deletion in the zebrafish genome and masked any observable effects on
the p0-p2 domain markers’ expression in bhlhe22 mutants. In order to address
potential genetic compensation in bhlhe22 mutants, mutant vs. wildtype transcriptome and proteome comparison with the use of RNA profiling and mass
spectrometry would help to identify a set of genes and proteins that are
upregulated in mutants but not in wildtypes.
Another way of testing the involvement of Bhlhe22 on Prdm12b function is
to design an in vitro assay where we can compare the fold reduction in luciferase
activity upon addition of Bhlhe22 construct along with Prdm12b construct in an in
vitro assay used in Figure 2.7. In this assay, the luciferase plasmid can be
designed in such a way that it harbors a Bhlhe22 binding site while the Prdm12b
constructs do not contain any GALDBD. If Prdm12b requires Bhlhe22 to bind
DNA effectively, Bhlhe22 sites might be sufficient for Prdm12b to bind upstream
of the luciferase plasmid and repress luciferase expression. This may further
prove that Prdm12b and Bhlhe22 act together to form a repressive complex. 108
A candidate approach to identify Prdm12b interactors may also require
additional potential interactors such as Dbx2. dbx2 is a homeodomain
transcription factor which binds DNA and is also expressed in the p1 domain.
The loss of dbx2 is implicated in the spinal cord patterning [144,147]. By using biochemical assays such as GST pull downs or coimmunoprecipitation, the potential interaction between Dbx2 and Prdm12b can be investigated.
EHMT2/G9a interaction
The Prdm family of proteins is closely related to SET domain- a catalytic
domain with histone methyl transferase (HMT) activity- proteins. This has led the
researchers to investigate whether Prdm members have intrinsic HMT activity.
Since Prdm12b appears to lack intrinsic methyltransferase activity (Figure 2.7C),
it would need to recruit additional factors to mediate its repressive effects. In
particular, Prdm1, 5, 6 and 12 have been shown to bind EHMT2/G9a – a H3K9
methyltransferase [45,48,62,137]. In the case of Prdm12, binding to EHMT2/G9a
is reportedly mediated by the second ZnF domain [89] and Prdm12 acts as a
G9a-dependent transcriptional repressor for V1 interneuron specification [93].
Considering this is the ZnF domain that is required for binding to Bhlhe22, I
examined this in further detail. I have also confirmed the Prdm12 and
EHMT2/G9a interaction but found that the ZnF domain is not required for such
binding. 109
Work in Xenopus has shown that Prdm12 along with a dominant negative
G9a fails to induce en1 and no longer represses nkx6.1 or dbx1 [93]. Additionally in zebrafish, it has been shown that the loss of g9a via morpholino is linked to disruptions in neurogenesis [148,149]. The question of involvement of
EHMT2/G9a in prdm12b-dependent p1 domain formation remains elusive.
Further investigation of G9a dependent Prdm12 repression for spinal cord
patterning- observed in Xenopus- is necessary to understand the involvement of
ehmt2/g9a in regulating eng1b expression in the p1 domain and V1 interneuron
formation together with prdm12b. Loss of ehmt2/g9a via morpholino can be
achieved in WT zebrafish embryos and test for any differences in eng1b
expression and p0-p2 domain markers. This experiment would not only show
whether ehmt2/g9a is important regulators of eng1 and V1 interneuron
determination, but would also indicate the in vivo interacting partner for prdm12b.
What are the regulatory relationships of a subset of genes in the p0-p2?
Dorsoventral patterning (D-V) of the neural tube is predominantly
established. The genes that are expressed in each progenitor domain are well
known. The transcription factors expressed in one particular progenitor domain
often repress neighboring domain markers to establish domain boundaries. In
particular, dbx2 (expressed in p0 and p1) and nkx6.1 (expressed in p1, p2, pMN 110 and p3) repress each other’s expression to establish the p0/p1 border [15-
17,144,147]. In addition, eng1b is repressed by nkx6.1 [15] (Figure 3.1).
Given that prdm12b acts as a repressor, I hypothesized that prdm12b represses nkx6.1 expression and that its effect on eng1b expression is an indirect outcome of nkx6.1 repression. In other words, nkx6.1 would expand into the p1 in prdm12b mutant animals, thus expanding the p2 domain and leading to loss of eng1b expression. This was supported in fish and frog systems where MO knockdown of prdm12b led to dorsal expansion of nkx6.1 [92,93]. Considering the cross-repressive nature of the regulatory relationship that governs domain boundaries (Figure 3.1), I also decided to look into the nkx6.1 mutants. Results from those studies revealed no noticeable changes in eng1b expression. Even in the absence of both nkx6.1 and prdm12b, loss of eng1b expression is not the consequence of nkx6.1-mediated expansion of the p2 domain (Figure 2.13F).
Given that our findings did not support a such cross-repressive interaction between prdm12b and nkx6.1, there is an alternative approach to clarify the regulatory relationships between prdm12b, nkx6.1 and eng1b. Driving mosaic eng1b expression in prdm12b mutant animals using heat-shock constructs would answer whether eng1b is sufficient to drive V1 interneurons and prdm12b must not be required for V1 interneuron formation other than driving eng1b expression.
To this end, prdm12b mutant animals can be injected with a construct where a heat-shock promoter drives expression of eng1b and GFP from a polycistronic mRNA. Since eng1b cannot be used as a marker for V1 interneurons due to its 111
Figure 3.1. Regulatory relationships of a subset of genes expressed in the p0-p2 domains.
Known interactions between gene pairs expressed in the p0-p2 domains are represented. The relationships between prdm12b/eng1b and prdm12b/nkx6.1 gene pairs are under question.
112
loss in prdm12b mutants, GFP could be a marker to determine whether V1 interneurons are formed. The projections of GFP positive cells can be assessed and compare to V1 interneuron morphology and projections via microscopy.
Since I could not find any evidence for a cross-repressive interaction
between prdm12b and nkx6.1 in our study (Figure 2.13G, Figure 2.14M-O), our
simplified model for regulatory relationships of p0-p2 genes (Figure 3.1) need to be addressed further via different epistatic genetic analyses. A recent publication
on the role of Dbx1 in the spinal cord development revealed that clearance of
Dbx1 in the p1 domain is critical for V1 interneuron differentiation [99]. Evx1 was
also shown to repress En1 (Eng1b) and to regulate spinal interneuron fate
decisions in mice [98]. In light of these facts, I realize that a complex network of
transcriptional activity specifies domain identities and boundaries along the
dorsoventral axis. Integration of prdm12b into a genetic pathway for p1 domain
formation and V1 interneuron differentiation would require more epistasis
analyses including dbx1, dbx2, evx1 and vsx2.
THE RELATIONSHIP OF GENE EXPRESSION THROUGH THE P0-P2
DOMAIN MARKERS DOES NOT INVOLVE SIMPLE DIRECT INTERACTIONS
Although there is considerable evidence for how eng1b is affected through
repressive interactions along the p0-p2 domain [15,98,99], formation of a network
integrating all affecting genes, including prdm12b, would be challenging and 113 needs additional examination. The questions of whether prdm12b directs eng1b expression via direct or indirect ways and whether prdm12b needs binding partners to target eng1b, remains elusive. One way of mapping prdm12b in a genetic pathway for the requirement of eng1b expression and V1 interneuron formation is to identify additional changes in global gene expression.
To further characterize the underlying cause of the change in eng1b expression, V1 interneuron loss and lethality in prdm12b mutants, single-cell
RNA-seq would provide information regarding all transcripts that are differentially expressed between wildtype and prdm12b mutants. Understanding which transcripts are affected by the loss of prdm12b would provide mechanistic insight into the role of prdm12b in regulating eng1b expression in the p1 domain. Single- cell techniques have recently been utilized to analyze the transcriptome of many different cell types due to their finetuning resolution on cell-to-cell variation
(reviewed in [150]). Since the p1 domain constitutes a very small area compared to the whole organism, preparation of tissue for RNA-seq to examine global changes in gene expression would be challenging with regards to tissue dissection. In addition, loss of eng1b is only observed in V1 interneurons not in the MHB or somites of prdm12b mutants. Since the effect of prdm12b knock-outs on eng1b is in very small scale in terms of overall size of the embryo, the choice of single-cell RNA-seq would be advantageous over standard RNA-seq in terms of the assessment of differences between cell types. Now that prdm12b mutants have shown embryonic lethality (Figure 2.4C) and the need to rely on 114
heterozygous in crosses for experimentation using prdm12b mutants, tissue preparation for next generation sequencing and selection of prdm12b mutants for
RNA-seq library from the heterozygous incross population prior to genotyping would be challenging. However, since prdm12b mutants have a behavioral phenotype (Figure 2.5), we could select prdm12b homozygous mutants prior to their genotyping based on their impaired touch-evoked response. The 4 dpf old embryos, which show an excessive C-bend upon a touch stimulus, can be selected for making an RNA-seq library followed by genotyping could follow for further confirmation. This way, the effect of prdm12b loss would not be diluted in the heterozygous incross population.
Another way of testing how prdm12b directs eng1b expression in the p1 domain is to identify interacting proteins in vivo. Instead of a candidate protein approach based on same expression domain with prdm12b, an unbiased assay like mass spectrometry would be a better choice for the search of interacting proteins. Recently, zebrafish research has advanced using mass spectrometry
[151,152]. This would require the generation of a CRISPR/Cas9 knock-in prdm12b fish line with GFP, FLAG, or Myc since we had no success in detecting endogenous prdm12b via available antibodies. Then endogenous prdm12b can be immunoprecipitated from the knock-in line using the corresponding antibody and mass spectrometry may be used to identify interacting proteins. However, further analysis of identified interactors via knockdown or knockout studies should follow in order to assess the effect on eng1b expression. 115
REMAINING QUESTIONS AND FUTURE DIRECTIONS OF THIS WORK
The work presented here clearly demonstrates a requirement for prdm12b for embryonic survival, eng1b expression, V1 interneuron formation, and a normal touch-evoked escape response. However, neither the mechanism nor the nature of the defect resulting from the loss of prdm12b are not yet fully clear. To further investigate the role of prdm12b, there are several questions to address.
What are the prdm12b direct targets?
To better understand Prdm12b function, we need to identify genes that are directly repressed by Prdm12b. Considering that Bhlhe22 is a Prdm12b binding partner via an in vitro approach, it is likely that Prdm12b direct targets carry
Bhlhe22 binding sites in order to ensure access of Prdm12b to their regulatory site.
There is one direct approach to determine prdm12b target genes. The genes likely to be repressed by Prdm12b might have a Bhlhe22 (CATATGNTNT) binding site in their enhancer/promoter regions. With the use of ChIP-seq for
Prdm12b or candidate approach, we could determine whether Prdm12b is present at those binding sites. A candidate gene approach would list the genes likely to be repressed by prdm12b as either expressed in p0 or in p2 but not in p1 domain. Such genes may be listed as nkx6.1, evx1, dbx1, and vsx2. These 116 would be candidates for prdm12b direct targets. ChIP-seq would be a global and unbiased approach for the search of prdm12b target genes. However, ChIP-seq has some limitations for applications with Prdm12b. First of all, there is currently no suitable Prdm12b antibody for zebrafish based on our own experience.
Second, injection of an epitope-tagged version of the protein, such as FLAG- tagged Prdm12b, into embryos would bring its own disadvantages. For example, the overexpression that usually results from such injections might have different manifestations versus endogenous prdm12b expression. To this end, generation of a knock-in line would be ideal to carry to ChIP-seq experiment in order to determine prdm12b direct target genes. I have attempted to generate three different CRISPR/Cas9 knock-in lines for prdm12b using FLAG, Myc and GFP; however, they all failed to result in stable lines carrying the desired epitope tag at the appropriate gene locus. Nevertheless, different single guide RNAs or different length and design of epitope tags could successfully generate CRISPR/Cas9 knock-in lines for prdm12b in the future. Using such knock-in lines and a candidate approach for genes likely to be repressed by prdm12b, we would potentially identify direct targets of prdm12b.
A recent study suggested that dbx1, nkx6.1 and nkx6.2 genes were shown to be directly repressed by Prdm12 using neuralized RA-treated animal caps overexpressing mouse Prdm12b [93]. Prdm12 binding sites were also observed upstream of post mitotic genes such as pax2, foxd3, lhx1/5 [93]. These results remain to be questionable at least in the context of zebrafish prdm12b. Since 117 prdm12b is only expressed in progenitor cells in zebrafish, and Prdm12 overexpression state does not necessarily recapitulate the endogenous prdm12b expression, further investigation remains to be necessary for such direct targets of prdm12b.
Do p1-derived neurons restrict Mauthner cells to fire only once?
prdm12b knockout animals have an abnormal escape response. While
WT fish respond to a head stimulus by performing a single large body bend (C bend) followed by moving away from the source of stimulus, prdm12b disruption leads to excessive C bends suggesting a defect in the Mauthner neuron circuitry.
Locomotion circuitry driving the Mauthner cell-dependent pathway is summarized as in Figure 3.2. Any defects along the transmission of neural input can manifest itself as a defective escape reflex. Even though Mauthner neuron morphology and the structure of trunk and tail musculature appear to be normal in prdm12b mutants (Figure 2.5E), electrophysiology of Mauthner neuron firing might be defective. The escape phenotype of prdm12b mutants is most readily explained by the Mauthner neuron firing repeatedly due to the lack of p1-derived glycinergic inhibitory interneurons. The excessive Mauthner neuron firing can be speculated as a lack of inhibitory feedback mechanism provided by V1 interneurons. In order to determine whether p1-derived neurons restrict Mauthner neurons to fire only once, electrophysiological recordings would address whether Mauthner neurons 118
fire repeatedly upon loss of prdm12b. Meanwhile, the involvement of glycinergic
inhibitory input into Mauthner neuron circuit could also be tested if p1-derived glycinergic V1 interneurons contact Mauthner cells. Since glycinergic neurons start to form in the zebrafish hindbrain around 24 hpf and make contact with
Mauthner neurons by 36 hpf [153,154], anti-glycine immunostaining, glyt2 in situ hybridization, or the glyt2:GFP transgenic line could help to assay whether prdm12b mutant fish lack glycinergic contacts with Mauthner neurons [153-155].
Furthermore, the loss of V1 interneurons can lead to rewiring of the remaining neurons in the locomotion circuitry. Such reconfiguration of neurons would almost certainly lead to abnormalities in terms of synapse formation and the transmission of electrical signals. Any changes in commissural axon projections would explain such reconfiguration of neurons. To test that, anti- neurofilament antibody aRM044 staining would be a good way to test commissural trajectories of reticulospinal Mauthner neurons and its homologs
MiD2 and MiD3 cells.
119
Figure 3.2. Schematic diagram depicting core elements of locomotion circuitry of distinct ventral interneuron classes with transcriptional identity of p1 derived V1 interneurons
[156].
120
CONCLUSION
In conclusion, this work establishes an essential role for prdm12b in neurogenesis in zebrafish. By generating germline mutants, I show that a loss of function prdm12b allele results in loss of eng1b expressing V1 interneurons, a subsequent defect in Mauthner cell-dependent locomotion, and ultimately embryonic lethality. In accordance with previous work, I have found that Prdm12b functions as repressor, and that this repressor activity requires an intact zinc finger domain [93,115]. Through in vitro studies, I have also shown that Prdm12b binds to both Bhlhe22 and EHMT2/G9a. The zinc finger domain of Prdm12b, which is essential for its repression activity, is necessary for binding to Bhlhe22 but not to EHMT2. By extrapolating our in vitro findings to in vivo settings, we find no evidence for the requirement of bhlhe22 in the formation of the p1 domain in zebrafish embryos. This suggests that the broad range of expression profiles of other bHLH transcription factors may play a role in compensating for the loss of bhlhe22 in neural tube patterning. Lastly, since Prdm12b acts as repressor
(Figure 2.7) and mutual repression between TFs expressed in neighboring domains reinforces domain boundaries during dorsoventral patterning (reviewed in [3], our analysis indicated that the loss of eng1b expression is not the manifestation of nkx6.1-mediated expansion of p2. Also, prdm12b mutants leave an undefined domain at the p1 position. In contrast to mouse studies where loss of nkx6.1 shifts ventral fates into adjacent dorsal fates [16], the regulation of 121 domain-specific gene expression might be species-dependent as in the case of zebrafish. Further genetic epistasis analyses would continue to provide insight into the complex regulatory relationship between the p1 and p2 domains.
APPENDIX: THE EFFECT OF PRDM12B AND BHLHE22 MISEXPRESSION
123
INTRODUCTION
prdm12b loss via MO is associated with the eng1b loss in both zebrafish and Xenopus [84,93]. I have further confirmed the necessity of prdm12b expression in order to drive eng1b in V1 interneurons in this work. This direct interaction between prdm12b and eng1b also needs to be tested in gain of function assays. Recent report suggested that prdm12b overexpression led to increase in en1 expression in Xenopus [93]. Moreover, misexpression of Bhlhb5
(Bhlhe22) in chick spinal cord led to the increase in En1 expression and loss of
Bhlhb5 via shRNA caused reduced En1 expression [136]. These two neural fate determining genes, prdm12b and bhlhe22, have a direct effect on V1 interneuron marker eng1.
METHODS
Zebrafish care
Zebrafish were handled as discussed in Chapter II.
Zebrafish embryonic injections
Embryos were collected from wild type matings. Embryos were then aligned on an 1% agarose mold and injected with 1-2ng of injection mix using a 124
borosil needle, micromanipulator, and dissecting microscope. For the injections
of prdm12b or bhlhe22 mRNA, a plasmid containing the full coding sequence of
prdm12b or bhlhe22 was in vitro transcribed. These mRNA were diluted in water
and phenol red to a final concentration of 200-400 ng/µl and injected into one-cell embryos either separately or together.
In situ RNA hybridization
Embryos were fixed in 4% paraformaldehyde (PFA) and stored in 100% methanol at -20°C. In situ RNA hybridization was performed as described [120] followed by a color reaction using NBT/BCIP in 10% polyvinyl alcohol. RNA probes for the genes eng1b were synthesized as previously described [118].
Embryos were dissected from the yolk and flat mounted in 80% glycerol for imaging on bridged coverslips or sectioned as described [121]. Images were captured using a Nikon Eclipse E600 microscope equipped with spot RT color camera (model 2.1.1). Images were imported into Adobe Photoshop and adjustments were made to contrast, levels, color matching settings and cropping only. All adjustments were made to the entire image.
125
RESULTS
Gain of function prdm12b and/or bhlhe22 does not affect eng1b expression
To characterize the function of prdm12b and bhlhe22 together, I next performed gain of function assays in zebrafish embryos. Zebrafish embryos overexpressing either prdm12b and bhlhe22 alone or together were assayed for eng1b expression via in situ. prdm12b overexpression did not result in misexpression of eng1b in neither hindbrain nor spinal cord (Figure A.1A). Only a few embryos showed reduced expression of eng1b (4/42) or misexpression of eng1b (6/42), which are not significant. On the other hand, bhlhe22 overexpression did not cause a change in eng1b expression at all. All embryos showed normal eng1b expression pattern (Figure A.1B). To investigate synergistic effect of prdm12b and bhlhe22 overexpression, I have injected both mRNAs into the same embryos and assay for eng1b expression. Most of the embryos overexpressing both mRNAs showed normal eng1b expression pattern.
Only small number of embryos show either reduced (2/38) or increased (4/38) eng1b expression (Figure A.1C). 126
127
Figure A.1. prdm12b and bhlhe22 overexpression has no overt effect on eng1 expression in zebrafish.
Wild type embryos were microinjected either 200 ng prdm12b (A), 200 ng bhlhe22 (B) or both 400 ng prdm12b and 200 ng bhlhe22 (C). eng1b expression in 24hpf embryos (A-C). Numbers in each panel indicate the fraction of animals with the specified phenotype. Brackets indicate r4, arrows mark V1 interneurons and arrowheads mark somites. MHB = midbrain–hindbrain boundary, HB = hindbrain and SC = spinal cord.
128
DISCUSSION
Genetic relationship between two genes is thought to be bi-directional. If
loss of one gene causes loss of another gene, expected outcome for
misexpression of driving gene would be ectopic expression of another gene. The
expected outcome for prdm12b overexpression in zebrafish embryos was to
observe ectopic expression of eng1b in both hindbrain and spinal cord. Even if
we did not see a causative relationship between loss of bhlhe22 and eng1b
expression, we expected to see a synergistic effect on reduced eng1b
expression upon loss of prdm12b and bhlhe22 misexpression together. However, misexpression of prdm12b, bhlhe22 alone or together did not result in significant change in eng1b expression contrary to Xenopus and chick. This could be
partially explained by species specific effect on gene expression. The regulatory
relationship of eng1b expressing V1 interneurons could involve many other
genes expressed in p1 or neighboring domains in neural tube.
129
REFERENCES
1. Del Pino I, Rico B, Marín O. Neural circuit dysfunction in mouse models of neurodevelopmental disorders. Curr. Opin. Neurobiol. 2018;48:174–82.
2. Kandel E. Principles of Neural Science, Fifth Edition. McGraw Hill Professional; 2013.
3. Jessell TM, Sanes JR. Development. The decade of the developing brain. Curr. Opin. Neurobiol. 2000;10:599–611.
4. Schmidt R, Strähle U, Scholpp S. Neurogenesis in zebrafish - from embryo to adult. Neural Dev. BioMed Central; 2013;8:3.
5. Rentzsch F, Bakkers J, Kramer C, Hammerschmidt M. Fgf signaling induces posterior neuroectoderm independently of Bmp signaling inhibition. Dev. Dyn. 2004;231:750–7.
6. Okuda Y, Ogura E, Kondoh H, Kamachi Y. B1 SOX coordinate cell specification with patterning and morphogenesis in the early zebrafish embryo. van Heyningen V, editor. PLoS Genet. 2010;6:e1000936.
7. Melton KR, Iulianella A, Trainor PA. Gene expression and regulation of hindbrain and spinal cord development. Front. Biosci. 2004;9:117–38.
8. Ogura Y, Sasakura Y. Switching the rate and pattern of cell division for neural tube closure. Neurogenesis (Austin). Taylor & Francis; 2016;3:e1235938.
9. van Straaten HW, Hekking JW, Wiertz-Hoessels EJ, Thors F, Drukker J. Effect of the notochord on the differentiation of a floor plate area in the neural tube of the chick embryo. Anat. Embryol. 1988;177:317–24.
10. Wilson L, Maden M. The mechanisms of dorsoventral patterning in the vertebrate neural tube. Developmental Biology. 2005;282:1–13.
11. Straaten HWM, Hekking JWM. Development of floor plate, neurons and axonal outgrowth pattern in the early spinal cord of the notochord-deficient chick embryo. Anat. Embryol. 1991;184:55–63.
12. Beattie CE, Hatta K, Halpern ME, Liu H, Eisen JS, Kimmel CB. Temporal Separation in the Specification of Primary and Secondary Motoneurons in Zebrafish. Developmental Biology. 1997;187:171–82. 130
13. Chiang C, Litingtung Y, Lee E, Young KE, Corden JL, Westphal H, et al. Cyclopia and defective axial patterning in mice lacking Sonic hedgehog gene function. Nature. Nature Publishing Group; 1996;383:407–13.
14. Briscoe J, Sussel L, Serup P, Hartigan-O'Connor D, Jessell TM, Rubenstein JL, et al. Homeobox gene Nkx2.2 and specification of neuronal identity by graded Sonic hedgehog signalling. Nature. Nature Publishing Group; 1999;398:622–7.
15. Vallstedt A, Muhr J, Pattyn A, Pierani A, Mendelsohn M, Sander M, et al. Different levels of repressor activity assign redundant and specific roles to Nkx6 genes in motor neuron and interneuron specification. Neuron. 2001;31:743–55.
16. Sander M, Paydar S, Ericson J, Briscoe J, Berber E, German M, et al. Ventral neural patterning by Nkx homeobox genes: Nkx6.1 controls somatic motor neuron and ventral interneuron fates. Genes Dev. Cold Spring Harbor Laboratory Press; 2000;14:2134–9.
17. Briscoe J, Pierani A, Jessell TM, Ericson J. A homeodomain protein code specifies progenitor cell identity and neuronal fate in the ventral neural tube. Cell. 2000;101:435–45.
18. Novitch BG, Chen AI, Jessell TM. Coordinate regulation of motor neuron subtype identity and pan-neuronal properties by the bHLH repressor Olig2. Neuron. 2001;31:773–89.
19. Liem KF, Tremml G, Roelink H, Jessell TM. Dorsal differentiation of neural plate cells induced by BMP-mediated signals from epidermal ectoderm. Cell. 1995;82:969–79.
20. Liem KF, Tremml G, Jessell TM. A role for the roof plate and its resident TGFbeta-related proteins in neuronal patterning in the dorsal spinal cord. Cell. 1997;91:127–38.
21. Lee KJ, Jessell TM. The specification of dorsal cell fates in the vertebrate central nervous system. Annu. Rev. Neurosci. Annual Reviews 4139 El Camino Way, P.O. Box 10139, Palo Alto, CA 94303-0139, USA; 1999;22:261–94.
22. Streit A, Berliner AJ, Papanayotou C, Sirulnik A, Stern CD. Initiation of neural induction by FGF signalling before gastrulation. Nature. Nature Publishing Group; 2000;406:74–8.
23. Diez del Corral R, Breitkreuz DN, Storey KG. Onset of neuronal differentiation is regulated by paraxial mesoderm and requires attenuation of FGF signalling. Development. 2002;129:1681–91. 131
24. Molotkova N, Molotkov A, Sirbu IO, Duester G. Requirement of mesodermal retinoic acid generated by Raldh2 for posterior neural transformation. Mech. Dev. 2005;122:145–55.
25. Alaynick WA, Jessell TM, Pfaff SL. SnapShot: Spinal Cord Development. Cell. 2011;146:178–178.e1.
26. Megason SG, McMahon AP. A mitogen gradient of dorsal midline Wnts organizes growth in the CNS. Development. 2002;129:2087–98.
27. Wine-Lee L, Ahn KJ, Richardson RD, Mishina Y, Lyons KM, Crenshaw EB. Signaling through BMP type 1 receptors is required for development of interneuron cell types in the dorsal spinal cord. Development. The Company of Biologists Ltd; 2004;131:5393–403.
28. Ericson J, Rashbass P, Schedl A, Brenner-Morton S, Kawakami A, van Heyningen V, et al. Pax6 controls progenitor cell identity and neuronal fate in response to graded Shh signaling. Cell. 1997;90:169–80.
29. Qiu M, Shimamura K, Sussel L, Chen S, Rubenstein JL. Control of anteroposterior and dorsoventral domains of Nkx-6.1 gene expression relative to other Nkx genes during vertebrate CNS development. Mech. Dev. 1998;72:77– 88.
30. Pierani A, Brenner-Morton S, Chiang C, Jessell TM. A sonic hedgehog- independent, retinoid-activated pathway of neurogenesis in the ventral spinal cord. Cell. 1999;97:903–15.
31. Pabst O, Herbrand H, Takuma N, Arnold HH. NKX2 gene expression in neuroectoderm but not in mesendodermally derived structures depends on sonic hedgehog in mouse embryos. Dev. Genes Evol. 2000;210:47–50.
32. Zhou Q, Anderson DJ. The bHLH transcription factors OLIG2 and OLIG1 couple neuronal and glial subtype specification. Cell. 2002;109:61–73.
33. Helms AW, Johnson JE. Specification of dorsal spinal cord interneurons. Curr. Opin. Neurobiol. 2003;13:42–9.
34. Chesnutt C, Burrus LW, Brown AMC, Niswander L. Coordinate regulation of neural tube patterning and proliferation by TGFbeta and WNT activity. Developmental Biology. 2004;274:334–47.
35. Bertrand N, Castro DS, Guillemot F. Proneural genes and the specification of neural cell types. Nat Rev Neurosci. Nature Publishing Group; 2002;3:517–30. 132
36. Huang S. Histone methyltransferases, diet nutrients and tumour suppressors. Nature Reviews Cancer. Nature Publishing Group; 2002;2:469–76.
37. Fog CK, Galli GG, Lund AH. PRDM proteins: important players in differentiation and disease. Bioessays [Internet]. 2012;34:50–60. Available from: http://www.ncbi.nlm.nih.gov/pubmed/22028065
38. Rea S, Eisenhaber F, O'Carroll D, Strahl BD, Sun ZW, Schmid M, et al. Regulation of chromatin structure by site-specific histone H3 methyltransferases. Nature. 2000;406:593–9.
39. Eom GH, Kim K, Kim S-M, Kee HJ, Kim J-Y, Jin HM, et al. Histone methyltransferase PRDM8 regulates mouse testis steroidogenesis. Biochemical and Biophysical Research Communications [Internet]. 2009;388:131–6. Available from: http://linkinghub.elsevier.com/retrieve/pii/S0006291X09015058
40. Hayashi K, Yoshida K, Matsui Y. A histone H3 methyltransferase controls epigenetic events required for meiotic prophase. Nature. Nature Publishing Group; 2005;438:374–8.
41. Derunes C, Briknarová K, Geng L, Li S, Gessner CR, Hewitt K, et al. Characterization of the PR domain of RIZ1 histone methyltransferase. Biochemical and Biophysical Research Communications [Internet]. 2005;333:925–34. Available from: http://www.ncbi.nlm.nih.gov/pubmed/15964548
42. Wu Y, Ferguson JE, Wang H, Kelley R, Ren R, McDonough H, et al. PRDM6 is enriched in vascular precursors during development and inhibits endothelial cell proliferation, survival, and differentiation. Journal of Molecular and Cellular Cardiology [Internet]. 2008;44:47–58. Available from: http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=PMC2683064
43. Pinheiro I, Margueron R, Shukeir N, Eisold M, Fritzsch C, Richter FM, et al. Prdm3 and Prdm16 are H3K9me1 methyltransferases required for mammalian heterochromatin integrity. Cell. 2012;150:948–60.
44. Kim K-C, Geng L, Huang S. Inactivation of a histone methyltransferase by mutations in human cancers. Cancer Res. 2003;63:7619–23.
45. Davis CA, Davis CA, Haberland M, Haberland M, Arnold MA, Arnold MA, et al. PRISM/PRDM6, a transcriptional repressor that promotes the proliferative gene program in smooth muscle cells. Molecular and Cellular Biology [Internet]. 2006;26:2626–36. Available from: http://mcb.asm.org/cgi/doi/10.1128/MCB.26.7.2626-2636.2006 133
46. Su S-T, Ying H-Y, Chiu Y-K, Lin F-R, Chen M-Y, Lin K-I. Involvement of histone demethylase LSD1 in Blimp-1-mediated gene repression during plasma cell differentiation. Molecular and Cellular Biology. American Society for Microbiology Journals; 2009;29:1421–31.
47. Ancelin K, Lange UC, Hajkova P, Schneider R, Bannister AJ, Kouzarides T, et al. Blimp1 associates with Prmt5 and directs histone arginine methylation in mouse germ cells. Nat Cell Biol [Internet]. 2006;8:623–30. Available from: http://www.ncbi.nlm.nih.gov/pubmed/16699504
48. Yu J, Angelin-Duclos C, Greenwood J, Liao J, Calame K. Transcriptional Repression by Blimp-1 (PRDI-BF1) Involves Recruitment of Histone Deacetylase. Molecular and Cellular Biology. 2000;20:2592–603.
49. Alliston T, Ko TC, Cao Y, Liang Y-Y, Feng X-H, Chang C, et al. Repression of bone morphogenetic protein and activin-inducible transcription by Evi-1. J. Biol. Chem. American Society for Biochemistry and Molecular Biology; 2005;280:24227–37.
50. Chittka A, Arevalo JC, Rodriguez-Guzman M, Pérez P, Chao MV, Sendtner M. The p75NTR-interacting protein SC1 inhibits cell cycle progression by transcriptional repression of cyclin E. J. Cell Biol. Rockefeller University Press; 2004;164:985–96.
51. Takahata M, Inoue Y, Tsuda H, Imoto I, Koinuma D, Hayashi M, et al. SKI and MEL1 cooperate to inhibit transforming growth factor-beta signal in gastric cancer cells. J. Biol. Chem. 2009;284:3334–44.
52. Yoshimi A, Goyama S, Watanabe-Okochi N, Yoshiki Y, Nannya Y, Nitta E, et al. Evi1 represses PTEN expression and activates PI3K/AKT/mTOR via interactions with polycomb proteins. Blood. American Society of Hematology; 2011;117:3617–28.
53. Hohenauer T, Moore AW. The Prdm family: expanding roles in stem cells and development. Development [Internet]. 2012;139:2267–82. Available from: http://www.ncbi.nlm.nih.gov/pubmed/22669819
54. Ren B, Chee KJ, Kim TH, Maniatis T. PRDI-BF1/Blimp-1 repression is mediated by corepressors of the Groucho family of proteins. Genes Dev. Cold Spring Harbor Laboratory Press; 1999;13:125–37.
55. Endo K, Karim MR, Taniguchi H, Krejci A, Kinameri E, Siebert M, et al. Chromatin modification of Notch targets in olfactory receptor neuron diversification. Nat Neurosci. Nature Publishing Group; 2011;15:224–33. 134
56. Izutsu K, Kurokawa M, Imai Y, Maki K, Mitani K, Hirai H. The corepressor CtBP interacts with Evi-1 to repress transforming growth factor beta signaling. Blood. 2001;97:2815–22.
57. Kajimura S, Seale P, Tomaru T, Erdjument-Bromage H, Cooper MP, Ruas JL, et al. Regulation of the brown and white fat gene programs through a PRDM16/CtBP transcriptional complex. Genes Dev. 2008;22:1397–409.
58. Nishikata I, Nakahata S, Saito Y, Kaneda K, Ichihara E, Yamakawa N, et al. Sumoylation of MEL1S at lysine 568 and its interaction with CtBP facilitates its repressor activity and the blockade of G-CSF-induced myeloid differentiation. Oncogene. Nature Publishing Group; 2011;30:4194–207.
59. Palmer S, Brouillet JP, Kilbey A, Fulton R, Walker M, Crossley M, et al. Evi-1 transforming and repressor activities are mediated by CtBP co-repressor proteins. J. Biol. Chem. 2001;276:25834–40.
60. Quinlan KGR, Nardini M, Verger A, Francescato P, Yaswen P, Corda D, et al. Specific recognition of ZNF217 and other zinc finger proteins at a surface groove of C-terminal binding proteins. Molecular and Cellular Biology. American Society for Microbiology Journals; 2006;26:8159–72.
61. Van Campenhout C, Nichane M, Antoniou A, Pendeville H, Bronchain OJ, Marine J-C, et al. Evi1 is specifically expressed in the distal tubule and duct of the Xenopus pronephros and plays a role in its formation. Developmental Biology. 2006;294:203–19.
62. Duan Z, Person RE, Lee H-H, Huang S, Donadieu J, Badolato R, et al. Epigenetic regulation of protein-coding and microRNA genes by the Gfi1- interacting tumor suppressor PRDM5. Molecular and Cellular Biology. 2007;27:6889–902.
63. Bard-Chapeau EA, Jeyakani J, Kok CH, Muller J, Chua BQ, Gunaratne J, et al. Ecotopic viral integration site 1 (EVI1) regulates multiple cellular processes important for cancer and is a synergistic partner for FOS protein in invasive tumors. Proc. Natl. Acad. Sci. U.S.A. National Academy of Sciences; 2012;109:2168–73.
64. Baudat F, Buard J, Grey C, Fledel-Alon A, Ober C, Przeworski M, et al. PRDM9 is a major determinant of meiotic recombination hotspots in humans and mice. Science. American Association for the Advancement of Science; 2010;327:836–40. 135
65. Chia N-Y, Chan Y-S, Feng B, Lu X, Orlov YL, Moreau D, et al. A genome- wide RNAi screen reveals determinants of human embryonic stem cell identity. Nature. Nature Publishing Group; 2010;468:316–20.
66. Delwel R, Funabiki T, Kreider BL, Morishita K, Ihle JN. Four of the seven zinc fingers of the Evi-1 myeloid-transforming gene are required for sequence-specific binding to GA(C/T)AAGA(T/C)AAGATAA. Molecular and Cellular Biology. American Society for Microbiology (ASM); 1993;13:4291–300.
67. Kuo TC, Calame KL. B lymphocyte-induced maturation protein (Blimp)-1, IFN regulatory factor (IRF)-1, and IRF-2 can bind to the same regulatory sites. J. Immunol. 2004;173:5556–63.
68. Chang JC, Meredith DM, Mayer PR, Borromeo MD, Lai HC, Ou Y-H, et al. Prdm13 mediates the balance of inhibitory and excitatory neurons in somatosensory circuits. Developmental Cell [Internet]. 2013;25:182–95. Available from: http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=PMC3644180
69. Ma Z, Swigut T, Valouev A, Rada-Iglesias A, Wysocka J. Sequence-specific regulator Prdm14 safeguards mouse ESCs from entering extraembryonic endoderm fates. Nat. Struct. Mol. Biol. Nature Publishing Group; 2011;18:120–7.
70. Seale P, Kajimura S, Yang W, Chin S, Rohas LM, Uldry M, et al. Transcriptional control of brown fat determination by PRDM16. Cell Metab. 2007;6:38–54.
71. Ross SE, McCord AE, Jung C, Atan D, Mok SI, Hemberg M, et al. Bhlhb5 and Prdm8 form a repressor complex involved in neuronal circuit assembly. Neuron [Internet]. 2012;73:292–303. Available from: http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=PMC3269007
72. Kajimura S, Seale P, Kubota K, Lunsford E, Frangioni JV, Gygi SP, et al. Initiation of myoblast to brown fat switch by a PRDM16-C/EBP-beta transcriptional complex. Nature. Nature Publishing Group; 2009;460:1154–8.
73. Katoh K, Omori Y, Onishi A, Sato S, Kondo M, Furukawa T. Blimp1 suppresses Chx10 expression in differentiating retinal photoreceptor precursors to ensure proper photoreceptor development. J. Neurosci. Society for Neuroscience; 2010;30:6515–26.
74. Brzezinski JA, Lamba DA, Reh TA. Blimp1 controls photoreceptor versus bipolar cell fate choice during retinal development. Development. 2010;137:619– 29. 136
75. Deng Q, Huang S. PRDM5 is silenced in human cancers and has growth suppressive activities. Oncogene. Nature Publishing Group; 2004;23:4903–10.
76. Meani N, Pezzimenti F, Deflorian G, Mione M, Alcalay M. The tumor suppressor PRDM5 regulates Wnt signaling at early stages of zebrafish development. Callaerts P, editor. PLoS ONE. Public Library of Science; 2009;4:e4273.
77. Nishikawa N, Toyota M, Suzuki H, Honma T, Fujikane T, Ohmura T, et al. Gene amplification and overexpression of PRDM14 in breast cancers. Cancer Res. American Association for Cancer Research; 2007;67:9649–57.
78. Dettman EJ, Justice MJ. The zinc finger SET domain gene Prdm14 is overexpressed in lymphoblastic lymphomas with retroviral insertions at Evi32. Hoheisel J, editor. PLoS ONE. 2008;3:e3823.
79. Dettman EJ, Simko SJ, Ayanga B, Carofino BL, Margolin JF, Morse HC, et al. Prdm14 initiates lymphoblastic leukemia after expanding a population of cells resembling common lymphoid progenitors. Oncogene. Nature Publishing Group; 2011;30:2859–73.
80. Boyer LA, Lee TI, Cole MF, Johnstone SE, Levine SS, Zucker JP, et al. Core transcriptional regulatory circuitry in human embryonic stem cells. Cell. 2005;122:947–56.
81. Seale P, Conroe HM, Estall J, Kajimura S, Frontini A, Ishibashi J, et al. Prdm16 determines the thermogenic program of subcutaneous white adipose tissue in mice. J. Clin. Invest. American Society for Clinical Investigation; 2011;121:96–105.
82. Seale P, Kajimura S, Spiegelman BM. Transcriptional control of brown adipocyte development and physiological function--of mice and men. Genes Dev. 2009;23:788–97.
83. Hanotel J, Bessodes N, Thélie A, Hedderich M, Parain K, Van Driessche B, et al. The Prdm13 histone methyltransferase encoding gene is a Ptf1a-Rbpj downstream target that suppresses glutamatergic and promotes GABAergic neuronal fate in the dorsal neural tube. Developmental Biology [Internet]. 2014;386:340–57. Available from: http://linkinghub.elsevier.com/retrieve/pii/S0012160613006817
84. Zannino DA, Sagerström CG. An emerging role for prdm family genes in dorsoventral patterning of the vertebrate nervous system. Neural Dev [Internet]. 2015;10:24. Available from: http://www.neuraldevelopment.com/content/10/1/24 137
85. Liu C, Ma W, Su W, Zhang J. Prdm14 acts upstream of islet2 transcription to regulate axon growth of primary motoneurons in zebrafish. Development [Internet]. 2012;139:4591–600. Available from: http://www.ncbi.nlm.nih.gov/pubmed/23136389
86. Olesnicky E, Hernandez-Lagunas L, Artinger KB. prdm1a Regulates sox10 and islet1 in the development of neural crest and Rohon-Beard sensory neurons. Genesis. John Wiley & Sons, Ltd; 2010;48:656–66.
87. Reid AG, Nacheva EP. A potential role for PRDM12 in the pathogenesis of chronic myeloid leukaemia with derivative chromosome 9 deletion. Leukemia. Nature Publishing Group; 2004;18:178–80.
88. Kolomietz E, Marrano P, Yee K, Thai B, Braude I, Kolomietz A, et al. Quantitative PCR identifies a minimal deleted region of 120 kb extending from the Philadelphia chromosome ABL translocation breakpoint in chronic myeloid leukemia with poor outcome. Leukemia. Nature Publishing Group; 2003;17:1313–23.
89. Yang C-M, Shinkai Y. Prdm12 is induced by retinoic acid and exhibits anti- proliferative properties through the cell cycle modulation of P19 embryonic carcinoma cells. Cell Struct. Funct. [Internet]. 2013;38:197–206. Available from: http://www.ncbi.nlm.nih.gov/pubmed/23856557
90. Kinameri E, Inoue T, Aruga J, Imayoshi I, Kageyama R, Shimogori T, et al. Prdm proto-oncogene transcription factor family expression and interaction with the Notch-Hes pathway in mouse neurogenesis. Hendricks M, editor. PLoS ONE [Internet]. 2008;3:e3859. Available from: http://dx.plos.org/10.1371/journal.pone.0003859
91. Sun X-J, Xu P-F, Zhou T, Hu M, Fu C-T, Zhang Y, et al. Genome-wide survey and developmental expression mapping of zebrafish SET domain- containing genes. Hoheisel J, editor. PLoS ONE [Internet]. 2008;3:e1499. Available from: http://dx.plos.org/10.1371/journal.pone.0001499
92. Zannino DA, Downes GB, Sagerström CG. prdm12b specifies the p1 progenitor domain and reveals a role for V1 interneurons in swim movements. Developmental Biology [Internet]. 2014;390:247–60. Available from: http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=PMC4030435
93. Thelie A, Thélie A, Desiderio S, Desiderio S, Hanotel J, Hanotel J, et al. Prdm12 specifies V1 interneurons through cross-repressive interactions with Dbx1 and Nkx6 genes in Xenopus. Development [Internet]. 2015;142:3416–28. Available from: http://dev.biologists.org/cgi/doi/10.1242/dev.121871 138
94. Senderek J, Reilly MM, Murphy SM, Kropatsch R, Michiue T, Windhager R, et al. Transcriptional regulator PRDM12 is essential for human pain perception. Nat Genet [Internet]. 2015;47:803–8. Available from: http://www.nature.com/doifinder/10.1038/ng.3308
95. Nagy V, Cole T, Van Campenhout C, Khoung TM, Leung C, Vermeiren S, et al. The evolutionarily conserved transcription factor PRDM12 controls sensory neuron development and pain perception. Cell Cycle [Internet]. 2015;14:1799– 808. Available from: http://www.tandfonline.com/doi/full/10.1080/15384101.2015.1036209
96. Lu DC, Niu T, Alaynick WA. Molecular and cellular development of spinal cord locomotor circuitry. Frontiers in Molecular Neuroscience. Frontiers; 2015;8:3003.
97. Lewis KE. How do genes regulate simple behaviours? Understanding how different neurons in the vertebrate spinal cord are genetically specified. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 2006;361:45–66.
98. Moran-Rivard L, Kagawa T, Saueressig H, Gross MK, Burrill J, Goulding M. Evx1 is a postmitotic determinant of v0 interneuron identity in the spinal cord. Neuron. 2001;29:385–99.
99. Pierani A, Moran-Rivard L, Sunshine MJ, Littman DR, Goulding M, Jessell TM. Control of interneuron fate in the developing spinal cord by the progenitor homeodomain protein Dbx1. Neuron. 2001;29:367–84.
100. Saueressig H, Burrill J, Goulding M. Engrailed-1 and netrin-1 regulate axon pathfinding by association interneurons that project to motor neurons. Development. 1999;126:4201–12.
101. Higashijima S-I, Higashijima SI, Masino MA, Mandel G, Fetcho JR. Engrailed-1 expression marks a primitive class of inhibitory spinal interneuron. J. Neurosci. [Internet]. 2004;24:5827–39. Available from: http://www.jneurosci.org/cgi/doi/10.1523/JNEUROSCI.5342-03.2004
102. Wenner P, O'Donovan MJ, Matise MP. Topographical and physiological characterization of interneurons that express engrailed-1 in the embryonic chick spinal cord. J. Neurophysiol. 2000;84:2651–7.
103. Alvarez FJ, Jonas PC, Sapir T, Hartley R, Berrocal MC, Geiman EJ, et al. Postnatal phenotype and localization of spinal cord V1 derived interneurons. J. Comp. Neurol. Wiley Subscription Services, Inc., A Wiley Company; 2005;493:177–92. 139
104. Li W-C, Higashijima S-I, Parry DM, Roberts A, Soffe SR. Primitive roles for inhibitory interneurons in developing frog spinal cord. J. Neurosci. 2004;24:5840– 8.
105. Benito-Gonzalez A, Alvarez FJ. Renshaw cells and Ia inhibitory interneurons are generated at different times from p1 progenitors and differentiate shortly after exiting the cell cycle. J. Neurosci. Society for Neuroscience; 2012;32:1156–70.
106. Gosgnach S, Lanuza GM, Butt SJB, Saueressig H, Zhang Y, Velasquez T, et al. V1 spinal neurons regulate the speed of vertebrate locomotor outputs. Nature [Internet]. 2006;440:215–9. Available from: http://www.ncbi.nlm.nih.gov/pubmed/16525473
107. Goulding M. Circuits controlling vertebrate locomotion: moving in a new direction. Nat Rev Neurosci [Internet]. 2009;10:507–18. Available from: http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=PMC2847453
108. Kimura Y, Satou C, Higashijima S-I. V2a and V2b neurons are generated by the final divisions of pair-producing progenitors in the zebrafish spinal cord. Development. 2008;135:3001–5.
109. Crone SA, Quinlan KA, Zagoraiou L, Droho S, Restrepo CE, Lundfald L, et al. Genetic ablation of V2a ipsilateral interneurons disrupts left-right locomotor coordination in mammalian spinal cord. Neuron. 2008;60:70–83.
110. Goulding M, Lanuza G, Sapir T, Narayan S. The formation of sensorimotor circuits. Curr. Opin. Neurobiol. 2002;12:508–15.
111. Kimmel CB, Patterson J, Kimmel RO. The development and behavioral characteristics of the startle response in the zebra fish. Dev Psychobiol. John Wiley & Sons, Ltd; 1974;7:47–60.
112. Granato M, van Eeden FJ, Schach U, Trowe T, Brand M, Furutani-Seiki M, et al. Genes controlling and mediating locomotion behavior of the zebrafish embryo and larva. Development. 1996;123:399–413.
113. Saint-Amant L, Drapeau P. Time course of the development of motor behaviors in the zebrafish embryo. J. Neurobiol. 1998;37:622–32.
114. Eaton RC, Lee RK, Foreman MB. The Mauthner cell and other identified neurons of the brainstem escape network of fish. Prog. Neurobiol. 2001;63:467– 85. 140
115. Matsukawa S, Miwata K, Asashima M, Michiue T. The requirement of histone modification by PRDM12 and Kdm4a for the development of pre-placodal ectoderm and neural crest in Xenopus. Developmental Biology [Internet]. 2015;399:164–76. Available from: http://linkinghub.elsevier.com/retrieve/pii/S0012160614006691
116. Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Dev. Dyn. 1995;203:253–310.
117. Montague TG, Cruz JM, Gagnon JA, Church GM, Valen E. CHOPCHOP: a CRISPR/Cas9 and TALEN web tool for genome editing. Nucleic Acids Res. 2014;42:W401–7.
118. Maurer JM, Sagerström CG. A parental requirement for dual-specificity phosphatase 6 in zebrafish. BMC Dev. Biol. BioMed Central; 2018;18:6.
119. Gagnon JA, Valen E, Thyme SB, Huang P, Akhmetova L, Ahkmetova L, et al. Efficient mutagenesis by Cas9 protein-mediated oligonucleotide insertion and large-scale assessment of single-guide RNAs. Riley B, editor. PLoS ONE. 2014;9:e98186.
120. Hauptmann G, Gerster T. Multicolor whole-mount in situ hybridization. Methods Mol. Biol. New Jersey: Humana Press; 2000;137:139–48.
121. Zannino DA, Appel B. Olig2+ precursors produce abducens motor neurons and oligodendrocytes in the zebrafish hindbrain. J. Neurosci. Society for Neuroscience; 2009;29:2322–33.
122. Ladam F, Stanney W, Donaldson IJ, Yildiz O, Bobola N, Sagerström CG. TALE factors use two distinct functional modes to control an essential zebrafish gene expression program. eLife. eLife Sciences Publications Limited; 2018;7:28.
123. Hatta K. Role of the floor plate in axonal patterning in the zebrafish CNS. Neuron. 1992;9:629–42.
124. Kok FO, Oster E, Mentzer L, Hsieh J-C, Henry CA, Sirotkin HI. The role of the SPT6 chromatin remodeling factor in zebrafish embryogenesis. Developmental Biology. 2007;307:214–26.
125. Ericson J, Thor S, Edlund T, Jessell TM, Yamada T. Early stages of motor neuron differentiation revealed by expression of homeobox gene Islet-1. Science. 1992;256:1555–60.
126. Tanabe Y, William C, Jessell TM. Specification of motor neuron identity by the MNR2 homeodomain protein. Cell. 1998;95:67–80. 141
127. Zannino DA, Sagerström CG, Appel B. olig2-Expressing hindbrain cells are required for migrating facial motor neurons. Dev. Dyn. Wiley-Liss, Inc; 2012;241:315–26.
128. McKeown KA, Moreno R, Hall VL, Ribera AB, Downes GB. Disruption of Eaat2b, a glutamate transporter, results in abnormal motor behaviors in developing zebrafish. Developmental Biology. 2012;362:162–71.
129. Lawson ND, Wolfe SA. Forward and reverse genetic approaches for the analysis of vertebrate development in the zebrafish. Developmental Cell. 2011;21:48–64.
130. Stainier DYR, Kontarakis Z, Rossi A. Making sense of anti-sense data. Developmental Cell. 2015;32:7–8.
131. Shalem O, Sanjana NE, Zhang F. High-throughput functional genomics using CRISPR-Cas9. Nat. Rev. Genet. Nature Research; 2015;16:299–311.
132. Doudna JA, Charpentier E. Genome editing. The new frontier of genome engineering with CRISPR-Cas9. Science. American Association for the Advancement of Science; 2014;346:1258096–6.
133. Jain RA, Bell H, Lim A, Chien C-B, Granato M. Mirror movement-like defects in startle behavior of zebrafish dcc mutants are caused by aberrant midline guidance of identified descending hindbrain neurons. J. Neurosci. 2014;34:2898– 909.
134. Liu KS, Fetcho JR. Laser ablations reveal functional relationships of segmental hindbrain neurons in zebrafish. Neuron. 1999;23:325–35.
135. Koh-Stenta X, Joy J, Poulsen A, Li R, Tan Y, Shim Y, et al. Characterization of the histone methyltransferase PRDM9 using biochemical, biophysical and chemical biology techniques. Biochem. J. [Internet]. 2014;461:323–34. Available from: http://biochemj.org/lookup/doi/10.1042/BJ20140374
136. Skaggs K, Martin DM, Novitch BG. Regulation of spinal interneuron development by the Olig-related protein Bhlhb5 and Notch signaling. Development. 2011;138:3199–211.
137. Győry I, Wu J, Fejér G, Seto E, Wright KL. PRDI-BF1 recruits the histone H3 methyltransferase G9a in transcriptional silencing. Nature Immunology. Nature Publishing Group; 2004;5:299–308.
138. Hutchinson SA, Cheesman SE, Hale LA, Boone JQ, Eisen JS. Nkx6 proteins specify one zebrafish primary motoneuron subtype by regulating late 142 islet1 expression. Development. The Company of Biologists Ltd; 2007;134:1671– 7.
139. Cheesman SE, Layden MJ, Ohlen Von T, Doe CQ, Eisen JS. Zebrafish and fly Nkx6 proteins have similar CNS expression patterns and regulate motoneuron formation. Development. The Company of Biologists Ltd; 2004;131:5221–32.
140. Stainier DY, Fouquet B, Chen JN, Warren KS, Weinstein BM, Meiler SE, et al. Mutations affecting the formation and function of the cardiovascular system in the zebrafish embryo. Development. 1996;123:285–92.
141. Weicksel SE, Gupta A, Zannino DA, Wolfe SA, Sagerström CG. Targeted germ line disruptions reveal general and species-specific roles for paralog group 1 hox genes in zebrafish. BMC Dev. Biol. BioMed Central; 2014;14:25.
142. Rossi A, Kontarakis Z, Gerri C, Nolte H, Hölper S, Krüger M, et al. Genetic compensation induced by deleterious mutations but not gene knockdowns. Nature. Nature Publishing Group; 2015;524:230–3.
143. Kok FO, Shin M, Ni C-W, Gupta A, Grosse AS, van Impel A, et al. Reverse genetic screening reveals poor correlation between morpholino-induced and mutant phenotypes in zebrafish. Developmental Cell. 2015;32:97–108.
144. Gribble SL, Nikolaus OB, Dorsky RI. Regulation and function of Dbx genes in the zebrafish spinal cord. Dev. Dyn. Wiley-Liss, Inc; 2007;236:3472–83.
145. Sorrells S, Toruno C, Stewart RA, Jette C. Analysis of Apoptosis in Zebrafish Embryos by Whole-mount Immunofluorescence to Detect Activated Caspase 3. Journal of Visualized Experiments. 2013.
146. Auer TO, Duroure K, De Cian A, Concordet J-P, Del Bene F. Highly efficient CRISPR/Cas9-mediated knock-in in zebrafish by homology-independent DNA repair. Genome Res. 2014;24:142–53.
147. Ma P, Zhao S, Zeng W, Yang Q, Li C, Lv X, et al. Xenopus Dbx2 is involved in primary neurogenesis and early neural plate patterning. Biochemical and Biophysical Research Communications [Internet]. 2011;412:170–4. Available from: http://linkinghub.elsevier.com/retrieve/pii/S0006291X11012940
148. Rai K, Jafri IF, Chidester S, James SR, Karpf AR, Cairns BR, et al. Dnmt3 and G9a cooperate for tissue-specific development in zebrafish. J. Biol. Chem. American Society for Biochemistry and Molecular Biology; 2010;285:4110–21. 143
149. Olsen JB, Wong L, Deimling S, Miles A, Guo H, Li Y, et al. G9a and ZNF644 Physically Associate to Suppress Progenitor Gene Expression during Neurogenesis. Stem Cell Reports. 2016;7:454–70.
150. Wang Y, Navin NE. Advances and applications of single-cell sequencing technologies. Mol. Cell. 2015;58:598–609.
151. Link V, Shevchenko A, Heisenberg C-P. Proteomics of early zebrafish embryos. BMC Dev. Biol. BioMed Central; 2006;6:1.
152. Baral R, Ngounou Wetie AG, Darie CC, Wallace KN. Mass spectrometry for proteomics-based investigation using the zebrafish vertebrate model system. Adv. Exp. Med. Biol. Cham: Springer International Publishing; 2014;806:331–40.
153. Moly PK, Hatta K. Early glycinergic axon contact with the Mauthner neuron during zebrafish development. Neuroscience Research. 2011;70:251–9.
154. Moly PK, Ikenaga T, Kamihagi C, Islam AFMT, Hatta K. Identification of initially appearing glycine-immunoreactive neurons in the embryonic zebrafish brain. Developmental Neurobiology. Wiley-Blackwell; 2014;74:616–32.
155. Cui WW, Low SE, Hirata H, Saint-Amant L, Geisler R, Hume RI, et al. The zebrafish shocked gene encodes a glycine transporter and is essential for the function of early neural circuits in the CNS. J. Neurosci. Society for Neuroscience; 2005;25:6610–20.
156. Grillner S, Jessell TM. Measured motion: searching for simplicity in spinal locomotor networks. Curr. Opin. Neurobiol. 2009;19:572–86.