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Negative valence neurons in the larval zebrafish pallium
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Negative valence neurons in the larval zebrafish pallium
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Content
NEGATIVE VALENCE NEURONS IN THE LARVAL ZEBRAFISH PALLIUM
By
Colton Dane Smith
A Dissertation Presented to the
FACULTY OF THE USC DORNSIFE COLLEGE OF LETTERS ARTS AND SCIENCES
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF NEUROSCIENCE
AUGUST 2024
Copyright 2024 COLTON DANE SMITH
ii
Epigraph
“The tree that never had to fight
For sun and sky and air and light,
But stood out in the open plain
And always got its share of rain,
Never became a forest king
But lived and died a scrubby thing.
The man who never had to toil
To gain and farm his patch of soil,
Who never had to win his share
Of sun and sky and light and air,
Never became a manly man
But lived and died as he began.
Good timber does not grow with ease,
The stronger wind, the stronger trees,
The further sky, the greater length,
The more the storm, the more the strength.
By sun and cold, by rain and snow,
In trees and men good timbers grow.
Where thickest lies the forest growth
We find the patriarchs of both.
And they hold counsel with the stars
Whose broken branches show the scars
Of many winds and much of strife.
This is the common law of life.”
- Douglas Malloch, Good Timber
iii
Acknowledgments
Wring an acknowledgments section is difficult as it could very well take up more space than the
entirety of this thesis. Alas, I don’t believe time permits me such an endeavor, so I will instead
attempt to begin settling my debts with small acknowledgments. Though you will likely say I
owe you nothing, I will forever owe you everything.
To my mother, my biggest fan and most avid supporter. I do not possess the words to express
how fortunate I am to have had your unwavering support throughout all these years. When I told
you I was going to become a neuroscientist, you never once questioned whether I had the
requisite intellectual ability or discipline for such an endeavor. You simply stated like you have
done with all my ventures, “Colton, once you’ve decided to do something, you will not stop until
you have accomplished your goal.” I wouldn’t be here without you. I love you.
To my fiancé, Joy, you have the patience of a saint and the kindness of one too. Knowing you
has redefined what I consider to be the meaning of life. Thank you for coming into my life at
exactly the right moment in time. You will forever be my Joy.
To my sister, to whom I have gotten closer with each passing day. We were raised in the same
garden. And by that, I mean to say that you and I will never be alone.
To my dear friend Q, you may think that your fate is in god’s hands, but I see how hard you
work on your own. If there is an all-powerful god, I choose to believe that he would only grant
blessings to the most hard-working and genuine of beings. If that is indeed the case, then your
cup of blessings is overflown. To my friend Mike, the countless hours spent talking with you
while playing video games has no doubt influenced who I am today. To my friend David, who
helped push me to be the best version of myself.
To Dr. Joseph LeDoux, many people know you for your work on the amygdala, but I know you
for your music. I find it funny that even though you’re a leading expert in the field that my thesis
topic addresses, we never once discussed it. I guess we were too busy having fun making music.
To Eric Morris, you’ve been an acting coach for more than 60 years, and your list of accolades
includes working with Jack Nicholson before he was famous, training Arnold Schwarzenegger,
and inspiring Johnny Depp. And yet, when a neuroscientist came knocking on your door, you
answered. Thanks for answering the door. You helped me to better engage with my creative side.
To Dr. Michael Angilletta, who knows the value of hard work and perseverance. You gave me
the wings to fly. To Dr. Jason Newbern, thank you for funding my master’s degree and giving
me a fighting chance. To Dr. Janet Neisewander, thank you for your guidance and support from
the beginning. To Dr. Carl Kesselman, thank you for having faith in me and pushing me to think
like a computer scientist. To Dr. Lawrence Krauss, your passion for communicating science to
the public has never ceased to inspire me. Thank you for the opportunity to work with the
Origins Project. To Dr. Irving Biederman, who even in his final days took the time to listen and
learn from his students. To Dr. Elyn Saks, thank you for giving me the chance to learn about how
we can apply scientific knowledge to help those who suffer from mental illness.
iv
To my committee, I genuinely feel that each of you has my best interest in mind. Dr. Scott
Fraser, your jovial attitude has been genuinely inspiring. I admire your ability to walk into any
room and be friends with anyone. Dr. Larry Swanson, I feel incredibly fortunate to have had you
on my thesis committee. Your experience in neuroanatomy has been invaluable and frankly
inspired me to delve deeper into the subject myself. Dr. Pat Levitt, thank you for being the head
of my committee and for your unwavering support. I admire you for always upholding the
highest standards of scientific rigor and integrity.
To my coworkers, Dr. Bill Dempsey, and Dr. Zhuowei Du, working with you in the early stages
of my PhD career was transformative. Your hard work and perseverance set me up for success
and I will forever be grateful for that. You two are the very definition of great coworkers. But
perhaps most importantly, you have become two of my friends.
To Dr. Don Arnold, thank you for allowing me to make my own mistakes and having enough
trust in me to work with minimal technical guidance. As far as intellectual guidance, you have
been invaluable. Over the years, I feel as though I have earned your respect and I feel honored to
be in this position. Your support has been invaluable to my intellectual growth, and I am honored
to have worked with you.
The work presented in this thesis was funded by the National Institutes of Health
(1R01OD019037-01 and 1U01NS122082-01)
v
Table of Contents
Epigraph.......................................................................................................................................... ii
Acknowledgments..........................................................................................................................iii
List of Figures................................................................................................................................ ix
Abstract......................................................................................................................................... xii
Chapter 1......................................................................................................................................... 1
Literature Review............................................................................................................................ 1
1.1 Introduction..................................................................................................................... 1
1.2 Fear ................................................................................................................................. 2
1.3 Emotional Learning ........................................................................................................ 3
1.4 Fear Conditioning ........................................................................................................... 6
1.6 The Zebrafish Pallium..................................................................................................... 7
1.6.1 Delineating the Boundaries of the Zebrafish Pallium................................................. 8
1.6.2 Where is the amygdala in zebrafish? ........................................................................ 11
1.6.3 The Zebrafish Pallial Amygdala (pAmy) ................................................................. 12
1.6.4 The Zebrafish Pallial Anterior Extended Amygdala ................................................ 13
1.6.5 Substance P (SP) in the Zebrafish Pallial Extended Amygdala................................ 14
1.7 Negative and Positive Valence Systems in The Brain.................................................. 16
Chapter 2....................................................................................................................................... 18
PNAS paper: ................................................................................................................................. 18
vi
Regional synapse gain and loss accompany memory formation in larval zebrafish .................... 18
2.1 Abstract............................................................................................................................... 18
2.2 Results................................................................................................................................. 19
2.2.1 Tail-Flick Classical Conditioning Protocol in Larval Zebrafish.................................. 19
2.2.2 Alterations in Pallial Neuronal Activity Correspond to Memory Acquisition ............ 21
2.3 Discussion........................................................................................................................... 25
Chapter 3....................................................................................................................................... 26
Noxious Stimuli: Exposing larval zebrafish to infrared laser heating and electric shock ............ 26
3.1 Abstract............................................................................................................................... 26
3.2 Introduction................................................................................................................... 27
3.3 IR Materials and Methods............................................................................................. 28
3.4 IR Results...................................................................................................................... 28
3.5 Electric Shock Materials and Methods......................................................................... 34
3.6 Electric Shock Results.................................................................................................. 34
3.7 Discussion........................................................................................................................... 40
Chapter 4....................................................................................................................................... 41
Threatening stimuli: exposing larval zebrafish to a full and partial looming stimulus ................ 41
4.1 Abstract......................................................................................................................... 41
4.2 Introduction................................................................................................................... 42
4.3 Full-Looming Stimulus Materials and Methods........................................................... 43
vii
4.4 Full-Looming Stimulus Results.................................................................................... 44
4.5 Partial-Looming Stimulus Materials and Methods....................................................... 50
4.6 Partial-Looming Stimulus Results................................................................................ 50
4.7 Discussion..................................................................................................................... 56
Chapter 5....................................................................................................................................... 57
Benign stimuli: Exposing larval zebrafish to physical vibration, light, and sound ...................... 57
5.1 Abstract......................................................................................................................... 57
5.2 Introduction................................................................................................................... 58
5.3 Tapping Stimulus Materials and Methods.................................................................... 58
5.4 Tapping Stimulus Results............................................................................................. 59
5.5 Light Stimulus Materials and Methods......................................................................... 65
5.6 Light Stimulus Results.................................................................................................. 65
5.7 Sound Stimulus Materials and Methods....................................................................... 71
5.8 Sound Stimulus Results................................................................................................ 71
5.9 Discussion..................................................................................................................... 77
Chapter 6....................................................................................................................................... 78
Elucidating the identity of the rostrolateral neurons in the pallium using Mapzebrain................ 78
6.1 Abstract......................................................................................................................... 78
6.2 Introduction................................................................................................................... 79
viii
6.3 Materials and Methods.................................................................................................. 79
6.3.1 Mapzebrain ............................................................................................................... 79
6.3.2 Tiam2a IR Laser Training......................................................................................... 82
6.3.3 Tiam2a pERK Immunohistochemisty....................................................................... 82
6.4 Results........................................................................................................................... 83
6.5 Discussion..................................................................................................................... 86
Chapter 7....................................................................................................................................... 88
Materials and Methods for Chapters 3-5 ...................................................................................... 88
7.1 Zebrafish husbandry and embryo/larval care................................................................ 88
7.2 Mounting Zebrafish in Caddie for Imaging on FlexSPIM............................................ 88
7.4 Arduino ......................................................................................................................... 94
7.5 Tg(βActin-NRSE::GCaMP6s)...................................................................................... 94
7.6 Data and Statistics......................................................................................................... 94
7.6.1 Calculating ΔF/F Traces, Area Under the Curve, and Statistics............................... 95
7.6.2 CaImAn..................................................................................................................... 98
Bibliography ............................................................................................................................... 102
ix
List of Figures
Figure 1.1 Schematic section labeled with topological terminology, drawing parallels
between the zebrafish pallium's four divisions and those of the mouse ................10
Figure 1.2 Molecular characteristics of the zebrafish amygdaloid complex ...........................15
Figure 2.1 Tail Flick Conditioning (TFC), a CC paradigm for larval zebrafish......................22
Figure 2.2 Neuronal activation within the rostrolateral pallium in response to the CS in
learner fish and to the US in naïve fish ..................................................................24
Figure 3.1 The regions of interest for this thesis .....................................................................30
Figure 3.2 The average fluorescence response of the zebrafish telencephalon in response
to infrared laser heating .........................................................................................30
Figure 3.3 Heat maps of the zebrafish telencephalon in response to IR laser heating ............31
Figure 3.4. Average brain region-specific activity traces before and after IR laser
heating....................................................................................................................32
Figure 3.5. The average activity of neuronal components before and after IR heating............33
Figure 3.6 The average fluorescence response of the zebrafish telencephalon in response
to electric shock .....................................................................................................36
Figure 3.7 Heat maps of the zebrafish telencephalon in response to electric shock................37
Figure 3.8 Average brain region-specific activity traces before and after electric shock........38
Figure 3.9 The average activity of neuronal components before and after electric shock ......39
Figure 4.1 The average fluorescence response of the zebrafish telencephalon in response
to a full-looming stimulus......................................................................................46
Figure 4.2 Heat maps of the zebrafish telencephalon in response to a full-looming
stimulus..................................................................................................................47
x
Figure 4.3 Average brain region-specific activity traces before and after a full-looming
stimulus..................................................................................................................48
Figure 4.4 The average activity of neuronal components before and after a full-looming
stimulus..................................................................................................................49
Figure 4.5 The average fluorescence response of the zebrafish telencephalon in response
to a partial-looming stimulus.................................................................................52
Figure 4.6 Heat maps of the zebrafish telencephalon in response to partial-looming.............53
Figure 4.7 Average brain region-specific activity traces before and after a
partial-looming stimulus........................................................................................54
Figure 4.8 The average activity of neuronal components before and after a
partial-looming stimulus........................................................................................55
Figure 5.1 The average fluorescence response of the zebrafish telencephalon in response
to vibration from tapping .......................................................................................61
Figure 5.2 Heat maps of the telencephalon in response to vibration from tapping .................62
Figure 5.3 Average brain region-specific activity traces before and after vibration from
tapping....................................................................................................................63
Figure 5.4 The average activity of neuronal components before and after vibration from
tapping....................................................................................................................64
Figure 5.5 The average fluorescence response of the zebrafish telencephalon in response
to a light turning on and off ...................................................................................67
Figure 5.6 Heat maps of the zebrafish telencephalon in response to a light turning on
and off ………….................................................................................................. 68
xi
Figure 5.7 Average brain region-specific activity traces before and after a light turning
on and off...............................................................................................................69
Figure 5.8 The average activity of neuronal components before and after in response
to a light turning on and off ...................................................................................70
Figure 5.9 The average fluorescence response of the zebrafish telencephalon in response
to the sound from an airhorn..................................................................................73
Figure 5.10 Heat maps of the zebrafish telencephalon in response to the sound from an
airhorn....................................................................................................................74
Figure 5.11 Average brain region-specific activity traces before and after the sound from
an airhorn ...............................................................................................................75
Figure 5.12 The average activity of neuronal components before and after the sound from
an airhorn ...............................................................................................................76
Figure 6.1 Tiam2a expression map on the Mapzebrain Atlas .................................................81
Figure 6.2 pERK expression in the transgenic Tiam2a zebrafish pallium following IR
laser exposure.........................................................................................................85
Figure 7.1 The schematic of the caddie used to house the zebrafish on the flexSPIM ...........90
Figure 7.2 Selective plane illumination microscopy................................................................92
Figure 7.3 Schematic detailing image acquisition throughput on flexSPIM experiments ......93
Figure 7.4 Example code for calculating ΔF/F, the area under the curve, and statistics.........97
Figure 7.5 CaImAn Parameters for Proper Signal Extraction ...............................................100
Figure 7.6. CaImAn Identified Component Example.............................................................101
xii
Abstract
To this day, the amygdala's functional and anatomical homologue in the zebrafish
remains a mystery. This is in part because the zebrafish, although a vertebrate by nature, has a
distinct neurodevelopmental plan, making it difficult to establish comparative homology to that
of the mammalian brain. Furthermore, our capacity to image the entire brain of the zebrafish is
currently restricted to developing larval zebrafish, making it difficult to understand how
information from larval zebrafish correlates with the neuroanatomical maps of adult zebrafish.
This thesis adopts the approach of exposing larval zebrafish to a wide variety of sensory
stimuli, ranging from harmful to not harmful, to identify amygdalar correlates within the
forebrain through the activity observed. Previous findings have shown increased pERK
activation in the rostrolateral pallium during fear conditioning, so this region has been designated
as a focal point for investigation.
Chapter 1 establishes the foundation for this work by examining the existing literature on
fear conditioning and the neuroanatomy of mammals, providing readers with the necessary
background to understand the potential analogs in zebrafish. Following that, I will provide an
overview of recent research on the zebrafish brain that advocates homology through molecular
and anatomical markers. Finally, I will delve into the complex interplay of the brain's valence
systems, creating a holistic context for the original research that follows.
Chapter 2 reviews prior experiments that motivated the current thesis topic. Chapter 3
details experiments designed to map neural activity in the zebrafish forebrain in response to
noxious stimuli. Chapter 4 expands to include the neural activity in response to life-threatening
stimuli. Chapter 5 contrasts these findings with the neural activity in response to benign stimuli.
This progression of experiments reveals a unique pattern of pallial activity in the larval zebrafish
xiii
that suggests functional specialization within the developing forebrain. To conclude, Chapter 6
presents work done to characterize the neurons of the rostrolateral pallium, potentially laying the
groundwork for future fear studies using larval zebrafish.
This thesis provides a much-desired understanding of the zebrafish forebrain, offering
insights into how stimuli are processed in the pallium that will hopefully serve as a guide to the
further identification of amygdala-like structures in the zebrafish and perhaps motivate more
interest in utilizing the zebrafish as a model organism for behavioral neuroscience.
1
Chapter 1
Literature Review
1.1 Introduction
In robotics there exists a set of rules a robot must obey. These are known as Asimovs
“Three Laws of Robotics”. These laws are as follows:
1. A robot may not injure a human being or, through inaction, allow a human being to come
to harm.
2. A robot must obey orders given it by human beings except where such orders would
conflict with the First Law.
3. A robot must protect its own existence as long as such protection does not conflict with
the First or Second Law.
These laws serve to guide the robot in such a way that it is subservient to mankind. All three
laws state in some capacity that robots are subservient to man, but the third law is the most
interesting. The third law states that a robot is only permitted to protect itself only if it does not
take priority over that of human life or human intention and what makes this so interesting is that
it is so counter to how many biological organisms behave. The third law is the opposite of selfpreservation.
Now let us imagine for a moment that a biological organism also has a set of laws it must
abide by. For the time being, I will provisionally call them “Smith’s Three Laws of Biology,”
2
acknowledging that these principles reflect intrinsic patterns observed in living organisms, rather
than novel discoveries of my own. They are as follows:
1. A biological organism must protect its own existence.
2. A biological organism must meet its physiological needs as long as such needs do not
conflict with the First Law.
3. A biological organism must propagate its genes so long as the act of doing so does not
conflict with the First Law or Second Law.
I am sure that my astute reader has realized that my first law is practically identical to Asimov’s
third law, with one large adjustment. Where my law differs is that self-preservation is paramount
to the biological organism, whereas not so with the robot. The discussion of the biological
substrates of self-preservation is central to what is to follow in the remainder of this thesis.
1.2 Fear
While discussions of self-preservation are commonly linked to discussions of fear, it
would be more accurate to say that the concepts are often used interchangeably, reflecting a
deeper misunderstanding of the nuanced relationship between the two. However, this assessment
is not to disparage the individual who conflates fear with self-preservation, but rather an
observation of the trickiness of words and the ever-pervasive threat of anthropomorphizing that
plagues our language and thoughts. For example, is it not an assumption to say that when a
mouse runs away from the shadow of a hawk this animal “feels” something? Or when we see a
school of fish scatter with the drop of a stone? Do we truly “know” that the animal “feels”
anything? Perhaps it is more a wish than a certainty. Joseph LeDoux, seen by many as the
world’s leading expert on the subject of fear, has raised important points regarding the language
3
we use to describe fear and anxiety, cautioning against the casual conflation of observed
behaviors with the presumed conscious experience in non-human subjects (LeDoux & Pine,
2016). However, since this is not a Wittgensteinian philosophical treatise on how the language
we use shapes our reality, I will keep these discussions to a minimum and will focus on
describing the experimental work and anatomical discoveries in this domain. That is not to say
that philosophical discourse on the emotional lives of animals holds no merit. I merely mean to
say that it is deficient without an accompaniment of the biology that brings it into being.
1.3 Emotional Learning
The standard and classic paradigms that are used to study fear and anxiety started
originally as paradigms through which to study emotional learning (Ledoux, 2000). That is to
say, the assays we use today were developed originally to understand how animals interact with
their environments and to get a better understanding of the neural mechanisms behind these
behaviors. This research was pioneered during the early twentieth century, a century which for
the scientific community was defined in part by luminaries such as Pavlov, Sherrington, Cannon,
Papez, and Hebb, who spearheaded research into the mechanical basis of animal behavior
(Ledoux, 2000). Their collective efforts were aimed at understanding how animals engage with
their surroundings and deciphering the neural mechanisms that govern their behaviors. One
could say that this was the beginning of a trend where we began to gain a greater appreciation of
how neurology dictated all the behaviors and ways in which we interacted with the world and
with one another, a role once attributed to the ethereal soul.
Though it was a culmination of the work done in the early 20th century that laid the
groundwork for our current understanding of animal behavior, it was Pavlov’s seminal work on
4
classical conditioning that is remembered the most. Through his research, Pavlov demonstrated
that by exposing animals to repeated pairings of a stimulus (an unconditioned stimulus, US) with
a neutral stimulus (a conditioned stimulus, CS), a CS can acquire affective meaning (Pavlov,
1927). Among this new affective meaning assigned to the conditioned stimulus are profound
physiological and behavioral changes including defensive behaviors, hypoalgesia, endocrine
response, and autonomic system arousal (Ledoux, 2000). This form of learning has been
observed across a diverse spectrum of organisms that are central to neuroscience research,
including but not limited to mice, rats, and humans, extending to more distantly related species
such as fish. Given the critical significance that this type of learning has for an organism, it's
unsurprising that the affective meaning learned in association with this conditioned stimulus
(CS) can endure for a lifetime (Gale et al., 2004; Mcallister et al., 1986). The durability of
learned responses may hold clinical significance and may give us insight into the myriad of
psychological disorders we humans endure, ranging from PTSD and anxiety to a myriad of other
psychological disorders.
A discussion on emotions is incomplete without discussing the brain regions that make
them possible, i.e. the neurological underpinnings. The first recorded experiment demonstrating
the role of a specific brain region responsible for processing emotional responses was done in
1888 by Brown and Schäfer (Brown & Schafer, 1888). The experiment involved the removal of
both temporal lobes of a rhesus monkey and a subsequent review of its behavior. The monkey
retained its basic sensory abilities yet appeared to lose the capacity to comprehend the
significance or meaning of various sensory inputs. Moreover, they observed a notable decrease
in the monkey's aggression and fear response, a finding later corroborated by Klüver and Bucy,
(Klüver & Bucy, 1938). Although these early studies involved the removal of too extensive an
5
area of brain tissue to accurately discern the specific contributions of each brain region within
the lesioned area, they laid the groundwork for understanding how the brain merges sensory
experiences with their emotional context. Importantly, these lesions encompassed a critical
structure that has since been the focus of extensive research: the amygdala.
But what is the significance of the amygdala and why does a further understanding of
how it functions concern us? Well, research has shown that humans with damaged amygdala
show deficits in fear conditioning, a subtype of classical conditioning that involves pairing a CS
with a noxious US such that over time the CS will begin to elicit defensive behaviors from the
organism (Bechara et al., 1995; Labar et al., 1995). Fear conditioning has been integral to
understanding how organisms learn to appropriately assign stimuli as dangerous and react
accordingly. Interestingly, another study found that subjects diagnosed with
psychopathy/antisocial personality showed significant volume reduction in the amygdala,
demonstrating the importance that the amygdala may play in determining our temperament
(Yang et al., 2009). The role that the amygdala plays in human temperament has even entered
public discussion centered on such figures as the professional rock climber Alex Honnold, who
climbs mountains without ropes and harnesses. In 2016, Dr. Jane Joseph at the Medical
University of South Carolina conducted an fMRI brain on Alex and found that he needed a much
higher level of stimulation than others to activate his amygdala. Findings like this raise the
interesting question of whether his amygdala has always been underactive or whether the
experiences of his life caused habituation to fearful stimuli. Like many questions on the topic of
nature vs nurture, I suspect the answer is both.
It's noteworthy to mention that the amygdala also plays a role beyond experiencing fear
itself. Specifically, individuals with damage to the amygdala encounter difficulties in interpreting
6
the nuances of emotional facial expressions and vocal tones, highlighting the amygdala's critical
role in processing emotional cues (Adolphs et al., 1995; Calder et al., 1996; Scott et al., 1997).
This suggests that the amygdala also plays a role in assessing and ascribing emotional valence.
1.4 Fear Conditioning
The amygdala is responsible for both the activation and expression of conditioned fear
responses (Adolphs & Anderson, 2018). Of the many distinct nuclei housed within the confines
of the amygdala, only three have been shown to be involved in fear conditioning: the central
nucleus (CeA), the basolateral nucleus (BLA), and the lateral nucleus (LA) (Adolphs &
Anderson, 2018). Numerous research methods have helped to build the standard model that
posits that the BLA is responsible for the initial learning of fear responses, while the CeA
mediates the display or output of the conditioned response (CR), as noted by a physical response
(Ledoux, 2000). In this framework, the CS and the US merge within the LA, enhancing the
synaptic activity in reaction to CS, and this information is sent to the CeA, which acts as the
amygdala's main conduit for output reactions (Adolphs & Anderson, 2018).
However, the internal wiring of the amygdala is much more complex than was once
assumed. New tools have been and will continue to be developed that make our old tools, and the
findings based on those tools, irrelevant. For example, recent optogenetic experiments have
shown that mice exhibited freezing behavior in their home cages after optogenetic activation of
Sf1+neurons in the dorsomedial ventromedial hypothalamus (Kunwar et al., 2015). This
evidence implies that freezing can be an unconditioned response, triggered in this case by direct
activation of the VMHdm. Furthermore, since these brain regions are also activated in natural
situations, like when an animal encounters a predator, it suggests that the brain can mediate
7
innate freezing responses to certain threats without the need for prior conditioning. To effectively
harness fear conditioning as a tool for understanding the nervous system's process of forming
associations with the environment, researchers should employ stimuli that are known to elicit
immediate freezing or flight reactions in naive mice from the very first exposure, such as the
overhead looming disk. This approach not only ensures the reliability of the experimental results
but also aligns closely with the natural behavioral mechanisms of the subjects under study.
1.6 The Zebrafish Pallium
The quest to define homologous or orthologous brain regions in fish has been
problematic, as their telencephalon is substantially different from that of mammals (Medina &
Abellán, 2009). In these fish, a developmental eversion causes a significant spatial
reorganization of the forebrain areas compared to other vertebrates (Mueller et al., 2004;
Nieuwenhuys, 2009; Northcutt, 2008; Rink & Wullimann, 2004). Despite years of research,
scientists have not yet been able to uniformly identify and map the pallial and subpallial regions
in even the most studied teleost fish, such as the zebrafish (Northcutt, 2008). There is also a lack
of distinctive markers, such as pax6 and reelin, that are typically used in mammals to help
delineate cortical regions at various developmental stages (Costagli et al., 2002; Wullimann &
Rink, 2001). Furthermore, the wealth of molecular and genetic expression research conducted on
zebrafish during their embryonic and larval phases has not yielded compelling evidence of
homologous structures (Mueller & Guo, 2009). Several hypotheses have been proposed to
explain eversion, but the lack of developmental data supporting these models means that there's
still no consensus on the precise anatomical structure of even well-recognized pallial homologies
8
in teleosts, including the amygdala, hippocampus, and cortex (Nieuwenhuys, 2009; Northcutt,
2008).
1.6.1 Delineating the Boundaries of the Zebrafish Pallium
Mueller et al. 2011 provide what is to my knowledge the most thorough approach to
unravel the structure of the zebrafish pallium (Mueller et al., 2011). Using molecular marker
staining on adult zebrafish brains against NADPHd and parvalbumin, they were able to delineate
four genuine pallial histogenetic units within the zebrafish pallium. Like the mammalian brain,
the zebrafish pallium comprises four distinct histogenetic divisions; the medial (Dm), lateral
(Dl), posterior (Dp), and central (Dc) zones (Mueller et al., 2011).
The distinctive staining of NADPHd activity clearly delineated four distinct pallial units
within the zebrafish pallium (Mueller et al., 2011). The lateral zone (Dl), situated in the
dorsolateral part of the pallium, was prominently marked in dark blue. In contrast, the central
zone (Dc), positioned at the heart of the pallium, exhibited only a few NADPHd-positive cells.
The medial zone (Dm), adjacent to Dl, and the posterior zone (Dp), situated below Dl, showed
no NADPHd-activity. The positioning of these pallial zones forms the basis of the topographical
naming system used for teleosts (Nieuwenhuys, 1963; Nieuwenhuys & Meek, 1990).
By examining consecutive sections from the rostral pole of adult zebrafish, Mueller et al.
were able to distinguish the topology of the major pallial zones more clearly. Of note were the
notable differences in the positioning of the pallial zones at the rostral pole compared to their
arrangement at mid-level sections of the telencephalon. In the rostral sections, the NADPHdpositive Dl zone, was positioned laterally relative to Dc zone, which showed sparse NADPHdpositivity, rather than adjacent to the Dm zone like in more mid-level sections. In these sections,
9
the Dc zone was vertically positioned between the Dm and Dl zones. Given this information
from both the rostral and caudal portions of the pallium, the researchers proposed that the Dc
zone be considered as the proper dorsal pallial division from the subpallium.
Moving caudally, the Dl zone begins to overlap the Dc zone (Mueller et al., 2011). These
observations suggest that the Dc zone undergoes a ventral displacement into the middle of the
pallium during development. Mueller et al hypothesize that the invagination process is likely
driven by the medially directed expansion of the Dl zone. Even more caudal, a pronounced
indentation begins to form between the Dl and medial (Dm) zones, which is known as the sulcus
ypsiloniformis (Y). The Y zone is a landmark used to distinguish the Dm zones from the other
dorsal zones.
Corroborating the pallial divisions established by the NADPHd staining was the
distribution of parvalbumin (Mueller et al., 2011). The Dl zone was characterized by numerous
parvalbumin-positive cells along with prominently labeled parvalbumin-positive fibers. On the
other hand, the Dc zone lacked parvalbumin-positive cells, but the zone did contain an
abundance of parvalbumin-positive fibers. In the Dl and Dc zones, the majority of parvalbuminpositive fibers originate from the anterior and posterior pallium. From the anterior direction,
parvalbumin-positive fibers form bundles at the boundary between the pallium and subpallium.
These bundles originate from the projection neurons located in the dorsal (Vd) and ventral (Vv)
nuclei of the subpallium found at the rostral pole of the telencephalon. At the posterior end, both
Dc and Dl receive parvalbumin-positive inputs via the lateral forebrain bundle, whose
projections were from nuclei within the preglomerular complex, which functions as a major
sensory relay center in the diencephalon (Mueller, 2012).
10
Based on this work, Mueller et al constructed an anatomical map that provides a
topological comparison of the zebrafish telencephalon with that of the mouse (Figure 1.1). This
model posits that the medial (Dm) zone in zebrafish aligns with the ventral pallium (VP) in mice,
the posterior (Dp) zone with the lateral pallium (LP), the central (Dc) zone with the dorsal
pallium, and the lateral (Dl) zone with the medial pallium (MP). The medial (Dm), lateral (Dl),
and posterior (Dp) zones of the zebrafish pallium correspond to the pallial amygdala,
hippocampus, and piriform cortex (Braford, 1995; Northcutt, 2006; Portavella et al., 2002).
Figure 1.1. A schematic section labeled with topological terminology, drawing parallels between
the zebrafish pallium's four divisions and those of the mouse. Figure and legend taken from
Mueller et al., 2011. Abbreviations: BLA, basolateral amygdala; Ctx, cortex; CP, caudate
putamen; EN, entopeduncular nucleus; GP, globus pallidus; Hip, hippocampus; lot, lateral
olfactory tract; LP, lateral pallium; MP, medial pallium; NT, nucleus taeniae; Sep, septum; V,
ventral telencephalon (subpallium); Vd, dorsal nucleus of the ventral telencephalon; VP, ventral
pallium; Vv, ventral nucleus of the ventral telencephalon; Y, sulcus ypsiloniformis.
11
Defining the Dm zone as the equivalent of the pallial amygdala aligns with findings from
behavioral studies (Mueller et al., 2011; Portavella et al., 2002, 2004), the observation of
ascending fibers from cholinergic cells in the basal forebrain, and the distribution of calretininpositive cells (Castro et al., 2006; Wullimann & Mueller, 2004). Similarly, the Dl zone's
identification as the teleostean hippocampus is corroborated by behavioral research and the
distribution of molecular markers (Castro et al., 2006; Mueller et al., 2011; Portavella et al.,
2002). According to Mueller et al, the posterior (Dp) zone is recognized as analogous to the
piriform cortex in amphibians and other tetrapods, distinct from the overlying Dl zone that is
marked by NADPHd and parvalbumin expression and neuropeptide Y (Braford, 1995; Castro et
al., 2006).
1.6.2 Where is the amygdala in zebrafish?
In the most recent and comprehensive study on the zebrafish amygdala to date, Porter &
Mueller et al. 2020 explored the zebrafish amygdala's structure through an integrative approach,
merging immunohistochemistry data with developmental information (Porter & Mueller, 2020).
Given that zebrafish exhibit a highly developed sense of smell, their approach drew inspiration
from studies done on the extended amygdala and the primary olfactory cortex, neuroanatomical
systems found in macrosmatic rodents, mammals endowed with a highly refined olfactory
system (De Olmos & Heimer, 1999; Swanson. & Petrovich, 1998).
Generally, the amygdala is acknowledged as an assembly of multiple regions that
coordinate to play an instrumental role in autonomic and emotional regulation (Swanson and
Petrovich, 1998). According to Porter and Mueller, the concept of the extended amygdala
expands the classical definition of the amygdala to include several subpallial regions: such as the
12
centromedial nuclei (CeA, or MeA in zebrafish), the basolateral amygdala (BLA), and segments
of the bed nucleus of the stria terminalis (BST). Of particular importance to this study, some of
the projections from the olfactory bulb integrate into the amygdala, an important anatomical
connection driving scent-driven behavior (Porter & Mueller, 2020).
By utilizing the distribution patterns of specific neuronal phenotypes and calcium-binding
proteins, such as calretinin and parvalbumin, Porter & Mueller delineated thirteen distinct
amygdaloid regions, sorted into areas that are primarily GABAergic and subpallial or primarily
glutamatergic and pallial (Mueller et al., 2006; Porter & Mueller, 2020; Puelles et al., 2000).
Porter & Mueller assert that their molecular profiling, along with previous research that has
demonstrated the structural, functional, and transcriptional characteristics of the pallium,
provides sufficient evidence to consider three distinct pallial regions within the zebrafish: the
pallial amygdala (DM), the dorsal pallium, and the medial pallium (Porter & Mueller, 2020).
1.6.3 The Zebrafish Pallial Amygdala (pAmy)
Using the vGlut2a:GFP transgenic line alongside the distribution of GABA, the
researchers pinpointed the ventralmost vGlut2a-GFP-positive cells of the pallial amygdala
(pAmy) as a defining feature of the dorsal aspect of the boundary between the pallium and
subpallium (PSB), situated next to the GABAergic, and isl1 free cell groups of the zebrafish
dorsal lateral ganglionic eminence (dLGE) (Figure 1.D1–G). The researchers assert that these
findings indicate the DM pallial region should no longer be viewed as a single area but should
instead be molecularly divided into two distinct areas: an anterior DM and a separate posterior
DM, which interestingly contains a cluster of Dm120a neurons that have been implicated in
associative fear learning (Lal et al., 2018). Porter & Mueller hypothesize that the emx3-negative
13
posterior Dm likely contains the posteromedial pallial nucleus (PMPa), a putative homologue of
the mammalian posteromedial cortical nucleus (PMCo). Interestingly, neurons positive for
vGlut2a:GFP are densely concentrated in a section of the lateral olfactory tract (nLOT) and
likely have connections with the Dm. This is important given that the PMCo in mammals is the
recipient of accessory olfactory bulb projections, as well as indirect connections from the
olfactory bulbs (Kevetter & Winans, 1981; Scalia & Winans, 1975)
1.6.4 The Zebrafish Pallial Anterior Extended Amygdala
In the zebrafish, the striatopallidum (Vd), septum (Vv), anterior central amygdaloid
nucleus (CeA), and the central amygdaloid nucleus (CeAl or Vc) are all characterized by dense
clusters of GABAergic neurons (Figure 1.2B-C) (Porter & Mueller, 2020). This is in stark
contrast to the sparse GABAergic labeling found in the posterior pallial regions including the
vGlut2a:GFP-labeled pallial amygdala (pAmy) (Porter & Mueller, 2020).
By comparing the expression of parvalbumin (Pv) and tyrosine hydroxylase (TH, a
dopaminergic neuron marker) with the patterns of the isl1:GFP transgenic line (which expresses
cell populations in the subpallium) the researchers found that the isl1:GFP-negative and GABApositive subpallial extended amygdala, was differentiated from the isl1:GFP-positive
striatopallidal areas (Vd). Located dorsally to the TH-positive dopaminergic cells, they found a
lack of isl1:GFP labeling that delineates most of the CeA (Figure 1.2D1-D2). They also found
that the CeA is identified by its lack of isl1 expression, positioning adjacent to the MeAa and the
pAmy, and by a lack of otpa, calretinin, and substance P expression (MeA markers).
To summarize, calretinin-positive neurons characterize the medial amygdala, while the
absence/sparse expression of neurons expressing calretinin is characteristic of the central
14
amygdaloid nuclei (Porter & Mueller, 2020). The calretinin expression pattern in zebrafish is
akin to that described in mammals (Wójcik, 2013).
1.6.5 Substance P (SP) in the Zebrafish Pallial Extended Amygdala
In mammals, the accessory olfactory bulb projects to the medial amygdala (MeA), which
is critical for reproductive behaviors (Abellán et al., 2013). Although it has been shown that
zebrafish lack accessory olfactory bulbs, they do possess pheromone-binding receptors in their
olfactory bulbs, essential for reproductive behaviors (Behrens et al., 2014). This prompted
Mueller et al to investigate if a homologue exists in zebrafish.
By assessing the distribution of substance P, a marker commonly used in mammals to
distinguish the MeA from the CeA, Porter and Mueller were able to detect two substance-Ppositive olfactory bulb tracts that target the medial extended amygdala. In contrast, the lack of
substance-P-positive fibers in the nLOT implies that the two tracts process different information,
with the nLOT processing primary olfactory and gustatory information (Porter & Mueller, 2020).
15
Figure 1.2. Molecular characteristics of the zebrafish amygdaloid complex. Figure and legend
taken from Mueller, 2020. The figure covers the precommissural telencephalon (A–E),
supracommissural, and postcommissural telencephalon (F–N). (R) shows a schematic of the
pallium and where the sections are taken from. Abbreviations: BST, bed nucleus of the stria
terminalis; BSTa, anterior division of BST; BSTm, medial division of BST; BSTpd, posterior
16
division of BST; Cantd, anterior commissure, pars dorsalis; Cantv, anterior commissure, pars
ventralis; CeA, central amygdala; CeAa, anterior division of CeA; CeAl, lateral (migrated)
division of CeA; CeAd, dorsal division of CeA; Dm, medial zone of D; DM, precomissural
(vGlut2a positive) territory of Dm; DP, dorsal pallium; MeA, medial amygdala; MeAa, anterior
division of MeA; MeAd, dorsal division of MeA; MeAp, posterior division of MeA (zebrafish);
MeAv, ventral division of MeA (zebrafish pendant to mouse MeApd); MP, medial pallium; nLOT,
nucleus of the lateral olfactory tract; PMPa, posteromedial pallial nucleus; Vd, dorsal zone of V;
Vdd, dorsalmost territory of V (=“extended dLGE”); VP, ventral pallium; Vv, ventral zone of V.
1.7 Negative and Positive Valence Systems in The Brain
As has been detailed in other sections of this literature review, viewing the amygdala as
the seat of fear is a simplified view. More recently, researchers have shown that valence
processing is often a brain-wide phenomenon, distributed across many nodes (Mukherjee et al.,
2018; Xiu et al., 2014). However, that is not to say that this large-scale distributed valence
processing is redundant across the entirety of the brain. There are specific nodes in valence
processing that work to decipher and properly relay stimuli to produce appropriate behavioral
responses. In fact, researchers have shown that valence assessment can be heterogeneous even
within subregions (Tye, 2018). For example, as one moves across the rostral-caudal/mediallateral gradient of the nucleus accumbens (NAc), evidence has shown that some different
neurons respond to either negative or positive stimuli (Tye, 2018).
The consensus in the field of emotion is that there is a myriad of mechanisms the nervous
system employs to encode valence (Tye, 2018). The first circuit motif is known as the labeled
lines motif. This motif features separate sensorimotor pathways for processing positive and
negative stimuli. Evidence for this motif can be found in the olfactory and gustatory systems that
distinguish appetizing and unappetizing food (Root et al., 2014; Tye, 2018; Wang et al., 2018).
The second circuit motif is the opposing components motif. In this motif, a circuit comprises
17
neurons with diverse functions within a specific anatomical projection (Tye, 2018). Evidence for
this motif has been found in the lateral hypothalamus (LH), where both glutamatergic and
GABAerbic neurons reside (Tye, 2018). The third motif is the divergent paths motif. This circuit
motif receives identical sensory inputs but diverges towards different downstream targets,
governed by synaptic weights (Tye, 2018). Evidence for this motif can be found in the amygdala
where a small subset of projection-defined neurons can facilitate both anxiogenic and anxiolytic
behaviors, with anxiolytic neurons projecting from the BLA to the CeA and anxiogenic neurons
projecting from the BLA to the hippocampus (Tye et al., 2011). The fourth and final motif is the
neuromodulatory gain motif. This circuit features the activation of metabotropic receptors that
prolong the plasticity timeframes (Tye, 2018). An example of this motif is where dopamine plays
a brain region-dependent role, mediating negative valence in the prefrontal cortex and positive
valence in the ventral striatum (Hsing-Chen Tsai, 2009; Phillips et al., 2003; Witten et al., 2011)
18
Chapter 2
PNAS paper:
Regional synapse gain and loss accompany memory formation in larval
zebrafish
Chapter modified from my lab’s work and publication:
William P. Dempsey, Zhuowei Du, Anna Nadtochiy, Colton D. Smith, Karl Czajkowski, Andrey Andreev, Drew N.
Robson, Jennifer M. Li, Serina Applebaum, Thai V. Truong, Carl Kesselman, Scott E. Fraser, and Don B. Arnold
2022. Regional synapse gain and loss accompany memory formation in larval zebrafish. PNAS.
2.1 Abstract
The prevailing theory in modern neuroscience that the formation of memories is
attributed to changes in synaptic connections was first proposed more than 100 years ago
(Mayford et al., 2012). However, identifying the structural modifications that underpin memory
still poses a significant challenge. While studies of long-term potentiation (LTP) in brain slices
have given scientists a plethora of information, our understanding of how synapses in a living
vertebrate are altered during memory formation remains limited (Engert, 1999; Maletic-Savatic,
1999). Therefore, despite many longitudinal imaging studies capturing experience-induced
alterations in the synapses of brain slices, only a handful of studies have directly observed
synaptic modifications taking place in the amygdala memory formation (Attardo et al., 2015;
Holtmaat & Svoboda, 2009)
In pursuit of a better understanding of associative memory formation, Dempsey et al.
2022 merged in vivo labeling with synaptic level imaging to chart the temporal synaptic
alterations within the intact brain of a living larval zebrafish before and after the process of
memory formation. The region of interest for this study was the zebrafish pallium as it is thought
19
to contain the homologue to the mammalian amygdala (Wullimann & Mueller, 2004). The
pallium’s location on the brain's surface, along with the high transparency of zebrafish larvae
enabled non-invasive, whole-brain imaging using SPIM microscopy.
In larval zebrafish that successfully formed memories after associative conditioning,
Dempsey et al. observed a significant increase in the number of synapses in the ventrolateral
pallium alongside a notable reduction in synapses in the dorsomedial pallium. Furthermore,
Dempsey et al. found evidence that neurons in the rostrolateral pallium, neurons which are a
central topic in this thesis, exhibited increased levels of pERK when exposed to the
unconditioned stimulus and in zebrafish learners exposed to the conditioned stimulus. These
findings indicate that conditioning-induced memory formation in zebrafish is linked to reciprocal
alterations in the number of synapses within the rostrolateral pallium and may involve negative
valence neurons. The contents of this thesis continue the investigation into how those areas of the
zebrafish brain process negative valence information.
2.2 Results
2.2.1 Tail-Flick Classical Conditioning Protocol in Larval Zebrafish
Dempsey, et al. 2022 designed a tail-flick conditioning (TFC) procedure to instill
associative memories in zebrafish aged 14 to 16 days post-fertilization. This method employs
heating from a near-infrared laser as the unconditioned stimulus (US) and illumination from
LED light as the conditioned stimulus (CS). The larval zebrafish heads are secured in agarose,
allowing free movement for the tail flick, which serves as the unconditioned response (UR) and
conditioned response (CR) as displayed in Figure 2.1A. A specialized behavioral setup was used
to perform the classical conditioning (CC) induction and evaluation across three distinct phases,
20
each with its own stimulus pattern: 1) habituation with CS alone (20 trials); 2) training with
overlapping CS and US (20 trials); and 3) testing with CS alone (5 trials), as shown in Figure
2.1B–D.
For imaging studies, fish were subjected to a fourth phase, reexposing them to the
training sequence for 10 additional trials to counter any possible extinction effects from the
testing phase (Figure 2.1B). Control groups were exposed to either the US, the CS, or no
stimulus (NS).
To verify learning, Dempsey, et al. 2022 measured tail-flicking. Baseline tail-flick rates
were established during the last five habituation trials pre-TFC, and random flicking was gauged
over five similar intervals without any stimulus post-training. The data is plotted in a cumulative
histogram (Figure 2.1E). Observing tail flicks during the five testing trials showed that 30% of
the fish flicked their tails at least three times in response to five CS presentations. Since none of
the control fish flicked more than twice, it was inferred that the minimum learning rate was 30%.
Fish that flicked in response to all five CS presentations during testing were labeled as
superlative learners (L, Figure 2.1C–E), those responding to three or four CS presentations were
termed partial learners (PL, Figure 2.1C–E), and those showing no response were deemed
nonlearners (NL).
As anticipated, fish that had undergone training exhibited increased tail flicking in
response to the US, while control fish that had not encountered the US showed negligible
flicking in the same timeframe (Figure 2.1F). Significantly, fish subjected to TFC had a CSinduced flick rate that was higher than that of the control group.
Learning during trace fear conditioning (TFC) was significantly inhibited when the fish
were exposed to APV, indicating that TFC learning relies on synaptic plasticity mediated by
21
NMDA receptors. Further experimentation to assess the potential for TFC learning to be
extinguished showed that fish classified as learners exhibited a decreased response to the CS
after 25 presentations without the unconditioned stimulus, with their response dropping to below
25% of the initial level. In contrast, the response levels of fish not re-exposed to the CS remained
largely unchanged from their initial response. These findings suggest that TFC learning in
zebrafish is dependent on NMDA receptor activity and is capable of extinction, which are key
characteristics of classical conditioning.
2.2.2 Alterations in Pallial Neuronal Activity Correspond to Memory Acquisition
Given that the dorsal pallium of the zebrafish is thought to contain the amygdala, it stands
to reason that pallial neurons would exhibit structural and functional alterations after TFC
Dempsey, et al. 2022. We assessed variations in pERK expression, an indicator of neuronal
activation, in response to the CS in zebrafish, for both learner and non-learner fish. Post-CS
presentation, fish demonstrated a pronounced increase in pERK within a specific anterolateral
region of the pallium (Figure 2.2A). The enrichment of pERK signaling was not observed fish
that didn’t learn, (Figure 2.2B). To delve deeper into pallial functionality, fish were subjected
naive fish to repeated US exposures. This resulted in heightened pERK levels in a specific area
of the anterolateral pallium, aligning with the CS-responsive region found in learner fish (Figure
2.2C). CS only exposed fish and fish not exposed to either the CS or US, exhibited little to no
pERK staining (Figure 2.2D-E). Therefore, the pallial response to the CS in learner fish mirrored
the response to the US in naive fish, demonstrating that after learning, the CS activated the same
rosrolateral area as the US.
22
Dempsey, et al. 2022 calculated the ratio of pERK labeling to a control region and found
that learner fish possessed a much higher ratio than non-learner and control fish. Similarly, US
exposed fish also displayed a marked increase in this ratio (Figure 2.2F). These findings indicate
that the very same neurons that respond to the US are the same neurons that respond to the CS
after training, suggesting these neurons are negative valence.
Figure 2.1. Tail Flick Conditioning (TFC), a CC paradigm for larval zebrafish.
Figure and legend taken from Dempsey et al, 2022.
(A) The head of the zebrafish undergoing TFC is encased in low-melt agarose, leaving the tail
free to move. The CS consists of light from a green LED; the US is heat produced by an NIR
laser. The CR is tail flicking. (B) Timeline of the TFC paradigm. (C) During training, fish are
exposed to both the CS and US. In response to the US, all fish flicked their tails vigorously
(black bars). Here, we display late-stage training rounds for three L and three NL fish. (D)
During the testing phase of TFC, the fish is exposed to the CS alone. L respond immediately
upon presentation of CS; NL do not. Here, early-stage testing rounds are shown. (E) Cumulative
23
histogram of the percentage of larval zebrafish that respond with tail flicking to the five CSs
presented during testing one or more times, two or more times, etc. (After TFC, inverted
triangles). The baseline flicking histograms from fish during the final rounds of Habituation
(Before TFC, gray triangles) or from fish assayed after TFC for five time windows when no
stimulus is present (Random, filled squares) are different. (F) The FR (fraction of time tail is
flicking) when US is presented during training are similar for L (n = 11 fish), PL (n = 6), NL (n =
11), and US only (US, n = 11) and different from the FR during the same time period for fish
exposed to CS only (n = 11) or NS(n = 11; ***P < 0.005, *P < 0.05, Kruskal–Wallis test). No
pairwise comparison between L, PL, NL, or US is significant except for L versus PL (*P < 0.05,
Kruskal–Wallis test). (G) The FR in response to the CS during testing is significantly different in
fish exposed to TFC versus control fish (***P < 0.0001, Kolmogorov–Smirnov test). Cumulative
probability distributions are shown. (H) The FR (averaged for all fish during testing) is
significantly reduced in fish exposed to 2-amino-5-phosphonovaleric acid (APV) during TFC
compared to control fish (n = 12 fish +APV, 19 fish APV; **P < 0.01, Mann–Whitney U test).
Each data point represents the response to a single presentation of the CS. (I) The FR diminishes
over 30 presentations of CS alone (extinction, n = 5) in a set of L fish by ∼75%. (J) FR before
and after extinction (Ext, t1 and t2)are significantly different (***P < 0.005, n = 5 as in H,
Mann–Whitney U test). FR at t1,t2 without extinction (n = 6) are not significantly different (P >
0.05, Mann–Whitney U test). Data available at https://doi.org/10.25551/1/1-1YZE (45).
24
Figure 2.2. Neuronal activation within the rostrolateral pallium in response to the CS in learner
fish and to the US in naïve fish. Figure and legend taken from Dempsey et al, 2022. (A)
Intense immunostaining of pERK in the pallium (magenta highlighted region, Inset)of an L
(learner) fish exposed to 5 CSs (control stimuli) following TFC (tail-flick-conditioning). The
strong signal in a rostrolateral region (yellow outline) of this optical section reveals regional
neuronal activation. Relatively less immunostaining is present in the medial pallium (cyan
outline). (B) A NL (non-learner) fish shows a lack of pERK staining in the rostrolateral region
(yellow outline) after exposure to 5 CSs in this equivalent optical section. (C) A naïve fish
reveals strong pERK staining in the same rostrolateral region (yellow outline) after exposure to
10 USs (unconditional stimulus). Equivalent optical section to those in A and B. (D) A naïve fish
exposed to 10 CSs does not show concentrated pERK labeling in the rostrolateral region (yellow
outline). Optical section equivalent to those in A–C. (E) A naïve fish not exposed to a CS or US
(NS) does not show concentrated pERK labeling in the rostrolateral region (yellow outline).
Optical section equivalent to those in A–D. (F) L and US-exposed naïve subjects show a
significantly higher lateral:medial pERK intensity ratio compared to NL and naïve untreated
subjects (*P < 0.02, ***P < 0.005, n = 5 fish per group, Kruskal–Wallis multiple comparison
25
test). White dashed lines mark the border of the pallium (midline = M) in A–E. (Scale bar for A–
E,20 μm.)
2.3 Discussion
The experiments in Dempsey, et al. 2022 reveal that neurons in the rostrolateral pallium shows a
heightened response (demonstrated by elevated pERK) to the US in naïve fish and to the CS in
learner fish. This suggests the rostrolateral pallium facilitates the initial formation and recall of
associative conditioning. Notably, this is the same region where Dempsey, et al. observed a
significant increase in synapse formation in learner fish suggesting a correlation between cellular
and synaptic activity in this region. These pERK experiments are supported by recent research
done in mice that show increased calcium activity in the amygdala before and after fear
conditioning (Grewe et al., 2017). The responses exhibited in this study by the rostrolateral cells
of the pallium may share homology with the aversive-response cells in the mouse's basolateral
amygdala (Namburi et al., 2015). However, in contrast to the dispersed distribution of aversive
cells in the mouse amygdala, the rostrolateral cells of the zebrafish pallium appear to be more
clustered, making them more amenable to scientific experimentation (Dempsey, et al. 2022).
26
Chapter 3
Noxious Stimuli: Exposing larval zebrafish to infrared laser heating and
electric shock
3.1 Abstract
It is known that the amygdala becomes active when an organism is exposed to painful stimuli
(Corder et al., 2019). Our prior research has identified a unique population of cells in the
rostrolateral pallium that showed an increase in pERK activity, a marker for active neurons, after
being exposed infrared laser heating. Though this result suggested a location to look for
activation in response to painful stimuli, it gives no information about its temporal component.
All experiments were done using 7 dpf larval zebrafish. Note that initial experiments were done
using 14 dpf larval zebrafish, as in Dempsey et al. However, we found that 7 dpf fish were better
able to tolerate the experimental paradigm and generated very similar data (not shown). We
exposed Tg(βActin-NRSE::GCaMP6s) zebrafish, positioned on a light sheet microscope, to an
infrared laser a total of five times while collecting images from which we later measured the
average response of ΔF/F. We also performed experiments using electric shock to determine
whether the activation patterns observed with the IR experiments were unique to laser heating or
universal for painful stimuli. For IR heating experiments, there was a dramatic bilateral increase
in the activity of rostrolateral neurons within the pallium. Furthermore, several other areas
including the habenula and a newly identified ventromedial area were also more active. When
exposed to electric shock, there was also a dramatic bilateral increase in the activity of
rostrolateral, habenula, and ventromedial areas within the pallium. These results provide
27
evidence that the rostrolateral neurons respond to both infrared heating and electric shock,
suggesting that they are sensitive to a variety of painful stimuli.
3.2 Introduction
Extensive research has been done into how the amygdala processes sensory information
from noxious stimuli leading to avoidance behaviors in mammals (Adolphs & Anderson, 2018).
For instance, the case of a neurosurgery patient, H.M., who had a bilateral medial temporal
lobectomy, demonstrated that injury to the basolateral amygdala (BLA) can lead to a condition
where harmful stimuli are recognized but fail to initiate avoidance behaviors (Hebben et al.,
1985). In contrast, damage to the somatosensory cortex impairs the capability to localize painful
stimuli and to quantify severity, while preserving avoidance behaviors suggesting that the BLA
may connect pain signals to feelings of discomfort and the choice of behaviors to mitigate such
experiences (Corder et al., 2019; Uhelski et al., 2012).
In contrast, how zebrafish process nociceptive stimuli is not fully understood. This
includes a lack of consensus on the anatomical location of the amygdala even in adult fish
(Mueller et al., 2011), let alone in larval zebrafish where most zebrafish research is conducted.
Because of the inverted developmental trajectory of zebrafish compared to mammals, it is
difficult to establish comparative homology based on anatomical location. Additionally, there are
no agreed upon markers of negative or positive valence neurons in the zebrafish like there are in
mice (Kim et al., 2016).
Our approach to this complex topic is to build from information gathered in Dempsey et
al, 2022 where we discovered a putative negative valence neuronal population clustered in the
rostrolateral zone of the pallium. We demonstrated using pERK that this area was highly active
28
after the zebrafish was exposed to an IR laser and highly active to a conditioned stimulus after
associative conditioning (Figure 2.2). In this chapter we will investigate the temporal dynamics
of the rostrolateral neuronal activation using GCaMP6s to determine whether this activation is
unique to the IR laser or extends to electric shock, and we will investigate how other areas in the
pallium respond to noxious stimuli.
3.3 IR Materials and Methods
Once the fish is prepped for imaging (refer to materials and methods found at the end of
this thesis for a more thorough description of fish preparation) it is positioned such that the
zebrafish’s left eye is hit by the infrared laser (NIR RLCO-980-1000-F laser, Roithner
Lasertechnik). After 1.5 hrs of acclimation, the fish is imaged for 5 min to establish a baseline
for neuronal activity. Then, while still imaging, the IR laser is turned on for 2 seconds before
being turned off. This is repeated 2 minutes later for a total of 5 trials. The experiment is then
terminated. Total experiment time is ~15-20min. After the experiment is finished, the fish is
released from the agarose and allowed to rest in a dish for one hour. Only fish that are still alive
and healthy after this waiting time are included in the analysis.
For further experimental details, methods, and code, refer to Chapter 7.
3.4 IR Results
Infrared stimulation activates three distinct areas of the telencephalon.
Figure 3.2 shows a representative image of where the Infrared (IR) laser was applied to
the zebrafish (Figure 3.2A-B). Refer to Figure 3.1 for reference to the anatomical locations
chosen for this analysis (Figure 3.1A-C). After presenting the laser stimulus to the zebrafish a
29
total of five times for the duration of the experiment, 2 min intervals, the average activation of
the brain is taken before and after the stimulus for six separate regions; left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). All six regions have varying levels of increased activation following the IR
stimulus, as can be seen in a representative fish (Figure 3.2B). Coronal and axial views of the
representative fish display the six distinct areas of activation after stimulation from another
perspective (Figure 3.3A). The change in fluorescence, ΔF/F, measured before and after
stimulation, 5 frames (6.65sec), shows sudden bilateral increases in the Hb, Rl, and Vm areas of
the telencephalon (Figure 3.4A). This increase in fluorescence was found to be statistically
significant across multiple fish (n=5) by comparing the area under the curve (AUC) of
fluorescence intensity before and after IR stimulation (lHb: p=0.0022, rHb: p=0.0061, lRl:
p=0.0098, lRl: p=0.0036, lVm: p=0.0017, rVm: p=0.0027; *p<0.05, **p<0.01, ***p<0.001 as
determined by one-way t-tests) (Figure 3.4B). Using CaImAn, individual neuronal components,
areas of activity detected over the duration of the experiment, were detected were extracted from
the brains (Figure 3.5A) The traces that accompany these components were found to exhibit a
range of activation and inhibition immediately following stimulus onset with this effect
diminishing over the following 20 seconds. These activity patterns are ordered using z-scores,
ranging from the highest (highly active) to lowest z-scores (inhibited activity) and are displayed
using a heat map (Figure 3.5B). In addition to the z-scored map, the ΔF/F traces for these
components are shown (Figure 3.5C).
30
Figure 3.1. The regions of interest for this thesis. Representative images of A) both habenula, B)
both rostrolateral areas, and C) both ventromedial areas and the size of the regions examined
(50µm x 50µm) and the depth from the top of the pallium for the “Average fluorescence
response of the zebrafish telencephalon” figures for Chapters 3-5. The blue line delineates the
borders of the pallium.
Figure 3.2. The response of neurons in the zebrafish telencephalon to infrared laser heating. A)
Configuration of an Infrared (IR) laser used to generate a noxious heat stimulus in a 7 dpf larval
zebrafish. B) The average brain response of several brain regions shown from representative fish
after being hit five times with an IR laser. The brain regions include: the left habenula (lHb),
right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and
right ventromedial (rVm). The dotted lines delineate the brain regions from the rest of the fish,
with the habenula circled within the brain, the rostrolateral regions delineated from the skin of
the fish, and the ventromedial regions delineated from the midline of the forebrain. The time
between each time point is 1.33 seconds. The black bars represent the time in which the there
was no stimulus, and the red bar indicates the time in which infrared laser heating was
administered to the fish.
31
Figure 3.3. Heat maps of the zebrafish telencephalon in response to infrared laser heating. A)
Coronal view of a representative fish emphasizing the change in rostrolateral and ventromedial
activity before and after laser heating. B) Axial view of a representative fish emphasizing the
change in rostrolateral, ventromedial, and habenular activity before and after laser heating.
Abbreviations: left habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right
rostrolateral (rRl), left ventromedial (lVm), and right ventromedial (rVm)
32
Figure 3.4. Average brain region specific activity traces before and after IR laser heating. A)
Brain region specific ΔF/F traces for the left habenula (lHb), right habenula (rHb), left
rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial
(rVm). B) Average area under the curve shown for each brain region, 5 frames (6.65sec) before
and after IR laser heating for multiple fish (n=5). lHb: p=0.0022, rHb: p=0.0061, lRl: p=0.0098,
lRl: p=0.0036, lVm: p=0.0017, rVm: p=0.0027; *p<0.05, **p<0.01, ***p<0.001 as determined
by one-way t-test.
33
Figure 3.5. The average activity of neuronal components before and after IR laser heating. A)
Representative images of three separate depths throughout the pallium shown with individual
neuronal components outlined. Depths from top to bottom; ~10um, ~30um, ~50um. B)
Individual neuronal components of a representative fish plotted such that each horizontal line
represents the z-score of the average ΔF/F before and after the IR stimulus. Each of the regions
are presented in the order of left habenula (lHb), right habenula (rHb), left rostrolateral (lRl),
right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial (rVm). C) The average
ΔF/F traces of individual neuronal components within each brain region from a representative
fish.
34
3.5 Electric Shock Materials and Methods
Once the fish was prepped for imaging (refer to materials and methods found at the end
of this thesis for a more thorough description of fish preparation) two 1-pin dual-male jumper
wires were secured onto the dive bar such that the pin connectors were positioned on or near
both sides of the fish, where the body meets the tail. They were positioned such that when
current is applied, it travels between the wires creating a circuit with the fish in between, causing
a small electric shock. After 1.5 hrs of acclimation, the fish is imaged for 5 min to establish a
baseline for neuronal activity. Then, while still imaging, an electric shock is applied to the fish
for 400ms before being turned off. This is repeated 2 minutes later for a total of 5 trials. The
experiment is then terminated. Total experiment time is ~15-20min. After the experiment is
finished, the fish is released from the agarose and allowed to rest in a dish for one hour. Only
fish that still alive and healthy after this waiting time are included in the analysis. Furthermore,
only fish that have no z-movement during brain imaging are included in the analysis to preserve
the integrity of these analyses.
For further experimental details, methods, and code, refer to Chapter 7.
3.6 Electric Shock Results
Electric shock activates three distinct areas of the telencephalon.
Figure 3.6 shows the configuration of electrodes used to deliver an electric shock of
500mA (0.5 Amps) through the 5V Arduino pin to the zebrafish for 400ms (Figure 3.6A).
Similar electric shocks were presented to the zebrafish, at 2min intervals, for a total of five times.
The average activation of the brain was measured before and after the stimulus for each of the
six regions: left habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral
35
(rRl), left ventromedial (lVm), and right ventromedial (rVm). All six regions show varying levels
of increased activation following the electric shock, as can be seen in a representative fish
(Figure 3.6B). Coronal and axial views of the representative fish display the six distinct areas of
activation after stimulation from another perspective (Figure 3.7A-B). The change in
fluorescence, ΔF/F, measured before and after stimulation, shows bilateral increases in the Hb,
Rl, and Vm areas of the telencephalon (Figure 3.8A). This increase in fluorescence was found to
be statistically significant across multiple fish (n=4) by comparing the area under the curve
(AUC) of fluorescence intensity before and after electric shock stimulation (4 frames (5.32sec)),
(lHb: p=0.0203, rHb: p=0.0407, lRl: p=0.0045, lRl: p=0.0174, lVm: p=0.0233, rVm: p=0.0199;
*p<0.05, **p<0.01, ***p<0.001 as determined by one-way t-test) (Figure 3.8B). Using CaImAn,
individual neuronal components were detected and extracted from the GCaMP images (Figure
3.9A). The traces that accompany these components were found to exhibit a range of activation
and inhibition. These activity patterns are ordered using z-scores, ranging from the highest
(highly active) to lowest z-scores (inhibited activity) and are displayed using a heat map (Figure
3.9B). In addition to the z-scored map, the ΔF/F traces for these components are shown (Figure
3.9C).
36
Figure 3.6. The average fluorescence response of the zebrafish telencephalon in response to
electric shock. A) Picture of an electric shock being applied to a zebrafish B). The average
response of several brain regions shown from one representative fish to an electric shock. The
brain regions include: the left habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right
rostrolateral (rRl), left ventromedial (lVm), and right ventromedial (rVm). The dotted lines
delineate the brain regions from the rest of the fish, with the habenula circled within the brain,
the rostrolateral delineated from the skin of the fish, and the ventromedial delineated from the
midline of the forebrain. The time between each time point is 1.33 seconds. The black bars
represent the time in which the there was no stimulus and the red bar indicates the time in which
electric shock was administered to the fish.
37
Figure 3.7. Heat maps of the zebrafish telencephalon in response to electric shock. A) Coronal
view of a representative fish emphasizing the change in rostrolateral and ventromedial activity
before and after electric shock. B) Axial view of a representative fish emphasizing the change in
rostrolateral, ventromedial, and habenular activity before and after electric shock. Abbreviations:
left habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left
ventromedial (lVm), and right ventromedial (rVm)
38
Figure 3.8. Average brain region specific activity traces before and after electric shock. A) Brain
region specific ΔF/F traces for the left habenula (lHb), right habenula (rHb), left rostrolateral
(lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial (rVm). B)
Average area under the curve shown for each brain region, 4 frames (5.32sec) before and after
before and after electric shock for multiple fish (n=4). lHb: p=0.0203, rHb: p=0.0407, lRl:
p=0.0045, lRl: p=0.0174, lVm: p=0.0233, rVm: p=0.0199; *p<0.05, **p<0.01, ***p<0.001 as
determined by one-way t-test.
39
Figure 3.9. The average activity of neuronal components before and after electric shock. A)
Representative images of three separate depths throughout the pallium shown with individual
neuronal components outlined. Depths from top to bottom; ~10um, ~30um, ~50um. B)
Individual neuronal components of a representative fish plotted such that each horizontal line
represents the z-score of the average ΔF/F before and after electric shock. Each of the regions
are presented in the order of left habenula (lHb), right habenula (rHb), left rostrolateral (lRl),
right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial (rVm). C) The average
ΔF/F traces of individual neuronal components within each brain region from a representative
fish.
40
3.7 Discussion
The experiments exposing larval zebrafish to infrared heating and electric shock have
revealed that: (1) The rostrolateral, and ventromedial neurons of the pallium, and habenular
neurons respond to both infrared heating, and electric shock, providing evidence that these areas
respond to a variety of noxious stimuli. (2) The rostrolateral neurons increase their activity above
baseline only temporarily following exposure to noxious stimuli and return to baseline within X
seconds (~10seconds). The rostrolateral area was previously found to respond to noxious heat
using pERK staining (Dempsey et al., 2022). In addition, two new areas were found to be highly
active following exposure to noxious stimuli; the habenula and the ventromedial pallium. The
increased activity in the habenula is not surprising given its critical roles in experiencedependent modification of fear response in zebrafish and in the selection of avoidance strategies
(Agetsuma et al., 2010; Amo et al., 2014; Okamoto et al., 2012). The increased activity in the
ventromedial pallium likely occurs as part of a subpallial network that relays information to the
rostrolateral neurons. Although the responses to the two noxious stimuli are qualitatively similar,
the response to electric shock demonstrates a notable delay of ~1.33 seconds in all three areas
compared with the response to heating (Fig.3.2B). This may indicate is possible that the
electrical stimulus is less aversive than the
In conclusion, the experiments in this chapter provide evidence that neurons in the
rostrolateral, ventromedial and habenular regions can respond to noxious stimuli. However, it
remains to be determined whether they respond only to painful stimuli, or whether they respond
to other threatening stimuli, or to neutral sensory stimuli.
41
Chapter 4
Threatening stimuli: exposing larval zebrafish to a full and partial looming
stimulus
4.1 Abstract
In Chapter 3, our investigation into the pallium's response to infrared laser heating not
only validated the results presented in Chapter 2, but also revealed that, alongside the
rostrolateral areas, the habenula and ventromedial regions of the pallium also exhibited
heightened activity. Furthermore, we discovered that the exposing zebrafish to electric shock
demonstrated an almost identical pallial response. However, this does not necessarily indicate
that the observed areas contain negative valence neurons; instead, they may be sensory neurons.
To determine whether these regions, particularly the rostrolateral areas, mediate negative valence
processing, we introduced other types of stimuli that are threatening, but not painful. Research
has demonstrated that zebrafish, like many other animals, are sensitive to looming stimuli, which
mimic the shadow of an approaching predator, prompting them to swim away in response
(Temizer et al., 2015).
Using the Tg(βActin-NRSE::GCaMP6s) zebrafish, positioned on a light sheet
microscope, we exposed zebrafish to a full looming stimulus, which consists of a circular
shadow that grows from 1 degree to 66 degrees (the full size of the projector screen) of the visual
field over 4 seconds concentrically. The final shadow, which is projected for 5 seconds, is large
enough to completely cover the entire zebrafish visual field. We exposed 7 dpf fish to the full
looming stimulus a total of five times while collecting images of GCaMP fluorescence with
which we later measured the average response using ΔF/F. We also performed separate
42
experiments using a partial-looming stimulus, which is identical to the full looming stimulus,
except that the final shadow covers only a fraction of the fish’s visual field (48 degrees) to see if
a looming stimulus that was slightly less threatening than a full looming stimulus would exhibit
similar activation patterns. There was a dramatic bilateral increase in the activity of rostrolateral
neurons, habenula, and ventromedial areas in response to the full looming stimulus. In contrast,
there was a striking absence of rostrolateral activation in the response to the partial looming
stimulus, even though the habenula and ventromedial regions were active. These results provide
evidence that the rostrolateral neurons respond to non-painful threatening stimuli, but not to all
threatening stimuli. The presence of rostrolateral activation in the full looming experiments and
an absence of rostrolateral activation in the partial looming experiments indicates that there may
be a threshold of threat that needs to be achieved before information reaches the rostrolateral
neurons.
4.2 Introduction
A major function of the visual system is to gather important information from the
environment that aid the nervous system in deciding and executing suitable behaviors. Some of
this information can indicate a potential threat to the organism and therefore needs to be
transmitted to the proper areas of the brain to initiate suitable avoidance behaviors. A looming
stimulus, a shadow, often signals a potential threat. For many species, including monkeys (Schiff
et al., 1962), humans (Ball & Tronick, 1971), rodents (Yilmaz & Meister, 2013), birds (Sun &
Frost, 1998), insects (De Vries & Clandinin, 2012), and fish (Temizer et al., 2015), the looming
stimulus triggers evolutionarily hard-wired defensive behaviors like freezing or fleeing.
43
Interestingly, the presence of the looming stimulus alone is not sufficient to prompt
organisms to react. Parameters such as the time to collision and the size on the retina are
important parameters that affect the likelihood the organism will reliably respond to the stimulus
(Temizer et al., 2015). Taking this into account, this chapter will explore whether a full-looming
stimulus and a partial-looming stimulus exhibit the same pattern of activation in the zebrafish
forebrain as do noxious stimuli.
Taking this into account, this chapter will explore whether a full-looming stimulus and a
partial-looming stimulus exhibit the same pattern of activation in the zebrafish forebrain
analogous to its reaction to noxious stimuli. To this end, we will employ two variations of the
looming disc: one that grows to completely obscures light and another that grows partially before
disappearing. We hypothesized that that if both stimuli are perceived as threatening, they will
elicit responses like those triggered by noxious stimuli. However, if both stimuli generate
comparable pallial activity patterns to that of Chapter 3, it could suggest that the implicated areas
are sensory in function, not negative valence. Conversely, if only one stimulus provokes a
reaction, it may be the sole fear-inducing stimulus. Should neither stimulus elicit a response, the
patterns observed in Chapter 3 may not reflect genuine fear but rather pain.
4.3 Full-Looming Stimulus Materials and Methods
Once the fish was prepped for imaging (refer to materials and methods found at the end
of this thesis for a more thorough description of fish preparation) a red-backlit projector screen
was positioned on the right side of the fish such that images and shapes were shown to the fish.
A very small circle (1 degree of the zebrafish visual filed) was maintained throughout the
duration of the experiment and is then at the beginning of each trial, expanded until it fully
44
covered the entirety of the projection screen (1 degree to 66 degrees of the visual field over 4
seconds), hence the name “full-looming stimulus”. After 1.5 hrs of acclimation, the fish was
imaged for 5 min to establish a baseline for neuronal activity (GCaMP fluorescence). Then,
while still imaging, a full-looming circle was expanded to cover the entirety of the projector
within (time to full cover = 500ms) before returning to its original size 5 seconds later. This was
repeated 2 minutes later for a total of 5 trials. The experiment was then terminated. Total
experiment time was ~15-20min. After the experiment was finished, the fish was released from
the agarose and allowed to rest in a dish for one hour. Only fish that were still alive and healthy
after this waiting time were included in the analysis. Furthermore, only fish that had no zmovement during brain imaging were included in the analysis to preserve their integrity.
For further experimental details, methods, and code, refer to Chapter 7.
4.4 Full-Looming Stimulus Results
Full-looming stimulus activated three distinct areas of the telencephalon, like the response
elicited from noxious stimuli.
Figure 4.1A shows a representative image of where a looming disc, starting from a small
circle and growing to cover the visual field, is shown to the zebrafish (Figure 4.1A) Refer to
Figure 3.1 for reference to the anatomical locations chosen for this analysis (Figure 3.1A-C.
After presenting the full-looming stimulus to the zebrafish a total of five times for the duration of
the experiment, 2min intervals, the average activation of the brain was measured before and after
the stimulus for six separate regions; left habenula (lHb), right habenula (rHb), left rostrolateral
(lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial (rVm). All six
regions had varying levels of increased activation following the full looming stimulus, as can be
45
seen in a representative fish (Figure 4.1B). Coronal and axial views of the representative fish
display the six distinct areas of activation after stimulation from another perspective (Figure
4.2A-B). The change in fluorescence, ΔF/F, measured 5 frames (6.65sec) before and after
stimulation, shows drastic, bilateral increases in the Hb, Rl, and Vm areas of the telencephalon
(Figure 4.3A). This increase in fluorescence was found to be statistically significant across
multiple fish (n=5) by comparing the area under the curve (AUC) of fluorescence intensity
before and after the full-looming stimulus (lHb: p=0.0049, rHb: p=0.0141, lRl: p=0.0054, lRl:
p=0.0044, lVm: p=<0.001, rVm: p=0.0011; *p<0.05, **p<0.01, ***p<0.001 as determined by
one-way t-test.) (Figure 4.3B). Using CaImAn, individual neuronal components were detected
were extracted from the GCaMP images (Figure 4.4A). The traces that accompany these
components were found to exhibit a range of activation. These activity patterns are ordered using
z-scores, ranging from the highest (highly active) to lowest z-scores (inhibited activity) and are
displayed using a heat map (Figure 4.4B). In addition to the z-scored map, the ΔF/F traces for
these components are shown (Figure 4.4C).
46
Figure 4.1. The average fluorescence response of the zebrafish telencephalon in response to a full
looming stimulus. A) Picture of a full looming disc being show to a zebrafish from a projector B)
The average brain response of several brain regions shown from representative fish after being
exposed to the full looming disc hit five times. The brain regions include: the left habenula (lHb),
right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and
right ventromedial (rVm). The dotted lines delineate the brain regions from the rest of the fish,
with the habenula circled within the brain, the rostrolateral delineated from the skin of the fish,
and the ventromedial delineated from the midline of the forebrain. The time between each time
point is 1.33 seconds. The black bars represent the time in which the there was nothing on the
projector screen and the red bar indicates the time in which the circle fully expanded and
remained in the FOV of the fish.
47
Figure 4.2. Heat maps of the zebrafish telencephalon in response to being exposed to a fulllooming disc. A) Coronal view of a representative fish emphasizing the change in rostrolateral
and ventromedial activity before and after the full looming disc. B) Axial view of a
representative fish emphasizing the change in rostrolateral, ventromedial, and habenular activity
before and after the full looming disc. Abbreviations: left habenula (lHb), right habenula (rHb),
left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial
(rVm)
48
Figure 4.3. Average brain region specific activity traces before and after being exposed to a fulllooming disc. A) Brain region specific ΔF/F traces for the left habenula (lHb), right habenula
(rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). B) Average area under the curve shown for each brain region, 5 frames
(6.65sec) before and after before and after being exposed to a full-looming disc for multiple fish
(n=5). lHb: p=0.0049, rHb: p=0.0141, lRl: p=0.0054, lRl: p=0.0044, lVm: p=<0.001, rVm:
p=0.0011; *p<0.05, **p<0.01, ***p<0.001 as determined by one-way t-test.
49
Figure 4.4. The average activity of neuronal components before and after being exposed to a fulllooming disc. A) Representative images of three separate depths throughout the pallium shown
with individual neuronal components outlined. Depths from top to bottom; ~10um, ~30um,
~50um. B) Individual neuronal components of a representative fish plotted such that each
horizontal line represents the z-score of the average ΔF/F before and after being exposed to a
full-looming disc. Each of the regions are presented in the order of left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). C) The average ΔF/F traces of individual neuronal components within each
brain region from a representative fish.
50
4.5 Partial-Looming Stimulus Materials and Methods
Once the fish was prepped for imaging (refer to materials and methods found at the end
of this thesis for a more thorough description of fish preparation) the same procedure was
performed as with the full looming stimulus, except that maximum shadow only partially
covered the projection screen, hence the name “partial-looming stimulus”. Here the stimulus
starts at 1 degree and grows to 48 degrees of the visual field over 300ms seconds, then remains
this size for 5 seconds before returning to its original size. The fish was then allowed to
acclimate for 1.5 hours and then imaged for 5 min to establish a baseline for neuronal activity.
Then, while still imaging, a partial-looming stimulus was projected (time to partial cover =
300ms) before returning to its original size 5 seconds later. This was repeated 2 minutes later for
a total of 5 trials. The experiment was then terminated. Total experiment time was ~15-20min.
After the experiment was finished, the fish was released from the agarose and allowed to rest in a
dish for one hour. Only fish that were still alive and healthy after this waiting time were included
in the analysis. Furthermore, only fish that had no z-movement during brain imaging were
included in the analysis.
For further experimental details, methods, and code, refer to Chapter 7.
4.6 Partial-Looming Stimulus Results
Partial-looming stimulus activated only two distinct areas of the telencephalon.
Figure 4.5A shows a representative image of where a looming disc, starting from a small
circle and growing to partially cover the visual field, was shown to the zebrafish (Figure 4.5A).
After presenting the partial-looming stimulus to the zebrafish a total of three times for the
duration of the experiment, 2-3min random intervals, the average activation of the brain was
51
measured before and after the stimulus for six separate regions; left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). Four regions, lHb, rHb, lVm, rVm, showed varying levels of increased
activation following the IR stimulus, while the lRl amd rRl showed no increase in activation, as
can be seen in a representative fish (Figure 4.5B). Coronal and axial views of the representative
fish displayed the six distinct areas of activation after stimulation from another perspective
(Figure 4.6A-B). The change in fluorescence, ΔF/F, measured 12 frames (~16sec) before and
after stimulation, shows, bilateral increases in the Hb and Vm areas of the telencephalon, but not
the Rl areas (Figure 4.7A). The increase in the fluorescence of Hb and Vm areas was found to be
statistically significant across multiple fish (n=7), while no statistically significant increase was
found for the Rl areas. This was determined by comparing the area under the curve (AUC) of
fluorescence intensity before and after the partial-looming stimulus (lHb: p=0.0026, rHb:
p=0.0179, lRl: p=0.6696, lRl: p=0.5356, lVm: p=<0.001, rVm: p=<0.001; *p<0.05, **p<0.01,
***p<0.001 as determined by one-way t-test. (Figure 4.7B). Using CaImAn, individual neuronal
components were extracted from the GCaMP images (Figure 4.8A). The traces that accompany
these components were found to exhibit a range of activation. These activity patterns are ordered
using z-scores, ranging from the highest (highly active) to lowest z-scores (inhibited activity) and
are displayed using a heat map (Figure 4.8B). In addition to the z-scored map, the ΔF/F traces
for these components are shown (Figure 4.8C).
52
Figure 4.5. The average fluorescence response of the zebrafish telencephalon in response to a
partial looming stimulus. A) Picture of a partial looming disc being show to a zebrafish from a
projector. B) The average brain response of several brain regions shown from representative fish
after being exposed to the partial looming disc hit five times. The brain regions include: the left
habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left
ventromedial (lVm), and right ventromedial (rVm). The dotted lines delineate the brain regions
from the rest of the fish, with the habenula circled within the brain, the rostrolateral delineated
from the skin of the fish, and the ventromedial delineated from the midline of the forebrain. The
time between each time point is 1.33 seconds. The black bars represent the time in which the
there was nothing on the projector screen and the red bar indicates the time in which the circle
expanded and remained in the FOV of the fish.
53
Figure 4.6. Heat maps of the zebrafish telencephalon in response to being exposed to a partiallooming disc. A) Coronal view of a representative fish emphasizing the change in rostrolateral
and ventromedial activity before and after the partial looming disc. B) Axial view of a
representative fish emphasizing the change in rostrolateral, ventromedial, and habenular activity
before and after the partial looming disc. Abbreviations: left habenula (lHb), right habenula
(rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm)
54
Figure 4.7. Average brain region specific activity traces before and after being exposed to a
partial-looming disc. A) Brain region specific ΔF/F traces for the left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). B) Average area under the curve shown for each brain region, 12 frames
(~16sec) before and after being exposed to a partial-looming disc for multiple fish (n=7). lHb:
p=0.0026, rHb: p=0.0179, lRl: p=0.6696, lRl: p=0.5356, lVm: p=<0.001, rVm: p=<0.001;
*p<0.05, **p<0.01, ***p<0.001 as determined by one-way t-test.
55
Figure 4.8. The average activity of neuronal components before and after being exposed to a
partial-looming disc. A) Representative images of three separate depths throughout the pallium
shown with individual neuronal components outlined. Depths from top to bottom; ~10um,
~30um, ~50um. B) Individual neuronal components of a representative fish plotted such that
each horizontal line represents the z-score of the average ΔF/F before and after being exposed to
a partial-looming disc. Each of the regions are presented in the order of left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). C) The average ΔF/F traces of individual neuronal components within each
brain region from a representative fish.
56
4.7 Discussion
The experiments done to expose larval zebrafish to a full-looming stimulus and a partiallooming stimulus have revealed several key findings: (1) The rostrolateral neurons of the pallium
robustly respond to a full-looming stimulus demonstrating that the neurons do not solely respond
to nociception. (2) The rostrolateral neurons do not activate in response to a partial-looming
stimulus demonstrating that the neurons do not respond all forms of stimuli. (3) Both forms of
looming stimuli exhibited activation of the habenula and ventromedial regions of the pallium.
It’s possible that an increase in the ventromedial areas of the pallium reflect one of two
scenarios: activation of excitatory networks, which leads to activation of the rostrolateral areas,
or activation of inhibitory neurons in the ventromedial areas, which suppresses activation of the
rostrolateral areas. Without more information about the neuronal makeup of this area, both
hypotheses are feasible. (4) The habenula is still active even if the rostrolateral neurons are not
active. This can be explained by the fact that light alone can activate neurons in the habenula
(Zhang et al., 2017).
In the next chapter, I will discuss how we extended our investigation to include other
forms of sensory information that are less threatening to test the hypothesis that the rostrolateral
neurons of the pallium are negative valence but have a high threshold for activation.
57
Chapter 5
Benign stimuli: Exposing larval zebrafish to physical vibration, light, and
sound
5.1 Abstract
In Chapter 3, we subjected larval zebrafish to noxious stimuli to identify responsive
structures within the pallium. Our investigations revealed bilateral activation in three regions,
notably including the rostrolateral neurons, which we hypothesize harbor negative valence
neurons. In Chapter 4, we introduced non-noxious but potentially threatening stimuli to discern
whether the rostrolateral neurons functioned as sensory rather than negative valence neurons. We
observed that exposure to a full-looming stimulus elicited a response remarkably similar to that
of the noxious stimuli. Intriguingly, exposure to a partial looming stimulus activated the
habenula and ventromedial regions without engaging the rostrolateral neurons. This finding not
only suggests that the rostrolateral region does not serve as a sensory processing center but also
indicates that various non-noxious, yet potentially life-threatening stimuli may activate this
region only when a certain threshold is surpassed.
Using the Tg(βActin-NRSE::GCaMP6s) zebrafish, we exposed the fish to three types of
stimuli; vibration, light, and sound. We exposed 7 dpf fish to these stimuli a total of five times
while collecting images of GCaMP fluorescence with which we later measured the average
response using ΔF/F. Vibration from tapping elicited a small increase in the activity of the right
habenula, alongside a larger increase in both ventromedial areas. Light also elicited a small
increase in the activity of the right habenula alongside a larger, increase in both ventromedial
areas. In contrast, in addition to a small increase in the activity of the right habenula and both
58
ventromedial areas, sound from and airhorn elicited activation of the left habenula. The most
striking result that came out of these experiments was a complete absence of rostrolateral
activation in response to all three forms of these benign stimuli. These data provide evidence that
the rostrolateral neurons do not respond to non-threatening stimuli. Like the results in Chapter 4,
these data provide further evidence that there is likely a threshold of threat that needs to be
achieved before information reaches the rostrolateral neurons.
5.2 Introduction
In this chapter, our goal is to explore the pallium's response to additional non-harmful,
yet potentially life-threatening stimuli. It is reasonable to hypothesize that vibrations, light, and
sound, while not inherently painful or evolutionarily ingrained threats like a looming shadow,
might still stimulate the rostrolateral regions if perceived as sufficiently threatening.
Alternatively, a limited reaction or complete absence of response to these stimuli could indicate a
high activation threshold for the rostrolateral region. In this chapter we will thoroughly
investigate the zebrafish pallial response to these three types of stimuli, aiming to unravel the
complexities of pallial activation in response to perceived threats.
5.3 Tapping Stimulus Materials and Methods
Once the fish was prepped for imaging (refer to materials and methods found at the end
of this thesis for a more thorough description of fish preparation) the fish is allowed to acclimate
for 1.5 hours and then imaged for 5 min to establish a baseline for neuronal activity. Then, while
still imaging, the experimenter tapped with a hand on the side of the chamber until a bout of tail
twitches was observed. This was repeated 2 minutes later for a total of 5 trials. The experiment
59
was then terminated. Total experiment time was ~15-20min. After the experiment was finished,
the fish was released from the agarose and allowed to rest in a dish for one hour. Only fish that
were still alive and healthy after this waiting time and that had no z-movement during brain
imaging were included in the analysis.
For further experimental details, methods, and code, refer to Chapter 7.
5.4 Tapping Stimulus Results
Tapping stimulus activated some telencephalic areas, but to a lesser extent than the looming disc.
Figure 5.1A shows a representative image of a zebrafish feeling the tapping on the side of
the imaging chamber. After the zebrafish was exposed to the tapping event a total of five times
for the duration of the experiment, 2min intertrial separation, the average activation of the brain
was measured before and after the stimulus for six separate regions; left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). Three regions, rHb, lVm, and rVm, showed varying levels of increased
activation following the IR stimulus, while the lHb, lRl amd rRl showed no increase in
activation, as can be seen in a representative fish (Figure 5.1B). Coronal and axial views of the
representative fish displayed the six distinct areas of activation after stimulation from another
perspective (Figure 5.2A-B). The change in fluorescence, ΔF/F, measured 7 frames (9.31sec)
before and after stimulation, showed weak activation of the rHb, no statistically significant
activation of the lHb, lRl or the rRl, and a statistically significant increase in the activation of the
lVm, and rVm (n=7) (Figure 5.3A). This was determined by comparing the area under the curve
(AUC) of fluorescence intensity before and after five tapping events (lHb: p=0.2302, rHb:
p=0.0416, lRl: p=0.9297, lRl: p=0.4581, lVm: p=0.0131, rVm: p=0.0072; *p<0.05, **p<0.01,
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***p<0.001 as determined by one-way t-test.) (Figure 5.3B). Using CaImAn, individual
neuronal components were extracted from the GCaMP images (Figure 5.4A). The traces that
accompany these components were found to exhibit a range of activation. These activity patterns
are ordered using z-scores, ranging from the highest (highly active) to lowest z-scores (inhibited
activity) and are displayed using a heat map (Figure 5.4B). In addition to the z-scored map, the
ΔF/F traces for these components are shown (Figure 5.4C).
61
Figure 5.1. The average fluorescence response of the zebrafish telencephalon in response to
vibration from tapping. A) Picture of a partial looming disc being show to a zebrafish from a
projector B) The average brain response of several brain regions shown from representative fish
after being exposed vibration from tapping five times. The brain regions include: the left
habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left
ventromedial (lVm), and right ventromedial (rVm). The dotted lines delineate the brain regions
from the rest of the fish, with the habenula circled within the brain, the rostrolateral delineated
from the skin of the fish, and the ventromedial delineated from the midline of the forebrain. The
time between each time point is 1.33 seconds. The black bars represent the time in which there
was no tapping and the red bar indicates the time in which the side of chamber was repeatedly
tapped.
62
Figure 5.2. Heat maps of the zebrafish telencephalon in response to being exposed to vibration
from tapping. A) Coronal view of a representative fish emphasizing the change in rostrolateral
and ventromedial activity before and after vibration from tapping. B) Axial view of a
representative fish emphasizing the change in rostrolateral, ventromedial, and habenular activity
before and after vibration from tapping. Abbreviations: left habenula (lHb), right habenula (rHb),
left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial
(rVm)
63
Figure 5.3. Average brain region specific activity traces before and after being exposed to
vibration from tapping. A) Brain region specific ΔF/F traces for the left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). B) Average area under the curve shown for each brain region, 7 frames
(9.31sec) before and after being exposed to vibration from tapping for multiple fish (n=7). lHb:
p=0.2302, rHb: p=0.0416, lRl: p=0.9297, lRl: p=0.4581, lVm: p=0.0131, rVm: p=0.0072;
*p<0.05, **p<0.01, ***p<0.001 as determined by one-way t-test.
64
Figure 5.4. The average activity of neuronal components before and after being exposed to
vibration from tapping. A) Representative images of three separate depths throughout the
pallium shown with individual neuronal components outlined. Depths from top to bottom;
~10um, ~30um, ~50um. B) Individual neuronal components of a representative fish plotted such
that each horizontal line represents the z-score of the average ΔF/F before and after being
exposed to vibration from tapping. Each of the regions are presented in the order of left habenula
(lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial
(lVm), and right ventromedial (rVm). C) The average ΔF/F traces of individual neuronal
components within each brain region from a representative fish.
65
5.5 Light Stimulus Materials and Methods
Once the fish was prepped for imaging (refer to materials and methods found at the end
of this thesis for a more thorough description of fish preparation) a red-backlit projector screen
was positioned on the right side of the fish such that light was visible to the fish. Then, a small
removable cardboard card was placed between the fish and the projector such that the light no
longer reached the fish, keeping the fish in complete darkness. The fish was allowed to acclimate
for 1.5 hours and then imaged for 5 min to establish a baseline for neuronal activity. Then, while
still imaging, the cardboard card was removed for 5 seconds to bathe the fish in light. After 5
seconds, the card was placed back in between the fish and projector to cover the light source.
This was repeated 2 minutes later for a total of 5 trials. The experiment was then terminated.
Total experiment time was ~15-20min. After the experiment was finished, the fish was released
from the agarose and allowed to rest in a dish for one hour. Only fish that were still alive and
healthy after this waiting time were included in the analysis. Furthermore, only fish with no zmovement during brain imaging were included in the analysis to preserve the integrity of these
analyses.
For further experimental details, methods, and code, refer to Chapter 7.
5.6 Light Stimulus Results
Light stimulus activates some telencephalic areas, but only after light turns off.
Figure 5.5A shows a representative image of a zebrafish being exposed to a light turning
on and then off. After the zebrafish was exposed to the light a total five times for the duration of
the experiment, 2min intertrial separation, the average activation of the brain was taken before
and after the stimulus for six separate regions; left habenula (lHb), right habenula (rHb), left
66
rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial
(rVm). Three regions, rHb, lVm, and rVm, showed varying levels of increased activation
following the light stimulus, while the lHb, lRl and rRl showed no increase in activation, as can
be seen in a representative fish (Figure 5.5B). Coronal and axial views of the representative fish
display the six distinct areas of activation after stimulation from another perspective (Figure
5.6A-B). The change in fluorescence, ΔF/F, measured 14 frames (18.62sec) before and after
stimulation, shows weak activation of the lHb, no statistically significant activation of the rHb,
lRl or the rRl, and a statistically significant increase in the activation of the lVm, and rVm (n=7)
(Figure 5.7A). This was determined by comparing the area under the curve (AUC) of
fluorescence intensity before and after five tapping events (lHb: p=0.0538, rHb: p=0.0249, lRl:
p=0.1348, lRl: p=0.1268, lVm: p=0.01, rVm: p=0.0376; *p<0.05, **p<0.01, ***p<0.001 as
determined by one-way t-test) (Figure 5.7B). Using CaImAn, individual neuronal components
were extracted from the GCaMP images (Figure 5.8A). The traces that accompany these
components were found to exhibit a range of activation. These activity patterns are ordered using
z-scores, ranging from the highest (highly active) to lowest z-scores (inhibited activity) and are
displayed using a heat map (Figure 5.8B). In addition to the z-scored map, the ΔF/F traces for
these components are shown (Figure 5.8C).
67
Figure 5.5. The fluorescence response of the zebrafish telencephalon in response to a light
turning on and off. A) Picture of a zebrafish being exposed a light turning on and off. B) The
average brain response of several brain regions shown from representative fish after in response
to a light turning on and off, for five trials. The brain regions include: the left habenula (lHb),
right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and
right ventromedial (rVm). The dotted lines delineate the brain regions from the rest of the fish,
with the habenula circled within the brain, the rostrolateral delineated from the skin of the fish,
and the ventromedial delineated from the midline of the forebrain. The time between each time
point is 1.33 seconds. The black bars represent the time in which the light was off and the red bar
indicates the time in which the light was on.
68
Figure 5.6. Heat maps of the zebrafish telencephalon in response to a light turning on and off A)
Coronal view of a representative fish emphasizing the change in rostrolateral and ventromedial
activity before and after a light turning on and off. B) Axial view of a representative fish
emphasizing the change in rostrolateral, ventromedial, and habenular activity before and after in
a light turning on and off. Abbreviations: left habenula (lHb), right habenula (rHb), left
rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial
(rVm)
69
Figure 5.7. Average brain region specific activity traces before and after being exposed to a light
turning on and off. A) Brain region specific ΔF/F traces for the left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). B) Average area under the curve shown for each brain region, 14 frames
(18.62sec) before and after being exposed to a light turning on and off for multiple fish (n=7).
lHb: p=0.0538, rHb: p=0.0249, lRl: p=0.1348, lRl: p=0.1268, lVm: p=0.01, rVm: p=0.0376;
*p<0.05, **p<0.01, ***p<0.001 as determined by one-way t-test.
70
Figure 5.8. The average activity of neuronal components before and after being exposed to a
light turning on and off. A) Representative images of three separate depths throughout the
pallium shown with individual neuronal components outlined. Depths from top to bottom;
~10um, ~30um, ~50um. B) Individual neuronal components of a representative fish plotted such
that each horizontal line represents the z-score of the average ΔF/F before and after being
exposed to a light turning on and off (Light on and off indicated by the red bar and black faded
region). Each of the regions are presented in the order of left habenula (lHb), right habenula
(rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). C) The average ΔF/F traces of individual neuronal components within each
brain region from a representative fish.
71
5.7 Sound Stimulus Materials and Methods
Once the fish was prepped for imaging (refer to materials and methods found at the end
of this thesis for a more thorough description of fish preparation), it was allowed to acclimate for
1.5 hours. and then imaged for 5 min to establish a baseline for neuronal activity. Then, while
still imaging, an air horn (1.4oz, Better Boat) was placed above the chamber and sounded. The
experimenter wore ear protection so as not to go deaf in this scientific pursuit. However, for one
trial the experimenter forgot he wasn’t wearing any ear protection and proceeded to deafen
himself for a small duration of time. This was repeated 2 minutes later for a total of 5 trials. The
experiment was then terminated. Total experiment time was ~15-20min. After the experiment
finished, the fish was released from the agarose and allowed to rest in a dish for one hour. Only
fish that were still alive and healthy after this waiting time were included in the analysis.
Furthermore, only fish that had no z-movement during brain imaging were included in the
analysis to preserve the integrity of these analyses.
For further experimental details, methods, and code, refer to Chapter 7.
5.8 Sound Stimulus Results
Air horn stimulus activates the habenula and deep medial areas of the telencephalon, but not the
rostrolateral areas.
Figure 5.9A shows a representative image of a zebrafish being exposed to an air horn
(Figure 5.9A). After the zebrafish was exposed to the airhorn a total five times for the duration of
the experiment, 2min intertrial separation, the average activation of the brain was taken before
and after the stimulus for six separate regions; left habenula (lHb), right habenula (rHb), left
rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right ventromedial
72
(rVm). Four regions, lHb, rHb, lVm, and rVm, showed varying levels of increased activation
following the light stimulus, while the lRl amd rRl show no increase in activation, as can be seen
in a representative fish (Figure 5.9B). Coronal and axial views of the representative fish display
the six distinct areas of activation after stimulation from another perspective (Figure 5.10A-B).
The change in fluorescence, ΔF/F, measured 13 frames (17.29sec) before and after stimulation,
shows activation of the bilateral habenula and ventromedial areas, with no activation of the lRl or
the rRl (n=4) (Figure 5.11A). This was determined by comparing the area under the curve (AUC)
of fluorescence intensity before and after five tapping events (lHb: p=0.0096, rHb: p=0.022, lRl:
p=0.7489, lRl: p=0.53, lVm: p=0.0394, rVm: p=0.0079; *p<0.05, **p<0.01, ***p<0.001 as
determined by one-way t-test. (Figure 5.11B). Using CaImAn, individual neuronal components
were extracted from the GCaMP images (Figure 5.12A). The traces that accompany these
components were found to exhibit a range of activation. These activity patterns are ordered using
z-scores, ranging from the highest (highly active) to lowest z-scores (inhibited activity) and are
displayed using a heat map (Figure 5.12B). In addition to the z-scored map, the ΔF/F traces for
these components are shown (Figure 5.12C).
73
Figure 5.9. The average fluorescence response of the zebrafish telencephalon in response to the
sound from an airhorn. A) Picture of a partial looming disc being show to a zebrafish from a
projector B) The average brain response of several brain regions shown from representative fish
after being exposed to the sound from an airhorn. The brain regions include: the left habenula
(lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial
(lVm), and right ventromedial (rVm). The dotted lines delineate the brain regions from the rest
of the fish, with the habenula circled within the brain, the rostrolateral delineated from the skin
of the fish, and the ventromedial delineated from the midline of the forebrain. The time between
each time point is 1.33 seconds. The black bars represent the time in which there was no sound
was and the red bar indicates the time in which the airhorn was sounded.
74
Figure 5.10. Heat maps of the zebrafish telencephalon in response to the sound from an airhorn.
A) Coronal view of a representative fish emphasizing the change in rostrolateral and
ventromedial activity before and after being exposed to the sound from an airhorn. B) Axial view
of a representative fish emphasizing the change in rostrolateral, ventromedial, and habenular
activity before and after being exposed to the sound from an airhorn. Abbreviations: left
habenula (lHb), right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left
ventromedial (lVm), and right ventromedial (rVm).
75
Figure 5.11. Average brain region specific activity traces before and after being exposed to the
sound from an airhorn. A) Brain region specific ΔF/F traces for the left habenula (lHb), right
habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and right
ventromedial (rVm). B) Average area under the curve shown for each brain region, 13 frames
(17.29sec) before and after being exposed to a light turning on and off for multiple fish (n=7).
lHb: p=0.0096, rHb: p=0.022, lRl: p=0.7489, lRl: p=0.53, lVm: p=0.0394, rVm: p=0.0079;
*p<0.05, **p<0.01, ***p<0.001 as determined by one-way t-test.
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Figure 5.12. The average activity of neuronal components before and after being exposed to the
sound from an airhorn. A) Representative images of three separate depths throughout the pallium
shown with individual neuronal components outlined. Depths from top to bottom; ~10um,
~30um, ~50um. B) Individual neuronal components of a representative fish plotted such that
each horizontal line represents the z-score of the average ΔF/F before and after being exposed to
the sound from an airhorn. Each of the regions are presented in the order of left habenula (lHb),
right habenula (rHb), left rostrolateral (lRl), right rostrolateral (rRl), left ventromedial (lVm), and
right ventromedial (rVm). C) The average ΔF/F traces of individual neuronal components within
each brain region from a representative fish.
77
5.9 Discussion
The experiments done to expose larval zebrafish to vibration, light, and sound have
revealed several key findings: (1) The rostrolateral neurons of the pallium do not respond to any
of the stimuli. These data provide further evidence that the rostrolateral neurons possess a high
activation threshold. (2) The light stimulus elicits a distinct pattern of neural activation in the
habenular ventromedial regions after the light turns off, but not when it turns on. This indicates
that ventrolateral and habenular areas possess the ability to distinguish specific aspects of stimuli
and likely belong to higher order networks within the brain. What is interesting about this
observation is that in Chapter 4, looming stimuli also decreases the overall luminance presented
to the zebrafish. The light stimuli experiments are similar in that by turning the light off, this also
decreases overall luminance. Both situations cause a striking increase in activity in the
ventromedial areas. Furthermore, while the fish is resting in the dark, this area is not active.
These observations suggest that the ventromedial area activates when there is a “decrease” in
luminence, not when there is an increase in luminance or persistent darkness. This may partially
be explained by the presence of dark photokinetic receptors in this area (Temizer et al., 2015).
However, the presence of these neurons doesn’t fully explain ventromedial activation since we
have shown that both forms IR heating and electric shock also elicited activation in this region.
(3) Tapping and light show statistically significant increases in activation of only the right
habenula. This is interesting because both stimuli are presented to the right side of the fish,
suggesting that information is sensory information is either being processed ipsilaterally once
received or is part of the upstream processing that occurs after initial sensory recognition,
perhaps part of the processing networks behavioral selection.
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Chapter 6
Elucidating the identity of the rostrolateral neurons in the pallium using
Mapzebrain
6.1 Abstract
In Chapter 2, I discussed how our previous work revealed that the rostrolateral pallium
that was highly active when a zebrafish was exposed to infrared laser heating and when it was
exposed to a conditioned stimulus after associative conditioning (Dempsey et al., 2022). In
Chapter 3, I extended the investigation to look at real-time activity changes of the zebrafish
pallium and uncovered the temporal dynamics of the rostrolateral neurons in response to noxious
stimuli. In Chapter 4, we discovered that the rostrolateral area only responded to one form of
looming stimuli, demonstrating that the area is not merely sensory processing and likely contains
negative valence neurons. In Chapter 5, we revealed that three more forms of stimuli, potentially
threatening but mostly benign, elicited no activity from the rostrolateral pallium, suggesting that
a certain threshold of threat must exist before eliciting activation from the rostrolateral pallium.
In this chapter, using the Mapzebrain Atlas we discovered that the transgenic Tg[Tiam2a:GFP]
zebrafish possesed GFP positive neurons in the rostrolateral pallium. To assess whether these
were the same neurons as the putative negative valence neurons discussed in previous chapters,
we exposed the Tg[Tiam2a:GFP] to infrared laser heating. We discovered that the GFP+ neurons
of fish that underwent IR laser heating heavily colocalized with pERK activity. These data
demonstrate that Tiam2a is a marker for negative valence neurons in zebrafish.
79
6.2 Introduction
Uncovering the identity of the negative valence rostrolateral neurons is difficult without
more advanced methods such as single-cell RNA sequencing. This technique would allow us to
isolate the cells, assess which genes are uniquely or highly expressed, and gain a better
understanding of what these cell types are, which would complement our data showing they are
negative valence. Unfortunately, the number of cells expressed in this region is limited, making
it very difficult to perform this type of analysis. Instead, we decided to take a different approach
to elucidate the identity of these cells. By using Mapzebrain, we hoped that among the hundreds
of expression maps for the larval zebrafish that we would find at least one map that labelled the
rostrolateral neurons. Furthermore, if such a map existed, there may also be a transgenic fish that
the map was based on. If we had that fish in our possession, we would be able to expose those
fish to noxious stimuli and see if the cells become active. With this, we would not only have a
marker for these neurons, but also have a way of identifying the anatomical location of these
neurons in adults. Knowing the location of these neurons in adult zebrafish would open up the
possibility of studying these neurons and their contribution to emotional learning at a later stage
in development.
6.3 Materials and Methods
6.3.1 Mapzebrain
To decipher how the zebrafish brain functions, it's essential to have a comprehensive
neuroanatomical map of the neurons involved. To address this need, researchers at the Max
Planck Institute, using the 6 days post-fertilization zebrafish brain, created an atlas of the brain
(Kunst et al., 2019). The atlas has been organized into cellular morphotypes, that quantitatively
80
characterizes neuronal shapes and their precise spatial positions in vivo. Additionally, more than
100 distinct transgene expression patterns have been integrated into this atlas, and that number
continues to grow as researchers around the world contribute with their own transgenic zebrafish
lines. With over 72 distinct brain areas identified, researchers can use the atlas to further their
own research endeavors into distinct brain region functioning and morphology (Kunst et al.,
2019).
Given the paucity of neuroanatomical information on the dpf7 zebrafish brain forebrain, I
used Mapzebrain to investigate our putative negative valence neurons within the rostrolateral
pallium. After painstakingly inspecting every single transgene map (723 maps), I found three
maps of interest, with only one available for purchase from the Zebrafish International Resource
Center (ZIRC). The map transgenic fish y264Et, with the construct Et(SCP1:GAL4FF), that
uniquely labelled Tiam2a positive cells within the rostrolateral pallium (Figure 6.1). I contacted
the vendor, had them fertilize WT eggs with the transgenic sperm, and raised the embryos until
they were adult breeding status.
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Figure 6.1. Tiam2a Expression Map on Mapzebrain Atlas. Here is a representative image of the
mapzebrain atlas. Using this atlas, zebrafish researchers can search through over 700 expression
pattern maps. This image shows in purple the y264Et[y264Tg] (Tg[Tiam2a:GFP]) expression
pattern map views of dorsal (left), transverse (top right), and coronal (bottom right).
82
6.3.2 Tiam2a IR Laser Training
Since the contents of this thesis are a continuation of the other work published in
this lab, this section has been adapted from Dempsey et al, 2022.
To begin, Tg[Tiam2a:GFP] zebrafish larvae were embedded in custom behavioral
training molds: a 60 mm culture dish filled with of 1% agarose dissolved in egg water, with a
10mm diameter circle of it is cut out from the center. The zebrafish was anesthetized and then
embedded in the 10mm diameter circle with 500 ml agarose + anesthetic (70 ppm MS-222 + 70
ppm Isoflurane). While the agarose solidified, the zebrafish’s position was parallel to the surface
of the chamber, at the top of the agarose surface, and was maintained using a single bristle brush.
After 5 min, 10 mL of sterilized husbandry system water was added to the chamber. Using fine
forceps, the tail of the zebrafish was carefully freed from the agarose. All free-floating agarose
was removed from the dish. The dish was then positioned under the objective of a custom-built
behavioral microscope (Dempsey et al., 2022) such that the left eye of the zebrafish was directly
under where the infrared laser spot would be when turned on (NIR RLCO-980-1000-F laser,
Roithner Lasertechnik).
An Arduino was programmed to deliver the infrared heating laser for two seconds, a total
of ten times, with an intertrial separation of 2 minutes. After the fish underwent ten trials, the fish
was then fixed in PFA for pERK immunohistochemistry.
6.3.3 Tiam2a pERK Immunohistochemisty
Since the contents of this thesis are a continuation of the other work published in
this lab, this section has been adapted from Dempsey et al, 2022.
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After exposing 7 dpf Tg[Tiam2a:GFP] larval zebrafish to the infrared laser ten times, zebrafish
were fixed in 4% PFA diluted in 1x PBS (pH 7.4), 70 ppm MS-222, 70 ppm Isoflurane, covered
in aluminum foil and kept at 4°C overnight. Then, fish were washed 3 times in 1x PBS (pH7.4)
and 0.25% PBST. After 3x PBST washes, the zebrafish were placed in 150 mM Tris HCl (pH 9)
for 5 min at room temperature, followed by 20 minutes at 65°C. Once this is complete, the
samples are cooled to room temperature and given 3x PBST washes. Zebrafish were then
permeabilized on ice for 45 minutes with 1x PBS, 0.05% Trypsin-EDTA diluted in 1x PBS (pH
7.4). After another 3x PBST wash cycle, the zebrafish were place for 1 hour at room temperature
in blocking buffer solution; (1x PBS, 1% bovine serum albumin, 2% DMSO, 2% NGS, 0.25%
Triton X-100). At a 1:300 dilution, zebrafish were stained with primary rabbit antiphosphorylated Erk1/2 antibodies (4696, Cell Signaling) in blocking buffer on a rotating
platform at 4°C overnight. After 5 PBST washes, the zebrafish were moved to blocking buffer
solution containing a 1:1000 dilution of Alexa Fluor 568-conjugated goat anti-rabbit secondary
antibodies and left overnight at 4°C on an orbital shaker in the dark. After 5 PBST washes,
samples were moved to PBS solution and imaged on a Zeiss LSM 780 confocal.
6.4 Results
Infrared laser heating results in increased pERK expression in Tiam2a positive neurons in the
rostrolateral pallium.
7 dpf Tg[Tiam2a:GFP] larval zebrafish were fixed and stained for pERK using
wholemount immunocytochemistry (Figure 6.2A, B) . A projection image of GFP and pERK
shows a striking lack of colocalization between the two markers showing that under control
conditions, there is little to no activation of neurons expressing Tiam2a. In contrast, when similar
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immunostaining was done to fish following exposure to heating with an IR laser, there was
dramatic overlap, with virtually every Tiam2a-postive neuron expressing pERK (Figure 6.2C, D)
To quantify this, a comparison was made between the fluorescence intensity of the pERK cells in
the Tiam2a cells with that of a background region within the fish. The ratio of pERK/background
was significantly higher in the IR Laser exposed group than the control group (p<0.001, one-way
t-test) (Figure 6.2E). These data demonstrate that the Tiam2a neurons are more active when
zebrafish are exposed to noxious stimuli.
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Figure 6.2. pERK Expression in Tg[Tiam2a:GFP] zebrafish pallium following Infrared Laser
Exposure. A) Representative image of the Tg[Tiam2a:GFP] control fish shown. The control fish
was not exposed to the infrared heating laser. B) Zoomed in image of control fish showing very
little colocalization of pERK expression with Tiam2a GFP+ neurons. C) Representative image of
Tg[Tiam2a:GFP] experimental fish shown. The experimental fish was exposed to the infrared
heating laser ten times. D) Zoomed in image of experimental fish showing high colocalization of
pERK expression with Tiam2a GFP+ neurons. E) Intensities ratios of pERK expressing Tiam2a
GFP+ neurons vs. background fluorescence. IR exposed fish (n=6) show a statistically
significant increase in pERK expression vs control fish (n=8) (p-value = <0.001; *p<0.05,
**p<0.01, ***p<0.001 as determined by one-way t-test).
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6.5 Discussion
Since the rostrolateral cells and the GFP+ cells of the Tg[Tiam2a:GFP] are in the same
anatomical location and both respond to noxious stimuli, we propose that Tiam2a is a genetic
marker for negative valence cells in the rostrolateral pallium. This discovery will provide
neuroscience researchers with a new way to understand the functioning of these cells for two
reason; 1) the cells are clustered together compare to the heterogenous distribution of cells in the
mammalian amygdala (Beyeler et al., 2018b) and 2) the zebrafish is much easier to work given
its size and morphological differences to that of the mammal. The Tg[Tiam2a:GFP] also
provides us with the opportunity to see where these cells reside in adults since we need to just
take sections later in development. This will not only allow researchers to find the cells but also
to possibly validate some of the theories on pallial development that are currently being
discussed in the field.
After looking through The Allen Brain Atlas we found that the Tiam2 gene is highly
expressed in the granule cell layer of the hippocampal dentate gyrus (not shown). This was a
surprisingly discovery, since we had been operating under the hypothesis that the rostro lateral
area was amygdalar in nature, given the presence of negative valence neurons. Perhaps this
assumption was ignorant to the possibility of negative valence neurons in other structures of the
brain. In fact, recent work has shown that the hippocampus segregates negative and positive
engrams (Shpokayte et al., 2022). If the rostrolateral area of the zebrafish is hippocampal, then
this may explain why we saw such a drastic rearrangement of synapses in this area after aversive
conditioning.
Regardless of what the rostrolateral area of the larval zebrafish pallium is homologous to
in mammals, amygdala or hippocampus, our work demonstrates the existence of clearly
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demarcated and clustered negative valence neurons. With this information, it is my hope that the
zebrafish field will continue to investigate this area and unravel the mechanisms with which
these cells contribute to emotional processing. By studying these cells, it may be possible for
researchers to uncover the underlying mechanisms of valence processing and further contribute
to our understanding of the nervous system and how it decodes the world around it.
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Chapter 7
Materials and Methods for Chapters 3-5
7.1 Zebrafish husbandry and embryo/larval care
Since the contents of this thesis are a continuation of the other work published in my lab,
this section is copied from Dempsey et al, 2022.
Larval and adult Casper mutant zebrafish were maintained in an in-house veterinary
facility in accordance with protocols approved by the University of Southern California (USC)
Institutional Animal Care and Use Committee (IACUC). All experiments described in this work
were approved by the USC IACUC. Embryos were kept in egg water (1.19 g NaCl, 0.377 g
CaSO4·2H2O, 265 μL methylene blue, filled up to 5 liters with ddH2O) at 28 °C in an incubator
with a 12 hr:11 hr light:dark cycle until 5-6 dpf, when they were transferred to tanks within the
husbandry facility. On the husbandry system, zebrafish were maintained on a 12 hr:12 hr
light:dark cycle. The air and water temperature in the zebrafish facility are maintained at 26-
28°C. Zebrafish larvae used in experiments (7-9 dpf) were transferred between the facility and
the experimental apparatus in zebrafish husbandry system water (HSW) and kept at 26-28°C
between experiments.
7.2 Mounting Zebrafish in Caddie for Imaging on FlexSPIM
Since the contents of this thesis are a continuation of the other work published in this lab,
this section has been adapted from Dempsey et al, 2022.
To prepare for behavioral experiments, zebrafish subjects are first embedded in a custom
behavioral training caddie generated by Protolabs. Specs: ABS-Like Translucent Clear
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(WaterShed), Normal Res, Natural, Stereolithography, X: 0.980in Y: 0.206in Z: 0.311in (Figure
7.1). The caddie is filled with 200uL of 1% (w/v) agarose (melting point ≤ 65°C, Millipore
Sigma) dissolved in egg water containing methylene blue, which suppresses fungal growth.
During agarose embedding and fluorescence imaging experiments, zebrafish were anesthetized
in a final concentration of 70 ppm MS-222 (Western Chemical) + 70 ppm Isoflurane (Phoenix),
an IACUC-approved combination that has been shown to enable fast recovery from anesthesia as
compared to using MS-222 alone (Huang WC, et al., 2010). The anesthetics are dissolved in
filter sterilized HSW. For all experiments involving agarose- embedding of the zebrafish, a 1.5%
(w/v) solution of low melting point agarose (Sea Plaque Agarose, Lonza) dissolved in filter
sterilized HSW with anesthetic is used. There was no bias in zebrafish sex for larval
experiments, as sex determination does not occur in zebrafish before the juvenile stage (~24 dpf)
(Uchida D, Yamashita M, Kitano T, & Iguchi T 2002).
As the agarose solidifies (before it completely solidifies at 5min), a hair stem (human hair
attached to a plastic rod) is used to position the zebrafish so that the body axis is parallel to the
surface of the chamber (normal swimming orientation). Once the agarose solidifies, the caddie is
placed into a 60mm culture dishes (Corning) and 10 mL of fresh filter sterilized husbandry
system water (HSW, which consists of instant ocean and baking soda dissolved in reverse
osmosis water to achieve pH 7.2 and a conductance of ~700 μS) is added. The agarose is then
shaved down with a razor such that only the head and body are encased. Before the anesthetic
wears off after the introduction of the fresh HSW solution (~30 seconds), the tail of the zebrafish
(just posterior to the swim bladder) is carefully freed from the agarose using fine forceps
(biology tip profile, Fine Science Tools). The rostral aspect of the zebrafish (anterior to the swim
90
bladder) is held firmly in place by the agarose. The caddie is then transferred to another 60mm
culture dish in preparation for transportation to imaging room.
Figure 7.1. The schematic of the caddie used to house the zebrafish during imaging on the
flexSPIM.
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7.3 Flexspim Light Sheet Microscope
After the zebrafish has been mounted into the custom Protolabs caddie it ready to be placed onto
the flexSPIM (Figure 7.2). The flexSPIM is a versatile, multi-laser twin-microscope system for
light-sheet imaging designed and built by our collaborators, Scott Fraser and Thai
Troung(Keomanee-Dizon et al., 2020). The caddie is securely fastened on the dive bar (DB,
figure 7.2) and placed into the chamber (SC) filled with 80ml of fresh filter sterilized husbandry
system water.
92
Figure 7.2. Selective plane illumination microscopy. A) Schematic for the imaging of the neural
activity larval zebrafish combined with behavior. B) A wide-field camera below the sample
chamber provides a view of the zebrafish body. Scale bar: 400 µm. (Keomanee-Dizon et al.,
2020).
93
Figure 7.3. Schematic detailing image acquisition throughput on flexSPIM experiments. The
objective position is programmed to follow as saw waveform (blue) such that the objective takes
a unidirectional path starting at the top of the fish pallium (0um) and transitioning through to the
most dorsal aspect of pallium (60um) at a frequency of 0.75hz. The brain camera follows a
square waveform (red) that instructs it to take 11 images per objective cycle. The HR/Tail
camera follows a square waveform (green) that instructs it to take 22 images per objective cycle
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7.4 Arduino
For the experiments described in Chapters 3-5, an Arduino unit ( https://www.arduino.cc/) and
the software from the same company are used to control the stimuli. For the IR experiments, a
simple program was designed to control the infrared laser (NIR RLCO-980-1000-F laser,
Roithner Lasertechnik) by turning it on and off for two seconds, every two minutes. For the
electricity experiments, a simple program was designed to allow the native Arduino 5V output to
be turned on and off for 400ms every two minutes.
7.5 Tg(βActin-NRSE::GCaMP6s)
To monitor cell activity in these experiments we used the transgenic zebrafish Tg(βActinNRSE::GCaMP6s). This zebrafish has GCaMP6s encoporated such that when calcium ions bind
to calmodulin, when the neuron becomes active, conformational changes occurs which lead to an
increase in the fluorescence intensity from the GFP. The βActin promoter allowed us to drive
widespread expression, and also to ensure expression levels remained visible during all stages of
development. Adding an NRSE (neuron-restrictive silencer element) to the construct, allowed us
to ensure expression was restricted to neurons and limited in other cell types.
7.6 Data and Statistics
Once the experiment is completed, the resulting files are three tif images: a 4d (XYZT) brain
series image, a 3d(XYT) heart series image, and a 3d(XYT) tail movement series image. Each of
these tif files are transformed into matrices in python using numpy, a popular library designed to
95
handle multi-dimensional arrays and matrices, for further analysis using scikit-image, a popular
open-source imaging library for python. Other packages are used as well.
7.6.1 Calculating ΔF/F Traces, Area Under the Curve, and Statistics
Listed below is the list of steps used to calculate ΔF/F from brain images (Figure 7.4). Full code
available in Jupyter Notebooks upon request.
1) Using imagej, a region of interest (ROI) was made over the following areas; left
habenula, right habenula, left rostrolateral, right rostrolateral, left ventromedial, and right
ventromedial.
2) From these ROIs, the Z-axis profile was calculated for each to extract intensity values of
the regions over the duration of the experiment. These values are saved in csv format.
3) The csv files are loaded into a custom python program, converted into pandas dataframes
for ease of use in python, and normalized.
4) Since there are five stimulus events, event regions are summed 45 frames before and 40
frames after (1min before and after). The baseline for is taken to be the 45 frames before
the stimulus is applied and the ΔF/F is calculated for the entirety of the 80 frames. The
values are then converted to percentages.
5) To calculate the area under the curve, a certain number of frames before and after the
stimulus are compared. The area under the curve before the stimulus is centered on zero
so that all fish can be properly compared before and after the stimulus events. Then, I
performed a paired sample t-test to test the hypothesis that the mean of the AUC of the
fish before the stimulus was greater than the AUC of the fish after the stimulus.
96
97
Figure 7.4. Example code for calculating ΔF/F traces, area under the curve, and statistics
98
7.6.2 CaImAn
Recent advancements in fluorescence microscopy, including the two-photon light-sheet
microscope used for the experiments in this thesis, have significantly improved our ability to
observe extensive brain regions in-vivo with enhanced temporal resolution. Using CaImAn, an
open-source calcium imaging tool (Giovannucci et al., 2019), I was able to automate many
crucial steps in data processing, such as motion correction and the identification of the neural
activity of individual neurons (Figure 7.5). CaImAn was trained and validated on a
comprehensive set of manual annotations from several experts across nine mouse two-photon
imaging datasets and found to have accuracy comparable to that of human experts. The CaImAn
library contains optimized methods and algorithms such that it can efficiently run a 2021
MacBook Pro Apple M1 Max Chip with 10‑Core CPU and 32‑Core GPU. Below are the steps
used;
1) Motion correction is performed, fixing any image blurring due to fish movement. Spatial
components are identified, and their fluorescent traces are extracted and deconvolved.
This is done for every single z-slice in the brain tif so that neurons can be extracted from
multiple regions.
2) Once neuronal components have been identified, the traces from the components from
multiple regions are identified from the regions demarcated on the average projection
image. These regions include the left habenula, right habenula, left rostrolateral, right
rostrolateral, left ventromedial, and right ventromedial.
3) Since there are five stimulus events, region-specific traces are summed 45 frames before
and 45 frames after (1min before and after). The baseline for is taken to be the 45 frames
99
before the stimulus is applied and the ΔF/F is calculated for the entirety of the 80 frames.
The values are then converted to percentages.
4) Z-scores are calculated for each of the neuronal components for the entirety of the 90
frames and plotted on a heat map with red being the highest z-score and blue being the
lowest z-score. For each of the neuronal components, the Z-scores for the five frames
after the stimulus are summed so that neuronal components can be ordered from most
active to lease active, from top to bottom.
5) Individual ΔF/F for neuronal components are then stacked from most active to least
active to provide another visual way of understanding the activity of the regions.
To properly identify neuronal components in an image, the parameters used during imaging
and must be properly input into CaImAn (Figure 7.5). These parameters include variables such
as the imaging rate and the decay time of the fluorescent activity reporter (GCaMP6S has a
decay time of 1.6seconds). As for the motion correction parameters, variables such as the patch
size for piece-wise motion correct and the overlap between patches must be specified and will
vary depending on the amount of motion artifacts present in the data. Finally, the specific
parameters for source extraction and deconvolution will depend on the size of the neurons in
pixels and the density of neurons within a specified patch size.
Once CaImAn is run, the program provides you with the found components for inspection
(Figure 7.6). Along with the trace signals taken from each of the found components in the movie,
the program will give you metrics such as the signal-to-noise ration (SNR) and CNN score (the
rank given by the network indicating its confidence that the found component is a neuron.
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Figure 7.5: CaImAn Parameters for Proper Signal Extraction
101
Figure 7.6. CaImAn Identified Component. A) Presented here is a single plane taken from the
brain of a living zebrafish using a 2Photon light sheet microscope. The image has been processed
using Caiman and presented here is a heat map of activity. B) This activity trace has been
extracted from a single component found using caiman. C) Caiman provides evaluation metrics
for each found component.
102
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Abstract (if available)
Abstract
To this day, the amygdala's functional and anatomical homologue in the zebrafish remains a mystery. This is in part because the zebrafish, although a vertebrate by nature, has a distinct neurodevelopmental plan, making it difficult to establish comparative homology to that of the mammalian brain. Furthermore, our capacity to image the entire brain of the zebrafish is currently restricted to developing larval zebrafish, making it difficult to understand how information from larval zebrafish correlates with the neuroanatomical maps of adult zebrafish.
This thesis adopts the approach of exposing larval zebrafish to a wide variety of sensory stimuli, ranging from harmful to not harmful, to identify amygdalar correlates within the forebrain through the activity observed. Previous findings have shown increased pERK activation in the rostrolateral pallium during fear conditioning, so this region has been designated as a focal point for investigation.
Chapter 1 establishes the foundation for this work by examining the existing literature on fear conditioning and the neuroanatomy of mammals, providing readers with the necessary background to understand the potential analogs in zebrafish. Following that, I will provide an overview of recent research on the zebrafish brain that advocates homology through molecular and anatomical markers. Finally, I will delve into the complex interplay of the brain's valence systems, creating a holistic context for the original research that follows.
Chapter 2 reviews prior experiments that motivated the current thesis topic. Chapter 3 details experiments designed to map neural activity in the zebrafish forebrain in response to noxious stimuli. Chapter 4 expands to include the neural activity in response to life-threatening stimuli. Chapter 5 contrasts these findings with the neural activity in response to benign stimuli. This progression of experiments reveals a unique pattern of pallial activity in the larval zebrafish that suggests functional specialization within the developing forebrain. To conclude, Chapter 6 presents work done to characterize the neurons of the rostrolateral pallium, potentially laying the groundwork for future fear studies using larval zebrafish.
This thesis provides a much-desired understanding of the zebrafish forebrain, offering insights into how stimuli are processed in the pallium that will hopefully serve as a guide to the further identification of amygdala-like structures in the zebrafish and perhaps motivate more interest in utilizing the zebrafish as a model organism for behavioral neuroscience.
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Smith, Colton
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Negative valence neurons in the larval zebrafish pallium
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Neuroscience
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2024-08
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