Gene Regulatory Mechanisms of Drosophila Embryonic Motor Neuron Development 

by 

Katherine Helena Fisher 

A dissertation accepted and approved in partial fulfillment of the  

requirements for the degree of 

Doctor of Philosophy  

in Biology 

Dissertation Committee:  

Dr. Karen Guillemin, Chair  

Dr. Chris Doe, Advisor  

Dr. Judith Eisen, Core Member  

Dr. Tory Herman, Core Member  

Dr. Matt Smear, Institutional Representative 

University of Oregon 

Summer 2025 



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© 2025 Katherine H. Fisher  
 This work is openly licensed via CC BY 4.0. 



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DISSERTATION ABSTRACT 

Katherine Helena Fisher 

Doctor of Philosophy in Biology 

Title: Gene Regulatory Mechanisms of Drosophila embryonic motor neuron development 

Abstract 

Neural progenitors give rise to distinct populations of neurons throughout development. 

Drosophila larval neural progenitors, neuroblasts (NBs), express temporal gradients of 

transcription factors and RNA-binding proteins to establish neuronal diversity. The function of 

temporal transcription factors (TTFs) is well-studied in larval neural development. Several 

factors are expressed early in larval development, including Imp and Chinmo, while other factors 

are expressed later, including Syp, Mamo, and Broad, and an additional TF, Sequoia, is 

expressed throughout larval development. While the gene regulatory network of these factors has 

been thoroughly characterized in larvae, little is known about their expression or function in 

embryonic CNS development. Here we characterize the expression of early and late temporal 

factors in embryonic development. We find that Imp and Sequoia are expressed in neuroblasts, 

with a gradient of low-to-high expression in aging neuroblasts, which is maintained in post-

mitotic neurons. Interestingly, the embryonic Imp gradient is opposite the larval Imp gradient. 

The embryonic Sequoia gradient also contrasts larval expression where no gradient is detected. 

Another larval early factor, Chinmo, is expressed in all post-mitotic neurons, but not in a 

gradient. The late factors Mamo, EcR, Syp, and Broad are not expressed in embryos, with the 

exception of sparse Broad expression. Loss-of-function experiments showed that Imp is required 

for Chinmo expression. Intriguingly, loss of Chinmo -- but not Imp -- derepresses Syp. Finally, 

we tested whether Imp and Chinmo are required for motor neuron identity or morphology. We 

found that Imp, Chinmo, and Sequoia do not have a role in specifying motor neuron identity, but 

Imp and Chinmo have a later function in promoting motor axon targeting to the correct body 

wall muscle, and Chinmo is also required to prevent ectopic motor neuron dendrite projections. 

Together, these results show that the temporal factors are regulated differently in embryos and 

larvae, and that Imp and Chinmo are required for proper motor neuron axon or dendrite 

projections. 



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This dissertation includes previously published and co-authored materials. 



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CURRICULUM VITAE  

NAME OF AUTHOR:  Katherine Helena Fisher 

GRADUATE AND UNDERGRADUATE SCHOOLS ATTENDED: 

University of Oregon, Eugene 
Indiana University Bloomington 

DEGREES AWARDED: 

Doctor of Philosophy, Biology, 2025, University of Oregon 
Bachelor’s of Science, Biology, 2019, Indiana University Bloomington 

AREAS OF SPECIAL INTEREST: 

Developmental Neurobiology 
Embryo Patterning 

PROFESSIONAL EXPERIENCE: 

Graduate Researcher, University of Oregon, 2023-2025 
Laboratory of Dr. Chris Q. Doe 

Graduate Researcher, University of Oregon, 2019-2023 
Laboratory of Dr. Daniel Grimes 

Graduate Teaching Assistant, University of Oregon, 2019-2020 

Undergraduate Researcher, Indiana University Bloomington, 2015-2019 
Laboratory of Dr. W. Daniel Tracey 

Summer Undergraduate Research Scholar, HHMI Janelia Research Campus, 2018 
Laboratory of Dr. Tzumin Lee 

NSF Summer Undergraduate Research Fellow, Marine Biological Laboratory, 2017 
Laboratory of Dr. Marko Horb 

Undergraduate Teaching Assistant, Indiana University Bloomington, 2017-2019 



 

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GRANTS, AWARDS, AND HONORS: 
 

National Institutes of Health National Research Service Award F31, Genetic Dissection 
of fluid flow signaling in Left-Right patterning of zebrafish (1F31HD108945), University 
of Oregon, 2022-2025 

 
 American Heart Association Predoctoral Fellowship, Genetic Dissection of fluid flow 

signaling in Left-Right patterning of zebrafish (#909113, declined), 2022-2024, 
University of Oregon 

 
 National Science Foundation Graduate Research Fellowship, Understanding 

differentiation: Investigation of CG11360 in neuronal development, Field of study: 
Life Sciences—Developmental Biology (#1842486), University of Oregon, 2019-2022 

 
 Adamson Family Award, University of Oregon, 2020 
 
 Institute of Neuroscience Scholar Award, University of Oregon, 2019-2021 
  
 Outstanding Honors Thesis Award, Indiana University Bloomington, 2019 
 
PUBLICATIONS: 
 

K.H. Fisher, S-L. Lai, C.Q. Doe. Imp and Chinmo are required for embryonic motor 
neuron axon and dendrite targeting. Biology Open, accepted. 2025. 
 
E.A. Bearce, B.T.B. Ricamona, K.H. Fisher, J.R. O’Hara-Smith, D.T. Grimes. 
Visualization and quantitation of spine deformity in zebrafish models of scoliosis by 
micro-computed tomography. STAR Protocols, Dec 7, 2023. Doi: 
doi.org/10.1016/j.xpro.2023.102739. 

 
S.E. Mauthner, K.H. Fisher, W.D. Tracey. The Drosophila gene smoke alarm regulates 
nociceptor-epidermis interactions and thermal nociception behavior. bioRxiv 2021 May. 
2021.05.12.44364; doi: doi.org/10.1101/2021.05.12.443649 

 
 
 



 

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ACKNOWLEDGMENTS 
 
I would like to first express my gratitude and appreciation to my advisor Dr. Chris Doe. 

Chris’s advisership has helped shape me into a capable and confident scientist and moreover has 

helped me grow into a confident and strong person in general. Chris was not only a scientific 

advisor but also as an advisor through an exceptionally challenging time in my life. Without 

Chris, I would not be writing this dissertation or graduating with my PhD. I will forever be 

grateful for the support he has given to me and thankful for caring about me as a scientist and 

also as human being.  

I thank the members of the Doe lab (Nathan Anderson, Elena Barth, Ben Brissette, Kasey 

Drake, Derek Epiney, Josmarie Graciani, Janet Hanawalt, Keiko Hirono, Sen-Lin Lai, Kristen 

Lee, Laurina Manning, Jordan Munroe, Gonzalo Morales Chaya, Peter Newstein, Heather 

Pollington, Megan Radler, Matalie Rico Carvajal, Austin Seroka, Rishi Sastry, Alanna Sowles 

and Chundi Xu) for welcoming me into their group. The Doe lab group is the most positive and 

nurturing group of people that I have worked with, and every member is willing to help with 

experiments, writing, and scientific discussion. Many days the Doe lab members uplifted and 

encouraged me to keep pursing my research and goals.  

I would like to thank the members of my committee, Dr. Karen Guillemin, Dr. Judith 

Eisen, Dr. Tory Herman, and Dr. Matt Smear. I thank them for their support and guidance through 

my transition into the Doe lab.  

Outside of the UO, I would like to thank my long-term mentors Stephanie Mauthner and 

Rosa Miyares. Stephanie and Rosa encouraged me to continue to pursue research and their 

influence over the years has helped me grow into a curious and rigorous scientist. I will forever be 

grateful for their time and dedication to my scientific pursuits. I would not be where I am today 

without them.  

For work in chapter I, I thank Sen-Lin Lai for collaborating on axon targeting and MCFO 

experiments, as well as help with troubleshooting and discussion. I thank Megan Radler and 

Josmarie Graciani for assistance with brain dissections. I thank Kristen Lee for helpful discussion 

on quantification methods.  



 

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For work in chapter III, I thank Martin Blum for Xenopus data. I also thank my undergrad 

trainee, Maisey Schering, for help with the generation of guides, injections, and scoring heart 

defects.  

I would like to thank the fly and fish communities for being an excellent source of 

resources for this work. Specifically, I thank Flybase for their immense resources on the genes 

studied in this project. I would also like to thank the Zebrafish Information Network (ZFIN) for 

their resources, specifically annotated expression data that made the L-R patterning screen 

possible. I extend a special thank you to the Bloomington Drosophila Resource Center (BDSC), 

located at my undergraduate institution, for providing fly stocks for my project and to the greater 

fly community. Antibodies from the Developmental Studies Hybridoma Bank (DHSB) were used 

in this study. I also thank Claude Desplan for providing antibody reagents.   

This investigation was supported by HHMI (awarded to Dr. Chris Doe), NIH 

5F31HD108945-02 (awarded to Katherine H. Fisher), NSF #1842486 (awarded to Katherine H. 

Fisher).  



 

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DEDICATION 

 

I dedicate this dissertation to my family. To my partner John, who I met in grad school, and who 

has since been my strongest source of support and encouragement. We have been through so much 

in this journey and you have always been by my side encouraging me and providing me endless 

love and support, for taking me to Ducks games, on trips and adventures big and small. I could not 

have made it through this without you. To my mom and dad, who have always shown up for me, 

time and time again, and are always available for light-hearted phone calls and laughs. To my 

sisters and their babies, for providing lightheartedness and fun when I come to visit and for visiting 

me in Oregon as well. To my dog Buster, who I have lost, you were my first baby and it was such 

an honor to take care of you and love you. To my reptiles, who inspire my love of science and 

nature.  

 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 



 

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TABLE OF CONTENTS 

Chapter Page 

I. IMP AND CHINMO ARE REQUIRED FOR EMBRYONIC MOTOR NEURON 
AXON/DENDRITE TARGETING ............................................................................. 13  

Introduction ............................................................................................................ 13 

 Results .................................................................................................................... 15 

Imp is expressed in a low-high temporal gradient in embryonic neurons ...........15 

The Imp temporal gradient is not established by the embryonic TTF cascade ...17 

Cross-regulation of Chinmo, Imp, and Syp in the embryonic CNS ....................17 

Imp does not specify motor neuron molecular identity .......................................19 

Chinmo is required for motor neuron axon and dendrite targeting .....................21 

Imp is required for motor neuron axon targeting .................................................21 

 Discussion .............................................................................................................. 22 

 Methods.................................................................................................................. 25 

II. CONTRIBUTION OF SEQUOIA TO EMBRYONIC MOTOR NEURON  
PATTERNING AND MORPHOLOGY...................................................................... 28 
 Introduction ............................................................................................................ 28 

 Results .................................................................................................................... 29 

 Sequoia is expressed in a low-to-high gradient in the embryonic CNS .......... 29 

 The Sequoia gradient does not interact with the TTF cascade ........................ 31 

 Sequoia is not sufficient to induce motor neuron axon phenotypes ................ 31 

 Discussion .............................................................................................................. 31 

 Methods.................................................................................................................. 32 

 Bridge ..................................................................................................................... 36 

III. GENETIC DISSECTION OF FLUID FLOW SIGNALING IN LEFT-RIGHT 
PATTERNING OF ZEBRAFISH ............................................................................... 37 
 Introduction ............................................................................................................ 37 

 Results .................................................................................................................... 38 

 Discovery of novel regulators of L-R patterning ............................................. 40 

 Fibrocystin regulates L-R patterning in somatic mutants in zebrafish and  

Xenopus ............................................................................................................ 41 

 Germline mutants do not phenocopy somatic mosaic mutants ........................ 42 



 

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 Discussion .............................................................................................................. 43 

 Methods.................................................................................................................. 44 

IV. DISCUSSION ........................................................................................................ 46 

APPENDICES ............................................................................................................. 47 

 SUPPLEMENT TO CHAPTER I .......................................................................... 47 

REFERENCES CITED ................................................................................................ 50



 

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LIST OF FIGURES 
 
Figure Page 
 
CHAPTER I 

1 Schematic of known and unknown roles for the larval temporal factors .............. 14 
2 Imp forms a low-to-high gradient in embryos. ...................................................... 16 
 
3 Cross regulation of Imp, Syp, and Chinmo in the embryonic VNC ...................... 18 

4 Imp and Chinmo are not required for motor neuron identity ................................. 19 

5 Chinmo is required for motor neuron axon and dendrite targeting ....................... 20 

6 Imp is required for motor neuron axon targeting ................................................... 22 

7 Schematic of embryonic and larval roles of neuronal temporal factors ................ 23 

CHAPTER II 

1 Sequoia is expressed in a low-to-high gradient ..................................................... 30 

2 The Sequoia gradient acts independent from the TTF cascade ............................. 31 

3 Sequoia overexpression is not sufficient to induce axonal phenotypes ................. 32 

CHAPTER III 

1 The L-R patterning system in zebrafish ................................................................. 37 

2 Reverse genetics screen to identify regulators of L-R patterning .......................... 39 

3 Fibrocystin is required for L-R patterning in zebrafish and Xenopus .................... 40 

4 pkhd1 expression in the Xenopus Left-Right organizer ......................................... 41 

  
  
  



 

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CHAPTER I 

 

IMP AND CHINMO ARE REQUIRED FOR EMBRYONIC MOTOR NEURON 

AXON/DENDRITE TARGETING 

 

Author Contributions 

Conceptualization: C.Q.D., K.H.F., S-L.L.; Methodology: K.H.F. S-L.L.; Formal analysis: 

K.H.F., S-L.L.; Investigation: K.H.F., S-L.L.; Writing – original draft: K.H.F.,C.Q.D; 

Visualization: K.H.F., S-L.L.; Supervision: C.Q.D.; Project administration: C.Q.D.; Funding 

acquisition: C.Q.D, K.H.F. 

Authors: Katherine H. Fisher, Sen-Lin Lai, and Chris Q Doe 

 

Introduction 

 The generation of distinct populations of neurons is an essential part of neuronal development. 

Neurons with diverse function, connectivity, and morphology are important for sensation of and 

generation of complex behaviors across the animal kingdom. Neural progenitors give rise to 

distinct populations of neurons throughout development. In both Drosophila and mammals, two 

distinct mechanisms are used to generate neuronal diversity. First, spatial patterning of the 

neuroectoderm generates molecularly distinct progenitor pools (mammals) or distinct individual 

progenitors (Drosophila neuroblasts, NBs)(Crews 2019; Erclik et al. 2017; Guillemot 2007; A. 

Sagner and Briscoe 2019). Second, each progenitor undergoes gene expression changes over 

time, a process called temporal patterning. Temporal patterning occurs in the mammalian spinal 

cord, cerebral cortex, and retina (Mattar and Cayouette 2015; Andreas Sagner et al. 2021; A. 

Sagner and Briscoe 2019), and in the Drosophila embryonic ventral nerve cord (VNC; analogous 

to the vertebrate spinal cord), larval central brain, and optic lobe (Doe 2017; El-Danaf, Rajesh, 

and Desplan 2023).  

 In Drosophila, temporal patterning occurs via two distinct mechanisms. In the embryonic 

VNC and optic lobe, a cascade of temporal transcription factors (TTFs) specifies neuronal 

temporal identity. In the VNC, the cascade is: Hunchback (Hb), Krüppel, Pdm1/2, Castor, and 

Grainy head (Pollington and Doe 2025). In addition, a TTF cascade occurs within the progeny of 

central brain Type II NBs, called intermediate neural progenitors (INPs), which sequentially 



 

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express the transcription factors Dichaete, Grainy head, and Eyeless over several rounds of cell 

division (Bayraktar and Doe 2013; Homem et al. 2013; Tang et al. 2022). Finally, distinct TTFs 

are used in the optic lobe (El-Danaf, Rajesh, and Desplan 2023). In all three regions of the CNS 

the concept is the same: each TTF in the cascade specifies one or a few specific neuronal and 

glial cell types.  

 A second mechanism of temporal patterning occurs in larval central brain neuroblasts, 

where opposing gradients of two RNA-binding proteins, IGF-II mRNA binding protein (Imp) 

and Syncrip (Syp), specify different neuronal identities based on the level of each protein (Figure 

1)(Guan et al. 2022; Z. Liu et al. 2015). In the mushroom body, Imp and Syp proteins are 

expressed in opposing gradients that cross-repress each other (Z. Liu et al. 2015). Imp is 

expressed early in a high-to-low gradient while Syp is expressed late in a low-to-high gradient. 

Knockout of Imp or Syp results in dramatic loss of early- or late-born mushroom body cell types 

respectively (Z. Liu et al. 2015). Regulation of translation of the transcription factor Chinmo 

adds an additional layer of neuronal diversity. Imp positively regulates Chinmo expression, while 

Syp represses Chinmo expression through binding of the 5’UTR (Z. Liu et al. 2015; Zhu et al. 

2006). Furthermore, low levels of Chinmo activate expression of Mamo, creating an additional 

layer of temporal diversity by generating a intermediate mushroom body cell type (L.-Y. Liu et 

al. 2019). Similarly,  in Type II central brain neuroblasts (Bello et al. 2008; Boone and Doe 

2008; Bowman et al. 2008), Imp and Syp are expressed in opposing gradients and Imp is 

required for early Type II neuron fates, while Syp is required for late-born fates (Ren et al. 2017; 

Syed, Mark, and Doe 2017). Chinmo is expressed early alongside Imp and both are repressed by 

Figure 1. Schematic of known and unknown roles for the larval temporal factors. On the right shows known 
roles of the indicated larval temporal factors over time; none of these factors has been investigated for a role in 
the embryonic ventral nerve cord (left). ALH, hours after larval hatching; AEL, hours after egg lay; St, 
embryonic stage. Timeline not to scale.  
 



 

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Syp midway through larval development (Ren et al. 2017). Two transcription factors E93 and 

Broad are expressed in the later half of larval development and function to specify late neuronal 

fates (Ren et al. 2017; Syed, Mark, and Doe 2017).  

 While the gene regulatory network of these factors has been thoroughly characterized in 

larvae, little is known about the role of these larval temporal factors -- Imp, Syp, Chinmo, 

Mamo, Broad, E93 -- in embryonic CNS development. What little is known suggests different 

mechanisms are used in embryonc versus larval neuroblasts: Castor is a late TTF in the 

embyonic CNS (Doe 2017), whereas it is an early TTF in larval neuroblasts (Dillon and Doe 

2024). Major unknown questions are: Are these larval factors expressed in the embryonic CNS? 

Do they form opposing Imp/Syp gradients? What is the relationship between the embryonic TTF 

cascade and Imp/Syp expression? Do they specify neuronal identity, or other aspects of CNS 

development? Here we address these questions, finding both similarities and differnces in the 

larval and embryonic Imp/Syp/Chinmo expression and function. Notably, we found that both 

Imp and Chinmo are required for proper embryonic motor neuron axon targeting to their proper 

muscle targets, and dendrites mistargeting within the CNS neuropil. 

 

Results  

 

Imp is expressed in a low-high temporal gradient in embryonic neurons  

In larval development, Imp is expressed in a high-to-low gradient in neuroblasts, and expression 

levels are inherited by the daughter cells (Islam and Erclik 2022). However, Imp expression has 

only been characterized at low resolution and via a GFP knock-in tagged Imp:GFP protein in the 

embryonic CNS (Adolph et al. 2009). Many open questions remain. Will the larval "late" factors 

lack embryonic expression? Will the "early" larval Imp gradient continue from embryo to larvae? 

In general, we want to determine the expression patterns of "late" larval factors in the embryo 

and test their function. We determined that Imp is expressed globally in the cytoplasm of 

embryonic neuroblasts (Figure 2A). Imp expression levels are consistent in NBs until stage 12, at 

which time they show an increase in Imp expression, as quantified in Figure 2B. In post-mitotic 

neurons, Imp and Chinmo are expressed pan-neuronally (Figure 2C). Imp expression is present 

throughout embryognesis, while Chinmo expression is detectable in neurons beginning at stage 

12. Quantification of early born, mid born, and late born Imp and Chinmo expression showed 



 

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that Imp is expressed in a low-to-high gradient, being low in deep, early-born neurons and high 

in superficial, late-born neurons (Figure 2C-D). Importantly, this is the opposite of the larval 

Figure 2. Imp forms a low-to-high gradient in embryos. (A-B) Imp forms a low-to-high gradient in aging 
embryonic neuroblasts. (A) Imp expression in Dpn+ neuroblasts at the indicated embryonic stages (left). 
Ventral view.  Scale bar = 5μΜ. (B) Quantification. n > 40 for each stage. (C-D) Imp forms a low-to-high 
gradient in aging embryonic neurons. (C) Imp and Chinmo expression in a cross-sectional (posterior) view, 
where older neurons are located in a deep layer and younger neurons are located in a more superficial layer. 
Chinmo is expressed but not in a gradient. Scale bar = 5μΜ. (D) Quantification. n > 40 for each stage. (E-F) 
Imp and Chinmo do not form gradients in the young-old U1-U5 motor neurons, identified by expression of 
the Eve transcription factor. (E) Imp, Chinmo, and Eve expression in stage 16 embryos. Ventral view. Scale 
bar = 5μΜ. (F) Quantification. n > 20 for each neuron. (G-H) Imp and Chinmo do not form gradients in the 
young-old EL1-EL10 interneurons, identified by lateral expression of the Eve transcription factor. (G) Imp, 
Chinmo, and Eve expression in stage 16 embryos. Ventral view. Scale bar = 5μΜ. (H) Quantification. n > 5 
for each neuron.  
 



 

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high-to-low gradient. Chinmo expression is relatively uniform across early to late born neurons 

with variation in expression levels in individual neurons (Figure 2C-D). Broad is expressed in a 

subset of neurons and has higher expression in thoracic segments; other late factors E93, Mamo, 

and Syp were not present in embryonic NBs or neurons (Supplemental Figure 1). 

To determine if the Imp and Chinmo have similar expression patterns in identified 

neurons with a known birth-order, we assayed the five U1-U5 from NB7-1; these represent 

early-born identified neurons (T. Isshiki et al. 2001; Seroka et al. 2020). We found that Imp and 

Chinmo were both expressed in U1-U5 MNs, but were not distributed in a gradient (Figure 2E-

F). We also assayed Imp and Chinmo in the ten Eve lateral neurons (EL1-EL10) from NB3-3 

(Tsuji, Hasegawa, and Isshiki 2008; Wreden et al. 2017); these include late-born identified 

neurons. Imp and Chinmo were both expressed in EL1-EL10 interneurons, but were not 

distributed in a gradient (Figure 2G-H). Overall, we conclude Imp is expressed in a low-high 

gradient whereas Chinmo has a more even distribution.  

 

The Imp temporal gradient is not established by the embryonic TTF cascade 

To determine if the Imp gradient is generated by the action of the known embryonic TTF cascade 

(Hb>Kr>Pdm>Cas), we used en-gal4 to misexpress UAS-hb -- which is known to stall the TTF 

cascade (T. Isshiki et al. 2001; Pollington and Doe 2025; Tran and Doe 2008) and assayed for 

loss of the Imp gradient. We found that Hb overexpression in the en-gal4 stripes had no effect on 

Imp expression in the embryonic CNS (Supplemental Figure 2). We conclude that the TTF 

cascade is not responsible for generating the observed Imp low to high gradient. 

 

Cross-regulation of Chinmo, Imp, and Syp in the embryonic CNS 

Chinmo and Imp show cross-regulation in the larval CNS (Figure 1, right). We wanted to know 

if these factors show the same or different modes of cross-regulation in the embryonic CNS. In 

wild type, Chinmo has relatively high expression; Imp has modest expression, and Syp is not 

detected in neurons, although there is expression in gut cells below the CNS and sporadically in 

a subset of glia (Figure 3A). In Imp mutant embryos, we observed loss of Chinmo expression but 

we saw no derepression of Syp (Figure 3B, quantified in G).  In chinmo1 mutant embryos, we 



 

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Figure 3. Cross regulation of Imp, Syp and Chinmo in the embryonic VNC. (A) Control stained for Chinmo, 
Imp, and Syp. (B) chinmo1 homozygous mutant stained for Chinmo, Imp, and Syp. (C) Chinmo overexpression 
(en-gal4 UAS-chinmo) stained for Chinmo, Imp, and Syp. (D) Imp7 homozygous mutant stained for Chinmo, 
Imp, and Syp. (E) Imp overexpression (en-gal4 UAS-Imp) stained for Chinmo, Imp, and Syp. (F) Syp 
overexpression (en-gal4 UAS-Syp) stained for Chinmo, Imp, and Syp. All panels show stage 16 embryos, ventral 
view, scale bar, 5μm. (G) Quantification. 
 



 

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observed loss of Imp expression and de-repression of Syp -- unlike in chinmo mutants (Figure 

3C, quantified in G). Note that we did not assay Syp mutant embryos because there is no 

expression of Syp in the wild type CNS (Figure 3A). In contrast, overexpression of Imp resulted 

in no change in Chinmo (Figure 3D, E, quantified in H), despite the overexpression of Imp 

flattening the Imp gradient (Supplemental Figure 3). Lastly, we assayed overexpression of 

Chinmo, and found no change in levels of Imp expression (Figure 3F, quantified in 3H). We 

conclude that (a) there are differences in Chinmo, Imp, and Syp cross-regulation in embryos 

compared to larval stages; and (b) importantly, loss of Imp and loss of Chinmo had different 

effects on Syp expression.  

 

Imp does not specify motor 

neuron molecular identity 

To begin our analysis of Imp 

and Chinmo function in 

embryos, we assayed their 

role in motor neuron 

molecular identity (this 

section) and motor neuron 

axon and dendrite targeting 

(next sections). We chose to 

analyze the well-

characterized U1-U5 motor 

neurons born from the first 

five divisions of NB7-1 

(Isshiki et al., 2001). In wild 

type, Eve is expressed in U1-

U5, Zfh2 is expressed in U2-

U5, Runt is expressed in U4 

(high level) and U5 (low 

level), and Cut is expressed 

in U5 (Figure 4A). We 

Figure 4. Imp and Chinmo are not required for motor neuron identity 
(A) Control; molecular markers show wild type U1-U5 motor neuron 
identity. Eve, U1-U5; Zfh2, U2 (low), U3-U5; Runt, U4 (high)-U5 (low); 
Cut (U5). Scale bar = 5μΜ (B) Imp7 homozygous mutant; molecular 
markers show wild type U1-U5 motor neuron identity. Eve, U1-U5; Zfh2, 
U2-U5; Runt, U4 (high)-U5 (low); Cut (U5). Scale bar = 5μΜ 
 



 

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observe the same molecular identity in Imp and chinmo null mutants (Figure 4B). We conclude 

that loss of Imp has no effect on the molecular identity of the U1-U5 motor neurons. 

 

Figure 5. Chinmo is required for motor neuron axon and dendrite targeting 
(A-C) Multicolor flip out to obtain single neuron labeling of the U1 or U2 motor neuron dendrites. Posterior 
view; midline, dashed line. (A) Control U1 or U2 motor neuron assayed in newly hatched larvae. Scale bar, 5 
μm. Note lack of midline crossing (white bracket). (B) chinmo mutant showing ectopic denrite arbors (yellow 
bracket) assayed in newly hatched larvae. Scale bar, 5 μm. Note the abnormal midline crossing (yellow 
bracket). (C) Quantitation. Genetics: NB7-1-KZ-Gal4, R57C10-Flp, UAS-MCFO, UAS-chinmo-RNAi. (D-F) 
Multicolor flip out to obtain single neuron labeling of the U3-U5 motor neuron dendrites. Posterior view; 
midline, dashed line. (D) Control U3-U5 motor neuron assayed in newly hatched larvae. Scale bar, 5 μm. Note 
lack of midline crossing (white arrowhead). (E) chinmo mutant showing ectopic denrite arbors contacting or 
crossing the midline (yellow arrowhead) assayed in newly hatched larvae. Scale bar, 5 μm. (F) Quantitation. 
Genetics: NB7-1-KZ-Gal4, R57C10-Flp, UAS-MCFO, UAS-chinmo-RNAi assayed in newly hatched larvae. 
(G-I) U1-U5 motor neurons were labeled with GFP; lateral view, dorsal, up. (G) Controls have extension of 
motor neurons to the dorsal muscle field. Scale bar, 20 μm. (H) After chinmo RNAi in motor neurons, their 
axons fail to project to the dorsal muscle field. Scale bar, 20 μm. z(I) Quantitation. Genetics: CQ-gal4, UAS-
myrGFP, UAS-chinmo-RNAi, late stage 17. 
 



 

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Chinmo is required for motor neuron axon and dendrite targeting  

Chinmo is not required for motor neuron molecular identity, but it may have a role in later events 

such as axon/dendrite morphogenesis, targeting, or connectivity. We assayed for U1-U5 axon 

targeting to the dorsal muscle field (U1/U2) or the lateral muscle field (U3-U5). In controls, we 

observed innervation of both dorsal and lateral muscle fields (Figure 5A; quantified in 5C). In 

contrast, chinmo RNAi expressed in U1-U5 neurons resulted in frequent failure to innervate the 

dorsal muscles (Figure 5B; quantified in 5C). We conclude that Chinmo is required for proper 

motor neuron-muscle connectivity.  

 Next, we used MCFO to specifically label individual U1-U5 motor neurons, and 

confirmed that control U1/U2 motor neurons were bipolar and had dentritic arbors distant from 

the midline in newly hatched larvae (Figure 5D; quantified in 5F). In contrast, chinmo RNAi 

expressed in U1-U5 neurons resulted in U1/U2 showing ectopic dendrite projections contacting 

or crossing the midline in newly hatched larvae (Figure 5D; quantified in 5F). Similar abnormal 

midline contacting/crossing was observed in the mono-polar U3-U5 motor neurons (Figure 

5G,H; quantified in 5I). We conclude that Chinmo is required to prevent ectopic motor neuron 

dendriting targeting. 

 

Imp is required for motor neuron axon targeting 

Imp is also not required for motor neuron molecular identity, but it may have a role in later 

events such as axon/dendrite morphogenesis, targeting, or connectivity. We assayed for U1-U5 

axon targeting to the dorsal muscle field (U1/U2) or the lateral muscle field (U3-U5). In controls, 

we observed innervation of both dorsal and lateral muscle fields (Figure 6A; quantified in 6C). In 

contrast, expression of Imp RNAi in U1-U5 neurons resulted in frequent failure to innervate the 

dorsal muscles (Figure 6B; quantified in 6C). We conclude that Imp is required for proper motor 

neuron-muscle connectivity.  



 

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Discussion   

We have found significant differences in the expression, regulation, and function between larval 

and embryonic stages (Figure 7). We observed the following differences: (1) Whereas larval NBs 

express opposing gradients of Imp and Syp (Z. Liu et al. 2015), in embryos only Imp is 

expressed while Syp is undetectable, and thus embryos have no role for Syp. (2) Whereas larval 

NBs show Imp activating Chinmo but not the opposite (Zhu et al. 2006), in embryos both Imp 

and Chinmo positively regulate each other. (3) Whereas larval neurons do not show Chinmo 

repressing Syp (Z. Liu et al. 2015), in embryos Chinmo clearly represses Syp, as chinmo mutants 

de-repress Syp in neurons. (4) Whereas larval NBs show pan-neuronal expression of the mid 

(Mamo) or late (E93, Broad) temporal factors (Ren et al. 2017; Syed, Mark, and Doe 2017), in 

embryos Mamo and E93 are not expressed in the CNS, and Broad is only detected in a small 

subset of neurons. 

 Knockdown of chinmo causes derepression of Syp broadly throughout the embryonic 

CNS, however knockdown of Imp has no effect of Syp levels, consistent with our data showing 

that Syp is not expressed in the embryonic CNS. Maternal expression of Imp in zygotic Imp7 

mutants may be sufficient to repress Syp, or maternal Imp is sufficient to activate sufficient 

levels of Chinmo to repress Syp throughout embryogenesis. The repression of Syp by Chinmo 

contrasts their relationship in larval development, where Syp represses chinmo, and Syp and Imp 

repress each other (Ren et al. 2017). 

Figure 6. Imp is required for motor neuron axon targeting. (A-C) Multicolor flip out to obtain single 
neuron labeling of the U1 or U2 motor neuron dendrites. Posterior view; midline, dashed line. (A) Control U1 
or U2 motor neuron assayed in newly hatched larvae. Scale bar, 5 μm. (B) Imp RNAi in U1-U5 neurons results 
in ectopic denrite arbors (yellow bracket) assayed in newly hatched larvae. Scale bar, 5 μm. (C) Quantitation. 
Genetics: NB7-1-KZ-Gal4, R57C10-Flp, UAS-MCFO, UAS-Imp-RNAi.  
 



 

23 
 

 

 Previous work has shown that Imp is required in larvae for Kenyon cell specification: 

knockdown of Imp or Syp alters the ratio of early-born vs late-born neuronal identity (Hamid et 

al. 2024; Z. Liu et al. 2015), in contrast, Imp and chinmo have no role in motor neuron 

specification, but rather they are both required in post-mitotic motor neurons for axon targeting 

to the correct body wall muscle. Similarly, Chinmo knock down in the NB7-1 lineage results in 

mis-targeting of motor neuron dendrites within the CNS neuropil. Unfortunately we were unable 

to test for a similar role of Imp in motor dendrites, as we were unable to generate MCFO-labeled 

neurons, perhaps due to Imp triggering NB7-1 apoptosis (Guan et al. 2022; Samuels et al. 2020). 

Similarly, at larval stages, loss of Imp does not alter expression of pMad, a marker of motor 

neurons (Boylan et al. 2008), suggesting that Imp is also dispensable in larvae for specification 

of motor neuron identity. Imp is an RNA-binding protein, and its RNA cargo have been defined 

in larval neuroblasts, where Imp plays a role in regulation of proliferation (Samuels et al. 2020; 

Yang et al. 2017); these RNAs are also good candidates for a role in embryonic dendrite and 

axon targeting. 

 Chinmo is required for generation of distinct cell types in larval mushroom body neurons 

(Z. Liu et al. 2015; Zhu et al. 2006). In contrast, we found that Chinmo is dispensable for 

neuronal identity, in U1-U5 motor neurons, but is required for motor neuron axon targeting and 

Figure 7. Schematic of embryonic and larval roles of neuronal temporal factors. On the right shows known 
roles of the indicated larval temporal factors over time; left expression and cross-regulation of the indicated 
temporal factors in the embryonic VNC. ALH, hours after larval hatching; AEL, hours after egg lay; St, 
embryonic stage. Timeline not to scale. Arrows, positive regulation; T bars, repressive regulation. Broad* 
indicates scattered neuronal expression; not pan-neuronal. 
 



 

24 
 

 

targeting of motor neuron dendrites. Interestingly, Syp is required in larval motor neurons where 

syp, along with msp300, is required for new synapse formation. Additionally, syp is required for 

synaptic plasticity in motor neurons (Titlow et al. 2020). Interestingly, we found that Chinmo is 

required to repress Syp expression in the embryonic CNS. Deprepression of Syp in chinmo1 

mutants may promote additional synapse formation or synaptic plasticity in motor neuron 

dendrites leading to ectopic neurite formation.  

There is a major difference in time scale between embryonic and larval neurogenesis: 

embryonic neurogenesis is completed in less than one day, whereas larval neurogenesis lasts 5 

days. The shorter time of embryonic neurogenesis may requre a more "hard-wired" mechanism 

such a TTF cascade that switches TTF expression approximately every hour (Pollington, Seroka, 

and Doe 2023), whereas longer larval neurogenesis may provide time to generate and respond to 

gradients of Imp and Syp RNA-binding proteins (Islam and Erclik 2022). We found that 

overexpression of Hb in NBs, which stalls the TTF cascade and prolongs Hb expression (T. 

Isshiki et al. 2001) does not alter the Imp gradient in post-mitotic neurons. Additionally, 

overexpression of Imp, creating levels similar to Imp expression in late-born neurons, does not 

alter Hb expression in NBs. These results suggest that Imp/Chinmo and the TTF cascade 

function in separate pathways to guide neuronal development; the TTF cascade is required for 

cell type identity and Imp/Chinmo are required for neurite targeting and morphology in 

embryonic development.   

In larval development, Imp regulates proliferation of neuroblasts through binding to myc 

mRNA (Samuels et al. 2020). The scarcity of ImpRNAi MCFO clones suggests that Imp may be 

required for NB survival and proliferation. We did not see evidence of NB loss in Imp7, however 

maternal Imp transcripts may be sufficient to prevent loss of NBs. Thus, our results show that 

Imp expression in NBs may be important for survival of NBs in the embryo, although further 

work needs to be done to confirm this role.  

Our finding that Imp and Chinmo both show failure of motor axons to target their correct 

body wall muscle target is similar to the Imp mutant phenotype described for larval motor 

neurons, where reduced Imp led to reduced motor neuron bouton number at the neuromuscular 

junction (Boylan et al. 2008). Previous work imaging movement of Imp:GFP in motor axons 

showed that Imp is trafficked bidirectionally (Boylan et al. 2008); this movement may be 

defective in Imp mutants and explain the reduced bouton formation in larval motor neurons. It 



 

25 
 

 

seems likely that Imp is also moving bidirectionally in embryonic motor neurons; this would be 

an interesting question to explore in the future.  

 

Methods 

Antibody staining 

For embryonic CNS imaging, embryos were transferred from apple caps into collection baskets 

and rinsed with dH2O. Embryos were dechorionated in 100% bleach (Clorox, Oakland, CA) for 

4min with gentle agitation. Dechorionated embryos were rinsed with dH2O for 30sec. Embryos 

were fixed 20mins in 2mL Eppendorf tubes containing equal volumes of heptane (Fisher 

Chemical, Eugene, OR, H3505K-4) and 4% PFA diluted in PEM [100mM PIPES pH6.95 

(Sigma, St. Louis, MO), 2mM EDTA pH8.0 (Sigma, St. Louis, MO) and 1mM MgSO4 (Sigma, 

St. Louis, MO). The lower fix layer was removed, and an equal volume of methanol was added 

to each tube. Tubes were then subject to vigorous agitation for 1 min in a step required for 

removing the vitelline membrane. Nearly all liquid was removed from the tubes, leaving the 

embryos. Embryos were rinsed in Methanol (Fisher Chemical, Eugene, OR, Lot# 206197, Cat. 

A412P-4) three times and stored at -20°C.  Embryos were washed three times for 5 minutes with 

rocking in 0.1% PBST (1xPBS/0.1% Triton-x 100). PBST was removed and embryos were 

blocked with 5% normal donkey serum (Jackson ImmunoResearch Laboratories, Inc., West 

Grove, PA) in PBST for 30 min at room temp with rocking. PBST was removed and antibody 

mixes in PBST were added and rocked overnight at 4°C. Primary antibody mixes were removed 

and embryos were washed for >15 minutes three times in PBST with rocking. PBST was 

removed and secondary antibody diluted in PBST was added. Embryos were rocked at room 

temperature for 2 hours or rocked overnight at 4°C. Embryos were washed for >15 minutes three 

times in PBST with rocking.  

 After washing off secondary, embryos were washed three times in PBS then mounted in 

lysine coverslips and dehydrated in an ethanol series (30%, 50%, 70%, 90%, 100%). Embryos 

were washed an additional time in 100% ethanol. Next, embryos were washed two times in 

xylenes (Sigma, St. Louis, MO), then mounted in DPX mounting media (Sigma, St. Louis, MO) 

and dried at room temperature for two days before imaging.  



 

26 
 

 

 For imaging axon targeting to muscles, embryos were transferred into 50% glycerol for 

20 minutes or until embryos have settled at the bottom of the tube. The 50% glycerol was 

removed, and 90% glycerol was added. Embryos were left at room temperature overnight to let 

them fully settle to the bottom of the tube before imaging.  

For MCFO analysis of dendrite targeting, embryos that were UAS-MCFO, UAS-ImpRNAi 

or chinmo-RNAi, CQ2-gal4 were collected over a 24-hour window then aged for 24 hours. 

Freshly hatched larval brains were dissected in HL3.1 (Feng, Ueda, and Wu 2004), fixed in 4% 

paraformaldehyde, and mounted on lysine coverslips. Brains were immunostained on coverslips 

as described above. Note that UAS-chinmoRNAi experiments generated ~3 hemisegments with 

labeled motor neurons, whereas UAS-ImpRNAi experiments generated a single labeled neuron in 

over 40 brains; why this genotype gave such a low number of labeled neurons despite identical 

treatment of chinmo RNAi and Imp RNAi is unknown. 

For MCFO analysis of axon targeting, embryos that were UAS-MCFO, UAS-ImpRNAi or 

chinmo-RNAi, NB7-1-gal4 were collected over a 24-hour window then fixed in 4% 

paraformaldehyde, and stage 17 embryos were immunostained on coverslips as described above.  

 

Primary and secondary antibodies 

Primary antibodies used were: chicken anti-GFP, 1:1000 (Aves Labs, Davis, CA); rabbit anti-

Imp 1:500 (MacDonald lab, UT Austin); rabbit anti-Syp 1:1000 Desplan Lab, NYU); rat anti-

Deadpan 1:20 (Abcam, Eugene, OR); mouse anti-Hunchback 1:200 (Abcam, Eugene, OR); 

mouse anti-Eve 1:100 (DSHB, Iowa City, Iowa); guinea pig anti-Chinmo 1:200 (Desplan Lab, 

NYU); rat anti-Zfh2 1:250 (Doe Lab); mouse anti-Broad 1:20 DSHB, Iowa City, Iowa; guinea 

pig anti-Mamo 1:200 (Desplan Lab, NYU); guinea pig anti-E93 1:500 (Doe lab); mouse anti-En, 

5mg/mL (DSHB, Iowa City, Iowa); mouse anti-Eve[2B8], 5mg/mL, (DSHB, Iowa City, Iowa); 

rabbit anti-Eve, 1:250 (Doe lab); rabbit anti-Hb, 1:200 (Tran and Doe 2008); guinea pig anti-

Runt, 1:1000 (Sullivan, Warren, and Doe 2019); rat anti-Tm1[MAC141], 1:500 (Abcam, 

Waltham, MA, USA); mouse anti-HA (Biolegend [901513]); chicken anti-V5 (Bethyl [A190-

218A]); rat anti-Flag (Novus [NBP1-06712]); rat anti-Ollas Novus [NBP1-06713]) 

 Secondary antibodies used were: DyLight 405, AlexaFluor 488, rhodamine RedTM-X, 

AlexaFluor 555, or Alexa Fluor 647-conjugated AffiniPureTM donkey anti-IgG (Jackson 



 

27 
 

 

ImmunoResearch, West Grove, PA, USA). The samples were mounted in 90% glycerol with 

Vectashield (Vector Laboratories, Burlingame, CA) or DPX (Sigma, St. Louis, MO).  

 

Confocal Microscopy 

Images were captured with a Zeiss LSM800, LSM 900 or LSM 900-Airyscan2 laser scanning 

confocal microscope with a z-resolution of 0.25 µm (Carl Zeiss AG, Oberkochen, Germany) 

equipped with an Axio Imager.Z2 microscope. A 40x/1.40 NA Oil Plan-Apochromat DIC m27 

objective lens and a 63x/1.40 Oil Plan-Apochromat DIC m27 objective lens and GaAsP 

photomultiplier tubes were used. Software program was Zen 2.3 (blue edition) (Carl Zeiss AG, 

Oberkochen, Germany). Images were processed using the open-source software FIJI or Imaris 

(Oxford Instruments plc, UK). Figures were assembled in Adobe Illustrator (Adobe, San Jose, 

CA). For each independent experiment, all samples were acquired using identical acquisition 

parameters.  

 

Motor neuron axon and dendrite imaging 

Genetics for Figure 5 and 6.  Dendrite: NB7-1-KZ, R57C10-Flp, UAS-MCFO with UAS-LacZ for 

controls or UAS-chinmo-RNAi or UAS-Imp-RNAi for experimental groups. Axon: CQ-gal4, UAS-

myrGFP, with UAS-LacZ for controls or UAS-chinmo-RNAi or UAS-Imp-RNAi for experimental 

groups. Embryo staging was done according to gut morphology, ensuring that both controls and 

experimentals were at the same age (late stage 17).  

 

Statistical analyses 

Statistics were computed using Prism (GraphPad, Boston, MA). All statistical tests used are 

listed in the figure legends. P-values are reported in the figures. n.s. = not significant, where 

p>0.05. Plots were generated using Prism (GraphPad). 

 

Figure production 

Images for figures were processed in FIJI. Figures were assembled in Adobe Illustrator 

(Adobe, San Jose, CA). Any changes in brightness or contrast were applied to the entire 

image. 



 

28 
 

 

CHAPTER II 

CONTRIBUTION OF SEQUOIA TO EMBRYONIC NEUROBLAST AND NEURON 

PATTERNING AND MORPHOLOGY 

 

Author Contributions 

Conceptualization: C.Q.D., S-L.L.; Methodology: K.H.F.; Formal analysis: K.H.F.; 

Investigation: K.H.F., S-L.L.; Writing – original draft: K.H.F.; Visualization: K.H.F.; 

Supervision: C.Q.D.; Project administration: C.Q.D.; Funding acquisition: C.Q.D, K.H.F. 

Authors: Katherine H. Fisher, Sen-Lin Lai, Chris Q. Doe 

 

Introduction 

The nervous system contains a multitude of specialized neuronal cell types that are critical for 

forming functional neuronal circuits. A single neuronal progenitor generates many differing cell 

types over time. In the drosophila embryo, sequential expression of transcription factors over 

time divides neurons into different temporal cohorts, with early born cells expression one set of 

transcription factors, and late born cells expressing a differing set of transcription factors 

(Takako Isshiki et al. 2001). Expression of this temporal transcription factor (TTF) cascade not 

only generates neuronal diversity but may also influence synaptic specificity and axonal 

innervation. Neurons have been shown to target their synapses in small domains with other 

neurons from the same temporal cohorts (Takako Isshiki et al. 2001; Mark et al. 2021). The 

mechanisms controlling synaptic specificity in the central nervous system (CNS) are not well 

understood. Does the TTF cascade solely control synaptic targeting domains or are other 

neuronally expressed factors involved? 

The pan-neuronally expressed gene, sequoia, is a candidate gene that may function in 

neuronal morphology and synaptic specificity. In the Drosophila retina, Sequoia is expressed in 

a gradient in photoreceptor cells, and this gradient is required for axons to project into the proper 

layer (Petrovic and Hummel 2008). Loss of sequoia results in multiple axons targeting to the 

same layer. Additionally, sequoia functions in Drosophila sensory neurons in the peripheral 

nervous system, called multidendritic (md) neurons, to regulate dendrite and axon 

morphogenesis. Loss of sequoia results in over-branching of dendrites in md neurons, but 

shortened axon projections (Brenman et al. 2001; Grueber, Jan, and Jan 2002).  



 

29 
 

 

 

We hypothesized that sequoia may function in the embryonic CNS to control neuronal 

morphology. Here we use the Drosophila embryo, specifically the NB 7-1 lineage, to determine 

how sequoia functions in synaptic patterning and tiling in motor neurons of the CNS. Key 

questions we address are: Is there a seq gradient in the embryonic CNS, and does this affect 

neuron morphology? Does seq interact with the TTF cascade? 

 

Results 

Sequoia is expressed in a low-to-high gradient in the embryonic CNS 

To determine if Sequoia is expressed in a gradient, we quantified pixel intensity of 

sequoia::GFP expression in neuroblasts from stage 9-12 of embryonic development. We 

validated that the sequoia::GFP line recapitulates Sequoia protein expression by staining with 

both GFP and a Sequoia antibody (data not shown). Sequoia expression is relatively uniform 

across neuroblasts, with a slight peak at stage 10 (Figure 1A-A’). Quantification of Sequoia in 

early-born Hunchback(Hb)+ cells and late-born Castor(Cas)+ cells showed that Sequoia 

expression in lower in early born cells, and higher in late born cells (Figure 1B-B’). Finally, 

quantifying Sequoia expression in the U motor neurons (UMNs) produced from the 7-1 lineage 

showed a low-to-high gradient of Seq over U1-U5 (Figure 1C-C’). These data confirm that 

Sequoia is expressed in a low-to-high gradient across neurons in the embryonic CNS both 

globally and in a specific lineage of sequentially born cells.  

 



 

30 
 

 

 

Figure 1. Sequoia is expressed in a low-to-high gradient. A) Sequoia expression in neuroblast at embryonic 
stage 9, 11, and 12. A’) Quantitation of Sequoia protein expression with a peak at stage 10. B) Sequoia expression 
in early-born Hb+ and late-born Cas+ cells across embryonic development from stage 12-16. B’) quantitation of 
Sequoia in Hb+ and late-born Cas+ cells. Sequoia expression is higher in late-born Cas+ cells than in early-born 
Hb+ cells at all quantified time points. C) Sequoia expression in UMNs from the 7-1 NB lineage. C’) Quantitation 
of Sequoia in U1-U5. Highest expression is seen in late-born U5 neurons.  



 

31 
 

 

 

The Sequoia gradient does not 

interact with the TTF cascade 

To determine if the Sequoia gradient 

interacts with the temporal 

transcription factor cascade, we 

overexpressed Sequoia in the UMNs 

and assessed TTF expression. 

Overexpression of Sequoia had no 

effect on the expression of Hb, Kr, or 

Cas (Figure 2). Normal expression of 

Hb was observed in U1-U2, Kr 

expression in U1-U3, and Cas 

expression in U5 at embryonic stage 

16.  

 

Sequoia is not required for motor neuron axon targeting 

Graded Sequoia expression is required in photoreceptor cells to properly pattern axons. We 

hypothesized that the Sequoia gradient in embryos may play a similar role. To test whether the 

Sequoia gradient affects neuronal morphology, we overexpressed sequoia in NB7-1 (7-1GAL4 

UAS-seq) to flatten the gradient in UMNs and assessed axon targeting. UMN axons target to 

specific muscles in the larval body wall and targeting is largely completed by the last stage of 

embryogenesis. Embryos showed no evidence of aberrant axon targeting (Figure 3). Further, we 

filleted third instar larvae to assess morphology of mature neuromuscular junctions and saw no 

obvious defects in NMJ morphology (Figure 3, scale bar 20µm).  

 

Discussion 

We observed that overexpression of sequoia in the neuroblast had no effect on the TTF cascade 

in NB7-1 progeny. This suggests that Sequoia expression levels have no effect on the expression 

Figure 2. The Sequoia gradient acts independent from the 
TTF cascade. A) In wild type embryos, U1-U2 express Hb and 
Kr, U3 expressed Kr, and U5 expressed Cas. B) Overexpression 
of Sequoia in UMNs does not change the TTF expression profile 
of these neurons. 



 

32 
 

 

of the TTF cascade. It 

would be interesting to 

alter the TTF cascade, 

through overexpression of 

Hb, to see if the Sequoia 

gradient is established by 

the sequential expression 

of the TTF cascade.  

As overexpression 

of Sequoia in NB7-1 did 

not alter the number of 

UMNs, we hypothesize 

that the Sequoia gradient 

is not dependent on the 

TTF cascade. If the TTF 

cascade is not required to 

establish the Sequoia gradient, then this may suggest that Sequoia is acting independent from the 

TTF cascade, and potentially as a regulator of synaptic or dendritic targeting rather than 

specifying cell identity. While no gross morphology differences in NMJs were observed with 

sequoia overexpression, further analysis of synapse number and morphology may reveal a more 

specific role for sequoia in motor neurons.  

Sequoia has been shown to be a negative regulator of md sensory neuron dendrites and a 

positive regulator of axon outgrowth in md neurons (Brenman et al. 2001; Grueber, Jan, and Jan 

2002). We did not see an effect of overexpression of Sequoia in motor neuron axons, but it is 

possible that the targeting of the dendrites to the neuropil may be affected. Further 

overexpression and loss-of-function experiments aimed at exploring dendritic tiling is needed to 

determine if Sequoia plays a role in dendrite growth outside of the PNS.  

 

Methods 

Fly stocks 

Figure 3. Sequoia overexpression is not sufficient to induce axonal 
phenotypes. A) Axon targeting of 7-1 UMNs labeled with FasII and 
targeting to dorsal muscle (Tm1) in late stage 17 embryos. WT axons 
extend to dorsal muscle 9. B) Overexpression of Seq results in axon 
targeting to muscle 9 similar to WT. scale bar = 20µm. C) WT NMJ labeled 
with HRP, C’) overlay with muscle cells (phalloidin). D) overexpression of 
seq does not affect morphology of the NMJ D’) overlay with muscle cells.  



 

33 
 

 

Stock Source or reference Identifier 

w; UAS-seq BDSC RRID:BDSC_9244 

w; UAS-LacZ BDSC RRID:BDSC_8529 

w; CQ2-Gal4 BDSC RRID:BDSC_7466 

NB7-1-KZ Doe lab N/A 

yw; seq::GFP BDSC RRID:BDSC_92639 

 

Antibody staining 

For embryonic CNS imaging, embryos were transferred from apple caps into collection baskets 

and rinsed with dH2O. Embryos were dechorionated in 100% bleach (Clorox, Oakland, CA) for 

4min with gentle agitation. Dechorionated embryos were rinsed with dH2O for 30sec. Embryos 

were fixed 20mins in 2mL Eppendorf tubes containing equal volumes of heptane (Fisher 

Chemical, Eugene, OR, H3505K-4) and 4% PFA diluted in PEM [100mM PIPES pH6.95 

(Sigma, St. Louis, MO), 2mM EDTA pH8.0 (Sigma, St. Louis, MO) and 1mM MgSO4 (Sigma, 

St. Louis, MO). The lower fix layer was removed, and an equal volume of methanol was added 

to each tube. Tubes were then subject to vigorous agitation for 1 min in a step required for 

removing the vitelline membrane. Nearly all liquid was removed from the tubes, leaving the 

embryos. Embryos were rinsed in Methanol (Fisher Chemical, Eugene, OR, Lot# 206197, Cat. 

A412P-4) three times and stored at -20°C.  Embryos were washed three times for 5 minutes with 

rocking in 0.1% PBST (1xPBS/0.1% Triton-x 100). PBST was removed and embryos were 

blocked with 5% normal donkey serum (Jackson ImmunoResearch Laboratories, Inc., West 

Grove, PA) in PBST for 30 min at room temp with rocking. PBST was removed and antibody 

mixes in PBST were added and rocked overnight at 4°C. Primary antibody mixes were removed 

and embryos were washed for >15 minutes three times in PBST with rocking. PBST was 

removed and secondary antibody diluted in PBST was added. Embryos were rocked at room 

temperature for 2 hours or rocked overnight at 4°C. Embryos were washed for >15 minutes three 

times in PBST with rocking.  

Embryos were transferred into 50% glycerol for 20 minutes or until embryos have settled 

at the bottom of the tube. The 50% glycerol was removed, and 90% glycerol was added. 



 

34 
 

 

Embryos were left at room temperature overnight to let them fully settle to the bottom of the tube 

before imaging.  

 

Larval Fillets 

Third instar wandering larvae were picked from crosses with a paintbrush and transferred into 

HL3.1(Feung et al., 2004). The cuticle was cut and pinned down to create a flat piece of cuticle 

and fixed in 4% paraformaldehyde. Fillets were washed in 0.2% PBST three times and then 

stained with primary and secondary antibodies. Samples were mounted in vectashield.  

 

Primary and secondary antibodies 

Primary antibodies used were: chicken anti-GFP, 1:1000 (Aves Labs, Davis, CA); rat anti-

Deadpan 1:20 (Abcam, Eugene, OR); mouse anti-Hunchback 1:200 (Abcam, Eugene, OR); 

rabbit anti-Castor 1:1000 (W. Odenwald Lab); mouse anti-Eve[2B8], 5mg/mL, (DSHB, Iowa 

City, Iowa); rabbit anti-Eve, 1:250 (Doe lab); rat anti-Tm1[MAC141],  

 Secondary antibodies used were: DyLight 405, AlexaFluor 488, rhodamine RedTM-X, 

AlexaFluor 555, or Alexa Fluor 647-conjugated AffiniPureTM donkey anti-IgG (Jackson 

ImmunoResearch, West Grove, PA, USA). The samples were mounted in 90% glycerol with 

Vectashield (Vector Laboratories, Burlingame, CA). 

 

Confocal Microscopy 

Images were captured with a Zeiss LSM800 or LSM 710 (Carl Zeiss AG, Oberkochen, 

Germany) equipped with an Axio Imager.Z2 microscope. A 40x/1.40 NA Oil Plan-Apochromat 

DIC m27 objective lens and a 63x/1.40 Oil Plan-Apochromat DIC m27 objective lens and 

GaAsP photomultiplier tubes were used. Software program was Zen 2.3 (blue edition) (Carl 

Zeiss AG, Oberkochen, Germany). Images were processed using the open-source software FIJI 

or Imaris (Oxford Instruments plc, UK). Figures were assembled in Adobe Illustrator (Adobe, 

San Jose, CA). For each independent experiment, all samples were acquired using identical 

acquisition parameters.   

 

Statistical analyses 



 

35 
 

 

Statistics were computed using Prism (GraphPad, Boston, MA). All statistical tests used are 

listed in the figure legends. *<0.0332, **<0.0021, ***<0.0002, otherwise P-values are reported 

in the figures. n.s. = not significant, where p>0.05. Plots were generated using Prism (GraphPad). 

 

Figure production 

Images for figures were processed in FIJI. Figures were assembled in Adobe Illustrator 

(Adobe, San Jose, CA). Any changes in brightness or contrast were applied to the entire 

image. 

 

  



 

36 
 

 

BRIDGE 

In chapters I and II, I established the gene regulatory network of the temporally expressed RNA-

binding proteins and transcription factors in drosophila motor neuron development.  

 

In the following chapter, I investigate another gene regulatory network controlling left-right 

patterning. This project, while conducted on a different signaling pathway, draws on the same 

principle of gene regulation and cell-cell signaling to achieve a target morphology.  

  



 

37 
 

 

CHAPTER III 

 

GENETIC DISSECTION OF FLUID FLOW SIGNALING IN LEFT-RIGHT 

PATTERNING OF ZEBRAFISH 

 

Author Contributions 

Conceptualization: D.T.G., K.H.F.; Methodology: K.H.F., M.B.; Formal analysis: K.H.F.; 

Investigation: Figure 2: K.H.F., M.S.; Figure 3 & 4: M.B.; Writing – original draft: K.H.F.; 

Visualization: K.H.F., M.B.; Supervision: D.T.G.; Project administration: D.T.G.; Funding 

acquisition: D.T.G., K.H.F. 

Author List: Katherine H. Fisher, Maisey Schering, Martin Blum, Daniel T. Grimes 

 

Introduction 

Cell-cell communication is crucial for patterning and morphogenesis. One mechanism of 

communication relies on extracellular fluid flows. Flows play critical roles in the kidney, 

vasculature, reproductive tracts, brain ventricles, and spinal canal (Freund et al. 2012). Aberrant 

flows cause developmental diseases including congenital heart disease (CHD) (Hoffman and 

Kaplan 2002), heterotaxy (Sutherland and Ware 2009), and hydrocephalus (Kothari 2014). 

Compared to chemical signaling pathways like Wnt or Hh, flow-induced pathways, including 

how flow signals are sent, sensed, and transduced, are poorly 

understood. Identifying new regulators of flow signaling and 

dissecting how they work will give insight into diseases 

caused by aberrant flow. 

The tractable left-right (L-R) patterning system in 

zebrafish (Grimes and Burdine 2017; Matsui and Bessho 

2012) is an ideal model for identifying components of the flow 

sensory pathway (Figure 1). In early embryonic development, 

asymmetric fluid flow in the zebrafish L-R organizer, called 

Kupffer’s vesicle (KV), breaks L-R symmetry (Kramer-

Zucker et al. 2005; Nonaka et al. 2002) (Figure 1A). KV is a 

transient spherical structure made of cells with motile cilia that 

Fig 1. The L-R patterning system 
in zebrafish. A) Symmetry is 
broken in Kupffer’s vesicle (KV). 
B) Asymmetric flow is generated 
by motile cilia in KV. C) Model of 
the flow sensory pathway, where 
Pkd1l1 and Pkd2 sense and 
transduce flow resulting in dand5 
repression. 



 

38 
 

 

generate fluid flow (Essner et al. 2005) (Figure 1B). Flow is stronger on the left side than the 

right. Cells in KV sense this asymmetric flow signal, which results in post-transcriptional 

repression of dand5 mRNA, the target of the flow signal, on the left side only (Hojo et al. 2007; 

Schweickert et al. 2010). It is hypothesized that the motile cilia in KV act as mechanosensory 

organelles to detect flow signals (McGrath et al. 2003; Shinohara and Hamada 2017; Yoshiba et 

al. 2012). In other left-right organizers, such as in mice and the African-clawed frog, specific 

cells on either side of the organizer, called sensory cells, contain non-motile cilia that are 

required for fluid flow detection (Schweickert et al. 2010). Additionally, the cytoskeleton also 

plays a role in symmetry breaking by creating subcellular chiral structures that influence 

laterality (McDowell et al. 2016). Asymmetric repression of dand5 mRNA by the flow signal 

leads to activation of the Nodal signaling pathway on the left but not the right side of KV. This 

asymmetric Nodal signal spreads to the left lateral plate mesoderm (LPM) and propagates 

anteriorly through the embryo (Long, Ahmad, and Rebagliati 2003; Shiratori and Hamada 2014).  

Symmetry breaking is dependent on the Polycystin protein, Pkd1l1 in mouse, medaka 

fish, zebrafish, and humans (Field et al. 2011; Grimes et al. 2016; Vetrini et al. 2016) (Figure 

1C). Additionally, Pkd2 is required for L-R patterning (Bisgrove et al. 2005; Pennekamp et al. 

2002). Pkd2 is a transient receptor potential ion channel that may mediate L-R patterning by flow 

induced Ca2+ spikes in flow sensory cells (Yuan et al. 2015). Pkd2 acts downstream of flow in 

the kidney, and forms a functional complex with the Pkd1l1 homolog, Pkd1 (Kamura et al. 

2011). Mutations in PKD2 and PKD1 cause autosomal dominant polycystic kidney disease 

(Harris and Torres 2009; Igarashi and Somlo 2007). While it is clear that these proteins are 

required for flow signaling in several contexts, it is not understood how they function.  

While we know that flow is critical for proper L-R patterning, very little is known about 

how the flow signal is sensed and transduced to ultimately result in asymmetric dand5 

repression. The zebrafish L-R patterning system, tractable to imaging and genetic manipulation, 

with a controllable flow input and quantifiable dand5 output, is ideal for discovering basic 

principles of flow signaling. 

 

Results 



 

39 
 

 

Sensation and transduction of the fluid flow signal is critical for establishing L-R asymmetry. 

However, few flow sensory pathway regulators are known. I performed a reverse genetic screen 

and discovered novel regulators of L-R patterning.  

 

Figure 2. Reverse genetics screen to identify regulators of L-R patterning. A) schematic of CRISPR-
CAS9 screen pipeline. Four single guides are generated in multiplex, then injected with Cas9 protein 
into one cell stage embryos, then embryos are raised for 24 hours at 28C. Heart tube positioning is then 
assessed. B) graph of percent abnormal heart jogging (middle or right) for each gene screened. C) a 
second set of guide RNAs was generated and injected for top hits to validate phenotypes. D) marveld1 
embryos displayed body shape defects indicated by the downward curve of the tail in addition to heart 
jogging defects. 



 

40 
 

 

Discovery of Novel Regulators of L-R Patterning. Using CRISPR-Cas9, I generated somatic 

mosaic (G0) mutants in zebrafish by targeting genes with a high concentration of four guide 

RNAs (gRNAs) (Wu et al. 2018) that target different regions of the same gene. With this 

technique, I generated loss-of-function-like phenotypes. To assess L-R patterning, I determined 

heart jogging, a reliable readout of L-R asymmetry, in embryos at 23 hours post fertilization 

(hpf) (Figure 2A). In wild-type embryos at 23 hpf, the heart is positioned to the left (Chen et al. 

1997), and in embryos with disrupted L-R asymmetry, heart positioning is disrupted (i.e. 

randomized between left and right, or sometimes positioned in the middle) (Baker, Holtzman, 

and Burdine 2008; 

Smith et al. 2008).   

I validated this 

method by targeting 

known regulators of L-

R patterning. Targeting 

dand5, pkd1l1, and 

cfap298, which are 

expressed in KV and 

have previously been 

implicated in L-R 

patterning (Bisgrove et 

al. 2005; Jaffe et al. 2016) resulted in 30-90% levels of abnormal heart laterality (Figure 2B). 

cfap298 also displayed a curved body axis, called curly-tail down, which phenocopied germline 

mutants (Jaffe et al. 2016). Additionally, we targeted a gene, kif6, that had no role in L-R 

patterning but has other visible body shape defects when knocked down (Buchan et al. 2014). 

These levels of heart defects, as well as those induced by Cas9 injection alone, served as a 

background for comparisons to novel genes. When assessing heart jogging, it is important to note 

that since the primary phenotype of L-R patterning mutants is randomization of laterality, 50% 

abnormalities could represent 100% penetrance of mutant phenotypes due to a randomization 

between left and right. 

To select a list of candidate genes, we searched through expression data available on the 

Zebrafish Information Network (ZFIN) database. Expression studies in zebrafish have annotated 

Fig 3. Fibrocystin is required for L-R patterning in zebrafish and Xenopus. 
B,D) double G0 pkhd1l1.1 and pkhd1l1.2 mutants display bilaterally symmetric 
L=R dand5 in KV at 10 somite stage. A,D,E) dand5 expression in the L-R 
organizer of stage 20 Xenopus is normally L<R, but morpholino knockdown of 
pkhd1 in Xenopus on the left, but not right side results in abnormal dand5 
expression (E). ***p<0.5, ns p>0.05, Pearson chai-square test (Bonferonni 
corrected). [Kontrolle = Control, links = left] 
 



 

41 
 

 

195 genes as expressed in KV as of 2021 (ZFIN). Of these 195 genes, 78 have previously 

identified roles in L-R patterning and/or motile cilia function, and 27 have known roles in motile 

cilia in contexts outside of L-R patterning, leaving 90 genes with no known role in L-R 

patterning or cilia motility/formation. We generated multiplexed gRNAs against each gene, 

injected guide sets with two replicates per gene, and assayed heart jogging. We observed a broad 

range of L-R patterning defects ranging from 15-50% heart laterality defects seen. To validate 

the phenotypes we observed and control for potential off-target effects of specific gRNAs, we 

selected several genes that had reproducibly high percent phenotypes to generate a second set of 

multiplexed gRNAs (Figure 2C).  

 

Fibrocystin regulates L-R patterning in somatic mutants in zebrafish and Xenopus 

pkhd1l1 encodes Fibrocystin, which is implicated in polycystic kidney disease (Harris and 

Rossetti 2004; Harris and Torres 2009). In zebrafish, Fibrocystin is encoded by tandem repeats, 

pkhd1l1.1 and pkhd1l1.2. I generated somatic mosaic mutants as described above and found L-R 

defects in mutants (Fig 2C). Targeting both repeats increased the levels of L-R defects. 

Additionally, we found that dand5 asymmetry was disrupted in Fibrocystin somatic mutants 

(Figure 3B-C). Wild-type AB embryos show stronger expression of dand5 on the right side. 

pkhd1ll double G0s embryos have right biased expression similar to WT but also have stronger 

expression on the left side as well, indicating decreased repression of dand5 on the left side.  

 In Xenopus, we found Pkhd1 (the ortholog of zebrafish pkhd1l1) is expressed in flow sensory 

cells in the L-R organizer (Figure 4). We targeted Pkhd1 with morpholinos and found L-R 

defects (Figure 3A). We specifically targeted 

cells on the left or right side of the embryo and 

found that knockdown of Pkhd1 on the right side 

had no effect on L-R patterning, whereas 

knockdown on the left side resulted in abnormal 

dand5 asymmetry (Figure 3D-E). Since the left 

side is where flow represses dand5, this suggests 

that Fibrocystin functions to activate dand5 

repression.  

Figure 4. pkhd1expression in the Xenopus 
Left-Right organizer. s=sensory cells. 



 

42 
 

 

 

Germline mutants do not phenocopy somatic mosaic mutants 

Upon a secondary screen through 16 genes, we selected four genes with reproducibly high 

phenotypes to generate whole-animal germline mutants for the following reasons: 

clstn1 is predicted to have calcium-binding activity. Therefore, Clstn1 may moderate L-R 

patterning through the Ca2+ spikes in flow sensory cells induced by Pkd2 (McGrath et al. 

2003; Yuan et al. 2015).  

eml2 is predicted to enable microtubule-binding activity and is associated with signal 

receptor binding, and thus influence L-R patterning by altering the cytoskeleton or acting 

as a signaling protein (McDowell et al. 2016).   

marveld1 is predicted to be involved in myelination. Intriguingly, marveld1 G0 mutants 

displayed axial defects in addition to L-R defects (Figure 2D). This suggests marveld1 

may impact L-R patterning the cilia level, as cilia-driven flow is required for axial 

straightening in zebrafish (Baker, Holtzman, and Burdine 2008; Kramer-Zucker et al. 

2005).   

Pkhd1l1 or Fibrocystin, which is encoded by two tandem repeats in zebrafish, pkhd1l1.1 

and pkhd1l1.2, is associated with autosomal recessive polycystic kidney disease 

(ARPKD) (Harris and Torres 2009), a disease with significant clinical overlap with 

autosomal dominant polycystic kidney disease (ADPKD) cause by Polycystin mutations. 

Therefore, Fibrocystin may moderate L-R patterning through interactions with the 

Polycystin proteins Pkd1l1 and Pkd2.  

Guides used to generate whole-animal germline mutants were selected from the 8 guides that had 

been used in the primary or secondary screen. To validate the CRISPR gRNAs selected to make 

germline mutants, I first injected two guides per gene at a high concentration along with Cas9 

and observed heart laterality. If guides produced L-R defects, then the pairs of guides were 

injected at a lower concentration to induce mutations. Mutations were confirmed by PCR and 

restriction enzyme digestion.  

 We raised injected embryos and performed subsequent outcrosses to generate F3 

homozygous adults. We bred pairs or groups of homozygous adults to generate homozygous 

mutant progeny and control for maternal effects. Surprisingly, we did not observe abnormal heart 



 

43 
 

 

jogging in clstn1, eml2, marveld1, or pkhd1l1 mutant embryos (data not shown). In clutches of 

30+ embryos, we scored low-to-near zero levels of heart jogging defects, matching wildtype 

levels of defects.  

 

Discussion 

Our findings pose conflicting phenotypic results between somatic mutants and whole 

animal mutants. In somatic mutants, L-R heart laterality defects happen at high frequency, but 

whole animal mutants had very low levels of L-R heart laterality defects. We hypothesize this 

may be due to a few factors. It is possible that L-R patterning defects in the heart are induced in 

somatic mutants due stress conditions. Mild cold stress has been shown to alter the movements 

of the dorsal forerunner cells (DFCs) that generate KV and altered DFC movement leads to 

disrupted KV formation and L-R patterning (Liu et al., 2022). Additionally, CRISPR-Cas9 

induced mutations in embryos generates mosaic embryos as the mutations are not always 

triggered as early as the one cell stage. Therefore, phenotypes induced in G0 mutants could be 

due to mosaicism in KV. Disruptions in varying amounts of cells in KV at random positions 

around the sphere may be enough to disrupt flow or flow sensation and induce L-R defects.  

Another possibility is differences in genetic compensation between somatic mutants and 

germline mutants. Reverse genetics screens with morpholinos (MO) in zebrafish, mice, and 

Arabidopsis have shown that there can also be differences between genetic mutants and gene 

knockdown, where genetic mutants display mild or no phenotypes, and morphants display more 

severe phenotypes (Daude et al. 2012; De Souza et al. 2006; Gao et al. 2015; Kok et al. 2015; 

Law and Sargent 2014; Smart and Riley 2013; Stainier, Kontarakis, and Rossi 2015). Previous 

work shown that deleterious mutations can induce upregulation of similar genes to compensate 

for the lack of the originally deleted gene, however, this is not detected in morphants with 

transcriptional or translational knockdown (Rossi et al. 2015). Further research has demonstrated 

that alleles that transcribe the mutated gene are able to induce genetic compensation, while true 

nulls that generate no transcript do not induce genetic compensation (El-Brolosy et al. 2019). 

Mutations were induced as early as possible within the first few exons of our germline mutants 

and thus transcriptional start sites likely are preserved. Therefore, genetic compensation could 

possibly explain the differences between G0 somatic mutants and germline mutants. It would be 



 

44 
 

 

interesting to understand if mutations induced through injection of multiplexed gRNAs is 

sufficient to trigger genetic compensation in zebrafish embryos.  

Our results show that Fibrocystin may function in the L-R patterning pathway, with 

evidence in both zebrafish and Xenopus. Fibrocystin localizes to primary cilia, the basal body, 

and cytoplasm in cell lines (Menezes et al. 2004; Wang et al. 2004; Ward 2003). In mouse 

kidney, it physically interacts with Pkd2 and loss of either causes kidney cysts (Kim et al. 2008). 

In KV, Pkd2 interacts and colocalizes to cilia with Pkd1l1 (Kamura et al. 2011). Transcriptomic 

analysis between zebrafish germline mutants and control siblings will be informative to 

determine if genetic compensation is masking a mutant phenotype.  

 

 
Methods 

Generation of germline mutants 

Embryos were injected with two single gRNA CRISPR guides at a concentration of 150µg/nl 

with Cas9 protein at the single cell stage. Embryos were raised and screened for mutations. I 

confirmed the guides induced mutations by PCR to amplifying around the cut site followed by 

restriction-enzyme digestion of the fragment. In embryos or fish where mutations were induced, 

the enzymes failed to digest the PCR fragment.  

F0 adults were outcrossed to ABC WT and embryos were screened for mutations. Adult fish that 

produced progeny with mutations were selected as founders. F1 progeny were raised and then 

outcrossed to ABC WT to isolate specific alleles. F2 heterozygous progeny were generated and 

then incrossed to generate F3 homozogous adults. F3 adults were incrossed to generate maternal-

zygotic mutant progeny for analysis.  

 

Statistical considerations for the L-R screen 

As heart positioning is nominal, square tests to determine significance. I performed power 

analyses to determine the number of embryos needed to assess significance. I assumed an 

instance of abnormality at 5% for controls and 66% for injected, based on the rates across 

validation experiments. From this, I calculated the sample size to be 26 individuals per group, 

with significance level alpha=0.05 and power=95%. In my screen, I performed two injection 



 

45 
 

 

replicates per gene, and injected at least 30 embryos per replicate, giving around 60 embryos per 

group, which is above the 26 required for statistical power.  

 
 
 

 
  



 

46 
 

 

CHAPTER IV 

 

DISCUSSION 

 

Overall, this work demonstrates a role for RNA-binding proteins and transcription factors in 

aspects of morphological identity of motor neurons in the Drosophila CNS. While we 

demonstrate roles for Imp and Chinmo in specifying morphological identity in motor neurons, 

we hypothesize that these genes may also have roles in morphological identity in interneurons. It 

would be interesting to understand if Imp and Chinmo play roles molecular identity in 

interneurons or if this is only observed in larval stages. Future work understanding the role of 

Imp, Chinmo, and Sequoia in interneurons will add to our knowledge or neuronal specification 

and identity more broadly.  

 We found that Imp and Chinmo knockdown caused opposing phenotypes in motor 

neuron axons and dendrites. While dendrites showed an increase in arborization with ectopic 

growth into the midline, axons showed decreased growth with axons not extending out to their 

target muscles. A similar phenotype has been seen in sequoia mutants in multidendritic sensory 

neurons. It would be interesting to understand how both over and under growth of axons and 

dendrites is achievable in the motor neurons. Further, understanding the behavior consequences 

of these morphological changes would be critical to understanding the functional role of these 

ectopic dendrites and improperly targeted axons. To understand the functional role of Imp and 

Chinmo, future work to know of these dendrites and axons for functional synapses will also be 

critical.  



 

47 
 

 

APPENDIX 
 

Appendix A. Chapter I supplementary figures 
 

 
 
Supplemental figure 1. Late larval proteins show little or no expression in embryos.  
Syp, Mamo, E93, are not expressed in the embryonic VNC. Br is expressed in embryonic 
neurons, but in a subset of cells rather than a gradient. Eve is shown as a fiduciary marker for a 
subset of post-mitotic motoneurons and interneurons. Stage 17 shown; anterior up; scale bar 
5μm. 
 
  



 

48 
 

 

Supplemental figure 2. The embryonic TTF cascade does not affect Imp levels 
(A-B) Overexpression of Hb does not alter Imp expression. (A) Hb is overexpressed in a stripe 
pattern via en-gal4 UAS-hb. Control, interstripe domain; Hb overexpression, stripe domain 
(brackets). (B) Quantification. n > 9 embryos. Ventral view. Scale bar = 5μΜ 
 
  



 

49 
 

 

Supplemental Figure 3. Imp overexpression can flatten the Imp gradient. 
(A) Imp overexpression in stripes (brackets) via en-gal4 UAS-Imp transgene. Ventral view, 
anterior up, scale bar, 10μm. (B) Imp overexpression leads to a flattening of the Imp gradient 
within the stripe domain.  
 
 
 
 
  



 

50 
 

 

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