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Neuroanatomy and neuropathophysiology of

volitional control of breathing

Pedro Henrique Trevizan Baú (BSc, MSc)

ORCID: 0000-0003-2915-3337

The Florey Institute of Neuroscience and Mental Health,

Faculty of Medicine, Dentistry and Health Sciences,

University of Melbourne,

Melbourne, VIC, Australia

Presents a thesis submitted in complete fulfilment of the requirements for the award

of the degree:

Doctor of Philosophy

June 2021

i

Abstract

Patients with neurodegenerative diseases, such as Parkinson’s, Alzheimer’s, and

frontotemporal dementia may present laryngeal motor dysfunction even during the

early stages of the disease progression. The larynx, also called the glottis, is the main

valve that regulates respiratory airflow. Moreover, the muscles of the glottis are

critically involved in the mediation of orofacial behaviours, such as vocalisation and

swallowing. For example, glottis prevents the aspiration of foods into the lungs during

swallowing. Furthermore, during speech, the vocal folds located within the glottis are

vibrated by the expiratory airflow to produce sound.

Post-mortem studies in patients with neurodegenerative diseases indicate that

neuronal degeneration within the brainstem respiratory network may be the cause of

laryngeal motor dysfunction. Not surprisingly, patients often present respiratory,

vocalisation, and swallowing malfunction. These clinical reports motivated my

research group to study a transgenic mouse model of neurodegeneration (i.e.,

tauopathy) in the brainstem respiratory network. Data from the respiratory

neurobiology team at the Florey found that similarly to clinical observations, this mouse

model presents laryngeal motor dysfunction, leading to respiratory and swallowing

impairments. Thus, the next step was to investigate whether tauopathy in the

brainstem can also be linked to impaired ultrasonic vocalisation (USV).

This working hypothesis was part of my first experimental study of my PhD (chapter

2). I found that the transgenic mouse model presented USV patterns similar to the

wild-type mice (control group), even during the late stage of diseases progression,

when many brainstem circuits have degenerated. However, it is crucial to note that

orofacial behaviours, including vocalisation, can be volitionally controlled by supra-

pontine brains regions, such as the cortex and the limbic system in the forebrain, and

I speculated that these control circuits for vocalisation might be able to compensate

for brainstem neurodegeneration. Surprisingly, the neuroanatomical pathways that

might be involved with volitional control of respiration during orofacial behaviours were

poorly defined.

Thus, the following experimental study (Chapter 3) examined the anatomical

projections from the forebrain to the brainstem respiratory network using a retrograde

tracing approach. I reported for the first time widespread descending cortical and

ii

subcortical descending projections to a subset of primary respiratory control nuclei.

Functionally, this neuroanatomical framework suggests that volitional information is

sent to the brainstem respiratory network to adapt breathing with orofacial behaviours,

such as vocalisation. Moreover, I found a highly redundant network connectome within

the brainstem respiratory network that could be implicated in the compensation of local

brainstem tauopathy (Chapter 4). Collectively, data from my thesis suggest the

presence of numerous pathways in the brain that may compensate for localised

neurodegeneration in the respiratory network of patients with neurodegenerative

diseases. In summary, my PhD thesis proposes a new neuroanatomical framework to

study behavioural adaptation of breathing in healthy and neurological diseases.

iii

Declaration

This is to certify that this thesis:

i) compromises only original work towards the PhD, except where indicated in

the preface,

ii) acknowledges all other used materials within the text,

iii) is fewer than 100,000 words in length, exclusive of tables, maps,

bibliography and appendices.

Signed:

_____________________________________

Pedro Henrique Trevizan Baú

June 2021

iv

Preface

None of the work presented in this thesis has been carried out prior to enrolment

in the degree or submitted for any other qualification.

The study presented in chapter 2 was published by Behavioural Brain Research

on September 2019 (DOI: 10.1016/j.bbr.2019.111916). Chapter 3 is a study that was

published by the Journal of Comparative Neurology in December 2020 (DOI:

10.1002/cne.25091). The study presented in chapter 4 was published by Brain

Research in April 2021 (DOI: 10.1016/j.brainres.2020.147255). For all these studies,

my supervisors and myself have conceived and designed the experiments. I have

conducted the experiments, as well as carried out experimental analysis, prepared

figures and tables and written the first versions of the manuscripts. All co-authors,

including my supervisors, contributed to interpretation of the data, reviewed drafts of

the manuscripts, and approved the final drafts before submissions. These manuscripts

only required moderate revisions before being accepted for publication. Part of chapter

5 (general discussion) is currently in preparation to be submitted as a review article.

My PhD research was supported by research grants from the National Health

and Medical Research Council of Australia (APP1165529 to Mathias Dutschmann &

Davor Stanic, and APP1078943 to Stuart Mazzone); and Australian Research Council

(Discovery project DP170104861 to Mathias Dutschmann). I was fully funded by

Melbourne Research Scholarship (University of Melbourne; 181858).

I acknowledge that this work was conducted on the traditional land of the Wurundjeri

People of the Kulin Nation. I pay my respect to their elders past, present and

emerging.

v

Publications

Trevizan-Baú, P., Dhingra, R. R., Burrows, E. L., Dutschmann, M., & Stanić, D.

(2019). Tauopathy in the periaqueductal gray, kölliker-fuse nucleus and nucleus

retroambiguus is not predicted by ultrasonic vocalization in tau-P301L

mice. Behavioural brain research, 369, 111916. DOI: 10.1016/j.bbr.2019.111916

Soliz, J., Schneider-Gasser, E. M., Arias-Reyes, C., Aliaga-Raduan, F., Poma-

Machicao, L., Zubieta-Calleja, G., Furuya, W. I., Trevizan-Baú, P., Dhingra, R.,

R., Dutschmann, M. (2020). Coping with hypoxemia: Could erythropoietin (EPO)

be an adjuvant treatment of COVID-19? Respiratory Physiology and

Neurobiology, 279, 103476. DOI: 10.1016/j.resp.2020.103476

Dutschmann, M., Bautista, TG, Trevizan-Baú, P., Dhingra, R. R., Furuya, W. I. (2021).

The pontine Kölliker-Fuse nucleus gates facial, hypoglossal, and vagal upper

airway related motor activity. Respiratory Physiology & Neurobiology, 284,

103563. DOI: 10.1016/j.resp.2020.103563

Trevizan-Baú, P., Dhingra, R. R., Furuya, W. I., Stanić, D., Mazzone S. B.,

Dutschmann, M. (2021a). Forebrain projection neurons target functionally

diverse respiratory control areas in the midbrain, pons and medulla oblongata.

Accepted for publication. Journal of Comparative Neurology, 1-22. DOI:

10.1002/cne.25091

Trevizan-Baú, P., Furuya W.I., Mazzone S.B., Stanić, D., Dhingra R.R., Dutschmann,

M. (2021b). Reciprocal connectivity of the lateral and ventrolateral columns of the

midbrain periaqueductal gray with core nuclei of the ponto-medullary respiratory

network in rat. Brain Research, 147255. DOI: 10.1016/j.brainres.2020.147255

vi

Furuya, W. I., Dhingra, R. R., Trevizan-Baú, P., McAllen, R. M., Dutschmann, M.

(2021). The role of glycinergic inhibition in respiratory pattern formation and

cardio-respiratory coupling in rats. Current Research in Physiology. DOI:

10.1016/j.crphys.2021.03.001

vii

Presentations

Conference presentations arising from this research:

Trevizan-Baú, P., Burrows, E. L., Dutschmann, M., Stanić, D. (2017). The impact of

tauopathy on ultrasonic vocalisation in a mouse model of frontotemporal

dementia with parkinsonism. Poster presented at the Central Cardiorespiratory

Control: future directions, Sydney, Australia.

Trevizan-Baú, P., Dhingra, R.R., Burrows, E. L., Dutschmann, M., Stanić, D. (2018).

Tauopathy in the periaqueductal gray, Kölliker-Fuse nucleus and nucleus

retroambiguus does not predict ultrasonic vocalisation disorder in Tau.P301L

mice. Poster presented at the Central Cardiorespiratory Control: future

directions, Melbourne, Australia.

Stanić, D., Trevizan-Baú, P., Burrows, E. L., Dhingra, R.R., Dutschmann, M. (2018).

The impact of tauopathy on ultrasonic vocalization in a mouse model of

frontotemporal dementia. Poster presented at the Experimental Biology, San

Diego, United States.

Trevizan-Baú, P., Dutschmann, M. (2019). Suprapontine monosynaptic projections to

the brainstem respiratory areas in rats. Poster presented at the Central

Cardiorespiratory Control: future directions, Auckland, New Zealand.

Trevizan-Baú, P., Dutschmann, M. (2019). Suprapontine monosynaptic projections to

the brainstem respiratory areas in rats. Poster presented at the Students of Brain

Research, Melbourne, Australia.

Trevizan-Baú, P., Dhingra, R.R., Dutschmann, M.. (2020). Descending forebrain

projections targeting respiratory control areas in the midbrain and brainstem of

rats. Poster presented at the Experimental Biology, San Diego, United States.

viii

Awards

Melbourne International Research Scholarship (2017-2021)

Melbourne International Fee Remission Scholarship (2017-2021)

Florey top-up Scholarship (2017-2020)

Florey Travel top-up (2017-2021)

ix

Acknowledgments

I owe a great thanks to Mathias Dutschmann for being my supervisor and guiding me

throughout this journey. I thank him very much for his tremendous support and

guidance. I also would like to thank Davor Stanic for his guidance and mentorship as

my co-supervisor. I am grateful to Song Yao and Emma Burrows for serving on my

advisory committee and for their valuable feedback during these years.

I acknowledge the University of Melbourne for awarding me with the International

Research Scholarship. I also thank the Florey Institute of Neuroscience and Mental

Health for the scholarship top-up and the travel money. I would also like to thank all

the Florey and University staff for their support.

Rishi Dhingra and Werner Furuya also deserve many thanks for their support and

feedback. I thank Rishi for helping me with the mathematical analysis and, thus,

improving my data analysis. I would like to thank Stuart Mazzone and his group for the

generous use of the brightfield microscope. I also thank Clare Parish and Lachlan

Thompson for the generous use of freezing microtome and microscopy facilities.

A huge and exceptional thanks to my wife, Flavia, for supporting me every day of my

life here in Australia. Your love, company, help, and great effort facilitated my

performance in my PhD. Also, I sincerely acknowledge my family and friends for all

their support. I cannot express enough gratitude for supporting my choices, including

my decision to move to Australia to start my PhD. I thank João Paulo for being a

fantastic brother and being patient with me. Your experience always helps me to do

and be better. And most important, an extraordinary thanks to mom and dad for

everything they do for me. They continue to inspire me to be and do better every day

of my life. I dedicate this thesis, as special gratitude, to them.

x

Table of Contents

Abstract ....................................................................................................................... i

Declaration ................................................................................................................. iii

Preface ....................................................................................................................... iv

Publications ................................................................................................................ v

Presentations ............................................................................................................ vii

Awards ..................................................................................................................... viii

Acknowledgments ...................................................................................................... ix

Table of Contents ....................................................................................................... x

List of figures ............................................................................................................. xv

List of Abbreviations ............................................................................................... xviii

Chapter 1 ............................................................................................... 1

General Introduction ............................................................................. 1

Lung ventilation and central control of breathing ........................................................ 2

Respiration is integrated with orofacial behaviours: a reflection on the respiratory-

vocalisation brain network .......................................................................................... 5

Neurodegenerative diseases affect voice and respiration ........................................ 10

Summary and experimental aims ............................................................................. 11

Chapter 2 ............................................................................................. 13

Tauopathy in the periaqueductal gray, Kölliker-Fuse nucleus and

nucleus retroambiguus is not predicted by ultrasonic vocalisation in

tau-P301L mice ................................................................................... 13

Abstract .................................................................................................................... 14

Introduction .............................................................................................................. 15

xi

Materials and Methods ............................................................................................. 17

Animals .................................................................................................................. 17

Experimental protocol for USV recordings ............................................................. 19

Immunohistochemistry ........................................................................................... 20

Tissue preparation ............................................................................................. 20

Experimental protocol ........................................................................................ 21

Primary antisera used ........................................................................................ 21

Image processing ............................................................................................... 22

Data analysis .......................................................................................................... 22

Ultrasonic Vocalisation ....................................................................................... 22

Linear discriminant analysis (LDA) ..................................................................... 23

Immunohistochemistry ....................................................................................... 23

Statistic .............................................................................................................. 24

Results ..................................................................................................................... 24

USV is similar in tau-P301L and wildtype FVB/N mice ........................................... 24

Repetitive recording reveals a decreased number of USVs emitted by symptomatic

tau-P301L and wildtype mice aged 8-9 months ..................................................... 28

Tauopathy in the midbrain periaqueductal gray, pontine Kölliker-Fuse nucleus and

medullary nucleus retroambiguus is not predictive of USV disorders. .................... 30

Discussion ................................................................................................................ 36

Conclusion ............................................................................................................... 41

Chapter 3 ............................................................................................. 42

Forebrain projections neurons target functionally diverse

respiratory control areas in the midbrain, pons and medulla

oblongata ............................................................................................ 42

Abstract .................................................................................................................... 43

Introduction .............................................................................................................. 44

Material and Methods ............................................................................................... 46

xii

Animals .................................................................................................................. 46

Surgery .................................................................................................................. 46

Tissue preparation and immunohistochemistry ...................................................... 48

Data analysis and image processing ...................................................................... 49

Results ..................................................................................................................... 51

Injections sites ........................................................................................................ 51

Cumulative distribution and laterality of descending forebrain projection ............... 54

Specific distribution of retrogradely CT-B labeled neurons in cortical and subcortical

brain regions following injections in the midbrain PAG ........................................... 56


brain regions following injections in the pontine KFn .............................................. 61


brain regions following injections in the medullary pre‐BötC and BötC .................. 64


brain regions following injections in the caudal raphé in the medulla oblongata .... 69

CT-B-labeled neurons associated with near-miss injections .................................. 71

Discussion ................................................................................................................ 72

Homeostasis, state-dependent or emotional modulation of respiration .................. 75

Volitional control of respiration ............................................................................... 77

Conclusions .............................................................................................................. 80

Chapter 4 ............................................................................................. 81

Reciprocal connectivity of the periaqueductal gray with the ponto-

medullary respiratory network in rat ................................................. 81

Abstract .................................................................................................................... 82

Introduction .............................................................................................................. 83

Material and Methods ............................................................................................... 85

xiii

Animals .................................................................................................................. 85

Surgery .................................................................................................................. 85

Tissue preparation and immunohistochemistry ...................................................... 86

Data analysis and image processing ...................................................................... 87

Results ..................................................................................................................... 88

Distribution of retrogradely labeled neurons in the PAG after CT-B injections in the

ponto-medullary respiratory nuclei ......................................................................... 89

Distribution of retrogradely labeled neurons in ponto-medullary respiratory nuclei

after CT-B injections in the lateral and ventrolateral columns of the PAG .............. 93

Summary of the afferent and efferent connectivity of the lateral and ventrolateral PAG

columns with the respiratory nuclei in the ponto-medullary brainstem ................... 96

Discussion ................................................................................................................ 98

Conclusion ............................................................................................................. 100

Chapter 5 ........................................................................................... 101

General Discussion .......................................................................... 101

Abstract .................................................................................................................. 102

Emerging concepts for functional neuroanatomy underlying the coordination of

volitionally evoked orofacial behaviour and breathing ............................................ 103

Pivotal role of postinspiratory control of glottal adduction in coordinating orofacial

behaviours and breathing ..................................................................................... 104

Corticobulbar and corticospinal tracts and their potential involvement in volitional

control of respiratory muscles .............................................................................. 108

The periaqueductal gray, a pre2motor area in the midbrain: an established interface

for volitional and emotion control of vocalisation .................................................. 112

The role of respiratory and premotor nuclei of the respiratory network during the

coordination of volitionally evoked orofacial behaviour and breathing .................. 114

Emotional control of respiration: a key role for the pre-Bötzinger complex? ........ 117

xiv

Neurologic diseases are tightly linked to impaired volitional control of orofacial

behaviour and breathing ...................................................................................... 119

Conclusions and future perspectives...................................................................... 124

References ........................................................................................ 127

xv

List of figures

Chapter 1

Figure 1.1. Schematic figure showing the phrenic nerve activity (PHN, green) and recurrent laryngeal

nerve activity (RLN, blue) during a respiratory cycle in mammals.………………………....5

Chapter 2

Figure 2.1. Representative spectrograms of USV sequences emitted during social male-female

interaction……………………………………………………………………………………...25

Figure 2.2. Ultrasonic vocalisation measurements of wildtype FVB/N (grey circles) and tau-P301L

(magenta triangles) mice across the asymptomatic (3-4 months), pre-symptomatic (5-6

months) and symptomatic (8-9 months) age groups…………………………………….…26

Figure 2.3. USV features do not predict the presence of tau-overexpression……………………...…28

Table 2.1. Linear discriminant model and accuracy……………………………………………….…...28

Figure 2.4. Age-dependent changes in the number of USVs/min in individual (A) wildtype FVB/N and

(B) tau-P301L mice…………………………………………………………………….……...30

Figure 2.5. The relationship between USVs emitted and tauopathy in 8-9 month-old tau-P301L and

FVB/N wildtype mice…………………………..………………………………………….…..31

Figure 2.6. PHF-Tau immunoreactivity (green) in the periaqueductal gray of 8-9 month-old tau-

P301L mice……………………………………………………………………...………….….32

Figure 2.7. PHF-Tau immunoreactivity (green) in the Kölliker-Fuse nucleus of 8-9 month-old tau-

P301L mice………………………………………………………………………………….....34

Figure 2.8. PHF-Tau immunoreactivity (green) in the nucleus retroambiguus of 8-9 month-old tau-

P301L mice…………………………………………………………………………...….…….35

Chapter 3

Table 3.1. Stereotaxic coordinates for CT-B microinjections………………………………..…….…..47

Figure 3.1. Schematic drawings (left) and photomicrographs (right) illustrate the anatomical location

of cholera toxin subunit B (CT‐B) microinjections in functionally diverse respiratory

control areas……………………………………………………………………………….…..53

Figure 3.2. Bar graphs of the total (a), ipsilateral (b) and contralateral (c) numbers of retrogradely

cholera toxin subunit B (CT‐B)‐labeled neurons in the forebrain after microinjections in

PAG, KFn, pre‐BötC, BötC, and caudal raphé nuclei……………………………………..55

Figure 3.3. Representative cholera toxin subunit B (CT‐B)‐labeled pyramidal neurons in Layer V/VI

of the insular cortex (a)……………………………………………………………….…….…56

Figure 3.4. Cholera toxin subunit B (CT‐B) microinjection in the PAG labeled neurons in various

cortical and forebrain regions………………………………………………………………...57

Figure 3.5. The rostrocaudal distribution of the numbers of retrogradely labeled neurons relative to

bregma (mm) in various cortical (a) and subcortical (b) brain regions following cholera

toxin subunit B (CT‐B) microinjections in the periaqueductal gray………………………..59

xvi

Figure 3.6. Cholera toxin subunit B (CT‐B) microinjection in the Kölliker‐Fuse nucleus labeled

neurons in various cortical and forebrain regions (paired photomicrographs: 1 is low and

2 is high magnification)………………………………………………………………..………62

Figure 3.7. The rostrocaudal distribution of retrogradely labeled neurons relative to bregma (mm) in

various cortical (a) and subcortical (b) brain regions following cholera toxin subunit B

(CT‐B) microinjections in the Kölliker‐Fuse nucleus…………………………………….…63

Figure 3.8. Cholera toxin subunit B (CT‐B) microinjection in the pre‐Bötzinger complex labeled

neurons in various cortical and forebrain regions (paired photomicrographs: 1 is low and

2 is high magnification)…………………………………………………………………..……65



(CT‐B) microinjections in the pre‐Bötzinger complex…………………………….………..66

Figure 3.10. The rostrocaudal distribution of the numbers of retrogradely labeled neurons relative to

bregma (mm) in various cortical (a) and subcortical (b) brain regions following cholera

toxin subunit B (CT‐B) microinjections in the Bötzinger complex…………………….…..67

Figure 3.11. Cholera toxin subunit B (CT‐B) microinjection in the caudal raphé nuclei labeled neurons

in various cortical and forebrain regions (paired photomicrographs: 1 is low and 2 is high

magnification)………………………………………………………………………..…….…..69



(CT‐B) microinjections into the caudal raphé nuclei………………………………………..70

Figure 3.13. Summary of the relative strength of descending projections arising from cortical (a) and

subcortical (b) areas in a connectivity map……...…………………………………….……74

Chapter 4

Figure 4.1. CT-B injections in the pontine Kölliker-Fuse nucleus, the medullary Bötzinger complex

and pre-Bötzinger complex, and the caudal raphé.XX…………………………………….89

Figure 4.9. Injections in the ponto-medullary target regions (KFn, BötC, pre-BötC, and caudal raphé)

resulted in CT-B-labeled neurons in the midbrain PAG……………………………..….….91

Figure 4.3. Projections from the midbrain PAG to the ponto-medullary respiratory control

nuclei…………………………………………………………………..……………….………92

Figure 4.4. CT-B injections in the midbrain PAG………………………………………………………..94

Figure 4.5. Projections from the ponto-medullary respiratory regions to the midbrain

PAG………………………………………………………………………………….…….…...95

Figure 4.6. Summary of the relative strength of projections arising from the midbrain PAG columns

(dmPAG, dlPAG, lPAG, and vlPAG; A), and the ponto-medullary regions (KFn, pre-BötC,

BötC, and the caudal raphé; B) in a connectivity map……………………………….……..97

https://www.sciencedirect.com/topics/neuroscience/periaqueductal-gray


xvii

Chapter 5

Figure 5.1. Respiratory motor activity is evoked by cranial and spinal nerves…………………...….107

Figure 5.2. Corticobulbar and corticospinal projections into the motor pools……………………...110

Figure 5.3. Top-down information from the forebrain to key brainstem respiratory control areas of the

pons and medulla oblongata might play critical roles adapting breathing with orofacial

behaviours and emotions……………………………………………………………………118

Table 5.1. Clinical studies have reported impaired orofacial motor behaviours in several

neurological diseases………………………………………………………………...……..123

xviii

List of Abbreviations

12N, hypoglossal nucleus

7N, facial nucleus

3V, third ventricle

4V, fourth ventricle

AbNA, abdominal nerve activity

Aca, anterior commissure

ACA, cingulate cortex

Amg, Amygdala

Aq, aqueduct (Sylvius)

BötC, Bötzinger complex

BT-FITC, biotinyl tyramide-fluorescein

C, cervical spinal nerves

C4, cervical spinal nerve 4

cc, corpus callosum

CPu, caudate putamen (striatum)

DEn, endopiriform nucleus

dlPAG, dorsolateral PAG

DMH, dorsomedial hypothalamus

dmPAG, dorsomedial PAG

E2, late expiratory phase

E/OV, ependymal and sublayer EV

ec, external capsule

exp, expiration

fmi, forceps minor of corpus callosum

FNA, facial nerve activity

FTD, frontotemporal dementia

FTDP-17, Frontotemporal dementia and Parkinsonism linked to Chromosome 17

GlyT2-KO, glycine transporter-2 knock-out

HNA, hypoglossal nerve activity

I, inspiratory phase

IC, internal capsule

IF, infralimbic cortex

xix

II, Layer 2 of cortex

III, Layer 3 of cortex

insp, inspiration

io, inferior olive

IX, glossopharyngeal nerve

KFn, Kölliker‐Fuse nucleus

L, lumbar spinal nerves

L1, lumbar spinal nerve 1

LDA, linear discriminant analysis

LH, lateral hypothalamus

lPAG, lateral PAG

LS, lateral septum

LV, lateral ventricle

mcp, middle cerebellar peduncle

MI, motor cortex

mlf, medial longitudinal fasciculus

MO5, motor trigeminal nuclei

NA, nucleus ambiguous

NRA, nucleus retroambiguus

NTS, solitary tract nucleus

och, optic chiasm

opt, optic tract

PAF, pulmonary airflow

PAG, periaqueductal gray

PHF-Tau, paired helical filament-tau

PHN, phrenic nerve activity

PI, postinspiratory phase

PL, prelimbic cortex

pre-BötC, pre-Bötzinger complex

PVN, paraventricular hypothalamus

py, pyramidal tract

pyx, pyramidal decussation

rf, rhinal fissure

xx

RLNA, recurrent laryngeal nerve activity

RMg, raphé magnus

ROb, raphé obscurus

RPa, raphé pallidus

RT, room temperature

scp, superior cerebellar peduncle

SGP, subglottal pressure

T, thoracic spinal nerves

TNA, trigeminal nerve activity

USV, ultrasonic vocalisation

V, Layer 5 of cortex

VI, Layer 6 of cortex

VII, facial nucleus

vlPAG, ventrolateral PAG

VMH, ventromedial hypothalamus

VRG, ventral respiratory group

X, vagus nerve

XII, facial motor nucleus

1

Chapter 1

General Introduction

2

Lung ventilation and central control of breathing

Respiration is a vital rhythmic motor function that maintains blood gas

homeostasis by controlling pulmonary oxygen uptake and excretion of carbon dioxide.

The underlying biomechanics of lung ventilation are related to distinct changes in

intrapulmonary pressure during the two alternating phases of inspiration and

expiration. During inspiration, contraction of the diaphragm increases intrapulmonary

volume and decreases intrapulmonary pressure, and the latter then propagates airflow

into the lungs. The subsequent expiration is often called ‘passive’ given that the force

for expiratory airflow out of the lungs is driven by the passive recoil of the expanded

lung and ribcage that generates the required increase in intrapulmonary pressure (for

review, see Bates, 2009). However, the term ‘passive’ for expiration is misleading

since maximal expiratory airflow does not occur during the early phase of expiration

when the recoil force would be greatest. Expiratory airflow is, in reality, actively

controlled by the so-called laryngeal valving mechanism. This involves the contraction

of laryngeal adductor (constrictor) muscles during the early phase of expiration (i.e.,

postinspiratory phase), which increases upper airway resistance by narrowing the

vocal folds and glottis aperture to limit expiratory airflow (Bartlett et al., 1973).

Therefore, respiration is not only determined by contractions of the diaphragm that

drive the respiratory cycle, but also requires the laryngeal valve to modulate airflow in

and out of the lungs. Because of the postinspiratory mediated glottal constriction

during early expiration, the respiratory cycle is divided into three primary phases:

inspiration, postinspiration and expiration (Dutschmann and Dick, 2012).

The 3-phase breathing pattern is generated by a central respiratory pattern

generator (rCPG) that is anatomically organized into a continuous cell column in the

brainstem, stretching from the rostral dorsolateral pons to the caudal ventrolateral

3

medulla, and is innervated by inhibitory and excitatory synapses (Dutschmann and

Dick, 2012; Smith et al., 2013). Because of the anatomical position of the rCPG, this

respiratory network is also called the lateral respiratory column. The synaptic

interactions between pontine and medullary respiratory neurons generate the

respiratory motor pattern that is in turn transmitted to cranial and spinal respiratory

motor pools. The respiratory motor neurons are located in the spinal cord and cranial

motor nuclei. The spinal motor neurons in the cervical (e.g., phrenic motor nucleus)

and thoracic spinal cord innervate respiratory pump muscles in the thorax and

abdomen, while the cranial motor pools innervate muscles of the oropharyngeal and

nasopharyngeal airway (Smith et al., 1990).

Specific respiratory motor activities can be expressed in a variety of motor

outputs. For instance, the phrenic nerve that innervates the diaphragm is the primary

motor output for inspiration (Fig. 1.1). However, cranial nerves are also activated

during inspiration. Inspiratory activity can be seen in the: 1) hypoglossal nerve, which

innervates the tongue; 2) trigeminal and facial nerves that innervate muscles of the

nasopharyngeal cavity; and 3) recurrent laryngeal nerve, which innervates the

laryngeal muscles (Fig. 1.1). During inspiration, these cranial nerves activate dilator

muscles across the upper airways to decrease airway resistance during inhalation.

During postinspiration, as described above, the recurrent laryngeal nerve drives the

activity of laryngeal constrictor muscles. At later stages of expiration, no nerve activity

is detected during normal breathing. However, when breathing at high tidal volumes

and frequencies, active expiratory activity can be expressed in abdominal expiratory

muscles as well as upper airway dilator pathways.

According to the current scientific view, the rCPG can be divided into

functionally different compartments that are associated with control of specific

4

respiratory motor activities either during inspiration, postinspiration or expiration

(Dutschmann and Dick, 2012; Feldman and Del Negro, 2006; Smith et al., 2013; Smith

et al., 1990). For example, the pre-Bötzinger complex (pre-BötC) has been postulated

as a local pacemaker circuit for the generation of the inspiratory rhythm (Feldman and

Del Negro, 2006). However, the generation of the inspiratory and expiratory motor

pattern requires synaptic interactions between all neurons of the rCPG. After the

initiation of inspiration, the generation of the sequential respiratory motor pattern

depends on reciprocal or mutual inhibitory/excitatory activity of neurons, forming an

oscillating activity that generates the 3-phase motor pattern. Thus, specific classes of

respiratory neurons are either excitatory or inhibitory. For example, a subclass of

excitatory inspiratory neurons in the pre-BötC initiates inspiration while a second

subclass of neurons inhibits all expiratory neurons during ongoing inspiration

(Feldman et al., 2003; Schwarzacher et a., 1995). A critical region mediating the

transition from inspiration to postinspiration is located at the rostral tip of the rCPG in

the pons. Previous work has shown that the postinspiratory phase is gated by the

Kölliker-fuse nucleus (KFn) and parabrachial nuclei. Synaptic blockade of these nuclei

causes delayed inspiratory termination and complete absence of postinspiratory

laryngeal adductor activity (Dutschmann and Herbert, 2006). Other compartments

such as the A5 cell group, parafacial respiratory group, the Bötzinger complex, and

the rostral and caudal ventral respiratory group, are seen to have specific roles in the

generation of expiration and contain an important population of neurons that relays the

central respiratory activity to the designated motor pools in the brainstem and spinal

cord (for review, see Dutschmann and Dick, 2012).

5

Fig 1.1. Schematic figure showing the phrenic nerve activity (PHN, green) and recurrent

laryngeal nerve activity (RLN, blue) during a respiratory cycle in mammals.

Phases of the respiratory cycle are highlighted as: dark grey for inspiration (I); light grey for

postinspiration (PI); and no shading for expiration (E). Note that RLN activity increases in PI.

Pontine control of the postinspiratory phase is of particular importance for the

mediation of upper airway reflexes (e.g., cough and sneeze), and orofacial motor

behaviours (e.g., vocalisation and swallowing), that all depend on the timely activation

of the laryngeal adductor muscles (Dutschmann and Dick, 2012; Dutschmann et al.,

2014). The following section discusses the role of the larynx and the laryngeal

adductors during postinspiration with focus on vocal sound production. Nevertheless,

it is essential to note that, as discussed above, laryngeal activation is critical for

modulating the expiratory airflow during numerous orofacial behaviours (e.g., sniffing

whisking, swallowing, etc.) that need be integrated with respiration.

Respiration is integrated with orofacial behaviours: a reflection on the

respiratory-vocalisation brain network

The larynx is the essential vocal organ that generates sound and pitch during

speech (Fitch, 2000; Fitch, 2006). Of particular importance are the laryngeal adductors

6

that have an essential role in breathing and are critically involved in the generation of

voice (Shiba et al., 1997). It is also crucial to note that vocalisation exclusively takes

place during the postinspiratory phase of the respiratory cycle (Dutschmann et al.,

2014).

The larynx connects the inferior part of the pharynx with the trachea and is also

the organ to which the vocal folds are attached internally, making the larynx essential

for mammalian vocalisation. The general physiological basis for the generation of

vocal behaviour has three components (Fitch, 2006). First is the modulation of

expiratory airflow, and to some extent, the inspiratory lung volume and capacity that

determines the duration of the subsequent expiratory vocal behaviour. The second

component comprises laryngeal muscle activity, and, in the third component,

associated supra-laryngeal movements of the jaws, lips, tongue, and soft palate

(Dutschmann et al., 2014; Fitch, 2006; Hage, 2010; Jürgens, 2000; Ludlow, 2005;

West and Larson, 1993). The laryngeal adductor muscles control the degree of glottal

narrowing by altering the stiffness of the vocal folds during the expiratory airflow as a

prerequisite for sound production. Once a sound is generated in the larynx, it can be

articulated and modulated by supra-laryngeal component of vocalisation.

The central pathways for the generation of vocalisation are not well understood.

However, there are some generally accepted concepts, such as the ‘forebrain-

midbrain-pons’ pathway. This pathway is hierarchically organised from the higher

levels of the brain to the pons, and controls the motor neurons associated with

vocalisation (Jürgens, 2002a,b). Studies in anencephalic children demonstrated their

inability to produce learned vocalisation, but they could still generate innate sound (for

review, see Gruber-Dujardin, 2010). In attempts to pinpoint essential brain regions for

vocalisation in cat, vocalisation could still be elicited following decerebration at the

7

forebrain-midbrain border. However, no vocalisation was recorded after decerebration

at midbrain-pons border, suggesting that the midbrain is essential for the generation

of vocalisation in mammals (Gruber-Dujardin, 2010).

The periaqueductal gray (PAG) has been considered a crucial area of the

midbrain for generating vocalisation because: 1) several studies have shown that

mammals elicit vocalisation when the PAG is electrically and pharmacologically

stimulated (Jürgens, 1994; Larson and Kistler, 1984); 2) PAG lesions lead to mutism

(Jürgens, 2002); 3) the PAG sends afferent projections to the nucleus retroambiguus

(NRA), which has premotor neurons connected with the laryngeal motor neurons

(Holstege, 1989; Jürgens, 2002); and 4) PAG neurons fire before and during

vocalisation behaviour (Düsterhöft et al., 2004). However, although the PAG seems to

play a critical role in generating vocalisation in mammals, it is still unknown how the

neural circuit for vocalisation is synchronised and organised with the neural control of

breathing. Moreover, the PAG is associated with survival and emotional behaviour

with vocalisation-correlated activity, such as cardiovascular changes, micturition,

receptive behaviour, pupil dilatation, and defensive posture and respiration (Holstege,

1989; Subramanian et al., 2008a). Thus, it is possible that vocalisation elicited by

PAG-stimulations could be an indirect effect of those behaviours. Even so, the

likelihood of the PAG being a relay station for vocalisation behaviour cannot be

excluded, and the question regarding this role should be: is the PAG involved in motor

coordination or gating function of vocalisation? Nevertheless, some studies are

indicating that the PAG does not change specific parameters of vocalisation, such as

frequency (Düsterhöft et al., 2004), but rather it modulates the intensity of sound and

triggers the initiation of sound, roles that would give the PAG a gating function

(Jürgens, 2000; Jürgens and Pratt, 1979).

8

Questions regarding the motor coordination of vocalisation and how sound

parameters are changed in mammals are not fully resolved. However, the respiratory

network seems to have a close relationship with neuronal networks responsible for

vocalisation behaviour. For instance, most of the neurons firing during expiration (i.e.,

rostral ventral respiratory group and Bötzinger complex) are also active during

vocalisation in anesthetized guinea pigs (Sugiyama et al., 2014). Additionally, the

interaction between respiration and vocalisation is well designed so that vocalisation

takes place during postinspiration, the early phase of controlled expiration (discussed

above; see Dutschmann et al., 2014). KFn neurons, which are important regulators of

the postinspiratory phase (Dutschmann and Dick, 2012), receive and send many

projections to the brainstem and spinal cord (Song et al., 2012). Therefore, it has been

hypothesized that the KFn may play an essential role in the fine control of vocalisation

via a proposed pathway between the PAG and the KF pathway (Dutschmann and

Dick, 2012). However, since many nuclei in the brainstem control breathing, it cannot

be excluded that other nuclei may also contribute to the motor control of vocalisation.

The relationship between respiration and vocalisation was reinforced recently by

Stanić and colleagues (Stanić et al., 2008), in which they observed high numbers of

Forkhead box protein P2 (FOXP2) immunoreactive nuclei in the brainstem areas

associated with breathing in rats, FOXP2 being a transcription factor associated with

the generation of vocalisation (Vargha-Khadem et al., 2005). Interestingly, the KFn

neurons showed the highest density of FOXP2 immunoreactivity compared to other

brainstem respiratory nuclei, suggesting that the pontine KFn is important for

vocalisation in mammals.

Besides the midbrain and pons controlling sound generation in mammals,

planning and audio-vocal learning vocalisation require activation of the motor cortex,

9

cerebellum, and limbic system (Jürgens, 2002). In the motor cortex, two areas have

been postulated to be essential for learned vocalisation in humans and non-human

primates: the anterior cingulate cortex and the motor cortex (Jürgens, 2002). Not only

primates, but a recent study in bats showed that genes associated with vocalisation

(e.g., FOXP1, FOXP2, and cntnaP2) are widely expressed in the bat brains, including

cortical and limbic brain regions that control vocal learning behaviour in mammals

(RodenasCuadrado et al., 2018). The brain network between these higher brain areas

is responsible for cognitive and behavioural motor commands (i.e., volitional

behaviour; Orem and Netick, 1986; Chang, 1992), but the descending pathways to the

phonatory motor neurons are still not entirely determined. Although the PAG receives

projections from cortical, limbic, thalamic, and hypothalamic areas, some studies have

argued that cortical neurons associated with volitional vocalisation project their axons

directly to the motor neurons, neglecting the respiratory-vocalisation network in the

brainstem (Butler, 2007; Corfield et al., 1998; Gandevia and Rothwell, 1987a,b;

Pouget et al., 2018).

The close relationship between vocalisation and respiration, and their

regulation by brainstem circuitry, may be highlighted by symptoms that occur in

neurodegenerative diseases. Many of these diseases in humans affect both

respiration and vocalisation behaviours, and symptoms involving respiration and

vocalisation are often present together in the early phase of the disease (discussed

below). Because of this link, and to better understand the neurobiology of neurological

diseases that affect respiration and vocalisation, it is necessary to study brain areas

involved in vocalisation, including the brainstem respiratory network.

10

Neurodegenerative diseases affect voice and respiration

The production of speech is a complex behaviour that involves many areas of

the central nervous system in mammals (as outlined in the previous section). However,

most studies concerned with voice and speech production in health and

neuropathology have focused on the forebrain, especially the cortical areas (Dronkers,

19996; Shu et al., 2005; Tarkka, 2001), thereby largely neglecting the brainstem areas

(discussed previously). However, it is well documented that the brainstem is affected

in neurodegenerative diseases (Eser et al., 2018). Moreover, it seems that

neurodegeneration that includes tauopathy (see below) may start in the spinal cord

and brainstem circuits and then spread to higher brain areas with disease progression

(Braak et al., 2002). Thus, the disease progression suggests that vocalisation areas

in the brainstem could be affected in the earlier stages of the disease, given that

speech disorder is one of the most common symptoms observed in patients with

tauopathy (Kent et al., 2000).

Tauopathy is the aggregation of misfolded and hyperphosphorylated tau protein

in the cytoplasm of the neurons (wolf, 2012), and the physiological role of tau protein

is the assembly of tubulin monomers into microtubules in axons (Binder et al., 1985)

and dendrites (Ittner et al., 2010). In tauopathy, tau protein cannot interact with

microtubules, resulting in impaired axonal transport mechanisms (Bramblett et al.,

1993). To understand the impact of tauopathy in the central nervous system, research

groups have been generating several animal models carrying tau-protein mutations

already identified in humans (Hutton et al., 1998). For example, aging mice carrying

mutated tau-protein showed upper airway dysfunctions, such as reduced

postinspiratory activity and airflow reduction (Dutschmann et al., 2010). Additionally,

tauopathy was observed in some areas of the central nervous system which are known

11

to play a critical role in the central control of respiration, such as the PAG, KFn,

Nucleus of Solitary Tract (NTS) axis and cervical and thoracic spinal cord

(Dutschmann et al., 2010). Indeed, many areas mentioned above could play a role in

orofacial (respiratory-related) behaviours such as vocalisation (Sugiyama et al., 2014).

Together with the idea that respiration and vocalisation are two behaviours with a close

connection and interaction in mammals, it has been hypothesised that mutations of

human tau-protein expressed in mice will also lead to vocalisation impairment, owing

to vocalisation requiring expiratory airflow through the larynx (Menuet et al., 2011b).

Summary and experimental aims

Respiration is modulated during orofacial motor behaviours, including

vocalisation and swallowing, as well as airway reflexes, such as cough and sneeze.

For example, during vocalisation, the vocal folds in the upper airway are vibrated by

the expiratory airflow, generating sound. During swallowing, glottal closure prevents

the aspiration of foods into the lungs. Moreover, evidence in the scientific literature

indicates that the neural control of orofacial motor behaviours is integrated within the

respiratory network areas in the brainstem. Additionally, clinical studies have shown

that patients with neurodegenerative diseases (Parkinson’s, Alzheimer’s, and

frontotemporal dementia) often develop impaired coordination of respiration with

orofacial motor behaviours that present as commonly observed vocalisation or

swallowing disorders. In order to develop a comprehensive understanding of the

neuropathophysiology of swallowing and speech in neurodegenerative diseases, it is

essential to decipher the neuroanatomical organization of the extended neural

networks that modulated and control breathing during orofacial behaviour. This thesis

aimed to investigate the extended neuroanatomical pathways that link cognitive and

behavioural commands with the primary respiratory network in the brainstem.

12

The first experimental chapter (chapter 2) of the present thesis investigates the

impact of tauopathy on the expression of ultrasonic vocalisation (USV) in a transgenic

mouse model of frontotemporal dementia that develops severe tauopathy in the

brainstem respiratory regions. For this study, I implemented a social vocalisation

paradigm of male-female interaction. My investigations showed that tauopathy in

brainstem respiratory control areas neither cause nor predict vocalisation disorders in

the investigated transgene mouse model.

Given that social vocalisation is thought to be volitionally initiated in the

forebrain, the second experimental chapter (chapter 3) aimed to identify cortical and

subcortical brain regions that connect to the brainstem respiratory control areas in rats

by using a retrograde tracer approach. Overall, chapter 3 establishes an anatomical

framework for the integration of volitional motor commands for orofacial behaviours

within critical respiratory control nuclei in the pons and medulla oblongata. In the third

experimental chapter (chapter 4), I investigated the connectivity pattern of the

identified primary targets of descending forebrain connections such as the midbrain

periaqueductal gray, the pontine Kölliker-Fuse nucleus, and the pre-Bötzinger

complex in the medulla. I identified a variety of redundant neural pathways that may

potentially functionally compensate of local brainstem tauopathy.

Chapter 5 (general discussion) provides a comprehensive summary of the

potential functional significance of neural pathways between higher brain centres and

the brainstem respiratory network that might be critically involved in cognitive and

behavioural commands for breathing and orofacial motor behaviours in health and

neurological diseases.

13

Chapter 2

Tauopathy in the periaqueductal gray, Kölliker-Fuse

nucleus and nucleus retroambiguus is not predicted by

ultrasonic vocalisation in tau-P301L mice

Pedro Trevizan-Baú, Rishi R. Dhingra, Emma L. Burrows+, Mathias Dutschmann,

Davor Stanić

The Florey Institute of Neuroscience and Mental Health, Discovery Neuroscience

theme, +Mental Health theme, University of Melbourne, Parkville, Victoria, 3010,

Australia

The following chapter was published by Behavioural Brain Research on September 2,

2019 (DOI: 10.1016/j.bbr.2019.111916). The full paper has been inserted as a thesis

chapter in accordance with The University of Melbourne guidelines.

14

Abstract

Upper airway and vocalisation control areas such as the periaqueductal gray (PAG),

kölliker-fuse nucleus (KF) and nucleus retroambiguus (NRA) are prone to developing

tauopathy in mice expressing the mutant human tau P301L protein. Consequently,

impaired ultrasonic vocalisation (USV) previously identified in tau-P301L mice at the

terminal disease stage of 8-9 months of age, was attributed to the presence of

tauopathy in these regions. Our aim was to establish whether the onset of USV

disorders manifest prior to the terminal stage, and if USV disorders are predictive of

the presence of tauopathy in the PAG, KF and NRA. USVs produced by tau-P301L

and wildtype mice aged 3-4, 5-6 or 8-9 months were recorded during male-female

interaction. Immunohistochemistry was then performed to assess the presence or

degree of tauopathy in the PAG, KF and NRA of mice displaying normal or abnormal

USV patterns. Comparing various USV measurements, including the number, duration

and frequency of calls, revealed no differences between tau-P301L and wildtype mice

across all age groups, and linear discriminant analysis also failed to identify separate

USV populations. Finally, the presence of tauopathy in the PAG, KF and NRA in

individual tau-P301L mice did not reliably associate with USV disorders. Our findings

that tauopathy in designated mammalian vocalisation centres, such as the PAG, KF

and NRA, did not associate with USV disturbances in tau-P301L mice questions

whether USV phenotypes in this transgenic mouse are valid for studying tauopathy-

related human voice and speech disorders.

15

Introduction

Tauopathy is a common hallmark of many neurodegenerative diseases,

including Alzheimer’s disease, corticobasal degeneration and frontotemporal

dementia (FTD) with Parkinsonism (Avila et al., 2004; Lee et al., 2001; Foster et al.,

1997; Ingram and Spillantini, 2002; Spillantini and Goedert, 1998). It is characterized

by the aggregation of misfolded and hyperphosphorylated tau protein in the cytoplasm

of neurons, which results in aberrant interaction of tau protein with microtubules,

impairment of axonal transport and changes in the morphological and functional

integrity of neuronal synapses and somas (Lee et al., 2001; Foster et al., 1997; Ingram

and Spillantini, 2002; Millecamps and Julien, 2013; Spillantini and Goedert, 1998).

Frontotemporal dementia and Parkinsonism linked to Chromosome 17 (FTDP-

17) commonly arises from mutations in the microtubule-associated protein tau gene.

Of the 53 currently known pathogenic mutations linked to FTDP-17, one of the most

common is the P301L mutation (Ghetti et al., 2015). In humans, this mutation is

associated with cognitive decline (Geschwind et al., 2001) and non-cognitive

symptoms, such as behavioural and motor disturbances (Ingram and Spillantini,

2002).

Besides these neurological impairments, speech disorders are common

symptoms in FTDP-17 patients (Bird et al., 1999; Reed et al., 1997; Swieten et al.,

1999). FTDP-17-related speech disorders often manifest early in tauopathy and

progress to mutism (Williams, 2006). Thus, brainstem laryngeal motor control

pathways may undergo tauopathy and neurodegeneration. Although brainstem

tauopathy is receiving increasing recognition in postmortem studies (Eser et al., 2018;

Parvizi et al., 1998; Rüb et al., 2001; Thal, 2002), how brainstem tauopathy contributes

16

to the development and progression of speech disorders in FTDP-17 and other

tauopathy-related disorders is not well understood.

Here, we have studied whether the emergence and progression of irregular

ultrasonic vocalisation (USV) in transgenic tau-P301L mice carrying the P301L

mutation (Terwel et al., 2005) is predictive of the presence of tauopathy in brainstem

vocalisation centres. Tau-P301L mice develop respiratory dysfunction primarily

associated with paradoxical vocal cord movement in the larynx (i.e., constriction of

vocal fold during inspiration; Maschka et al., 1997; Patterson et al., 1974) at around 9

months of age, during the terminal phase of disease progression (Dutschmann et al.,

2010; Menuet et al., 2011a). Considering that the production of ultrasonic vocalisation

(USV) in rodents requires laryngeal airflow modulation via laryngeal adductors

(Dutschmann et a.., 2014; Laplagne, 2018; Riede, 2018; Riede, 2013), reports that

tau-P301L mice become mute at the terminal stage of disease progression (Menuet

et al., 2011b) are not surprising.

Irregular USV production and mutism in tau-P301L mice has been correlated with

tauopathy in the periaqueductal gray (PAG), Kölliker-Fuse nuclei (KF) and nucleus

retroambiguus (NRA) (Menuet et al., 2011b), regions of the midbrain and brainstem

that have been designated as vocalisation centres in mammals (Bandler and Carrive,

1988; Boers et al., 2002; Dutschmann and Dick, 2012; Holstege, 1989; Jürgerns and

Richter, 1986; Jürgerns, 2004; Jürgerns, 2002; Larson and Kistler, 1984; Yajima et al.,

1980; Zhang et al., 1995). Importantly, contrary to other mouse models of tauopathy

incorporating the P301L mutation (e.g., the rTg(tauP301L)4510 mouse model; Ramsden

et al., 2005), tau-P301L mice develop tauopathy predominantly in the midbrain,

brainstem and spinal cord (Dutschmann et al., 2010; Menuet al., 2011a,b; Terwel et

al., 2005; Terwel et al., 2008). Thus, it was concluded that mutism in tau-P301L mice

17

is closely associated to non-cognitive dysfunction of the laryngeal control circuit

(Menuet et al., 2011b).

The present study aimed to identify whether deficits in USV production are an

early predictor for disease onset that could serve as an indicator of tauopathy-related

phenotype progression, and a potential pre-clinical diagnostic for frontotemporal

dementia (Ryan, 2018). We investigated whether USV deficits in male tau-P301L mice

were associated with tauopathy in the PAG, KF and NRA using a previously described

experimental model of social male-female interaction (Bozdagi et al., 2010; Castellucci

et al., 2016; Castellucci et al., 2018; Hanson and Hurley, 2012; Oddi et al., 2014;

Portfors, 2007; Scattoni et al., 2011; Yang et al., 2012; Yang et al., 2013; Warren et

al., 2018). USV production in tau-P301L mice was examined at the following three age

periods: 1) 3-4 months, during which tau-P301L mice are asymptomatic, with no

deficits or tauopathy reported at this age (Terwel et al., 2005; Dutschmann et al., 2010;

Boekhoorn et al., 2006); 2) 5-6 months, a period prior to symptom onset in tau-P301L

mice that include respiratory-related deficits and the emergence of tauopathy from 7

months of age, but no apparent irregular locomotor activity (Terwel et al., 2005;

Dutschmann et al., 2010); and 3) 8-9 months, a symptomatic stage in which the

majority of tau-P301L suffer motor-related behavioural disturbances and present with

tauopathy (Terwel et al., 2005; Dutschmann et al., 2010; Lewis et al., 2000).

Materials and Methods

Animals

USVs were examined using 25 male wildtype FVB/N mice and 56 male

transgenic tau-P301L mice expressing the longest human tau isoform with the P301L

mutation (tau-4R/2N-P301L) under control of the mouse thy1 gene promoter (Terwel

et al., 2005). A further 40 female wildtype FVB/N mice were used during social male-

18

female interaction with male FVB/N wildtype or tau-P301L mice. Animals from the

same strain were group housed (4-5 mice per cage) and separated by gender and age

prior to testing, and singly housed 1h prior to the USV session. All animals were

maintained under a 12:12 h light/dark cycle, with free access to lab chow (Ridley

Corporation Limited) and tap water. All experiments followed protocols approved by

the Florey Institute of Neuroscience and Mental Health Animal Ethics Committee and

performed in accordance with the National Health and Medical Research Council of

Australia code of practice for the use of animals for scientific purposes.

USV recordings of tau-P301L and wildtype FVB/N mice were made from 3 age

groups in order to compare potential USV phenotypes at different stages of disease

progression. In search of an early phenotype in tau-P301L mice relating to

vocalisation, individual male mice aged 3-4 months (tau-P301L, n=23; FVB/N wildtype,

n=12) were used. At 5-6 months of age, USVs were recorded using n=35 male tau-

P301L and n=16 male FVB/N wildtype mice. Of these, USVs were recorded from n=24

tau-P301L and n=13 FVB/N wildtype mice during their first male-female USV recording

session, while the remainder of USVs (n=11 tau-P301L; n=3 FVB/N wildtype) were

obtained from mice used previously when aged 3-4 months. At 8-9 months of age,

USVs were recorded using n=31 male tau-P301L and n=11 male FVB/N wildtype

mice. Of these, USVs were recorded from n=9 tau-P301L mice during their first male-

female USV recording session, while the remainder of USVs (n=22 tau-P301L; n=11

FVB/N wildtype) were obtained from mice used previously when aged 5-6 or 3-4

months. Note that at both 5-6 months and 8-9 months, a student’s t-test revealed no

difference in the number of USVs produced by tau-P301L mice: 1) during their first

male-female USV recording session; and 2) from which USVs were recorded

previously when aged 5-6 or 3-4 months (data not shown). Similarly, at 5-6 months,

19

no difference was found when comparing the number of USVs produced by FVB/N

wildtype mice during their first male-female USV recording session and FVB/N

wildtype mice from which USVs were recorded previously when aged 3-4 months (data

not shown). In addition, repeated measurement analysis of USV production was

performed using male tau-P301L (n=10) and wildtype (n=20) mice when aged 5-6 and

8-9-months.

Experimental protocol for USV recordings

Social male-female interaction (Bozdagi et al., 2010; Castellucci et al., 2016;

Castellucci et al., 2018; Hanson and Hurley, 2012; Oddi et al., 2014; Portfors, 2007;

Scattoni et al., 2011; Yang et al., 2012; Yang et al., 2013; Warren et al., 2018) was

used to evoke USV in tau-P301L and corresponding wildtype mice that were not

sexually naïve. Given that the oestrous phase of female mice can influence USV

production during male-female interactions (Hanson and Hurley, 2012), female FVB/N

mice in the pre-oestrus and oestrus phase were used in all male-female interaction

sessions. The oestrous state of female mice was assessed using vaginal lavage (0.1-

0.2 ml, 0.9% saline) within 12 h of the USV recording session. The lavage was placed

onto a slide and dried (1-2 h, 37 ºC). Vaginal epithelial cells were then counter-stained

with Toluidine Blue O (Sigma-Aldrich, St. Louis, MO) and the oestrous cycle state was

determined (Cora et al., 2015). Both male and female mice were experienced in social

male-female interaction prior to participating in USV recording sessions. These

experiences were performed in an identical manner to the USV recording sessions, so

that the mice were habituated to the testing enclosure.

Prior to commencing the USV recording session, a male tau-P301L or FVB/N

mouse was placed in a clean cage with fresh litter and transferred to the recording

room to acclimatize for 1h before starting the recording session. At the start of the

20

recording session, a female wildtype FVB/N mouse was placed into the cage with the

male mouse. USVs of both male and female mice (see data analysis) were recorded

for 5 min. In rare cases (5%) where no USVs were detected during the male-female

interaction, the male mouse was used for an additional recording session during which

it interacted with a different female FVB/N mouse. Each female wildtype FVB/N mouse

was used twice a day for social male-female interaction. USVs were recorded using a

condenser ultrasound microphone UltraSoundGate CM16/CMPA (Avisoft Bioacustics,

Glienicke, Germany) and the Avisoft-RECORDER USGH software. The microphone

was suspended 15-20 cm over the centre of each cage for optimal recording of USVs.

All data were stored for offline analysis using Avisoft-SASLab Pro.

Immunohistochemistry

Tissue preparation

No later than 1 week after their USV recording session, mice were deeply

anaesthetized using sodium pentobarbitone (Ilium Pentobarbitone, Troy Laboratories,

Smithfield, NSW, Australia, 100 mg/kg i.p.) and perfused through the heart with 20 ml

Ca2+-free Tyrode’s buffer (37 ºC), followed by 20 ml 4% paraformaldehyde (Sigma-

Aldrich) containing 0.2% picric acid (Sigma) diluted in 0.16 M phosphate buffer (Merck

KGaA, Darmstadt, Germany; pH 7.2, 37 ºC) and 50 ml of the same fixative at 4ºC, the

latter for ~5 min. The brains were dissected and postfixed in the same fixative for 90

min at 4ºC, and finally immersed for 72 h in 15% sucrose dissolved in 0.1 M phosphate

buffer (pH 7.4) containing 0.01% sodium azide (Sigma) and 0.02% bacitracin (Sigma).

Coronal sections were cut using a freezing microtome (Leica Biosystems SM2010 R)

at a thickness of 40 µm, and stored in a cryoprotectant solution [30% v/v ethyleneglycol

(Merck); 15% w/v sucrose; 35% v/v 0.1M phosphate buffer; 35% v/v distilled H2O], at

−20°C.

21

Experimental protocol

Sections were washed in 0.01 M PBS and incubated for 24 h at room

temperature (RT) with a mouse anti-human paired helical filament-tau monoclonal

antibody (1:16,000 diluted from original concentration of 200 µg/ml; Thermo Fisher

Scientific, Scoresby, Vic, Australia; Product No. MN1020; Lot No. OA178994) diluted

in PBS containing 0.3% Triton X-100 and 0.5% BSA (Sigma). To visualize the

immunoreactivity, sections were processed using a commercial kit (TSA+, NEN Life

Science Products, Inc., Boston, MA), as described previously [53,54]. Briefly, sections

were washed in PBS followed by TNT buffer (0.1 M TRIS-HCl, pH 7.5; 0.15 M NaCl;

0.05% Tween20 Sigma), incubated with TNB blocking solution (TNT with 0.5% Dupont

blocking powder) for 1 hr, and incubated with donkey anti-mouse/horseradish

peroxidase conjugate (Jackson ImmunoResearch Laboratories, West Grove, PA)

diluted 1:750 in TNB for 1h at RT. Sections were then washed in TNT buffer and

incubated in a biotinyl tyramide-fluorescein (BT-FITC) conjugate diluted 1:100 in

amplification diluent for 10 min. Sections were then washed 3x in 15 min in TNT, and

2x in 10 min in 0.01 M PBS, mounted onto slides coated with 0.5% gelatin (Sigma)

and 0.05% chromium (III) potassium sulphate dodecahydrate (Merck), and

coverslipped using a fluorescent mounting medium (ProLong Glass with NucBlue,

Invitrogen, Eugene, Oregon).

Primary antisera used

Paired helical filament-Tau. Mouse anti-human paired helical filament-tau (PHF-

Tau) monoclonal antibody (clone AT8), raised against partially purified phosphorylated

human PHF-tau (Thermo Fisher Scientific, Scoresby, Vic, Australia; Product No.

MN1020; Lot No. OA178994). This antibody recognizes the phosphatase-sensitive

PHF-tau epitopes containing the phosphorylated Ser202/Thr205 residues, with a

22

molecular weight of approximately 79 kDA (Thermo Fisher). We detected no PHF-Tau

immunoreactivity in FVB/N wildtype mice (n=3).

Image processing

After processing, sections were examined using a Zeiss Observer.Z1

microscope (Oberkochen, Germany), equipped with a dark field condenser and epi-

polarization, epifluorescence with appropriate filter combinations, and objective lenses

of x10 (LD A-Plan, N.A. 0.25), and x20 (Plan-Apochromat, N.A. 0.8). Photographs

were taken using a AxioCam ICc5 digital camera (Zeiss), using ZEN 2012 (blue

edition) software (Zeiss).

Digital images from the microscopy were slightly modified to optimize for image

resolution, brightness and contrast using Adobe Photoshop CS6, version 13, software

(Adobe Systems Inc., San Jose, CA), so as to best represent the

immunohistochemistry observed at the microscope.

Data analysis

Ultrasonic Vocalisation

USV spectrograms were generated using a 256-bin Fast Fourier transform

applied to sliding windows with a 50% overlap. USVs were first automatically detected

by thresholding the total power at a given time point (Avisoft). Subsequently, USV calls

were examined manually to assure that all calls were detected and analysed correctly.

In order to compare our results with a previous study conducted in the same mice

model (Menuet et al., 2011b), the following USV parameters were assessed over the

5 min period of male-female interaction: 1) number of USVs per minute; 2) duration of

each USV (milliseconds); 3) total duration of all USVs during the 5 minute period of

male-female interaction (seconds per minute); 4) peak frequency of all USVs (KHz);

5) peak USV amplitude of all USVs (dB)/power; 6) frequency bandwidth; and 7) latency

23

to the first emitted USV after the female mouse was placed into the recording chamber

(log) (Menuet et al., 2011b; Scattoni et al., 2011; Yang et al., 2012; Castellucci et al.,

2016).

In a previous assessment of the number of USVs emitted by male or female mice

as they interacted, 84.5% of USVs were generated by male mice (Warren et al., 2018).

Thus, analysis of our data is based on the assumption that the majority of USVs were

generated by male mice during the male-female interaction.

Linear discriminant analysis (LDA)

Next, we tested whether USV features (call duration, frequency, bandwidth and

power) were predictive of the presence of tau overexpression by using linear

discriminant analysis (LDA) to perform supervised dimensionality reduction on the

dataset of all USV calls in this study (NWT=31,250; NTau-P301L = 74,134; Fig. 2.3). LDA

finds a projection of the data that best separates features belonging to known groups,

in our case, genotypes. Thus, if USV features are predictive of the presence of

tauopathy, LDA should find the function that best classifies USVs generated by

wildtype or tau-P301L mice. LDA was applied to the dataset using standard routines

in the scikit-learn (Pedregosa et al., 2011). To measure the confidence intervals (95%)

associated with LDA model parameters and to assess model classification accuracy,

we performed bootstrapped cross validation of the LDA model using 60% of the

dataset for training and 40% for testing model accuracy in 10,000 bootstrap-shuffled

LDA models (Table 1) (Mittal et al., 2018a,b).

Immunohistochemistry

Semi-quantitative comparative analyses for the expression of PHF-tau

immunoreactive cell bodies was performed on tau-P301L and wildtype mice aged 8-9

months. The severity of tauopathy, or density of PHF-tau immunoreactive cell bodies,

24

in the regions examined was categorized as: severe/high (+++); moderate (++);

weak/low (+); and absent immunoreactivity (-). Representative examples of these

categories are presented in Fig. 2.5B-D. Note that PHF-tau immunoreactivity was not

present in FVB/N wildtype mice, nor in tau-P301L mice prior to the age of 7 months

(data not shown) (Terwel et al., 2005). Thus, a potential relationship between USV

phenotype and PHF-tau immunoreactivity was only possible to examine in tau-P301L

animals aged 8-9 months.

Statistic

Since Kolmogorov-Smirnov tests indicated that our data were normally

distributed, we implemented a one-way analysis of variance (ANOVA) followed by a

post hoc Bonferroni multiple comparison test to determine statistical significance of

USV measurements between age-groups and genotypes.

A paired t-test compared age-dependent changes in the USV phenotype of

individual tau-P301L and wildtype FVB/N mice at 5-6 and 8-9 months. Values are

given as mean ± standard deviation (SD) and p-values less than 0.05 were accepted

to be statistically significant. All statistical analyses were performed using GraphPad

Prism, version 7.02 (GraphPad Software; San Diego, CA).

Results

USV is similar in tau-P301L and wildtype FVB/N mice

Overall, mice from both genotypes emitted similar USV patterns during social

male-female interaction at all age-groups (Fig. 2.1). Even symptomatic tau-P301L

mice that developed end-stage motor dysfunction (Fig. 2.1B; n=5) emitted USV

patterns similar to wildtype mice (Fig. 2.1A) and asymptomatic/pre-symptomatic tau-

P301L mice.

25

Figure 2.1. Representative spectrograms of USV sequences emitted during social male-female

interaction.

(A) 9-month-old wildtype male FVB/N mouse enclosed with a female wildtype FVB/N mouse; (B) 9-

month-old tau-P301L mouse enclosed with a female wildtype FVB/N mouse. Note that the tau-P301L

mouse presented with end-stage gait disturbance and was still able to produce wildtype-like USV

sequences during the 5 min of male-female interactions.

Further analysis of USV parameters included an examination of the: 1) number

of USVs per∙min (Fig. 2.2A); 2) mean duration of each individual USV (Fig. 2.2B); 3)

total time of all USVs during the 5 min period of male-female interaction (Fig. 2.2C);

peak frequency of all USVs (KHz, Fig. 2.2D); peak amplitude of all USV calls (dB, Fig.

2.2E); and latency to the first USV emitted by male mice after the female mouse was

placed into the recording chamber (Fig. 2.2F). ANOVA revealed no difference in any

of these USV parameters between wildtype and tau-301L mice at any of the three time

points examined (Fig. 2.2).

26

Figure 2.2. Ultrasonic vocalisation measurements of wildtype FVB/N (grey circles) and tau-

P301L (magenta triangles) mice across the asymptomatic (3-4 months), pre-symptomatic (5-6

months) and symptomatic (8-9 months) age groups.

(A) Number of USVs per minute; (B) mean duration of each USV in milliseconds; (C) total duration of

all USVs emitted during the 5 min male-female interaction, in seconds per minute; (D) peak frequency

in kilohertz; (E) peak amplitude in decibels; and (F) latency to emit the first call. Note that a log-

transformation was applied in (F) for better visualisation of the data distribution. All values are expressed

as mean ± SD.

Because individual mice from both wildtype and tau-P301L cohorts displayed

USV measurements that appeared to be outliers (see Fig. 2.2), we used linear

discriminant analysis (LDA) applied to the dataset of all USVs to further assess

whether USV features were predictive of the presence of tau overexpression. LDA

finds a linear function that best discriminates two or more independent, Gaussian-

distributed classes. Thus, if USV features were predictive of tauopathy, we would

expect that wild-type versus Tau-P301L mice would be well separable in the LDA-

defined feature space. In Fig. 2.3A, we show the scatter-plot matrix of USV features

used to train LDA models: USV duration, peak frequency, frequency bandwidth and

27

power. In all dimensions, individual USV features were largely overlapping in their

original feature space. Applying LDA as a dimensionality reduction technique (using

all data for LDA training) showed that USVs from wildtype and tau-P301L mice also

overlapped in an LDA-defined feature space suggesting that USV features were not

predictive of the presence of tau-overexpression (Fig. 2.3B). Further, to derive the 95%

confidence intervals of the LDA model and to assess its classification accuracy, we

performed bootstrapped cross validation of the LDA model by generating 10,000 LDA

models each trained on a different subset (60%) of the original dataset. LDA model

parameters and classification accuracy are shown in Table 2.1. Bootstrapped LDA

model parameters (Table 1) were similar to those of the overall LDA model (Fig. 2.3B).

Moreover, the LDA classification accuracy was 69.636%, which simply reflected the

distribution of wildtype versus tau-P301L mice in the dataset. Taken together, the USV

data generated from male-female interaction cannot be clustered, supporting our

overall conclusion that the USV production of both genotypes is not different. This is

the case even in mice at the symptomatic stage (8-9 month), where general motor

symptoms (partial hind limb clasping and gait disturbance) are prominent.

28

Figure 2.3. USV features do not predict the presence of tau-overexpression.

To test whether USV features were predictive of the presence of tau-overexpression, we used linear

discriminant analysis to perform supervised dimensionality reduction/classification of USV features

(syllable duration, peak frequency, frequency bandwidth and syllable power). (A) This scatterplot matrix

shows the distributions of all USV features produced by wildtype (blue; 31,250 USVs) or tau-P301L

(red; 74,134 USVs) mice. While most parameters showed overlapping distributions, frequency

bandwidth appeared bi-modal. Therefore, we applied linear discriminant analysis to assess whether

there existed a linear discriminant that could accurately classify wildtype versus tau-P301L mice. (B)

The distribution of linear discriminant scores is shown for all wildtype and tau-P301L USVs. Like their

representation in the original 4-dimensional feature space, the LD scores of wildtype and tau-P301L

mice overlapped suggesting that USV features were not predictive of the mouse genotype.

Table 2.1. Linear discriminant model and accuracy.

βsyllable duration βmean frequency βfrequency bandwidth βsyllable power Accuracy (%)

2.865 × 10-3 ±

1.696 × 10-6

(2.862 × 10-3 –

2.868 × 10-3)

4.317 × 10-2 ±

1.148 × 10-5

(4.314 × 10-2 –

4.319 × 10-2)

-1.264 × 10-2 ±

4.003 × 10-6

(-1.265 × 10-2 –

-1.263 × 10-2)

1.160 × 10-2 ±

7.167 × 10-6

(1.158 × 10-2 –

1.161 × 10-2)

69.636 ±

1.697 × 10-2

(69.632 –

69.639)

Repetitive recording reveals a decreased number of USVs emitted by

symptomatic tau-P301L and wildtype mice aged 8-9 months

29

Since our observation of the lack of genotype and/or age-related deterioration in

the generation of USV during male-female interaction is in contrast to a previous report

(Menuet et al., 2011b), we also performed experiments with tau-P301L and wildtype

mice in which USVs emitted during male-female interaction were first recorded at the

age of 5-6 months, and repeated at 8-9 months. Analysis of age-related and genotype-

related USV in this manner revealed a decreased number of USVs in both

symptomatic tau-P301L (p = 0.037) and wildtype FVB/N (p = 0.014) mice aged 8-9

months versus the number of USVs they generated when aged 5-6 months (Fig. 2.4).

When assessing other USV parameters, comparisons between tau-P301L or wildtype

mice when aged 4-5 months, and subsequently at 8-9 months, also revealed

diminished total duration of all USVs during the 5 min period of male-female interaction

and peak amplitude/power (data not shown). In addition, peak frequency and latency

to first USV emitted after the female mouse was placed into the recording chamber

were reduced in 8-9 month-old tau-P301L mice, but not wildtype FVB/N mice (data not

shown).

30

Figure 2.4. Age-dependent changes in the number of USVs/min in individual (A) wildtype FVB/N

and (B) tau-P301L mice.

USVs during male-female interaction were first recorded at the age of 5-6 months, and repeated at 8-9

months (paired t-test, * p = 0.014, ** p = 0.037).

Tauopathy in the midbrain periaqueductal gray, pontine Kölliker-Fuse

nucleus and medullary nucleus retroambiguus is not predictive of USV

disorders.

We next verified the severity of tauopathy in tau-P301L mice and the density of

PHF-tau immunoreactive neurons in the: 1) PAG, KF and NRA - designated midbrain-

brainstem vocalisation areas; and 2) forebrain, brainstem and spinal cord, as a general

measure of the distribution of tauopathy in the central nervous system of the tau-

P301L mice examined. Immunohistochemistry was performed on 8-9 month-old

symptomatic tau-P301L mice (n=14) in which partial hindlimb clasping and gait

disturbances were obvious, and in which the number of USVs emitted during male-

female interaction was either hypoactive (0-35 USVs per minute), hyperactive (>260

USVs per minute), or average (90-205 USVs per minute). Similarly, the number of

USVs emitted by 8-9 month-old wildtype FVB/N mice on which immunohistochemistry

was performed (n=3), ranged from 15-250 USVs per minute (Fig. 2.5).

31

Figure 2.5. The relationship between USVs emitted and tauopathy in 8-9 month-old tau-P301L

and FVB/N wildtype mice.

(A) The number of USVs emitted by tau-P301L mice during male-female interaction, and severity of

PHF-tau immunoreactivity in the periaqueductal gray (PAG), Kölliker-Fuse nucleus (KF) and nucleus

retroambiguus (NRA), brainstem, spinal cord and forebrain. Scoring criteria for the severity of tauopathy

is based on a 4 point scale of the density of PHF-tau immunoreactive neurons: severe/high (+++),

moderate (++), weak/low (+) and absence of immunoreactive neurons (-). (B-D) Representative

photographs of PHF-tau immunoreactivity, illustrating the scoring criteria for the severity of tauopathy

in tau-P301L mice: (B) +, low tauopathy; (C) ++, moderate tauopathy; (D) +++, severe tauopathy.

Abbreviations: Aq, aqueduct (Sylvius); KF, Kölliker-Fuse nucleus; mcp, middle cerebellar peduncle;

NRA, nucleus retroambiguus; PAG, periaqueductal gray; scp, superior cerebellar peduncle. Scale bar

in B = 200 µm, applies B-D.

32

Figure 2.6. PHF-Tau immunoreactivity (green) in the periaqueductal gray of 8-9 month-old tau-

P301L mice.

(A, B) Absent or low density of PHF-Tau immunoreactive cell bodies in the PAG of a tau-P301L mouse

that displayed (A) average-hyperactive (animal 4, Fig. 2.5) or (B) hypoactive (animal 12, Fig. 2.5) USV

emission during male-female interaction. (C, D) Severe tauopathy, and high density of PHF-tau

immunoreactive cell bodies in the PAG of a tau-P301L mouse that displayed (C) hyperactive (animal

16, Fig. 2.5) or (D) hypoactive (animal 15, Fig. 2.5) USV emission during male-female interaction.

Abbreviations: Aq, aqueduct (Sylvius); PAG, periaqueductal gray. Scale bar in A = 500 µm, applies A-

D.

In tau-P301L mice, no relationship was found between tauopathy in the PAG

(Figs. 2.5-2.6), KF (Figs. 2.5, 2.7) or NRA (Figs. 2.5, 2.8) and the observed USV

33

phenotype. Of the tau-P301L mice (n=6) in which severe-moderate tauopathy was

present in the PAG (Figs. 2.5, 2.6C-D) and KF (Figs. 2.5, 2.7C-D), and moderate-weak

tauopathy was detected in the NRA (Figs. 2.5, 2.8C-D), only one (animal 15, Fig. 2.5)

of these six mice was silent during male-female interaction. The USV phenotype in the

remaining five tau-P301L mice was either: average (animals 9-11, Fig. 5) or

hyperactive (animals 16-17, Fig. 2.5). Contrasting these observations, of the eight tau-

P301L mice in which no PHF-tau immunoreactivity was found in the PAG (Figs. 2.5,

2.6A-B), KF (Figs. 2.5, 2.7A-B) and NRA (Figs. 2.5, 2.8A-B), three animals displayed

a hypoactive USV phenotype during male-female interaction (animals 12-14, Fig. 2.5).

The number of USVs emitted by the remaining five animals was in the average range

(animals 4-8, Fig. 2.5). As expected, PHF-Tau immunoreactivity was not detectable in

any WT mice (animals 1-3, Fig. 2.5).

34

Figure 2.7. PHF-Tau immunoreactivity (green) in the Kölliker-Fuse nucleus of 8-9 month-old tau-

P301L mice.

(A, B) Absent or low density of PHF-tau immunoreactive cell bodies in the KF of a tau-P301L mouse

that displayed (A) average (animal 4, Fig. 2.5) or (B) hypoactive (animal 12, Fig. 2.5) USV emission

during male-female interaction. (C, D) Severe tauopathy, and high density of PHF-tau immunoreactive

cell bodies in the KF of a mouse that displayed (C) hyperactive (animal 16, Fig. 2.5) or (D) hypoactive

(animal 15, Fig. 2.5) USV emission during male-female interaction. Abbreviations: KF, Kölliker-Fuse

nucleus; mcp, middle cerebellar peduncle; scp, superior cerebellar peduncle. Scale bars: A = 200 µm,

applies A-D.

35

Figure 2.8. PHF-Tau immunoreactivity (green) in the nucleus retroambiguus of 8-9 month-old

tau-P301L mice.

(A, B) Absent or low density of PHF-tau immunoreactive cell bodies in the NRA of a tau-P301L mouse

that displayed (A) average (animal 4, Fig. 2.5) or (B) hypoactive (animal 12, Fig. 2.5) USV emission

during male-female interaction. (C, D) Severe tauopathy, and high density of PHF-tau immunoreactive

cell bodies in the NRA of a mouse that displayed (C) hyperactive (animal 16, Fig. 2.5) or (D) hypoactive

(animal 15, Fig. 2.5) USV emission during male-female interaction. Abbreviations: 12N, hypoglossal

nucleus; NRA, nucleus retroambiguus; pyx, pyramidal decussation. Scale bars: A = 500 µm, applies A-

D.

36

Discussion

The present study investigated USV in male tau-P301L mice, a mouse model of

tauopathy in human frontotemporal dementia (Terwel et al., 2005). We examined USV

at the following 3 stages of disease progression, based on the progressive motor

impairment that occurs in tau-P301L mice over time: tau-P301L mice aged 3-4

months, when animals are asymptomatic (Terwel et al., 2005; Dutschmann et al.,

2010); 5-6 months, a pre-symptom onset period prior to the emergence of tauopathy

and motor deficits (Terwel et al., 2005); and 8-9 months, a symptomatic stage when

motor-related disturbances are obvious and tauopathy is present (Terwel et al., 2005).

Overall, analysis of our USV recordings from tau-P301L or wildtype FVB/N mice

did not identify significant changes in any of the USV parameters examined at any of

these time points or stages of disease progression. Only a repeated assessment of

USV in tau-P301L mice at pre-symptomatic versus symptomatic ages showed a

significantly reduced number of USVs in the symptomatic cohort, an observation also

detected in wildtype FVB/N mice. These latter age-related results are in agreement

with a previous study that reported a severe decrease in USV emission, and mutism

of male tau-P301L mice aged 8-10 months (Menuet et al., 2011b).

Our data assumes the majority of USVs were emitted by male mice during the

male-female interaction, an assumption supported by previous work reporting that 82-

85% of all USVs were generated by male mice as they freely interacted with female

mice (Warren et al., 2018; Neunuebel et al., 2015; Heckman et al., 2017). Further

support that the majority of USVs are derived from male mice comes from studies

utilizing devocalised male mice, a condition induced by sectioning of the inferior

laryngeal nerve. In this experiment setting, no USVs were detected from female mice

during interaction with devocalized male mice (Warburton et al., 1989; Sugimoto et al.,

37

2011). On the other hand, a recent study has shown that when only one partner in a

male-male rat interaction can vocalize, the vocal partner increases its call rate (Burke

et al., 2018). Therefore, it remains unclear whether female FVB/N mice may have

increased their USV emission in response to a hypoactive male mouse and therefore

could have masked a potential USV phenotype of our male Tau-P301L mice.

However, additional test from our experiments showed a lack of USV emission from

female mice following the removal of the male mouse from the male-female

experimental setting, while conversely, after removal of the female mouse, USV

production by the male mouse continued (data not shown). Thus, we still remain

confident that the majority of USVs analysed in our study were produced by male mice.

In previous work, mutism of tau-P301L was found to correlate with severe

tauopathy in designated midbrain/brainstem vocalisation centres, such as the PAG

and KF (Menuet et al., 2011b). However, in our tau-P301L mice, analysis of PHF-Tau

immunoreactivity (a standard marker that identifies tauopathy at the motor-

symptomatic stage of disease progression) failed to reveal any relationship between

irregular USV and tauopathy in the midbrain PAG, pontine KF and medullary NRA.

The reasons underlying this discrepancy between our results of those of Menuet et al.

(2011b) are potentially multi-factorial.

Vocalisation is a complex behaviour influenced by numerous innate and

environmental factors (Granon et al., 2018), which could have influenced the outcome

of our study. Thus, subtle differences in the general housing conditions, such as the

level of environment enrichment, can influence USV production in genetically

homogeneous mice (Brenes et al., 2016). Nevertheless, the overall USV pattern

(average call duration, frequency band, etc.) and amount of USV activity of tau-P301L

and FVB/N mice in the present study is consistent with previous reports (Menuet et

38

al., 2011b; Oddi et al., 2014; Yang et al., 2013). Moreover, the USV activity in younger

male tau-P301L reported by Menuet et al. (2011b) was almost identical to our study.

Contrary to our experimental protocol, the female mice used previously by

Menuet et al. (2011b) were not assessed for their phase in the oestrus cycle, despite

evidence that the female oestrus stage has strong influence on the motivation of a

male rodent to emit USV (Barthelemy et al., 2004; Hanson and Hurley, 2012; McGinnis

and Vakulenko, 2003). Thus, the general motivation of male tau-P301L mice to emit

USV may have been significantly stronger in our study. While such external biological

factors may have contributed to the different outcome of both studies, it is important

to note that similar differences have been observed in other transgene mouse strains.

Previous investigations of USV phenotypes in Shank3 mutant mice, a mouse model

of autism spectrum disorder, identified an impaired USV phenotype (Bozdagi et al.,

2010), while a second group could not confirm these results (Yang et al., 2012).

Similarly, the absence or presence of a USV phenotype was reported by two

independent studies that investigated USV in a mouse model of autism spectrum

disorder that carries a null mutation in the neuroligin4 gene (Ey et al., 2012; Jamain et

al., 2008).

Surprisingly, our results show that even aberrant USV in individual mice did not

predict the presence of tauopathy in the PAG, KF and NRA. A potential explanation

for the discrepancy between our findings and previous results where tauopathy in the

PAG and KF was identified (Menuet et al., 2011b) could be the use of antibodies

derived from alternative sources. Previously, AT8 and AT100 monoclonal antibodies

directed against phosphorylated human protein tau at epitopes pS198/pS202/pS205

and pT231/pS235 were used (Menuet et al., 2011b), while results from our study were

derived from a monoclonal AT8 antibody targeting phosphorylated human PHF-tau at

39

epitopes pS202/pT205. Despite this variation, our observation of prominent PHF-tau

immunoreactivity in the spinal cord and brainstem of all analysed tau-P301L mice at

the motor-symptomatic age of 8-9 months is consistent with previous studies

(Dutschmann et al., 2010; Menuet et al., 2011a,b; Terwel et al., 2005; Terwel et al.,

2008), and provides evidence that the PHF-tau antibody used in our study is a robust

marker of tauopathy. This is further validated by our observation of the lack of PHF-

tau immunoreactivity in wildtype mice, and no, or minimal immunoreactivity in

asymptomatic or pre-symptomatic tau-P301L mice (data not shown). Thus, our data

showing no relationship between aberrant USV and PHF-tau immunoreactivity

suggest that tauopathy, especially in the PAG-NRA axis, a critical motor pathway for

vocalisation (Zhang et al., 1995; Jürgens and Richter, 1986; Holstege, 1989; Oka et

al., 2008; Gerrits and Holstege, 1996), or the KF, an important pre-motor area for

laryngeal adductor control (Dutschmann and Dick, 2012; Dutschmann and Herbert,

2006), are not predictive of USV disorders in individual tau-P301L mice.

Possible implications of these findings are that neither the PAG, KF nor NRA are

important for the mediation of USV. In particular, the PAG is well-established as a

common final pathway for vocalisation in mammals (Jürgens and Richter, 1986;

Jürgens, 1994; Jürgens, 2002a,b; Larson and Kistler, 1984; Holstege, 1989; Jürgens

and Pratt, 1979; Düsterhöft et al., 2004), including humans (Esposito et al., 1999), that

do not utilize USV for communication. Thus, the role of the PAG in the mediation of

USV in rodents could be controversial. Against this notion, lesions of the dorsal PAG

in rat completely abolished fear- and pain-related 22kHz vocalisations (Leman et al.,

2003). While in adult mice, USVs at slightly higher frequencies of 30kHz are produced

under fear-induced (Ko et al., 2005), and restraint stress (Chabout et al., 2012)

conditions, and 37kHz USVs have been reported in response to chronic pain (Kurejova

40

et al., 2010), the pathways involved in mediating such USVs are not well understood.

Furthermore, whether a PAG-dependent deficit of fear- and pain-related vocalisation

occurs in tau-P301L mice requires further investigation. In relation to the mediation of

50kHz USVs in both mouse and rat, the role of the PAG in is presently unclear,

although lesions or stimulation of components of the central reward circuits, and

particularly the ventral tegmental area, was shown to modulate or reduce 50kHz USV

in rat (Burgdorf et al., 2007). Thus, rodent 50kHz USV could be mediated by different

vocalisation circuits that do not essentially require the functional integrity of the PAG,

KF or NRA.

A repertoire of at least ten mouse-specific USV categories have been classified,

including complex, harmonics, two-syllable, frequency steps, upward and downward

USVs (Scattoni et al., 2008). Our results, which failed to identify significant changes

between wildtype FVB/N mice and tau-P301L mice at varying degrees of disease

progression, were based on data that did not discriminate between USV categories

(see e.g. the presence of complex and frequency steps USVs in both wildtype and tau-

P301L mice in Fig. 1.1). Thus, the potential that specific categories of USV are affected

in tau-P301L mice remains a possibility, and is an avenue for further investigation.

This may be important in the context of the identified links between classes of USVs

and discrete respiratory patterns (Castellucci et al., 2018), and the aberrant breathing

patterns and shortening of the expiratory interval reported in tau-P301L mice aged 7-

8 months (Dutschmann et al., 2010).

Finally, the fact that a USV phenotype was not confirmed in our study does not

mean that tau-P301L mice do not develop laryngeal dysfunction that is expressed, for

example, during stationary breathing (Dutschmann et al., 2010; Menuet et al., 2011b).

Indeed, a recent study has shown that glycine transporter-2 knock-out (GlyT2-KO)

41

mice develop paradoxical vocal cord movement in the larynx (Hülsmann et al., 2018),

a general phenotype that strongly overlaps with the respiratory deficits in the tau-

P301L mouse strain (Dutschmann et al., 2010; Menuet et al., 2011b). Interestingly,

the emission of USV in GlyT2-KO was also indistinguishable from USVs of the

associated wildtype mice (Hülsmann et al., 2018). Importantly, Hülsmann et al. (2018)

identified compensatory mechanisms for the unchanged USV phenotype in GlyT2-KO

mice. The wildtype mice produced USV during the early expiratory phase (or

postinspiratory phase) when critical laryngeal adductor activity modulates expiratory

airflow and when vocalisation usually takes place (Dutschmann et al., 2014). In

contrast, GlyT2-KO mice shifted their USV calls to late expiration and used forced

(abdominal) expiratory airflow to vocalize (Hülsmann et al., 2018). Thus, further

investigation of similar mechanistic compensation for tauopathy in laryngeal motor

pathways in tau-P301L mice is required to understand common voice and speech

disorders associated with tauopathy-related neurodegenerative disease.

Conclusion

Contrary to a previous study, our present experiments did not reveal a significant

USV disorder in male tau-P301L mice during an experimental protocol of male-female

interaction, even at late stages of progressing tauopathy (8-9 months of age). The

different outcome compared to the previous study might originate in the stronger

stimulus for male USV, which were triggered in our study by the presence of female

mice in oestrous. The fact that tauopathy in the midbrain PAG, pontine KF and

medullary NRA in tau-P301L mice did not reliably associate with irregular USV in

individual mice also raises the question of whether USV phenotypes in this transgenic

mouse model are valid for studying tauopathy-related human voice and speech

disorders.

42

Chapter 3

Forebrain projections neurons target functionally diverse

respiratory control areas in the midbrain, pons and medulla

oblongata

Pedro Trevizan-Baú1, Rishi R. Dhingra, 1Werner I. Furuya1, Davor Stanic1, Stuart B.

Mazzone2, Mathias Dutschmann1

1 The Florey Institute of Neuroscience and Mental Health, The University of Melbourne,

Parkville, Victoria 3010, Australia.

2 Department of Anatomy and Neuroscience, The University of Melbourne, Parkville,

VIC 3010, Australia.

The following chapter was published by Journal of Comparative Neurology on

December 19, 2020 (DOI: 10.1002/cne.25091). The full paper has been inserted as a

thesis chapter in accordance with The University of Melbourne guidelines.

43

Abstract

Eupnea is generated by neural circuits located in the ponto‐medullary brainstem, but

can be modulated by higher brain inputs which contribute to volitional control of

breathing and the expression of orofacial behaviours, such as vocalisation, sniffing,

coughing, and swallowing. Surprisingly, the anatomical organization of descending

inputs that connect the forebrain with the brainstem respiratory network remains poorly

defined. We hypothesized that descending forebrain projections target multiple

distributed respiratory control nuclei across the neuroaxis. To test our hypothesis, we

made discrete unilateral microinjections of the retrograde tracer cholera toxin subunit

B in the midbrain periaqueductal gray (PAG), the pontine Kölliker‐Fuse nucleus (KFn),

the medullary Bötzinger complex (BötC), pre‐BötC, or caudal midline raphé nuclei. We

quantified the regional distribution of retrogradely labeled neurons in the forebrain 12–

14 days postinjection. Overall, our data reveal that descending inputs from cortical

areas predominantly target the PAG and KFn. Differential forebrain regions innervating

the PAG (prefrontal, cingulate cortices, and lateral septum) and KFn (rhinal, piriform,

and somatosensory cortices) imply that volitional motor commands for vocalisation are

specifically relayed via the PAG, while the KFn may receive commands to coordinate

breathing with other orofacial behaviours (e.g., sniffing, swallowing). Additionally, we

observed that the limbic or autonomic (interoceptive) systems are connected to

broadly distributed downstream bulbar respiratory networks. Collectively, these data

provide a neural substrate to explain how volitional, state‐dependent, and emotional

modulation of breathing is regulated by the forebrain.

44

Introduction

Studies on volitional control of breathing, which were mostly performed in

humans, indicate that corticospinal pathways bypass the respiratory network of the

brainstem and directly modulate respiratory motor pools in the spinal cord

(Butler, 2007; Corfield et al., 1998; Gandevia and Rothwell, 1987a,b; Pouget et

al., 2018). In mammals, corticospinal pathways that specifically target spinal

respiratory motor pools have been demonstrated (Rikard‐Bell et al., 1985). However,

descending anatomical pathways that target nuclei of the primary respiratory rhythm

and pattern generating network remain poorly defined despite functional evidence that

activity within the respiratory network is modulated by behavioural commands

(Chang, 1992; Orem and Netick, 1986). Cognitive, emotional, and behavioural motor

commands modulate respiratory activities during a variety of volitional orofacial

behaviours including vocalisation (speech), swallowing, chewing, coughing, sneezing,

emotional sighing, and sniffing (Davis et al., 1996; Deschênes et al., 2012;

Jean, 2001; Laplagne, 2018; Li et al., 2020; Ludlow, 2015; McElvain et al., 2018;

Moore et al., 2013; Moore et al., 2014; Nonaka et al., 1999; Sherwood et al., 2005).

Many of these orofacial behaviours depend on the recruitment of upper airway

muscles that regulate airway patency during inspiration and expiration (Dutschmann

and Paton, 2002). Control of expiratory airflow during the postinspiratory phase of

respiration is essential for the mediation of vocalisation, swallowing and expulsive

expiratory behaviour (Dutschmann et al., 2014). Since the primary motor networks that

control postinspiration are distributed in the brainstem (Dhingra, et al., 2019a,b;

Dhingra et al., 2020), we hypothesized that crucial descending forebrain projections

target respiratory control nuclei above the spinal cord. In addition, we assumed that

orofacial behaviours that involve augmented inspiratory activity (e.g., sighing, sniffing)

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45

may also recruit brainstem nuclei that generate and/or modulate the inspiratory

rhythm, instead of overwriting ongoing respiratory activity with corticospinal motor

commands. This idea is supported by a recent study showing supra‐pontine inputs to

the pre‐Bötzinger complex (pre‐BötC), the brain area that generates the inspiratory

rhythm (Yang et al., 2020).

To test our working hypothesis, we used the retrograde tracer cholera toxin

subunit B (CT‐B) to characterize the topography of descending monosynaptic

projections from the forebrain to five anatomically distinct respiratory areas in the

midbrain and ponto‐medullary brainstem that have established function in the

modulation or generation of respiratory activity. The descending forebrain connectivity

of the periaqueductal gray (PAG) was analysed because it has a profound role in the

modulation of respiration during vocalisation and defensive behaviours

(Dampney, 2015; Faull et al., 2019; Subramanian, Balnave, and Holstege, 2008;

Subramanian and Holstege, 2014; Subramanian, Huang, and Balnave, 2008; Zhang

et al., 1994), but has no breath‐by‐breath role in respiratory rhythm and pattern

generation (Farmer et al., 2014). Because the PAG has established connectivity with

various forebrain nuclei (Dampney et al., 2013), it also serves as an important control

for the analysis of the additional respiratory brain areas studied. The pontine Kölliker‐

Fuse nucleus (KFn) was investigated because it regulates the inspiratory–expiratory

phase transition and is involved in the breath‐by‐breath formation of the respiratory

motor pattern (Caille et al., 1981; Dutschmann and Herbert, 2006; Mörschel and

Dutschmann, 2009; Smith et al., 2007; Wang et al., 1993). Moreover, the KFn has

major implications in the control of laryngeal adductor function during breathing

(Dutschmann and Herbert, 2006) and orofacial behaviours (Dutschmann and

Dick, 2012). The pre‐BötC was chosen for its essential role in inspiratory rhythm

















46

generation (Del Negro et al., 2018; Feldman and del Negro, 2006; Smith et al., 1991)

and the neighbouring BötC was targeted because of its proposed function as an

essential part of respiratory rhythm generating circuit (Burke et al., 2010; Marchenko

et al., 2016; Smith et al., 2013). Finally, descending forebrain inputs to nuclei of the

caudal raphé were analysed since these serotonergic neurons have important

neuromodulatory action on the respiratory motor pattern (Besnard et al., 2009; Hodges

and Richerson, 2010; Holtman et al., 1986; Lindsey et al., 1998; Richter et al., 2003).

Material and Methods

Animals

Adult Sprague–Dawley rats of either sex (n = 28, weight range: 280–350 g)

were used for this study. All animals were housed under a 12:12 h light/dark cycle,

with free access to lab chow (Ridley Corporation Limited, Australia) and water.

Experiments followed protocols approved by the Florey Institute of Neuroscience and

Mental Health Animal Ethics Committee and performed in accordance with the

National Health and Medical Research Council of Australia code of practice for the

use of animals for scientific purposes.

Surgery

For tracer microinjections, rats were initially anesthetized with isoflurane (5%

vol/vol in oxygen). After mounting in a stereotaxic apparatus (TSE systems, Bad

Homburg, Germany), anesthesia was maintained with isoflurane (~2% in oxygen) via

a nose cone. After the rats were placed with the skull in a flat position in the stereotaxic

frame, a craniotomy was performed. Surgeries were performed under an aseptic

technique (iodine antiseptic solution). A midline incision exposed the skull between

bregma and the interaural line, and a small burr hole was drilled according to

coordinates defined relative to bregma (Table 1). Using a 1 μl Hamilton syringe (25 s‐












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47

gauge needle), 150 nl of 1% CT‐B (1 mg/ml; Invitrogen, OR) was pressure injected

unilaterally in the following brain nuclei: PAG (dorsolateral and ventrolateral

columns; n = 5, plus 3 near‐miss injections), KFn (n = 5, plus 2 near‐miss injections),

BötC (n = 3, plus 1 near‐miss injection), pre‐BötC (n = 3, plus 1 near‐miss injection),

and caudal raphé nuclei (raphé pallidus [RPa], raphé magnus [RMg], and raphé

obscurus [ROb]; n = 3, plus 2 near‐miss injections). The total volume was injected at

a rate of 20 nl/min, and CT‐B injections were made on the left side of the brain. To

avoid bleeding after injection through the transverse sinus for medullary targets (for

BötC, pre‐BötC, and caudal raphe injections), the syringe was angled and inserted

from a more rostral location to reach the specific coordinates (for details, see

Table 3.1). After the injection, the syringe remained in the brain tissue for at least 10

min and was withdrawn at 1 mm/min to minimize non‐specific spread of the tracer in

brain tissue along the injection tract (Finkelstein et al., 2000). Immediately after

surgery, animals received 3 mg/kg of the anti‐inflammatory drug Meloxicam (Tory‐

Ilium, NSW, Australia; 5 mg/ml, delivered subcutaneously). The animals were allowed

to recover for 12–14 days before transcardial perfusion.

Table 3.1. Stereotaxic coordinates for CT-B microinjections.

Brain

regions

Stereotaxic coordinates (mm)

micropipette angle4 Rostrocaudal1 Mediolateral2 Dorsoventral3

Kölliker-Fuse -9.0±0.1 +2.6 -6.5 0°

Pre-Bötzinger -9.5 +2.0 -9.5 22°

Bötzinger -9.0 +2.0 -9.5 22°

Caudal raphé -8.7 0.0 -9.4 22°

Periaqueductal

gray

-8.0±0.5 +0.8 -4.8±0.5 0°

1relative to Bregma; 2relative from midline; 3from dorsal surface of the brain; 4relative

to vertical.


48

Tissue preparation and immunohistochemistry

Twelve to fourteen days after injection, the rats were deeply anesthetized with

sodium pentobarbitone (Ilium Pentobarbitone, Troy Laboratories, Smithfield, NSW,

Australia, 100 mg/kg i.p.) and transcardially perfused with 60 ml Ca2+‐free Tyrode's

buffer (37°C), followed by 60 ml 4% paraformaldehyde (PFA, Sigma‐Aldrich)

containing 0.2% picric acid (Sigma) diluted in 0.16 M phosphate buffer (Merck KGaA,

Darmstadt, Germany; pH 7.2, 37°C) and finally, an additional 300 ml PFA/picric acid

solution at 4°C. Brains were removed from the skull and post‐fixed in PFA/picric acid

solution for 90 min at 4°C. Next, the brains were immersed for 72 h in 0.1 M phosphate

buffer (pH 7.4) containing 15% sucrose, 0.01% sodium azide (Sigma) and 0.02%

bacitracin (Sigma) for cryoprotection. A small incision from the olfactory bulbs to the

brainstem was made in the hemisphere contralateral to the CT‐B injection site to allow

for subsequent measurements of the ipsilateral and contralateral distribution of

projection neurons (Fig 3.2). After post fixation and cryoprotection, brains were rapidly

frozen using liquid carbon dioxide. Finally, brains we stored at −80°C until

cryosectioning.

Serial coronal sections (40 μm thickness) of the entire brain (from the olfactory

bulb to the spino‐medullary junction) were cut using a cryostat (Leica CM1850, Leica

Microsystems), and stored in a cryoprotectant solution (30% vol/vol ethylene glycol

[Merck]; 15% wt/vol sucrose; 35% vol/vol 0.1 M phosphate buffer; 35% vol/vol distilled

H2O) at −20°C. Brain sections were cut serially, and sections processed for

immunohistochemistry were separated by 120 μm. Freely floating sections were

washed in 0.01 M PBS, followed by incubation in hydrogen peroxide for 20 min to block

endogenous peroxidase activity. Next, sections were incubated for 24 h at room

temperature (RT) with a goat anti‐CT‐B antibody (List Biological Laboratories, CA;

49

catalogue No #112; 1:10,000) diluted in PBS containing 0.3% Triton X‐100 and 0.5%

BSA (Sigma). The goat anti‐CT‐B primary antibody recognizes the B‐subunit of

cholera toxin. Sections were then washed in 0.01 M PBS and blocked with 5% normal

donkey serum in 0.01 M PBS for 1 h at RT. Sections were immediately incubated in

the corresponding secondary antibody (anti‐sheep biotin, Jackson ImmunoResearch

Laboratories, West Grove, PA; 1:500 in 0.01 M PBS) for 1 h at RT. Next, sections were

washed in 0.01 M PBS and incubated in an ABC kit (Vectastain Elite ABC‐HRP Kit)

for 1 h at RT. Finally, sections were washed in 0.01 M PBS and subsequently

incubated in diaminobenzidine (DAB) substrate (1:10, Roche Diagnostics, Mannheim,

Germany) for 4 min, followed by an incubation in 1% H2O2 for 4 min (Stanic et

al., 2003). Sections were then washed ×3 in 0.01 M PBS and mounted on slides

coated with 0.5% gelatin (Sigma) and 0.05% chromium (III) potassium sulfate

dodecahydrate (Merck), and left to dry overnight. Slides were coverslipped using DPX

(Sigma‐Aldrich).

Data analysis and image processing

We used a bright‐field microscope (Leica Biosystems) to identify and document

the location of the midbrain and brainstem injection sites. Injection sites were

photographed and plotted on semischematic drawings of the respective sections

containing the PAG, KFn, BötC, pre‐BötC, and caudal raphé nuclei (Fig 3.1). We used

the following criteria to classify injections of the adjacent BötC and pre‐BötC nuclei in

the medulla. The center of the BötC injection was located 0.0–0.5 mm to caudal edge

of the facial motor nucleus (VII) where the medial longitudinal fasciculus (mlf) in the

midline is ventral‐dorsally prolonged till the fourth ventricle (4V), whereas the center

of the pre‐BötC injection sites was located 0.5–0.8 mm to caudal edge of the facial

motor nucleus where the mlf is less ventral‐dorsally prolonged in the midline. Another


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landmark for the pre‐BötC location was the presence of the hypoglossal nucleus (XII)

in the dorsomedial region of the section, which is not present in the BötC sections

(Fig 3.1c).

Representative images were taken using a digital camera (Leica DFC7000T)

mounted to the microscope (Leica DM6B LED). Adobe Photoshop CC19 software

(Adobe Systems Inc., San Jose, CA) was used to assemble representative images,

and to optimize brightness and contrast of the digital images to best represent sections

viewed under the microscope.

We examined sections separated by 120 μm across the neuroaxis of the entire

forebrain. We quantified in forebrain areas with significant and consistent numbers of

retrogradely CT‐B‐labeled neurons (>n = 3) for all experimental cases. Near‐miss

injections served as controls. CT‐B labeled cell bodies were recognized by the

presence of black punctate granules in the neuronal cell bodies (somata) and

processes, and the absence of immunoreactive cell nuclei. The background staining

of neurons was characterized by light and diffuse DAB staining of somata including

the cell nuclei. The specific location and distribution of retrogradely labeled forebrain

projection neurons in cortical and subcortical areas are classified according to the rat

brain atlas of Paxinos and Watson (2007).

We used a one‐way analysis of variance (GraphPad Prism, version 7.02;

GraphPad Software; San Diego, CA) followed by Tukey's multiple comparison test to

determine the statistical significance of the total, ipsilateral and contralateral numbers

of CT‐B‐labeled neurons between the brainstem target areas (i.e., PAG, KFn, pre‐

BötC, BötC, and caudal raphé nuclei; Fig 3.2). Values are given as mean ± SEM

and p‐values less than .05 were considered statistically significant.




51

Results

Injections sites

Microinjections of the retrograde tracer CT‐B were anatomically confined to

discrete injection sites in all target areas (see representative photographs, Fig 3.1) or

around the target areas (“near‐miss injections”). The latter are illustrated as black

crosses in the semischematic drawings (Fig 3.1, left panel).

CT‐B injection sites in columns of the midbrain PAG were confined to the vlPAG

(n = 4) and the dlPAG (n = 1; Fig 3.1a). The rostrocaudal levels of the injections were

identified by the size and location of the aqueduct (Bandler et al., 1991; Carrive, 1993).

Three injection sites were centered in the rostral PAG (Cases #1–3), whereas the

other two cases (#4, 5) were located more caudally. We also report three near‐miss

injections, which were localized ventrolaterally to the PAG (Fig 3.1a, black x's).

CT‐B injections in the pontine KFn, located ventral to the lateral tip of the

superior cerebellar peduncle (scp), were restricted to the rostral and intermediate

regions of the KFn (Cases #6–10, Fig 3.1b), and strongly overlap with the core circuitry

of pontine respiratory group in rodents (Dutschmann and Herbert, 2006). Two near‐

miss injections were found slightly ventrocaudally to the pontine KFn (Fig 3.1b, black

x's).

Anatomical locations of the medullary pre‐BötC (Cases #11–13) and BötC

(Cases #14–16, Fig 3.1c) were localized ventral to the nucleus ambiguus (NA) and

caudal to the facial nucleus (VII) in the medulla oblongata. Two near‐miss injections

were located dorsolaterally to the NA (Fig 3.1c, black x's).

Injection sites for Cases #17–19 were confined to the caudal midline raphé

(Fig 3.1d), which is subdivided in three regions: ROb, RPa, and RMg. Injections in the

caudal raphé were located rostral to the BötC/pre‐BötC brainstem sections. Case #17













52

was predominantly localized to the RPa. Case #18 was more rostral to Case #17 and

localized to the RPa/RMg. Case #19 was localized to the RMg/ROb. Two near‐miss

injections were located lateral to the caudal midline raphé (Fig 3.1d, black x's).


53

Figure 3.1. Schematic drawings (left) and photomicrographs (right) illustrate the anatomical

location of cholera toxin subunit B (CT‐B) microinjections in functionally diverse respiratory

control areas.

54

The periaqueductal gray (a), the Kölliker‐Fuse nucleus (b), the pre‐Bötzinger and Bötzinger complexes

(c), and the caudal raphé nuclei (d). Left: Schematic drawings depict the location and dimensions of all

CT‐B injections in the brainstem target areas relative to bregma. In this and in Figures 3.2, 3.5, 3.7, 3.9,

3.10, and 3.12, colors code specific on‐target injections; and black x's indicate near‐miss, off‐target

injections (see results). Right: Photomicrographs of representative injection sites. 4V, fourth ventricle;

BötC, Bötzinger complex; KFn, Kölliker‐Fuse nucleus; mcp, middle cerebellar peduncle; io, inferior

olive; mlf, medial longitudinal fasciculus; NA, nucleus ambiguus; NTS, solitary tract nucleus; PAG,

periaqueductal gray; pre‐BötC, pre‐Bötzinger complex; py, pyramidal tract; RMg, raphé magnus; rob,

raphé obscurus; RPa, raphé pallidus; scp, superior cerebellar peduncle; XII, facial motor nucleus. Scale

bars: 1 mm (schematic drawings on the left); 200 μm (representative photographs on the right).

Cumulative distribution and laterality of descending forebrain projection

Quantitative analysis of the total numbers of CT‐B‐labeled neurons following

injections in the midbrain PAG, pontine KFn, medullary pre‐BötC, BötC, or the caudal

raphé revealed that the highest number of CT‐B‐labeled cells in the forebrain projected

monosynaptically to the PAG (p < .05; Fig. 3.2). Injections in the pontine KFn resulted

in the second highest number of CT‐B labeled neurons in the forebrain, followed by

the medullary pre‐BötC, and the caudal raphé. The lowest numbers of CT‐B‐labeled

neurons were observed after injections in the medullary BötC.

Analysis of the ipsilateral and contralateral distribution of the CT‐B‐labeled

neurons demonstrates that the majority of the CT‐B‐labeled neurons after PAG and

KFn injections were located in the ipsilateral hemisphere (PAG, 83.1 ± 3.1%; KFn, 82.5

± 4.6%). For injections in the pre‐BötC and BötC, we respectively observed 69.8

± 6.5% and 60.7 ± 6.0% of CT‐B‐labeled neurons were located ipsilaterally.

CT‐B‐labeled cell bodies in cortical regions were almost exclusively found in

the Layer V/VI, and displayed the characteristic morphology of cortical pyramidal

neurons (Fig. 3.3). It is also important to note that CT‐B‐labeled neurons in the

hippocampus, olfactory bulb, basal ganglia, and cerebellum were not detected in any

experiments reported in this study.






55

Figure 3.2. Bar graphs of the total (a), ipsilateral (b) and contralateral (c) numbers of retrogradely

cholera toxin subunit B (CT‐B)‐labeled neurons in the forebrain after microinjections in PAG,

KFn, pre‐BötC, BötC, and caudal raphé nuclei.

In (a‐c), color‐coded circles (experimental cases) align with those Figure 3.1; and zero laterality for

caudal raphé nuclei. All values are expressed as mean ± SEM. The greatest number of CT‐B

retrogradely labeled forebrain neurons were detected from injections in the PAG compared to those in

the KFn, pre‐BötC, BötC, and caudal raphé nuclei (one‐way analysis of variance [ANOVA] followed by

Tukey's multiple comparison test, *p < .05). BötC, Bötzinger complex; KFn, Kölliker‐Fuse nucleus; PAG,

periaqueductal gray; pre‐BötC, pre‐Bötzinger complex.


56

Figure 3.3. Representative cholera toxin subunit B (CT‐B)‐labeled pyramidal neurons in Layer

V/VI of the insular cortex (a).

(b,c) Photomicrographs show labeled neurons at progressively higher magnifications. (a) The

photomicrograph at the lowest magnification shows the specificity of retrograde labeling in the context

of the neighboring cells and structures. (b) At highest magnification, the morphology of labeled neurons

(black arrowheads) is more apparent. (c) At a slightly higher magnification than that in (b), the dentrites

(red arrowheads) of the pyramidal neurons are visible. II, Layer 2 of cortex; III, Layer 3 of cortex; V,

Layer 5 of cortex; CPu, caudate putamen (striatum); ec, external capsule; rf, rhinal fissure. Scale bars:

a = 200 μm; b = 100 μm; and c = 50 μm.

Specific distribution of retrogradely CT-B labeled neurons in cortical and

subcortical brain regions following injections in the midbrain PAG

Figure 3.4 shows representative images of retrogradely labeled neurons in the

cortical and subcortical brain regions following PAG injections. Figure 3.5 illustrates

the rostrocaudal gradients of all CT‐B‐labeled neurons in relation to bregma.



57

Figure 3.4. Cholera toxin subunit B (CT‐B) microinjection in the PAG labeled neurons in various

cortical and forebrain regions.

In this and Figures 3.6, 3.8, 3.6, 3.8, and 3.11, we present the paired photomicrographs of the labeled

neurons. 1) The collage on the left is at low magnification to show the specificity of the retrolabeling in

the context of the neighboring structures. Dashed‐line boxes in 1 outline the area of collage in 2. 2) The




58

collage on the right is at high magnification to show the morphology of the labeled cells. Red arrowheads

point to representative labeled neurons. The following areas had CT‐B‐labeled neurons: (a1‐2) the

cingulate cortex (ACA); (b1‐3) motor and prelimbic cortices (MI, PL); (c1‐2) infralimibic (IL); (d1‐2) lateral

septum (LS); (e1‐2) amygdala (Amg); (f1‐2) paraventricular hypothalamus (PVN); (g1‐2) insular cortex

(IC); (h1‐2) endopiriform nucleus (DEn); (i1‐2) claustrum (CI); (j1‐2) dorsomedial (DMH); (k1‐2)

ventromedial (VMH); (l1‐2) lateral hypothalamus (LH); and (m1‐2) preoptic area (PO). 3V, third ventricle;

aca, anterior commissure; cc, corpus callosum; CPu, caudate putamen (striatum); E/OV, ependymal

and sublayer EV; ec, external capsule; fmi, forceps minor of corpus callosum; IC, internal capsule; LV,

lateral ventricle; mlf, medial longitudinal fasciculus; och, optic chiasm; opt, optic tract; rf, rhinal

fissure. Scale bars on this and Figures 3.6, 3.8, 3.6, 3.8, and 3.11: 200 μm (low‐magnification images);

50 μm (high‐magnification images).




59

Figure 3.5. The rostrocaudal distribution of the numbers of retrogradely labeled neurons relative

to bregma (mm) in various cortical (a) and subcortical (b) brain regions following cholera toxin

subunit B (CT‐B) microinjections in the periaqueductal gray.

The location of the specific injection sites are represented by color‐codes, which are the same as

Figure 3.1. Black lines represent the mean number of CT‐B‐labeled neurons detected throughout the

rostrocaudal levels for each cortical and subcortical brain region. DMH, dorsomedial hypothalamus; LH,

lateral hypothalamus; PAG, periaqueductal gray; PVN, paraventricular hypothalamus; VMH,

ventromedial hypothalamus.

We observed the largest number of CT‐B‐labeled neurons in the prefrontal

cortex (PFC), including the dorsal (Figure 3.5a1; prelimbic, 412 ± 51 neurons/case



60

[n/c]) and ventral PFC (Figure 3.5a2 infralimbic, 198 ± 40 n/c). In prelimbic cortex, CT‐

B‐labeled neurons occupied the rostral and intermediate levels (bregma: from +4.0 to

+1.5 mm). In infralimbic cortex, CT‐B‐labeled neurons were predominantly located at

rostral levels (bregma: from +4.0 to +2.5 mm). We also observed a large number of

CT‐B‐labeled neurons in motor cortex (Figure 3.5a3; 192 ± 99 n/c; bregma: from +4.0

to 0 mm). Substantial numbers of CT‐B‐labeled neurons in cingulate cortex were

observed after injections in the rostral PAG (Case #1, 336 neurons; Case #2, 648

neurons), whereas injections in the caudal region of the midbrain PAG revealed

smaller numbers of CT‐B labeled‐neurons in the cingulate cortex (Case #3, 41

neurons; Case #4, 33 neurons; Case #5, 0 neurons) (Fig 3.5). In addition, we also

observed a considerable number of CT‐B‐labeled neurons in the insular cortex

(Figure 3.5a5; 172 ± 88 n/c, bregma: from +4.5 to +1.5 mm), particularly after injections

in the rostral vlPAG (Case #2, 272 neurons; Case #3, 469 neurons). Caudal injections

in the vlPAG (Cases #4–5) revealed fewer numbers of CT‐B‐labeled neurons in the

insular cortex (Case #4, 14 neurons; Case #5, 106 neurons), whereas no CT‐B‐

labeled neurons were observed after injections in the rostral dlPAG (Case #1). A small

number of CT‐B‐labeled neurons were found in the rhinal (50 ± 20 n/c; bregma: from

−3.18 to −7.3 mm) and endopiriform cortices (Cases #1–5, 37 ± 4 n/c; Fig 3.5a6). CT‐

B‐labeled neurons in the endopiriform were restricted to the rostral level of this nuclei

(bregma: from −0.7 to +2.4 mm). Finally, smaller numbers of monosynaptic projections

to the PAG were observed from the somatosensory cortex (2 ± 1 n/c; bregma: from

+1.8 to +1.3 mm).

Substantial numbers of monosynaptic projections to the PAG were also

observed from subcortical regions, such as the claustrum, amygdala, lateral septum,

and hypothalamus (Fig 3.4 and 3.5). Injections in the PAG revealed high numbers of








61

retrogradely labeled neurons in the amygdala (146 ± 72 n/c; bregma: from −1.0 to

−2.6 mm), whereas a smaller number of labeled neurons were observed after

injections in the dlPAG (Case #1, 5 neurons). Injections in the rostral PAG revealed a

large number of CT‐B‐labeled neurons in the lateral septum (Case #1, 123 neurons;

Case #2, 140 neurons; bregma: from +0.6 to +1.8 mm), whereas injections in caudal

PAG (Case #4, 17 neurons; Case #5, 2 neurons) yielded smaller numbers of

descending projection neurons from the lateral septum. Hypothalamic nuclei showed

high numbers of CT‐B‐labeled neurons in all cases (Figure 3.5b1,b2,b3,b5,b8).

Overall, the highest numbers of retrogradely CT‐B‐labeled neurons were observed in

the ventromedial hypothalamus (VMH, 486 ± 159 n/c; bregma: from −1.7 to −4.6 mm),

followed by the preoptic area (PO, 288 ± 83 n/c; bregma: from −1.2 to +0.5 mm), lateral

hypothalamus (LH, 280 ± 88 n/c; bregma: from −1.2 to −3.84 mm), dorsomedial

hypothalamus (DMH, 72 ± 36 n/c; bregma: from −2.5 to −3.9 mm), and paraventricular

hypothalamic nucleus (PVN, 14 ± 8 n/c; bregma: from −1.0 to −2.0 mm). Finally, a

small number of CT‐B‐labeled neurons were observed in the claustrum after injections

in the rostral or caudal PAG (40 ± 2 0 n/c; bregma: from +2.2 to +1.6 mm). Overall, the

distribution of forebrain projections to the midbrain PAG is consistent with the literature

(Dampney et al., 2013; see discussion).


subcortical brain regions following injections in the pontine KFn

Figure 3.6 shows representative images of retrogradely CT‐B‐ labeled neurons in the

cortical and subcortical regions that project to the KFn. Figure 3.7 depicts the

rostrocaudal gradients of CT‐B‐labeled neurons.





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Figure 3.6. Cholera toxin subunit B (CT‐B) microinjection in the Kölliker‐Fuse nucleus labeled

neurons in various cortical and forebrain regions (paired photomicrographs: 1 is low and 2 is

high magnification).

63

The following areas had CT‐B‐labeled neurons: (a1‐2) motor cortex (MI); (b1‐2) prelimbic cortex (PL);

(c1‐2) infralimibic (IL); (d1‐2) somatosensory cortex (S1); (e1‐2) amygdala (Amg); (f1‐2) dorsomedial

hypothalamus (DMH); (g1‐2) insular cortex (IC); (h1‐2), endopiriform nucleus (DEn); (i1‐2), rhinal cortex

(Rh); (j1‐2) claustrum (CI); (k1‐2) paraventricular hypothalamus (PVN); (l1‐2) ventromedial (VMH). 3V,

third ventricle; aca, anterior commissure; CPu, caudate putamen (striatum); E/OV, ependymal and

sublayer EV; ec, external capsule; fmi, forceps minor of corpus callosum; IC, internal capsule; mlf,

medial longitudinal fasciculus; och, optic chiasm; opt, optic tract; rf, rhinal fissure. Scale bars: 200 μm

(low‐magnification images) and 50 μm (high‐magnification images).

Figure 3.7. The rostrocaudal distribution of retrogradely labeled neurons relative to bregma

(mm) in various cortical (a) and subcortical (b) brain regions following cholera toxin subunit B

(CT‐B) microinjections in the Kölliker‐Fuse nucleus.




64

rostrocaudal levels for each cortical and subcortical brain region. DMH, dorsomedial hypothalamus;

KFn, Kölliker‐Fuse; LH, lateral hypothalamus; PVN, paraventricular hypothalamus; VMH, ventromedial

hypothalamus.

Quantitative analysis of monosynaptic projections to the KFn revealed

considerable numbers of CT‐B‐labeled neurons in infralimbic (60 ± 21 n/c; bregma:

from +3.7 to +2.3 mm), rhinal (146 ± 63 n/c; bregma: from −2.3 to −7.5 mm),

endopiriform (60 ± 22 n/c; bregma: from +3 to +0.1 mm), somatosensory (63 ± 25 n/c;

bregma: from +2.5 to +0.1 mm), insular (64 ± 15 n/c; bregma: from +1.4 to −2.5 mm),

prelimbic (29 ± 5 n/c; bregma: from +4.0 to +3.7 mm), and motor cortices (29 ± 4 n/c;

bregma: from +4.5 to +1.4 mm). In contrast to PAG injections, we did not detect CT‐

B‐labeled neurons in the cingulate cortex following injections in the KFn.

Monosynaptic projections to the KFn were also found from subcortical areas

such as the claustrum, amygdala and various hypothalamic nuclei. In contrast to PAG

injections, no neurons were found in the lateral septum after CT‐B injection in the KFn.

The highest numbers of retrogradely labeled neurons were found in the LH (225 ± 88

n/c; bregma: from −1.44 to −3.6 mm), followed by the VMH (112 ± 33 n/c; bregma:

from −2.7 to −3.6 mm), DMH (47 ± 8 n/c; bregma: from −2.5 to −3.9 mm), PO (34 ± 12

n/c; bregma: from +0.5 to −1.2 mm), and PVN (29 ± 4 n/c; bregma: from −1.3 to

−2 mm). We also observed CT‐B‐labeled neurons in the claustrum (Fig 3.7b7; 56 ± 1

n/c; bregma: from +2.2 to −2.4 mm) and amygdala (23 ± 7 n/c; bregma: from −1.2 to

−2.6 mm).


subcortical brain regions following injections in the medullary pre‐BötC and

BötC


65

Figure 3.8 shows representative images of retrogradely CT‐B‐labeled neurons

in the cortical and subcortical regions following CT‐B injections in the pre‐BötC.

Figure 3.9 (pre‐BötC injections) and 3.10 (BötC injections) illustrate the rostrocaudal

gradients of CT‐B‐labeled neurons in relation to bregma.

Figure 3.8. Cholera toxin subunit B (CT‐B) microinjection in the pre‐Bötzinger complex labeled



The following areas had CT‐B‐labeled neurons: (a1‐2) motor cortex (MI); (b1‐2) insular (IC); (c1‐2)

amygdala (Amg); (d1‐2) paraventricular hypothalamus (PVN); (e1‐2) preoptic area (PO); and (f1‐2)

lateral hypothalamus (LH). 3V, third ventricle; och, optic chiasm; pre‐BötC, pre‐Bötzinger complex; rf,

rhinal fissure. Scale bars: 200 μm (low‐magnification images) and 50 μm (high‐magnification images).



66



(CT‐B) microinjections in the pre‐Bötzinger complex.


Figure 3.1. Black lines represent the mean numbers of CT‐B‐labeled neurons detected throughout the


lateral hypothalamus; pre‐BötC, pre‐Bötzinger complex; PVN, paraventricular hypothalamus; VMH,



67

Figure 3.10. The rostrocaudal distribution of the numbers of retrogradely labeled neurons

relative to bregma (mm) in various cortical (a) and subcortical (b) brain regions following cholera

toxin subunit B (CT‐B) microinjections in the Bötzinger complex.



rostrocaudal levels for each cortical and subcortical brain region. BötC, Bötzinger complex; DMH,

dorsomedial hypothalamus; LH, lateral hypothalamus; PVN, paraventricular hypothalamus; VMH,


The highest number of descending projection neurons to the pre‐BötC was

observed in the motor cortex (50 ± 21 n/c; bregma: from +5 to +1.2 mm), followed by


68

the insular (36 ± 9 n/c; bregma: from +3.4 to +1.3 mm), somatosensory (14 ± 3 n/c;

bregma: from +2 to +1.1 mm), infralimbic (5 ± 2 n/c; bregma: from −3.2 to −2.7 mm),

rhinal (4 ± 2 n/c; bregma from −5.3 to −4.9 mm), prelimbic (2 ± 2 n/c; bregma: from

+3.8 to +3.3 mm), and endopiriform cortices (1 ± 1 n/c; bregma: from +1.7 to +1.4 mm).

Similar to KFn CT‐B injections, we did not detect labeled neurons in the cingulate

cortex after injections in the medullary pre‐BötC.

In subcortical regions (Fig 3.9), we observed monosynaptic projections to the

pre‐BötC from amygdala (59 ± 33 n/c; bregma: from −1.4 to −2.4 mm), claustrum

(11 ± 5 n/c; bregma: from +0.5 to −0.7 mm), and hypothalamic nuclei, including the LH

(89 ± 25 n/c; bregma: from −1.4 to −3.6 mm), PO (45 ± 34 n/c; bregma: from +0.3 to

−1.4 mm), PVN (39 ± 10 n/c; bregma: from −2 to −1 mm), DMH (35 ± 19 n/c; bregma:

from −2.7 to −3.9 mm), and VMH (7 ± 5 n/c; bregma: from −2.9 to −3.4 mm).

By contrast, CT‐B injections in the BötC of the ventral respiratory column

revealed substantially fewer numbers of descending projection neurons in cortical

brain regions (p = 0.0003). We observed CT‐B‐labeled neurons projecting to BötC in

the insular (31 ± 28 n/c; bregma: from +3.7 to +1.3 mm), endopiriform (3 ± 2 n/c;

bregma: from +1.9 to +1.2 mm), motor (1 ± 1 n/c; bregma: +3.3 mm), infralimbic (1 ± 1

n/c; bregma: +3.0 mm), somatosensory (1 ± 1 n/c; bregma: +1.6 mm), rhinal (1 ± 1 n/c;

bregma: from −5.1 to −5.4 mm), and prelimbic cortices (1 ± 1 n/c; bregma: +3.6 mm).

Similarly, only small numbers of CT‐B‐labeled neurons were detected in subcortical

regions following BötC injections: amygdala (21 ± 21 n/c; bregma: from −1.7 to

−2.2 mm), LH (3 ± 2 n/c; bregma: from −2.4 to −2.6 mm), PO (1 ± 1 n/c; bregma:

−0.5 mm), PVN (1 ± 1 n/c; bregma: −1.5 mm), and claustrum (1 ± 1 n/c; bregma:

−0.2 mm). Finally, CT‐B‐labeled neurons in lateral septum, DMH and VMH were never

observed after injections in the medullary BötC.


69


subcortical brain regions following injections in the caudal raphé in the medulla

oblongata

Figure 3.11 shows representative images of retrogradely CT‐B‐labeled

neurons in cortical and subcortical regions following CT‐B injections in the caudal

raphé nuclei of the medulla oblongata. Figure 3.12 illustrates the quantitative analysis

of CT‐B‐labeled neurons along their rostrocaudal gradients in relation to bregma. In

accordance with published literature (Hermann et al., 1997), CT‐B injections in the

caudal raphé showed variable numbers of labeled neurons in the forebrain, which is

dependent whether the injections were centered in the RMg, ROb, or RPa. For

example, Case #17 (injection in the caudal RPa) revealed the highest number of CT‐

B‐labeled neurons in the cortex (Fig 3.12).

Figure 3.11. Cholera toxin subunit B (CT‐B) microinjection in the caudal raphé nuclei labeled



The following areas had CT‐B‐labeled neurons: (a1‐2) motor cortex (MI); (b1‐2) dorsomedial

hypothalamus (DMV); (c1‐2) lateral hypothalamus (LH); and (d1‐2) preoptic area (PO). 3V, third

ventricle; aca, anterior commissure; DMH, dorsomedial hypothalamus; LH, lateral hypothalamus; MI,





70

motor cortex; och, optic chiasm; PO, preoptic nucleus; VMH, ventromedial hypothalamus. Scale bars:

200 μm (low‐magnification images) and 50 μm (high‐magnification images).



(CT‐B) microinjections into the caudal raphé nuclei.




lateral hypothalamus; PVN, paraventricular hypothalamus; VMH, ventromedial hypothalamus.


71

CT‐B injections in the caudal raphé revealed a modest number of monosynaptic

projections from cortex. The highest numbers of descending projection neurons were

observed in the motor (66 ± 26 n/c; bregma: from +5 to +0.3 mm), followed by the

insular (59 ± 13 n/c; bregma: +4.5 to +1.7 mm), prelimbic (20 ± 12 n/c; bregma: from

+5 to +1.9 mm), endopiriform (14 ± 6 n/c; bregma: from +2.6 to +1.2 mm), infralimbic

(9 ± 9 n/c; bregma: from +3.2 to +2.2 mm), somatosensory (4 ± 3 n/c; bregma: from +3

to +1.6 mm), and cingulate cortices (4 ± 4 n/c; bregma: from +2.3 to +1.8 mm). No CT‐

B‐labeled neurons were detected in the rhinal cortex followed by injections in any

caudal raphé nucleus.

In subcortical regions, descending connectivity outside the hypothalamus was

absent or weak (e.g., claustrum, amygdala, and lateral septum). However, high to

moderate numbers of projection neurons were observed in various hypothalamic

nuclei. The highest numbers of CT‐B‐labeled neurons were in the DMH (125 ± 60 n/c;

bregma: from −2.5 to −3.7 mm), followed by the LH (91 ± 64 n/c; bregma: from −1.4 to

−3.1 mm), and PO (84 ± 49 n/c; bregma from +0.2 to −1 mm). The number of CT‐B‐

labeled neurons in the PVN (12 ± 8 n/c; bregma: from −1.8 to −1.3 mm) and VMH (3

± 1 n/c; bregma: from −2.2 to −2.7 mm) were smaller compared to the aforementioned

hypothalamic nuclei.

CT-B-labeled neurons associated with near-miss injections

Analysis of near‐miss injection sites (n = 9) presented far fewer labeled neurons

compared to injections that were centered in the target areas. For instance, injections

placed ventrolateral to the PAG presented far fewer labeled neurons in the forebrain.

Only small numbers of CT‐B‐labeled neurons were observed, predominantly in the

insular, rhinal, and motor cortices, but not in hypothalamic areas (data not shown). In

contrast, near‐miss injections ventrocaudally to the KFn revealed modest numbers of

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CT‐B‐labeled neurons in the cortex, but we still identified robust numbers of projection

neurons in the hypothalamus (e.g., LH) and midbrain PAG (data not shown). The later

illustrates that the caudal extension of the KFn remains connected to key nuclei of the

autonomic nervous system in the forebrain, but lacks major cortical inputs. Finally,

near‐miss injections placed in the reticular formation outside the pre‐BötC, BötC, and

caudal raphé target areas resulted in almost complete absence of forebrain projection

neurons, although labeled cells were still observed in the KFn and PAG (data not

shown). Thus, the near miss injections underline the specificity of the descending

projection patterns in the investigated respiratory nuclei (as well as the specificity of

CT‐B antibody used in this study).

Discussion

We identified widespread monosynaptic descending projections from the

forebrain to all respiratory control areas investigated suggesting that forebrain‐evoked

modulations of respiration do not simply bypass brainstem circuits, but instead, are

likely coordinated with spontaneous respiratory network activity. Descending

projection neurons that target designated respiratory control areas in the midbrain and

brainstem were restricted to a variety of cortical areas, the claustrum, lateral septum,

various hypothalamic nuclei and the amygdala. Examination of the distribution of

retrogradely labeled neurons across the entire axis of the forebrain failed to detect any

descending projection neurons in some major neural systems of the forebrain, such

as the thalamus, hippocampus, olfactory bulb, or basal ganglia.

The topographical organization of detected monosynaptic descending forebrain

projections is summarized in a network connectivity graph (Fig 3.13). The graph shows

the relative proportion of projections from a given cortical or subcortical area to each

respiratory control nuclei investigated. The general distribution of descending


73

projection neurons located in subcortical areas, such as the hypothalamus or

amygdala, reveals a broad connectivity pattern with the downstream targets. These

subcortical projections are discussed in the context of their putative role in

homeostasis, state‐dependent, and emotional modulation of breathing

(Section: Homeostasis, state-dependent or emotional modulation of respiration). In

terms of cortical projections to the brainstem respiratory control areas, we have

detected some inputs targeting the medullary regions, such as the pre‐BötC (similarly

to Yang et al., 2020). However, the present study implies that retrogradely labeled

neurons predominantly target the midbrain PAG and pontine KFn. The only exception

is the insular cortex which also provides an equal proportion of descending inputs to

all investigated respiratory control areas. The long‐range forebrain‐brainstem

projection neurons detected in this study were exclusively located in the deepest

layers of cortex. The role of these cortex‐brainstem projections is discussed in the

context of volitional control of breathing (Section: Volitional control of respiration).


74

Figure 3.13. Summary of the relative strength of descending projections arising from cortical (a)

and subcortical (b) areas in a connectivity map.

The weight of connecting lines are proportional to the normalized maximal number of cholera toxin

subunit B (CT‐B)‐labeled neurons found in the specific source of descending projections.

75

Finally, among all downstream brainstem targets investigated, the BötC shows

the least amount of forebrain connectivity, despite its designated key function in

respiratory pattern formation (Burke et al., 2010; Marchenko et al., 2016; Smith et

al., 2013). Given that the neighboring pre‐BötC in the medulla has a vital role on

breathing rhythm generation (Del Negro et al., 2018; Feldman and del Negro, 2006;

Smith et al., 1991) and the generation of sighs (Li et al., 2016a), it is not surprising that

this brain area receives also receives considerable cortical inputs.

Homeostasis, state-dependent or emotional modulation of respiration

The dense descending connectivity of visceral sensory areas of the insular

cortex with respiratory control areas confirm previous anatomical studies (Grady et

al., 2020; Saper, 1982; Sato et al., 2013), and are in line with several functional studies

that have associated the insular cortex with cardiorespiratory modulation (Aleksandrov

et al., 2000; Ruggiero et al., 1987; Yasui et al., 1991), which may be associated with

adaptation of breathing to interoceptive states (Verdejo‐Garcia et al., 2012). The

present study also confirms well‐documented descending projections of hypothalamic

nuclei targeting the PAG, KFn, pre‐BötC, BötC, and caudal raphé (Geerling et

al., 2010; Peyron et al., 1998; Yang et al., 2020). It can be speculated that the

hypothalamic projections to the brainstem regions may control autonomic functions

other than respiration. For example, there is strong evidence in the literature that the

DMH sends stress‐related information to the caudal raphé to regulate autonomic

functions such as body temperature and energy metabolism (Kataoka et al., 2014;

Sarkar et al., 2007; Tupone et al., 2011). However, it is also well‐documented that a

range of respiratory patterns linked to homeostasis, state dependency (sleep–wake),

and stress, are mediated by hypothalamic subnuclei (for review, see Fukushi et





















76

al., 2019), suggesting that respiration adapts to interoceptive states as well as

emotional and stress‐related information.

The insular cortex and hypothalamus are part of the widely distributed limbic

system that includes the amygdala, and the piriform, rhinal, and prefrontal cortices.

Because emotions have a profound influence on respiratory activity (Harper et

al., 1984; Holstege, 2014; Homma and Masaoka, 2008; Onimaru and Homma, 2007;

Subramanian and Holstege, 2014), it is not surprising that all aforementioned areas

have direct descending projections to several respiratory control areas investigated in

the present study (Fig 3.13). While it was previously postulated that the midbrain PAG,

with its known function in respiratory modulation in relation to fear and defensive

behaviour (Carrive, 1993; Carrive et al., 1988; Carrive et al., 1997; Depaulis et

al., 1989; Subramanian and Holstege, 2014), is the primary interface that links

emotion with breathing, the present study illustrates that similar descending inputs also

target nuclei of the primary respiratory rhythm and pattern generating network in the

brainstem. Significant inputs from the amygdala, and the prelimbic and infralimbic and

rhinal cortices to the pontine KFn and medullary pre‐BötC, with their specific function

in controlling respiratory phase transitions (Dutschmann and Herbert, 2006) and

rhythm generation (Del Negro et al., 2018; Feldman and del Negro, 2006; Smith et

al., 1991), suggests that the synaptic output of the widely distributed limbic system

also connects to a similarly distributed respiratory network spanning from the midbrain

to medulla oblongata. Considering recent evidence suggesting that action/motor

encoding involves cell assemblies whose members are distributed in many areas

beyond the cortex, including the PAG (Steinmetz et al., 2019), the present study

supports the working hypothesis that the precise encoding of homeostatic, state‐


















77

dependent or emotional breathing patterns may be partially outsourced to the

respiratory network.

Volitional control of respiration

In contrast with previous suggestions that cortical motor commands for the

respiratory system may be mediated via corticospinal pathways that connect the

cortex with primary respiratory motor pools (Gandevia and Rothwell, 1987a,b; Pouget

et al., 2018; Rikard‐Bell et al., 1985), our study identifies distinct descending pathways

that connect cortical output neurons with specific respiratory premotor nuclei

(Fig 3.13a). For instance, descending inputs from cortical areas of the frontal lobe,

including the PFC (prelimbic and infralimbic cortices) and the cingulate cortex, have

the strongest connectivity with the lateral and ventrolateral columns of the PAG.

Indeed, our data show strong cortical projections to the vlPAG and confirm a previous

study that reported significant inputs from the motor, prelimbic, infralimbic, cingulate,

insular, endopiriform, and rhinal cortices (Floyd et al., 2000). However, our study

detected significant numbers of CT‐B positive neurons in the somatosensory cortex,

which were not reported previously. Among all cortical and subcortical areas with

descending projections to respiratory control areas, the cingulate and lateral septum

almost exclusively target the PAG (Fig 3.13a). The neural networks composed of the

cingulate and septal nuclei, including the PFC (area of Brocca), harbor the primary

synaptic network for the generation of speech in humans and vocalisation in mammals

(Hage and Nieder, 2016; Holstege and Subramanian, 2016; Jürgens, 2009). In line

with the specific descending projection pattern that links the PAG to volitional motor

commands for vocalisation, stimulation of the midbrain PAG (particularly the lPAG and

vlPAG) evokes vocalisation in animals (Bandler and Carrive, 1988; Jürgens and

Richter, 1986; Larson, 1985; Zhang et al., 1994). Thus, the present study supports the














78

hypothesis that the midbrain PAG is the “final common pathway” for vocalisation in

mammals (Düsterhöft et al., 2004; Holstege, 1989; Jürgens, 1994; Jürgens, 2002a,b;

Jürgens and Richter, 1986; Subramanian and Holstege, 2009). However, given that

recent studies using optogenetic approaches have defined forebrain inputs to the PAG

that play critical roles in predator and prey behaviours (for review, see Franklin, 2019),

we cannot assume that the descending inputs to the PAG found in the present study

necessarily control volitional respiration or vocalisation. Nevertheless, it is well‐known

that the PAG is a crucial region in the midbrain that coordinates cardiorespiratory

responses to behaviours that are initiated in forebrain circuits (Bandler et al., 2000;

Benarroch, 2012; Dampney et al., 2013).

Studies in humans, however, have shown that the midbrain PAG is not

activated during other voluntarily controlled orofacial behaviours such as coughing

(Mazzone et al., 2011) and swallowing (Zald and Pardo, 1999). Moreover, the PAG

neither possesses direct synaptic connection to cranial and spinal motor respiratory

pools (Dampney et al., 2013; Holstege, 1989; Holstege, 2014; Subramanian and

Holstege, 2009; Zhang et al., 1995), nor is it part of the primary rhythm and pattern

generating circuit (Farmer et al., 2014). Thus, the midbrain PAG, as a mediator for

modulation or reconfiguration of respiratory activity during orofacial behaviours,

arousal or emotion, requires additional downstream synaptic interactions with the

ponto‐medullary brainstem respiratory network.

The PAG shares the highest‐level reciprocal connectivity with the KFn when

compared to other medullary nuclei of the primary respiratory network (BötC, pre‐

BötC, or caudal raphé; data not shown, Trevizan‐Baú et al., 2020). Since both nuclei

receive dense innervation from the claustrum, a general cortico‐thalamic integration




















79

and output nucleus (Dillingham et al., 2017), it is possible that the PAG and the KFn

may mediate various volitional motor commands in concert.

However, respiratory functions of the KFn in the mediation of the inspiratory–

expiratory phase transition (Caille et al., 1981; Mörschel and Dutschmann, 2009;

Smith et al., 2007; Wang et al., 1993) and gating of the respiratory cranial nerve

activities (Bautista and Dutschmann, 2014; Dutschmann and Herbert, 2006), including

in the branch of the facial nerve that innervates the nostrils and whisker pads

(Dutschmann et al., 2020), prime the KFn as a target for volitional motor commands

for orofacial behaviour. For instance, laryngeal adduction and the coordination of

facial, vagal, and hypoglossal nerve activity are prerequisites for the mediation of

swallowing (Bautista and Dutschmann, 2014; Dick et al., 1993) and sniffing (Cobos

Pallares et al., 2016; Semba and Egger, 1986). The present study now identifies

significant descending input from the piriform and rhinal cortices to the KFn,

suggesting that the KFn nucleus might receive major primary olfactory information

(Blazing and Franks, 2020; Chapuis et al., 2013; Leitner et al., 2016). Additional

substantial descending inputs to the KFn originate from the somatosensory barrel

cortex, implying that the KFn also receives tactile sensory information from the whisker

pads (Petersen, 2019; Woolsey and van der Loos, 1970). In summary, because the

KFn gates facial and upper‐airway respiratory motor activity (Dutschmann et al., 2020)

and also receives cortical mono‐synaptic tactile and olfactory sensory inputs, it is

tempting to postulate that the KFn mediates volitional motor commands to coordinate

breathing activity with sniffing and whisking. However, although significantly weaker,

parallel descending projections from rhinal and somatosensory cortices to the pre‐

BötC may determine the frequencies of whisking and sniffing (Kleinfeld et al., 2014).

Thus, the coordination of an integrated behaviour of breathing, sniffing and whisking



















80

may require the synaptic interactions between the tightly connected pre‐BötC and KFn

(Tan et al., 2010; Trevizan‐Baú et al., 2020; Yang and Feldman, 2018).

Conclusions

In conclusion, the descending projection pattern of neurons of the widely

distributed limbic network (i.e., amygdala, rhinal cortex, endopiriform, and the PFC)

connects to a similarly distributed network of respiratory control neurons from the

midbrain to ponto‐medullary brainstem. However, it is likely that specific volitional

motor commands for vocalisation are mediated via the midbrain PAG, while the

coordination of whisking, sniffing, and swallowing with breathing activity is mediated

by inputs that target the KFn. Among the medullary respiratory nuclei, the pre‐BötC

and caudal raphé receive limited descending input from the cortex (e.g., motor cortex),

and therefore may have some additional function in the mediation or synaptic priming

of volitional motor commands.




81

Chapter 4

Reciprocal connectivity of the periaqueductal gray with the

ponto-medullary respiratory network in rat

Pedro Trevizan-Baú1, Werner I. Furuya1, Stuart B. Mazzone2, Davor Stanic1, Rishi R.

Dhingra1, Mathias Dutschmann1

1 The Florey Institute of Neuroscience and Mental Health, The University of Melbourne,

Parkville, Victoria 3010, Australia.

2 Department of Anatomy and Neuroscience, The University of Melbourne, Parkville,

VIC 3010, Australia.

The following chapter was published by Brain Research on January 27, 2021 (DOI:

10.1016/j.brainres.2020.147255). The full paper has been inserted as a thesis chapter

in accordance with The University of Melbourne guidelines.

82

Abstract

Synaptic activities of the periaqueductal gray (PAG) can modulate or appropriate the

respiratory motor activities in the context of behaviour and emotion via descending

projections to nucleus retroambiguus. However, alternative anatomical pathways for

the mediation of PAG-evoked respiratory modulation via core nuclei of the brainstem

respiratory network remains only partially described. We injected the retrograde tracer

Cholera toxin subunit B (CT-B) in the pontine Kölliker-Fuse nucleus (KFn, n = 5),

medullary Bötzinger (BötC, n = 3) and pre-Bötzinger complexes (pre-BötC; n = 3), and

the caudal raphé nuclei (n = 3), and quantified the descending connectivity of the PAG

targeting these brainstem respiratory regions. CT-B injections in the KFn, pre-BötC,

and caudal raphé, but not in the BötC, resulted in CT-B-labeled neurons that were

predominantly located in the lateral and ventrolateral PAG columns. In turn, CT-B

injections in the lateral and ventrolateral PAG columns (n = 4) produced the highest

numbers of CT-B-labeled neurons in the KFn and far fewer numbers of labeled

neurons in the pre-BötC, BötC, and caudal raphé. Analysis of the relative projection

strength revealed that the KFn shares the densest reciprocal connectivity with the PAG

(ventrolateral and lateral columns, in particular). Overall, our data imply that the PAG

may engage a distributed respiratory rhythm and pattern generating network beyond

the nucleus retroambiguus to mediate downstream modulation of breathing. However,

the reciprocal connectivity of the KFn and PAG suggests specific roles for synaptic

interaction between these two nuclei that are most likely related to the regulation of

upper airway patency during vocalisation or other volitional orofacial behaviours.

83

Introduction

The midbrain periaqueductal gray (PAG) plays a critical role in integrating

autonomic function with numerous behavioural responses such as pain, threat,

vocalisation, emotion, and micturition (Bandler et al., 2000; Benarroch,

2012; Dampney et al., 2013). For example, it is well-reported that stimulation of the

lateral and ventrolateral cell columns of the midbrain periaqueductal gray (lPAG and

vlPAG, respectively) can elicit behavioural modulations and appropriations of ongoing

respiratory activity, in particular during vocalisation (Jürgens and Pratt, 1979; Jürgens,

1994; Jürgens, 2009; Larson and Kistler, 1984; Magoun et al., 1937). In addition,

these cell columns of the PAG have function in the mediation of cardio-respiratory

effects associated with emotions (Carrive et al., 1997; Fanselow, 1991; Walker and

Carrive, 2003). Thus, the PAG is seen as the primary interface that connects cortical

and limbic forebrain circuits with autonomic networks in the brainstem (Dampney et

al., 2013; Subramanian and Holstege, 2014).

The neuroanatomical framework for PAG-evoked modulation of cardio-

respiratory activity is linked to descending PAG projections that target the nucleus

retroambiguus (NRA) in the caudal medulla oblongata (Holstege, 1989; Holstege et

al., 1997; Vanderhorst et al., 2000). In turn, the NRA projects to motoneurons of

the nucleus ambiguus (Vanderhorst et al., 2001) and to various pools of respiratory

motoneurons in the ventral horn of the spinal cord (Holstege et al., 1997; Vanderhorst

and Holstege, 1995). Therefore, the PAG-NRA pathway is considered essential for

PAG-mediated volitional and emotional modulation of breathing (Faull et al., 2019;

Subramanian et al., 2008a,b; Subramanian and Holstege, 2014). However, while the

NRA has a role in respiratory rhythm generation (Jones et al., 2012), it does not

contribute to the generation of the respiratory motor pattern (Jones and Dutschmann,


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2016), which depends on synaptic interactions between the rhythmogenic pre-

Bötzinger complex (pre-BötC; Feldman and Del Negro, 2006; Smith et al., 1991) in the

rostral medulla, and pontine circuits (Jones and Dutschmann, 2016). Given that the

PAG is not an integral part of the respiratory pattern generating circuit (Farmer et al.,

2014), the mediation of respiratory modulation via the PAG-NRA pathway may not

bypass the ponto-medullary rhythm and pattern generating network. Indeed, various

anterograde and retrograde tracing studies have reported that descending PAG

projections also target broader pontine and medullary brainstem areas, including

the pontine tegmentum (Holstege, 1991), the rostral ventrolateral medulla (Carrive et

al., 1988; Carrive et al., 1989; Carrive and Bandler, 1991; Yasui et al., 1990), and the

caudal raphé nuclei (Beitz et al., 1983; Cowie and Holstege, 1992; van Bockstaele et

al., 1991). However, many of these studies were performed in cat and were often

focused on general PAG pathways linked to cardiorespiratory regulation during

defense behaviour. The specific connectivity with core nuclei of the brainstem

respiratory rhythm and pattern generating network were often not specifically

investigated, except in the work of Krout et al. (1998) who reported a modest amount

of anterograde-labeled fibers in the Kölliker-Fuse nucleus (KFn), a core nucleus of the

pontine respiratory group (Dutschmann and Dick, 2012).

To develop a better understanding of the anatomical frameworks for the PAG-

mediated modulation of breathing in the context of emotion and behaviour, we

specifically assessed the relative strength of the reciprocal connectivity of the lPAG

and vlPAG with core nuclei of the respiratory network such as the Bötzinger complex

(BötC), pre-BötC, KFn, and the caudal raphé nuclei using a retrograde tracing

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Material and Methods

Animals

Adult Sprague-Dawley rats of either sex (n = 25, weight range: 280–350 g)

were used. Rats were housed under a 12:12 h light/dark cycle, with free access to lab

chow (Ridley Corporation Limited, Australia) and water. All experimental procedures

were approved by the Florey Institute of Neuroscience and Mental Health Animal

Ethics Committee (AEC 18–074) and performed in accordance with the National

Health and Medical Research Council of Australia code of practice for the use of

animals for scientific purposes.

Surgery

Details of the full surgical procedures are published in a recent study (Trevizan-

Baú et al., 2021a). In brief, after craniotomy, a 1 µl Hamilton syringe (25 s-gauge

needle) filled with 1% CT-B (1 mg/mL; Invitrogen, OR, USA) was used to pressure

inject 150nL of the retrograde tracer (20 nL/min) unilaterally (left side of the brain) in

the KFn, BötC, pre-BötC, caudal raphé nuclei, or PAG. We used the following

coordinates: KFn: −9 mm relative to bregma (A/P), 2.6 mm lateral to the midline (M/L),

and 6.5 mm ventral to the dorsal surface of the brain (D/V); BötC: −9 mm A/P (angled

at 22° backwards), 2 mm M/L, and 9.5 mm D/V; pre-BötC: −9.5 mm A/P (angled at 22°

backwards), 2 mm M/L, and 9.5 mm D/V; caudal raphé nuclei: −8.7 mm A/P (angled

at 22° backwards), 0 mm M/L , and 9.4 mm D/V; and PAG: −8 mm A/P, 0.8 mm M/L,

and 4.8 mm D/V. Injections in the medullary regions (i.e., BötC, pre-BötC, and caudal

raphé), required an adaptation and the syringe was angled at 22° backwards from

vertical to avoid bleeding from superficial cerebral blood sinuses. After injections were

completed, the syringe remained in the brain tissue for 10 min and was withdraw at



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1 mm/min to minimize non-specific spread of the tracer along the injection tract

(Finkelstein et al, 2000).

Tissue preparation and immunohistochemistry

The full experimental protocols for the immunohistochemistry are published

in Trevizan-Baú et al. (2021a). In brief, 12–14 days post-injections, rats were

transcardially perfused with paraformaldehyde (PFA, Sigma-Aldrich) containing

0.2% picric acid (Sigma) diluted in 0.16 M phosphate buffer (Merck KGaA, Darmstadt,

Germany; pH 7.2, initially at 37 °C and subsequently at 4 °C). After postfixation with

PFA/picric acid solution for 90 min at 4 °C, the brains were immersed for 72 h in 0.1 M

phosphate buffer (pH 7.4) containing 15% sucrose, 0.01% sodium azide (Sigma) and

0.02% bacitracin (Sigma) for cryoprotection. Brains were divided into two part at

bregma level of −5 mm, rapidly frozen using liquid carbon dioxide, and stored at −80 ͦC

until sectioning. The caudal part of the brain was entirely sectioned with a cryostat

(Leica CM1850, Leica Microsystems; coronal sections of 40 µm thickness) and stored

in a cryo-protectant solution [30% v/v ethyleneglycol (Merck); 15% w/v sucrose; 35%

v/v 0.1 M phosphate buffer; 35% v/v distilled H2O] at − 20 °C. Importantly, before

sectioning the brain, a small superficial incision was made from the midbrain to the

brainstem to distinguish the ipsilateral and contralateral sides of the brain relative to

the injection site. Free-floating sections were washed in 0.01 M PBS, followed by

incubation in hydrogen peroxide for 20 min and incubated for 24 h at room

temperature (RT) with a goat anti-CT-B antibody (1:10,000 List Biological

Laboratories, CA, USA). Then, sections were washed in 0.01 M PBS, blocked with 5%

normal donkey serum (NDS) in 0.01 M PBS for 1 h at RT, incubated in the

corresponding secondary antibody (anti-sheep biotin, Jackson ImmunoResearch

Laboratories, West Grove, PA; 1:500 in 0.01 M PBS) for 1 h at RT, and subsequently



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incubated in an ABC kit (Vectastain® Elite ABC-HRP Kit) for 1 h at RT. Finally,

sections were incubated in diaminobenzidine hydrochloride (DAB) substrate (1:10,

Roche Diagnostics Mannheim, Germany) for 4 min, followed by 1% H2O2 incubation

for 4 min. After several washing steps, the sections were then mounted on slides and

cover-slipped.

Data analysis and image processing

To identify and document the location of the midbrain and brainstem injection

sites, we used a brightfield microscope (Leica Biosystems). Brain sections with the

CT-B injections were counterstained (see immunohistochemistry protocol; previous

section), photographed, and represented on a schematic drawing (Figs. 2.1B; 2.3A).

Additional information regarding the distribution and location of the injection sites are

detailed in our previous study (Trevizan-Baú et al., 2021a). For instance, we were able

to differentiate the location of CT-B injections in the ventrolateral medulla (i.e., pre-

BötC and the BötC) by observing the so-called anatomical landmarks of the medullary

sections. For example, the hypoglossal nucleus (XII) in the dorsomedial region of the

medullary sections is present in the pre-BötC coronal sections, but not in the BötC

sections. Another critical anatomical landmark for differentiating injections in the BötC

nucleus is the medial longitudinal fasciculus (mlf) in the midline of the medulla. In the

BötC coronal sections, mlf is ventral-dorsally prolonged till the fourth ventricle (4 V) of

the section, while mlf is less ventral-dorsally prolonged in the pre-BötC sections.

For analysis of the retrogradely CT-B-labeled cells, we quantified the number

of CT-B-labeled neurons detected in the brain region of interest of all experimental

cases. We analysed sections separated by 120 µm (every 3rd 40 µm-sections). CT-

B-labeled neurons were recognized by the presence of black punctate granules in the

neuronal cell bodies (somata) and dendrites, and the absence of stained cell nuclei

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(for details, see Trevizan-Baú et al., 2021a). Cell counts were analysed with GraphPad

Prism (version 7.02; GraphPad Software; San Diego, CA). Data were analysed by

One-way ANOVA followed by Tukey’s multiple comparison test to compare the

number of CT-B-labeled neurons between the target regions. To determine statistical

significance between the numbers of CT-B-labeled neurons in the ipsilateral and

contralateral hemisphere, data were analysed by paired t-test (parametric). All values

are presented as mean of the number of CT-B-labeled neurons per experimental

case ± standard error of the mean (SEM), and p-values<0.05 were considered

statistically significant.

A digital camera (Leica DFC7000T) mounted to the microscope (Leica DM6B

LED) was used to take representative images. To best represent sections viewed

under the microscope, we used the Adobe Photoshop CC19 software (Adobe Systems

Inc., San Jose, CA) to optimize brightness and contrast of the digital images, as well

as to assemble representative images.

Results

We microinjected the retrograde tracer CT-B (150 nL) unilaterally in the pontine

KFn (n = 5), the medullary BötC (n = 3), the pre-BötC (n = 3), and the

midbrain PAG (n = 4); and on the midline into the caudal raphé (n = 3). CT-B injections

were anatomically confined to discrete injection sites in the brainstem target areas

(see representative photographs, Fig. 4.1A & 4.4). Note that the same tracer injections

were reported in a most recent study from our laboratory that performed an extensive

analysis of descending forebrain inputs to these brainstem nuclei (Trevizan-Baú et al.,

2021a).






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Figure 4.1. CT-B injections in the pontine Kölliker-Fuse nucleus, the medullary Bötzinger

complex and pre-Bötzinger complex, and the caudal raphé.

Photomicrographs of representative injections (A) and schematic drawings (B), based on atlas

of Paxinos and Watson (2007), depicting the location of all CT-B injections in the KFn, BötC, pre-BötC,

and caudal raphé relative to bregma. The coloring delineates the rostro-caudal tracer spread for specific

on-target injections. Off-target injections are represented by black X’s. Levels caudal to bregma are

given as an approximate guide relating to the atlas of Paxinos and Watson (2007). Abbreviations:

4 V = fourth ventricle; BötC = Bötzinger complex; KFn = Kölliker-Fuse nucleus; mcp = middle

cerebellar peduncle; io = inferior olive; mlf = medial longitudinal fasciculus; NA = nucleus ambiguus;

NTS = solitary tract nucleus; PAG = periaqueductal gray; pre-BötC =, pre-Bötzinger complex;

py = pyramidal tract; RMg = raphé magnus; ROb = raphé obscurus; RPa = raphé pallidus;

scp = superior cerebellar peduncle; VII = facial nucleus. Scale bars: 200 µm (representative

photographs); 1 mm (schematic drawings).

Distribution of retrogradely labeled neurons in the PAG after CT-B

injections in the ponto-medullary respiratory nuclei





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CT-B injections in the pontine KFn resulted in a mean total of 448 (±121) labeled

neurons per case (n/c; counted every 3rd section) in the series of PAG sections

analysed. Quantification of the ipsilateral and contralateral distributions of these

labeled neurons revealed a significant ipsilateral dominance of descending PAG

projections targeting this region (394 ± 109 vs. 55 ± 15n/c; p < 0.05) (Fig. 4.2). After

CT-B injections into the medullary pre-BötC, we observed a mean of 303 ± 98 labelled

cells in the PAG, which showed a clear tendency towards an ipsilateral PAG projection

to this region (247 ± 84 vs. 56 ± 16n/c). Caudal raphé injections resulted in a mean of

353 ± 109 labelled cells in the PAG and, because injections were on the midline of the

medulla, we observed similar numbers of labeled neurons in both the left and right

sides of the PAG (data not shown). Strikingly, no CT-B-labeled neurons were detected

following tracer injection in the BötC (Fig. 4.2). Analysis of the distribution of CT-B-

labeled neurons in specific longitudinal cell columns of the PAG, which include the

dorsomedial (dmPAG), dorsolateral (dlPAG), lateral (lPAG), and ventrolateral (vlPAG)

columns (Bandler et al., 1991; Carrive, 1993), revealed that the majority of the

projection neurons were located in the lPAG and vlPAG (Fig. 4.3A and B). Although

CT-B-labeled neurons were observed throughout the rostro-caudal extension of the

PAG columns (i.e., rostral, intermediate, and caudal PAG), the majority of the

projection neurons for all experimental cases were located in the intermediate and

caudal aspects of the respective PAG columns (Fig. 4.3C).







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Figure 4.2. Injections in the ponto-medullary target regions (KFn, BötC, pre-BötC, and caudal

raphé) resulted in CT-B-labeled neurons in the midbrain PAG.

Total (circle), ipsilateral (triangle), and contralateral (square) distribution of labeled neurons in the PAG

after CT-B injections in the KFn, BötC, pre-BötC, or caudal raphé. Analysis of the ipsilateral and

contralateral distribution of CT-B-labeled neurons revealed a significant ipsilateral dominance of

descending PAG projections targeting the KFn (p < 0.05), and a tendency of ipsilateral PAG projections

to the pre-BötC. Note that for the caudal raphé injections, however, we only present the total number

of CT-B-labeled neurons in the PAG. Given that injections in the caudal raphé were on the midline of

the medulla, we observed similar numbers of CT-B-labeled neurons in the left and right hemisphere of

the PAG (data not shown).


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Figure 4.3. Projections from the midbrain PAG to the ponto-medullary respiratory control nuclei.

A: Representative images of CT-B-labeled neurons in the ipsilateral hemisphere of a PAG coronal

section (bregma −8.0 mm): i) the photomicrograph at the lowest magnification shows the retrograde

labeled neurons in the context of the PAG columns; ii) at higher magnification (20×), CT-B-labeled

neurons (red arrowheads) in the lPAG and vlPAG are shown; and iii) at 40x, CT-B-labeled neurons (red

arrowheads) are more evident. This representative image of CT-B-labeled cells in the PAG was

produced by a CT-B injection in the pre-BötC (green injection site, schematic drawing of panel B of Fig.



93

4.1). Level caudal to bregma is given as an approximate guide relating to the atlas of Paxinos and

Watson (2007). B: Distribution of the total (circle), ipsilateral (triangle), and contralateral (square)

numbers of CT-B-labeled neurons in the dmPAG, dlPAG, lPAG, and vlPAG after CT-B injections in the

pontine KFn, pre-BötC, or caudal raphé. Overall, we observed a predominance of CT-B-labeled neurons

in the lateral and ventrolateral columns of the PAG. All values are expressed as mean of the number of

CT-B-labeled neurons per experimental case. C: Illustration of the rosto-caudal spread of retrogradely

labeled neurons in specific PAG columns. Abbreviations: Aq = aqueduct; BötC = Bötzinger complex;

KFn = Kölliker-Fuse nucleus; dlPAG = dorsolateral PAG; dmPAG = dorsomedial PAG; lPAG = lateral

PAG; PAG = periaqueductal gray; vlPAG = ventrolateral PAG; pre-BötC = pre-Bötzinger

complex. Scale bars: 100 µm (i); 50 µm (ii); 25 µm (iii).

We had some off-target (i.e., near miss) CT-B injections (n = 6) in the ponto-

medullary brainstem. These injections revealed small numbers CT-B-labeled neurons

in the midbrain PAG (data now shown) compared to the on-target injections. For

instance, two off-target injections that were slightly caudal and ventral to the KFn, but

still in the dorsolateral pons (Fig. 4.1B), presented far fewer CT-B-labeled neurons in

the midbrain PAG. Similarly, we observed that off-target injections in the medulla, such

as two injections slightly lateral to the caudal raphé as well as two injections lateral to

the BötC/pre-BötC, resulted in small numbers of CT-B-labeled neurons in the PAG

compared to the on-target medullary injections.

Distribution of retrogradely labeled neurons in ponto-medullary respiratory

nuclei after CT-B injections in the lateral and ventrolateral columns of the PAG

The locations of CT-B injections in the lPAG and vlPAG are illustrated in Fig.

4.4. Overall, our analysis revealed the highest number of CT-B-labeled neurons in the

KFn (Fig. 4.5A1-4), in comparison to the medullary BötC, pre-BötC, or caudal raphé

(p < 0.05; Fig. 4.5A5-8, B). Specifically, injections in the PAG revealed on average

48 ± 6n/c in the KFn, versus only 6 ± 1n/c in the pre-BötC, 5 ± 2n/c in the BötC, or

1 ± 1n/c in the caudal raphé. Analysis of the laterality of retrogradely neurons in the

pontine KFn indicated the highest number of CT-B-labeled neurons located










94

ipsilaterally (p < 0.05; Fig. 4.5B). We detected 32 ± 3 neurons/case (n/c) in the

ipsilateral KFn and 15 ± 4n/c in the contralateral KFn. In the medullary pre-BötC, we

observed a tendency for higher number of CT-B-labeled neurons in the ipsilateral

hemisphere. We detected 5 ± 1n/c located ipsilaterally and 1 ± 1n/c contralaterally

(Fig. 4.5B). Finally, in the BötC, we detected 2 ± 1n/c in the ipsilateral hemisphere,

and 3 ± 1n/c in the contralateral hemisphere (not significant; Fig. 4.5B). The rostro-

caudal distribution of retrogradely labelled brainstem neurons is summarized in Fig.

4.5C.

Figure 4.4. CT-B injections in the midbrain PAG.

Photomicrograph of a representative CT-B injection in the midbrain PAG (A) and schematic drawings

(B), based on atlas of Paxinos and Watson (2007), depicting the location of all CT-B injections in the

PAG. In this figure, colors code the rostro-caudal tracer spread for specific on-target injections. Levels

caudal to bregma are given as an approximate guide relating to the atlas of Paxinos and Watson (2007).

Abbreviations: Aq = aqueduct; dlPAG = dorsolateral PAG; dmPAG = dorsomedial PAG; lPAG = lateral

PAG; PAG = periaqueductal gray; vlPAG = ventrolateral PAG. Scale bars: 500 µm (representative

photograph); 400 µm (schematic drawings).










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Figure 4.5. Projections from the ponto-medullary respiratory regions to the midbrain PAG.

A: Representative images of CT-B labeled neurons in the KFn (A1-4), pre-BötC (A5-6), and caudal

raphé (A7-8). The photomicrograph on the right (A2, A4, A6, & A8) are of boxed regions marked in A1,

A3, A5 and A7, respectively, to show the CT-B-labeled cells. These representative images of CT-B-

labeled cells were produced by a CT-B injection in the vlPAG (purple injection site, schematic drawing

of panel B of Fig. 4.4). Levels caudal to bregma are given an as approximate guide relating to the atlas

of Paxinos and Watson (2007). B: Total (circle), ipsilateral (triangle), and contralateral (square) cell

counts of CT-B-labeled neurons in the KFn, BötC, pre-BötC, and the caudal raphé after injections in

the PAG. We observed a greater number of CT-B-labeled neurons in the KFn, compared to pre-BötC,





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BötC, and caudal raphé. Additionally, our analysis revealed the highest number of CT-B-labeled

neurons in the ipsilateral KFn (p < 0.05). In the medullary pre-BötC, we observed a tendency of higher

number of CT-B-labeled neurons ipsilaterally, and no laterality was detected for the BötC. Note that we

only present the total number of CT-B-labeled neurons in the caudal raphé because this medullary

region is located in the midline of the medulla. All values are expressed as mean of the number of CT-

B-labeled neurons per experimental case. C: Illustration of the rostro-caudal distribution of retrogradely

labeled neurons in relation to bregma. Abbreviations: 4 V = fourth ventricle; BötC = Bötzinger complex;

io = inferior olive; KFn = Kölliker-Fuse nucleus;; mcp = middle cerebellar peduncle; mlf = medial

longitudinal fasciculus; NA = nucleus ambiguus; NTS = solitary tract nucleus; pre-BötC = pre-Bötzinger

complex; py = pyramidal tract; scp = superior cerebellar peduncle; VII = facial nucleus. Scale bars:

400 µm (A; A1, A3, A5, & A7); 100 µm (A2, A4, A6, & A8).

Note that relative proportion of efferent versus afferent projections is an order

of magnitude different. This could be in part because the entirety of a PAG column

was not covered by a single CT-B injection, whereas most of the ponto-medullary

structures could be covered by a single CT-B injection since that they are smaller.

Given that the injection volumes were of a consistent volume, and were targeted

toward the caudal aspect of the PAG in the case of afferent projections (see Fig. 4.4),

their relative proportions should still be a reliable measure for comparison.

Importantly, we did not observe CT-B-labeled neurons in the ponto-medullary

respiratory regions (i.e., KFn, pre-BötC, BötC, and caudal raphé) after injecting slightly

lateral to the lPAG and vlPAG (off target injection; data not shown).

Summary of the afferent and efferent connectivity of the lateral and

ventrolateral PAG columns with the respiratory nuclei in the ponto-medullary

brainstem

The topographical organization of the descending and ascending PAG

projections is summarized in a network connectivity graph (Fig. 4.6). The graph shows

the relative proportion of efferent projections from the midbrain PAG columns to each





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ponto-medullary target nuclei (Fig. 4.6A), and the afferent inputs to the PAG arising

from the downstream targets (Fig. 4.6B). The functional implications of the reciprocal

connectivity of the PAG in the context of volitional control of breathing are discussed

below.

Figure 4.6. Summary of the relative strength of projections arising from the

midbrain PAG columns (dmPAG, dlPAG, lPAG, and vlPAG; A), and the ponto-medullary regions

(KFn, pre-BötC, BötC, and the caudal raphé; B) in a connectivity map.

The weight of connecting lines are proportional to the normalized maximal number of CT-B-labeled

neurons found in the specific source of descending projections.




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Discussion

The midbrain PAG is thought to play a critical role in modulating breathing via

inputs to the ponto-medullary respiratory network. However, our understanding of the

anatomical connectivity of the PAG with core nuclei of the respiratory network remains

vague. The present study identified neurons in the PAG that target either the pontine

KFn, the caudal raphé, or the medullary pre-BötC, but not the BötC. It is likely that

these descending PAG neurons are important for the volitional modulation or

appropriation of respiratory motor activities. Several studies reported that stimulations

of the PAG (lPAG and vlPAG, especially) evoke strong respiratory modulations and,

in particular, trigger vocalisations (Davis and Zhang, 1996; Farmer et al., 2014;

Hayward et al., 2003; Huang et al., 2000; Jürgens and Pratt, 1979; Jürgens,

1994; Jürgens, 2009; Larson and Kistler, 1984; Magoun et al., 1937; Subramanian et

al., 2008b; Subramanian and Holstege, 2013; Zhang et al., 1994; Zhang et al.,

2007; Zhang et al., 2009). Furthermore, changes in respiratory activity (i.e., an

increase in the respiratory frequency) can also be correlated with defensive

behaviours (e.g., fight and flight, freezing, aggressive or escape actions) that are

elicited by PAG stimulation (Bandler, 1982; Bandler et al., 1985; Bandler and

Depaulis, 1991; Carrive et al., 1997; Fanselow, 1991; Hayward et al., 2003; Walker

and Carrive, 2003). The present study identified approximately equal numbers of PAG

neurons projecting to the KFn, pre-BötC, and caudal raphé nuclei, which have different

functions in respiratory pattern formation (KFn; see Dutschmann and Herbert,

2006; Mörschel and Dutschmann, 2009; Smith et al., 2007), rhythm generation (pre-

BötC; see Feldman and Del Negro, 2006; Smith et al., 1991; Wenninger et al., 2004),

and modulation of the excitability of respiratory network neurons (caudal raphé;

see Hodges and Richerson, 2010; Lindsey et al., 1998; Richter et al., 2003). Taken






































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together, our anatomical finding suggests that the PAG may engage a widely

distributed network of brainstem nuclei to modulate respiratory motor pattern.

It has been proposed that the PAG might be the final common pathway

mediating changes in the eupneic respiratory motor pattern in the context of various

volitional behaviours and emotions (Subramanian and Holstege, 2014), including

vocalisation (Düsterhöft et al., 2004; Holstege, 1989; Jürgens and Richter,

1986; Jürgens, 1994; Jürgens, 2002a,b; Subramanian and Holstege, 2009).

However, the proposed descending pathway that connects the PAG output with

laryngeal and spinal respiratory motor neurons was solely based on PAG projections

to the nucleus retroambiguus (NRA) in the caudal medulla (Holstege, 1989; Holstege

et al., 1997; Vanderhorst et al., 2000). In the PAG-NRA hypothesis, the NRA

subsequently acts as an interface that distributes PAG signals to laryngeal motor

neurons of the nucleus ambiguus (Vanderhorst et al., 2001) and various respiratory

motor pools in the spinal cord (Holstege et al., 1997; Vanderhorst and Holstege, 1995),

bypassing the respiratory rhythm and pattern generating network in the ponto-

medullary brainstem, and is thought to be critical for PAG-mediated modulation of

breathing, particularly during vocalisation (Subramanian and Holstege, 2009; Zhang

et al., 1995). However, recent work from our laboratory has shown that the eupneic

respiratory motor pattern is generated by an anatomically widely-distributed brainstem

network (Dhingra et al., 2019a,b; Dhingra et al., 2020). Consistent with these

observations, the present study shows that the lPAG and vlPAG send multiple

descending projections to these widely-distributed and functionally diverse brainstem

respiratory control nuclei. For instance, in line with a previous study, the PAG receives

afferents from the respiratory rhythm generating circuits of the pre-BötC (Yang and

Feldman, 2018), and, therefore, may receive information regarding the respiratory
























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rhythm. However, compared to pre-BötC, we observed that the KFn has significantly

stronger afferent inputs to the PAG. Thus, the lPAG and vlPAG columns reciprocally

connected with the KFn, which also receives substantial afferent input from the NRA

(Jones et al., 2016).

In a recent study of the descending forebrain projections targeting the

brainstem nuclei important for breathing control, we reported efferent cortical

projections predominantly target the PAG and KFn (Trevizan-Baú et al., 2021a). The

reciprocal connectivity of the PAG and NRA with the KFn seems a coherent

anatomical structure for the mediation of volitional respiratory modulation (e.g.,

vocalisation and sniffing; see Trevizan-Baú et al., 2021a), which requires control of

expiratory airflow via glottal adduction during the postinspiratory phase (Dutschmann

et al., 2014). In contrast, changes in inspiratory frequency associated with emotion

(Homma and Masaoka, 2008) or behaviour (Faull et al., 2019; Subramanian et al.,

2008a,b; Subramanian and Holstege, 2014) might be mediated via forebrain to PAG

projections (this study; Trevizan-Baú et al., 2021a) targeting the pre-inspiratory rhythm

generating circuit of the pre-BötC (Subramanian and Holstege, 2013).

Conclusion

The midbrain PAG projects to the ponto-medullary respiratory network (i.e.,

pontine KFn, medullary pre-BötC, and the caudal raphé), and receives ascending

projections from those respiratory control areas, especially from the KFn. The

reciprocal PAG-KFn connectivity provides an anatomical framework for the regulation

of upper-airway patency during the mediation of breathing modulation in the context

of volitional and emotional behaviours.













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Chapter 5

General Discussion

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Abstract

Breathing is the only vital function that can be volitionally controlled in addition to its

modulation and adaptation via reflexive viscero- and somatosensory feedback

mechanisms and emotions. Neuroanatomical tract tracing studies have identified

patterns and origins of descending forebrain projections that target the midbrain

periaqueductal gray and nuclei of the respiratory rhythm and pattern generating

network in the brainstem, specifically including the pontine Kölliker-Fuse nucleus and

the pre-Bötzinger complex in the medulla oblongata. This review discusses the

potential functional implications of the forebrain-brainstem connectivity that could

underlie the volitional control of orofacial behaviours and their coordination with

breathing.

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Emerging concepts for functional neuroanatomy underlying the coordination of

volitionally evoked orofacial behaviour and breathing

A neural network in the pons and medulla oblongata generates the eupneic

respiratory rhythm and motor pattern. The synaptic mechanism and operational

principles of respiratory rhythm and pattern generation were extensively reviewed

elsewhere (Champagnat et al., 2009; Dutschmann and Dick 2012; Feldman 1986;

Feldman and Del Negro, 2006; Feldman et al., 2013; Mörschel and Dutschmann,

2006; Onimaru et al., 2009; Ramirez and Baertsch, 2018; Richter and Smith, 2014;

Richer, 1982; St-John, 1998; Smith et al., 2009; Smith et al., 2013). The primary

activity of pattern of the respiratory network is further shaped by viscerosensory

feedback from the lungs (pulmonary stretch receptors; see Kubin et al., 2006;

Widdicombe, 1982) or by oxygen sensors of the carotid body (Guyenet, 2013; Spyer

and Gourine, 2009; Wilson and Teppema, 2016). In addition, stimulation of viscero-

and somatosensory afferents can trigger a variety of orofacial behaviours and

expulsive airway reflexes that recruit respiratory motor activity and require the

modulation of ongoing respiratory network activity during vocalisation, swallowing,

chewing, coughing, sneezing, sighing, sniffing and whisking (Davis et al., 1996;

Nanoka et al., 1999; Jean 2001; Ludlow, 2005; Sherwood et al., 2005; Deschênes et

al., 2012; Moore et al., 2013; Moore et al., 2014; Laplagne, 2018; McElvain et al.,

2018; Li et al., 2020). All aforementioned orofacial behaviours and expulsive reflexes,

except sneezing, which cannot be consciously controlled (Songu and Cingi, 2009), are

also initiated by emotional and cognitive commands arising from neural circuits in the

forebrain.

The anatomical frameworks underlying the cognitive and emotional modulation

of respiratory motor activities during volitionally evoked orofacial behaviours are

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starting to emerge. This review discusses the potential functional implications of

descending projections that connect supra-pontine brain regions with orofacial and

respiratory motor pools. In particular, we highlight recent neuroanatomical discoveries

that suggest that volitional motor commands target pre2motor in the midbrain and

premotor nuclei of the respiratory network in the pontomedullary brainstem to

coordinate volitional orofacial behaviour and respiration. The last sections of this

review emphasize the point that impaired orofacial behaviour and its coordination with

ongoing respiration are general symptoms in a wide-selection of neurological

disorders ranging from neurodegenerative diseases to autism spectrum disorders.

Pivotal role of postinspiratory control of glottal adduction in coordinating

orofacial behaviours and breathing

The respiratory motor cycle is divided into three distinct phases: inspiration,

postinspiration (early expiration), and late expiration. During eupnea (normal

respiration), respiratory motor activity are present in all cranial motor nerves that also

mediate appropriate orofacial behaviours (Fig.1a). Motor pools in spinal cord drive the

respiratory pump muscles such as the diaphragm during inspiration and abdominal

expiratory neurons during eupnea and under high metabolic demands or chemical

drive that require active expiration.

The postinspiratory phase, as the centre phase of the respiratory motor cycle,

holds a key position for the coordination of orofacial behaviour and breathing. During

eupnea, the contraction of glottal adductor (constrictor) muscles breaks the expiratory

airflow generated by the passive recoil force of the inflated lungs and extended ribcage

to control the expiratory airflow profile during the postinspiratory phase of the

respiratory motor cycle (for details, see Dutschmann et al., 2014). The dynamic

modulation of upper airway patency is critically involved in respiratory-related

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behaviours such as vocalisation, sniffing and swallowing. The vocal sound is

exclusively generated during postinspiration, when the expiratory airflow vibrates the

constricted vocal folds of the larynx (Riede, 2011). Although sniffing is usually referred

to as an inspiratory behaviour linked to the contraction of diaphragm (Moore et al.,

2013; Kleinfeld et al., 2014), initial glottal constriction during sniffing (Mukai, 1989;

Lawson et al., 1991; Pérez de Los Cobos Pallares et al., 2016) is required for the

initiation and generation of characteristic high nasal flow pressures of sniff; and

restriction of inspiratory and expiratory airflow is essential to maintain blood gas

homeostasis. The latter is crucial for rodents that generate sniffing frequencies of up

to 12Hz (Wesson et al., 2008). Feeding behaviour includes mastication and

swallowing since the pathways for air and food cross in the pharynx (Matsuo and

Palmer, 2009). During swallowing, the postinspiratory activity inhibits respiration and

constriction of the epiglottis prevents pulmonary aspiration (Bautista et al., 2014;

Bautista and Dutschmann, 2014; Dutschmann et al., 2014) during the propulsion of

food from the oral cavity into the stomach (Nishino et al., 1985; Selley et al., 1989).

Increased glottal resistance is also seen during mastication (Fontana et al., 1992) and

the masseter masticatory, as main masticatory muscle (Hylander et al., 1992),

interestingly also receives postinspiratory drive during eupnea (Fig. 1; St-John, 1998).

Thus, the central pre2motor and premotor circuits that generate and control

postinspiratory motor activity during breathing also have key function for mediating

reflex or volitionally evoked motor behaviour.

The neural circuit for the generation and modulation of upper airway patency

during inspiration and postinspiration (controlled expiration) appears to be

anatomically widely distributed across the ponto-medullary brainstem (Dhingra et al.,

2019a,b; Dhingra et al., 2020). Nevertheless, several nuclei may have a more

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significant role in the generation and shaping of respiratory motor pattern than others

(Dhingra et al., 2021). Key premotor areas for the initiation and generation of the

postinspiratory phase include: i) the pre-Bötzinger complex (pre-BötC) in the rostral

ventrolateral medulla, which is better known for its role in inspiratory rhythm generation

(Smith et al., 1991; Feldman and Del Negro, 2006; Del Negro et al., 2018), but was

recently shown to also have a significant function in the generation of eupneic

postinspiratory motor activity (Dhingra et al., 2019a,b; Dhingra et al., 2020); ii) the

adjacent Bötzinger complex (BötC), which is considered an essential medullary region

that controls the initiation of postinspiration (Burke et al., 2010; Smith et al., 2013;

Marchenko et al., 2016), and, for instance, synaptic interactions between the pre-BötC

and BötC are considered to reflect the respiratory core circuit for the generation of the

eupneic respiratory motor pattern in many computational or conceptual models (Rybak

et al., 2007; Smith et al., 2013; Richter and Smith, 2014; Marchenko et al., 2016;Del

Negro et al., 2018); iii) the pontine the Kölliker-Fuse nucleus (KFn), which plays a

critical role in gating eupneic respiratory activity in cranial nerves (vagus, hypoglossal,

facial nerves; Dutschmann et al., 2020), in particular the vagal postinspiratory activity

in the efferent vagus (Dutschmann and Herbert 2006). In addition, the KFn, as part of

the pontine pneumotaxic centre (Markwald, 1888; Lumsden 1923), is critical for the

mediation of the inspiratory-postinspiratory phase transition (Morschel and

Dutschmann, 2009; Dutschmann and Dick, 2012) and thus most likely for the initiation

and gating of orofacial behaviour which almost exclusively occurs during the

postinspiratory phase.

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Figure 5.1. Respiratory motor activity is evoked by cranial and spinal nerves.

(A) Cranial and spinal motor neurons innervate respiratory muscles in the upper airway, oronaso-

pharyngeal and thoracic cavities, and abdomen. The primary control areas of the brainstem respiratory

network, which is anatomically organised from the dorsolateral pons to the caudal ventrolateral medulla

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(represented in dark gray), control respiratory activity via the trigeminal (in cyan), facial (in magenta),

hypoglossal (in blue), vagus (in red), phrenic (in green), and abdominal (in orange) motor pools. (B)

Schematic representation of respiratory motor outputs, subglottal pressure and upper airway airflow in

relation to the three phases of the respiratory cycle (I, inspiratory phase; PI, postinspiratory phase; and

E2, late expiratory phase) during eupneic breathing, active expiration, vocalisation, and sniffing.

Inspiratory motor activity is evoked by phrenic motoneurons (PNA, in green) that innervate diaphragm,

the main inspiratory pump muscle. However, facial (FNA, in red), hypoglossal (HNA, in blue) and

recurrent laryngeal (RLNA: a branch of the vagus nerve, in red) nerves present pre-inspiratory and

inspiratory discharge patterns, facilitating oronaso-pharyngeal airway passage during inspiration. In

addition to pre-I and I motor activity during eupneic breathing, hypoglossal nerve shows i) expiratory

motor discharge to decrease subglottal pressure during active expiration (Britto and Moraes, 2016); and

ii) postinspiratory activity to modulate the tongue during vocalisation. The recurrent laryngeal nerve

plays a critical role on upper airway modulation during respiration and orofacial behaviours. Laryngeal

motor activity (i.e., glottal adduction and abduction) modulates expiratory airflow during postinspiration.

The modulation of expiratory airflow is critical for numerous orofacial behaviours, such as vocalisation

(Riede, 2011; Sirotin et al., 2014) and sniffing (de los Cobos Pallares et al., 2016). For vocalisation, the

modulation of expiratory airflow during postinspiration vibrates the vocal folds located within the glottis.

During sniffing (nasal ventilation), glottal constriction prevents lung ventilation. The abdominal nerve

presents postinspiratory motor activity (AbNA, in orange) in eupneic breathing and late expiratory

activity during active expiration. Additionally, USV emission (vocalisation) requires abdominal nerve

discharge during postinspiration (Lancaster and Speakman, 2001). Trigeminal motor activity (TNA, in

cyan) is also important for facilitating airway passage during early expiration in eupneic breathing as

well as during late expiration for active expiration. Note that subglottal pressure (SGP) is related to

airway resistance and pulmonary airflow (PAF). Abbreviations: 7N = facial nucleus; 12N = hypoglossal

nucleus; C4 = cervical spinal nerve 4; exp = expiration; insp = inspiration; L1 = lumbar spinal nerve 1;

MO5 = motor trigeminal nuclei; NA = nucleus ambiguous; USV = ultrasonic vocalisation.

Corticobulbar and corticospinal tracts and their potential involvement in

volitional control of respiratory muscles

Breathing is tightly coordinated during the mediation of consciously and

reflexively controlled orofacial behaviours. Forebrain circuits can volitionally control

respiration and orofacial behaviours (e.g., cortex; Pouget et al., 2018) and are also

influenced by interoceptive signals (see Faull et al., 2019) and emotions (see Homma

and Masaoka., 2008). Human and animal studies proposed that volitional control of

the respiration could be directly mediated via cortical modulation of spinal and cranial

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motor activity, bypassing the primary respiratory network in the brainstem (Butler,

2007; Corfield et al., 1998; Gandevia and Rothwell, 1987a,b; Pouget et al., 2018).

Neuroanatomical studies have extensively shown that the motor cortex

(Kuypers, 1958; Shriver and Noback, 1967; Rikard-Bell, et al., 1985; Yasui et al., 1985;

Sreenivasan, et al., 2015) and regions of the prefrontal cortex (e.g., infralimbic; Hurley

et al., 1991) send monosynaptic inputs to the brainstem cranial motor pools, such as

the trigeminal, facial, hypoglossal, and nucleus ambiguus, as well as the spinal cord

motor pools in the cervical, thoracic, and lumbar segments (Fig.2). Descending

corticobulbar projections specifically originate from areas of the motor cortex linked to

control of whisker (vibrissae) movements (Grinevich et al., 2005) and laryngeal motor

control (Simonyan and Horwits, 2011). Consequently, these long-range descending

projections of layer V pyramidal neurons (Kuypers, 1958; Rikard-Bell, et al., 1985;

Shriver and Noback, 1967; Yasui et al., 1985) were suggested to play essential roles

for the mediation of volitional orofacial motor behaviours such as whisking, swallowing

and vocalisation.

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Figure 5.2. Corticobulbar and corticospinal projections into the motor pools.

Pyramidal neurons of layer V in the motor cortex project to the respiratory motor pools in the brainstem

and spinal cord. These corticobulbar and corticospinal tracts were postulated to mediate the contraction

of cranial and abdominal respiratory muscles during voluntary orofacial motor behaviours, such as

chewing, sniffing, whisking, vocalisation and swallowing. Abbreviations: 7N = facial nucleus; 12N =

hypoglossal nucleus; C = cervical spinal nerves (e.g. phrenic); L = lumbar spinal nerve (e.g.

iliohypgastric nerve); MO5 = motor trigeminal nuclei; NA = nucleus ambiguous; T = thoracic spinal

nerves (e.g. intercostal nerves).

Given that layer V pyramidal neurons are the primary excitatory output neurons

in the motor cortex (DeFelipe and Farinas, 1992; Bekkers, 2011), it was postulated

that they directly control the contraction of respiratory muscles in the oronaso-

pharyngeal (e.g., nasalis and tongue) and thorax and abdomen (e.g., diaphragm and

internal obliques). However, the generation and timing of sequential contractions of a

variety of anatomically and functionally different muscles and their coordination with

respiration may require not only excitatory, but also inhibitory synaptic mechanisms.

Particularly in humans, it has been thought that the motor cortex control respiratory

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muscles (including orofacial muscles) during voluntary motor behaviours, presumably

bypassing the brainstem (Benecke et al., 1988; Gandevia and Rothwell, 1987a,b;

Pouget et al., 2018). However, some studies using transcranial magnetic stimulation

technique show that there is no evidence of intracortical inhibition to coordinate facial

muscle activity, and, thus, proposing that volitional information initiated in the motor

cortex might be relayed in the brainstem (Kobayashi et al., 2001; Ginatempo et al.,

2019). Studies in animals have shown that inhibitory and excitatory pre-motor neurons

in the brainstem network coordinate orofacial motor activity during whisking and

sniffing (Bellavance et al., 2010; Bellavance et al., 2017; Furuta et al., 2008; Moore et

al., 2013;), as well as chewing (Nakamura and Katakura, 1995), and swallowing (Saito

et al., 2003; Bautista and Dutschmann, 2014). Therefore, functional data rather

support the notion that brainstem premotor networks are indispensable for the

patterning of muscle contraction during orofacial behaviours and even more so when

orofacial behaviours need to be coordinated and timed with ongoing breathing activity

(Moore et al., 2013; Moore et al., 2014; Barlow et al., 2010; Dutschmann et al., 2014).

The same concepts also apply to orofacial behaviours that are less directly intertwined

with respiration such as the control of the whisker movements or mastication. A recent

functional and neuroanatomical study has shown that corticobulbar projections

neurons engage a premotor networks in the brainstem (trigeminal pre-motor nuclei) to

control vibrissae and jaw movements (Lindsay et al., 2019).

In summary, orofacial motor networks often directly interact to produce

coordinated motor pattern engaging distributed networks across the entire neuro axis

(Dickinson, 2006). This is especially true for oromotor network (Barlow et al., 2010;

Kleinfeld et al., 2014; McElvain et al., 2018) in the mammalian brain. While the

existence of cortical motor CPGs are considered (Yuste et al., 2005), this concept still

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lacks clear experimental evidence to our knowledge. Thus, the synaptic interactions

between multiple orofacial motor network and the pontomedullary respiratory circuit in

the brainstem are essential for the generation and patterning of coordinated orofacial

motor behaviours. We suggest that the descending cortical inputs to cranial orofacial

and respiratory motor pools may rather prime the synaptic inputs from orofacial-

respiratory pre2motor and premotor networks than mediating these motor outputs. In

order to develop an understanding of cross-modal and cross-network facilitation of the

coordination of orofacial-respiratory behaviours, it is necessary to consider the relay

of descending forebrain motor commands within pre2motor areas such as the

periaqueductal gray (PAG) in the midbrain and respiratory and orofacial pre-motor

circuits of the brainstem (see below).

The periaqueductal gray, a pre2motor area in the midbrain: an established

interface for volitional and emotion control of vocalisation

An established brain area that relays descending forebrain inputs to autonomic

and respiratory networks in the brainstem is the midbrain periaqueductal gray (PAG),

and, therefore, the PAG was postulated to interface behavioural and emotional motor

commands to adapt cardio-respiratory activity during a wide-range of behaviours

(Bandler et al., 2000; Benarroch, 2012; Dampney et al., 2013; Faull et al., 2019).

Observations drew major interest that pharmacological and/or electrical stimulation of

the PAG can elicit vocalisations (Magoun et al., 1937; Jürgens and Pratt, 1979; Larson

and Kistler, 1984; Zhang et al., 1994, 2007, 2009; Davis and Zhang, 1996; Jürgens,

1994, 2009) and/or profound modulation of cardio-respiratory activity (Huang et al.,

2000; Hayward et al., 2003; Subramanian et al., 2008a,b; Subramanian and Holstege,

2013; Farmer et al., 2014). The hypothesis of the PAG serving as an essential

interface between the forebrain and the autonomic centers in the brainstem is strongly

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supported by the presence of descending projections from the cortex (e.g., motor,

insular, somatosensory, cingulate, piriform, and rhinal cortices) and limbic brain

regions (e.g., amygdala, lateral septum, hypothalamus, claustrum) (Floyd et al., 2000;

Dampney et al., 2013; Trevizan-Baú et al., 2021a) to the PAG. In turn, the PAG sends

descending inputs to the nucleus retroambiguus (NRA) at the caudal end of the

medulla oblongata. The NRA projects to the respiratory motor pools of the nucleus

ambiguus (NA) and the spinal cord (Holtsge, 1989; Holstege, 1997; Vanderhost, et al.,

2000). Because of the connectivity of NRA with cranial and spinal respiratory motor

pools, some authors have argued that PAG-NRA projections could be a ‘final common

pathway’ for the adaptation of respiration with behaviours (Düsterhöft et al., 2004;

Holstege, 1989; Holstege, 2009; Jürgens, 1994; Jürgens, 2002a,b; Jürgens and

Richter, 1986, Subramanian and Holstege, 2013). However, the PAG also sends

inputs to pre-motor nuclei of the respiratory network in the pons (e.g., inputs to the

KFn) and medulla oblongata (e.g., inputs to the ventral respiratory group, overlapping

with the pre-BötC and BötC) (Carrive et al., 1988, 1989; Carrive and Bandler, 1991;

Holstege, 1991; Trevizan-Baú et al., 2021b; Yasui et al., 1990;). Thus, given that

neither the NRA nor the PAG play a critical role in generation of the eupneic respiratory

motor pattern (Farmer et al., 2014; Jones et al., 2016), PAG projections to the ponto-

medullary respiratory nuclei suggests that volitional motor commands as well as

emotional and interoceptive signals that are initiated in the forebrain and relied in the

PAG are sent to the primary respiratory network in the pons and medulla in order to

modulate behavioural breathing (Fig. 3). Interestingly, the ponto-medullary respiratory

nuclei (KFn, pre- BötC, and BötC) also send ascending inputs back to the midbrain

PAG (Dutschmann and Dick, 2012; Trevizan-Baú et al., 2021b). In addition, the NRA

as the designated major relay for autonomic-respiratory output of the PAG has

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significant descending projections the KFn (Jones et al., 2016), but not the pre-BötC,

which support a role of the KFn as main pontine interface for the coordination of

orofacial behaviours including vocalisation (see below).

In summary, the reciprocal connectivity among anatomically and functionally

diverse brainstem respiratory regions in the midbrain, pons, and medulla might play a

critical role in coordinating and adapting respiration with orofacial behaviours.

The role of respiratory and premotor nuclei of the respiratory network

during the coordination of volitionally evoked orofacial behaviour and breathing

Recent anatomical studies investigated the connectivity of the forebrain with

the pontine (KFn) and the designated medullary core circuit for respiratory rhythm and

pattern generation (BötC and pre-BötC). Whilst hypothalamic nuclei connect with a

distributed brainstem autonomic network, including the BötC/pre-BötC and KFn

(Geerlin et al., 2010; Peyron et al., 1998; Trevizan-Baú et al., 2021a), our recent

tracing study and others report that the origins of descending projection targeting the

BötC/pre-BötC and KFn have more selective connectivity patterns compared to the

hypothalamic projections.

An interesting observation of a comparative analysis of descending cortical

projections of cortical layer V pyramidal neurons was the virtual absence of cortical

connectivity with the BötC Trevizan-Baú et al., 2021a). This finding was surprising

since the BötC is considered an essential element of respiratory pattern formation

within the medullary core circuit (Burke et al., 2010; Marchenko et al., 2016; Smith et

al., 2013) and, thus, would have been expected to be a target for cortical inputs during

volitionally evoked orofacial behaviour. Contrary, other critical pre-motor nuclei of the

pontomedullary network such as the pontine KFn and pre-BötC have significant




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connectivity with various cortical areas. Thus, it appears that cortical commands for

volitional orofacial motor behaviour may target selective neural network nodes in the

respiratory network to coordinate ongoing respiration with volitional motor commands.

The cortical projections arising from various cortical areas have connectivity

with the pre-BötC (Fig. 3A; Yang et al., 2020; Trevizan-Baú et al., 2021a). Although

the strength of the connectivity is overall weak, the more robust connectivity of the pre-

BötC is with M1 motor cortex. The M1 motor cortex constitutes the laryngeal cortex

and specifically controls the cricoarytenoid muscle (Espadaler et al., 2012; Simonyan,

2013), as the main laryngeal adductor that controls the expiratory airflow during

postinspiration (see Fig.1), supports the notion that the pre-BötC is involved in the

coordination of orofacial behaviour with ongoing expiration during vocalisation and

swallowing that require glottal adduction. Overall, a similar picture of broad cortical

connectivity is also emerging for the pontine KFn (Fig. 3B). However, the KFn retains

strong connectivity with specific cortex areas such as the rhinal, piriform and

somatosensory barrel cortices (Trevizan-Baú et al., 2021a). Thus, the KFn receives

cortical inputs that convey olfactory and tactile sensory information (Petersen, 2019;

Woolsey and van der Loos, 1970) to coordinate sniffing and whisking with breathing.

While premotor areas for sniffing, whisking and respiration are identified in the

medullary reticular formation (Moore et al., 2013 McElvain et al., 2018), the KFn has

descending projections to all pre-motor areas, including the pre-BötC as designated

key integrative neural node for the coordination of orofacial behaviour and breathing

(Moore et al., 2013; Kleinfled et al. 2014; Deschênes et al., 2016), as well as the lateral

column of the facial motor nucleus (Fig.2 from Dutschmann and Dick, 2012). These

elegant conceptual frameworks for the coordination of respiration with sniffing and

whisking are based on the assumption that sniffing is an inspiratory behaviour.

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However, as discussed before, sniffing might be initiated in the postinspiratory phase

(Mukai, 1989; Lawson et al., 1991; Pérez de Los Cobos Pallares et al., 2016) to restrict

pulmonary airflow and to generate high intranasal pressure and nasal ventilation that

is required for odor detection. Functional data further support such a role for the

pontine KFn on the control and initiation of the medullary oscillators activation for

sniffing and whisking. The KFn gate the inspiratory/pre-inspiratory facial and

postinspiratory vagal motor activity (Dutschmann and Herbert, 2006, Dutschmann et

al., 2020), which implies a major function for the coordination of orofacial muscles

during respiration and most likely during the mediation of orofacial behaviour and

expulsive reflexes (Dutschmann and Dick, 2012). A second observation that supports

our hypothesis is that blockade of the KFn triggered spontaneous swallowing activity

(Bautista et al., 2014), which supports the notion that the KFn, as part of the only

unconditional motor oscillator in the brain, may inhibit all other conditional oscillators

for orofacial behaviour during eupneic breathing. Overall, the selective connectivity of

the KFn with the rhinal, piriform and somatosensory barrel cortices (Trevizan-Baú et

al., 2021a) and all downstream orofacial pre-motor area and oscillatory networks in

the medulla oblongata strongly suggest that pre2motor and premotor neurons of the

KFn may have that coordinates breathing with sniffing and whisking. Moreover, the

KFn gates the respiratory activity of hypoglossal nerve (Bautista and Dutschmann

2014) and postinspiratory activity of the masseter (St John and Bedsoe, 1985), which

also has a critical function during vocalisation and speech. Since mandibular and

tongue movements are essential components for vocalisation, the KFn may also play

an essential role in the coordination of respiration and vocalisation (Dutschmann and

Dick, 2012). Finally, KFn blockade abolished the protective laryngeal adduction during

sensory sequential swallowing (Bautista and Dutschmann, 2014) and, thus,

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descending volitional motor commands for swallowing may also initially target the KFn,

given that glottal constriction has to proceed a save swallowing sequence.

In summary, we propose that the KFn interface for volitional orofacial motor

commands may serve as a master clock for the initiation as well as timing and

synchronisation of orofacial behaviour with respiration. However, the proposed KFn

master clock might be embedded into an extended neural node of the densely

connected PAG-KFn neuro axis (see above; Trevizan-Baú et al., 2021b) to coordinate

volitionally evoked orofacial-respiratory behaviours.

Emotional control of respiration: a key role for the pre-Bötzinger complex?

In addition to cortical inputs targeting the brainstem respiratory network to

control orofacial behaviours volitionally, we must also regard that emotional states

processed in the limbic system influence in particular respiration (for review, see

Homma and Masaoka, 2008). Expression of emotional states in orofacial behaviour

might be restricted to vocalisation, and, indeed, a recent study in humans identified 24

different emotions that can be conveyed during vocalisation (Cowen et al., 2019).

The emotional modulation of vocalisations is supposed to be mediated by the

extensive connectivity of the PAG with the limibic system (Holstege, 1989; Holstege,

2014; Subramanian and Holstege, 2014). However, anatomical tracing studies have

shown that brain regions of the limbic system send monosynaptic projections to the

nuclei of the ponto-medullary brainstem (Yang et al., 2020; Trevizan-Baú et al.,

2021a). Interestingly, the pre-BötC seems to receive a larger number of projection

neurons from the amygdala, as a critical region of the limbic system, compared to

other nuclei of the ponto-medullary respiratory control areas (Trevizan-Baú et al.,

2021a). It is well known that the amygdala processes numerous emotional responses

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such as fear and anxiety (Davis, 1992), and functional studies showed that respiratory

activity is altered by electrical stimulation of the amygdala (Dlouhy et al., 2015; Harper

et al., 1984). However, these direct amygdala-pre-BötC projections are unlikely to be

involved in the emotional modulation of vocalisations during expiration because of the

primary function of the pre-BötC in inspiratory rhythm generation (Feldman and Del

Negro, 2006). Nevertheless, these projections could convey triggers for sigh

generation (Li et al., 2016) or hyperventilation during panic attacks (Garssen et al.,

1996).

Figure 5.3. Top-down information from the forebrain to key brainstem respiratory control areas

of the pons and medulla oblongata might play critical roles adapting breathing with orofacial

behaviours and emotions.

Schematic sagittal view of the brain highlighting the higher brain information processed in forebrain

regions that send monosynaptic projections to pontine KFn (A) and medullary pre-BötC (B).

Monosynaptic projections from the rhinal and piriform (Pir) cortices might send olfactory information (in

green) to the KFn and pre-BötC, but the KFn receives the densest projections from the rhinal and

piriform cortices. Olfactory information to the brainstem respiratory network is critical to adapt breathing

with sniffing. Motor planning information (in blue), such as learned vocalisation and volitional

swallowing, is processed in the prefrontal cortex (PFC; including the motor cortex) and was thought to

119

adapt breathing with volitional behaviours by direct inputs to the cervical and thoracic motor pools in

the spinal cord, presumably bypassing the brainstem respiratory network. However, recent studies have

shown that the motor cortex and regions of the PFC send projections to the midbrain PAG (receives

densest projections) as well as the KFn and pre-BötC. Somatosensory cortex (S1/S2) processes tactile

sensory information (in cyan), which is crucial for sniffing and whisking, especially in rodents.

Somatosensory information is sent to the KFn and pre- BötC, but the KFn receives the densest inputs

from somatosensory cortex. Other forebrain regions constitute the so-called limbic system, such as the

amygdala, hypothalamus, PFC, and insular, rhinal and piriform cortices. The PAG seems to receive the

densest projections from the limbic system, however limbic information is also sent to key brainstem

respiratory of the pons and medulla.

Neurologic diseases are tightly linked to impaired volitional control of

orofacial behaviour and breathing

In humans, volitional motor behaviours include several orofacial-respiratory

motor activities such as speech (vocalisation), swallowing, sniffing (odor detection),

and mastication. Higher brain regions generate and process volitional motor

commands that activate the brainstem network, including the respiratory control areas,

to elicit orofacial-respiratory behaviours. As discussed above, the brainstem

respiratory network plays a fundamental role in mediating and coordinating volitional

information during orofacial motor behaviours. However, in neurological diseases,

these volitional descending pathways targeting the brainstem respiratory network may

have degenerated, and, therefore, patients with neurodegenerative (e.g., Alzheimer’s

and Parkinson’s diseases) and neurodevelopmental (e.g., autism spectrum, e.g. Rett

syndrome) diseases, as well as patients with brain injuries (e.g., stroke and brain

cancer) often present orofacial motor impairments, which include dysfunctions in

speech, swallowing, sniffing, mastication, and in the vocal cord coordination as well

as sleep-disordered breathing (Table 1).

It is extensively reported in the clinical literature that patients with neurological

diseases present orofacial motor disturbances linked to respiratory motor impairment

120

(Brown, 1994; Nogués et al., 2002; Polkey et al., 1999; Table 1). Respiratory

impairments in neurological diseases are mostly correlated with upper airway motor

dysfunctions, specifically oro-pharyngeal and laryngeal motor impairments (Herer et

al., 2001; Izquierdo-Alonso et al., 1994; Sabaté et al., 1996). A recent postmortem

study has shown that patients with neurodegenerative diseases present neuronal

degeneration in several brainstem regions (i.e., reticular formation in the clinical

literature; Eser et al., 2018). Because most of the respiratory premotor and motor

neurons are located in the brainstem, it can be speculated that orofacial motor

impairments are linked to neuronal degradation in the brainstem of these patients. In

order to fully understand how neuronal degradation affects orofacial-respiratory

behaviours in humans, studies using animal models are required to elucidate and

understand pathological mechanisms that underlie a large variety of neurological

diseases.

The availability of animal models for a neurologic disease of the autism

spectrum or neurodegeneration is based on human mutations that can be in transgene

mouse models. However, the impact of tauopathy, beta-amyloid, alpha synuclein, or

mutation of the gene for Huntington in the mouse model is usually studied in the

context of cognitive decline or loss of motor coordination. Surprisingly, only a handful

of studies tried to evaluate potential impairments of orofacial behaviour, despite their

prevalence in human patients (see Table 1). Overall, a handful of studies have

reported either respiratory-related paradoxical vocal cord movement or ultrasonic

vocalisation (USV) deficits as a consequence of neuronal degradation (Dutschmann

et al., 2010; Grant et al., 2014; Holtzman et al., 1996; Menuet et al., 2011b). For

example, Grant and colleagues (2014) investigated the USV phenotype of a mouse

model of Parkinsonism with widespread overexpression of alpha-synuclein in the

121

brain. Although the authors observed USV deficits, the mice were still able to emit

USV, and no clear phenotype that would relate to severe speech and voice disorders

seen in human patients was identified. Another study that used a mouse model of

Down syndrome demonstrated a delay in USV development during the postnatal

period, but the mice could emit normal USV pattern in adulthood (Holtzman et al.,

1996). Moreover, Menuet et al. (2011b) investigated a mice model (tau-P301L mice)

that presented tauopathy (neurodegeneration) predominantly in the midbrain and

brainstem regions and progressive USV deficits and disease end-stage mutism.

However, our recent study using the same animal model (tau-P301L mice) failed to

replicate their observation (Trevizan-Baú et al., 2019), although tau-P301L mice still

presented respiratory upper airway motor dysfunctions (Dutschmann et al., 2010) and

suggesting that tauopathy even in critical nuclei of the vocalisation (e.g., PAG and

KFn) can be compensated. A recent publication by Hülsmann et al. (2019) studied

USV emissions in a mouse model of progressive loss of glycinergic synaptic inhibition.

Despite the predicted vocal cord dysfunction caused by reduced glycinergic inhibition

(Dutschmann and Paton, 2002; Furuya et al., 2021), their study revealed no

impairment of USV emission. Interestingly, the study identified a compensatory

mechanism by demonstrating that after the loss of glycinergic inhibition, mice started

to emit USV during late expiration instead of the normal USV transmission during

postinspiration. Thus, these mice apparently learned to compensate for impaired

postinspiratory control of the vocal folds by emitting USV in a different respiratory

phase of the respiratory motor cycle.

Taken all together, experimental models replicating human neuropathologies

might help to understand the cause of orofacial-respiratory motor disturbances in

patients with neurologic diseases. However, most of the investigations showed that

122

transgenic mice can still evoke normal orofacial behaviours such as vocalisation even

when several brain regions are affected by neurodegeneration. This is raising the

question of whether i) transgene mouse models based on rare human mutations are

useful animal models since most neurodegenerative are sporadic and have no clear

cause or ii) neurodegeneration in key neuronal pathways that mediate reflexes and

volitional orofacial behaviours can be compensated by redundant network

mechanisms across the entire brain axis. A better understanding of how orofacial

motor pathways collapse in neurologic disease requires further studies to elucidate

the functional anatomy of orofacial behaviour across the entire brain axis.

123

Table 5.1. Clinical studies have reported impaired orofacial motor behaviours in several

neurological diseases.

Impaired orofacial motor control

Disease classifications

Voice/vocalisation disorder:

• Vocal cord paresis or paralysis (VCP)

• Spasmodic dysphonia

• (SD)

• Apraxia (A)

• Mutism (M)

• Dysarthria (D)

Neurodegenerative: Alzheimer’s disease (Cera et al., 2013 A;

Gerstner et al., 2007 A); Parkinson’s disease (Ho et al., 1998 A;

Streifler et al., 1984 A); Amyotrophic lateral sclerosis (Roth et al.,

1996 SD); Motor neurons disease (Lillo and Hodges, 2009 A);

Frontotemporal dementia (Lillo and Hodges, 2009 A); Huntington’s

disease (Hartelius et al., 2003 D)

Neurodevelopmental: Autism with selective mutism (Windsor et

al., 1994 M; Panasiuk, 2019 M)

Brain injury: Stroke (Kothare et al., 1998 M; Vidovic et al., 2011

A); Brain tumours (Schenke-Reilly, 2018 A)

Swallowing disorder:

• Dysphagia (D)

• Aspiration (A)

Neurodegenerative: Alzheimer’s disease (Corriea et al., 2010 D; Wada et al., 2001 A); Parkinson’s disease (Potulska et al., 2003 D; Akbar et al., 2015 A); Cerebral palsy (Arvedson, 2013 D; Krigger, 2006 A); Frontotemporal dementia (Langmore et al., 2007 D); Huntington’s disease (Kagel and Leopold, 1992 D; Hamakawa et al., 2004 A&D); Motor neurons disease (Leighton et al., 1994 D) Neurodevelopmental: Rett Syndrome (Isaacs et al., 2003 D; Motil et al., 2012 D) Brain injury: Stroke (Johnson et al., 1993 A&D; Sellars et al., 1999 D); Brain tumours (Rubenstein and Schenke-Reilly, 2018 D)

Impaired sniffing/olfaction:

• Odor identification (ID)

• Odor discrimination (DIS)

• Odor threshold dysfunction (OTD)

• Reduced sniff airflow (RSA)

Neurodegenerative: Alzheimer’s disease (Förster et al., 2010 OTD; Murphy, 1999 ID DIS); Parkinson’s disease (Sobel et al., 2001 RSA; Haehner et al., 2011 ID DIS; Rahayel et al., 2012 ID DIS OTD); Motor neuron disease (Hawkes et al., 1998 ID DIS) Neurodevelopmental: Tourette (Kronenbuerger et al., 2018 ID DIS OTD) Brain injury: Idiopathic intracranial hypertension (Bershad et al., 2014 OTD)

Mastication disorders

• Reduced muscle function (RMF)

Neurodegenerative: Alzheimer’s disease (Kieser et al., 1999 RMF); Parkinson’s disease (Eadie and Tyrer, 1965; Kieser et al., 1999 RMF); Huntington’s disease (Hamakawa et al., 2004 RMF) Neurodevelopmental: Rett Syndrome (Motil et al., 2012) Brain injury: Stroke (Kim and Han, 2005)

Vocal cord dysfunction

• Vocal cord paresis or paralysis (VCP)

• Vocal cord atrophy (VCA)

Neurodegenerative: Amyotrophic lateral sclerosis (Graaff et al., 2009 VCP; Isozaki et al., 1996 VCP) Parkinson’s disease (Stelzig et al., 1999 VCA) Neurodevelopmental: Williams-Beuren syndrome (Koren et al., 2015 VCP; Vaux et al., 2003 VCP) Shy-Drager syndrome (Williams et al., 1979 VCP) Brain injury: Stroke (Venketasubramanian et al., 1999 VCP; Sellars et al., 1999 VCP)

Sleep-disordered breathing:

• Stridor (S)

• Nocturnal hypoventilation (NH)

• Obstructive sleep apnea (OSA)

Neurodegenerative: Alzheimer’s disease (Gaig and Iranzo, 2012 OSA); Parkinson’s disease (Gaig and Iranzo, 2012 OSA); Amyotrophic lateral sclerosis (Gaig and Iranzo, 2012 S&NH); Frontotemporal dementia (Lillo and Hodges, 2009 A); Huntington’s disease (Banno et al., 2005 OSA) Neurodevelopmental: Autism spectrum disorder (Accardo and Malow, 2015 OSA) Brain injury: Stroke (Dyken and Im, 2009 OSA)

124

Conclusions and future perspectives

The works of my thesis showed that local brainstem tauopathy does not affect

volitional ultrasonic vocalisation (USV) linked to social male-female interaction,

despite the presence of tauopathy in key vocalisation areas in the midbrain

(periaqueductal gray), pons (Kölliker-Fuse nucleus) and medulla oblongata (nucleus

retroambiguus). This outcome of my extensive studies of social USV was surprising

and could be partially explained by the activation of redundant network connections

that may compensate local tauopathy to allow for the undisturbed mediation of USV

during social interaction. However, my thesis did not further investigate whether other

types of USV could be affected by brainstem tauopathy. To further clarify the role of

brainstem tauopathy in clinically highly relevant speech and voice disorders, it would

be of major interest to study reflex evoked USVs that can be triggered by fear and

painful stimuli. A previous study showed that midbrain PAG lesioning in rat abolished

fear-related USV (Leman et al., 2003). Therefore, it can be speculated that tauopathy

in the PAG might impair reflex evoked USVs, but not volitional vocalisation. Future

studies need to address this question.

In my second and third experimental studies (chapters 3 & 4), I investigated

descending neuroanatomical pathways controlling behavioural adaption of breathing

in the context of vocalisation and other orofacial behaviours (e.g., swallowing, sniffing

and whisking). I reported numerous forebrain regions projecting to the primary

respiratory control nuclei in the brainstem. Moreover, I found strong reciprocal

connectivity within the brainstem respiratory network, which might play a critical role

in adapting breathing during forebrain initiated volitional motor commands.

Following the significant amount of neuroanatomical data provided in chapter 3

& 4 of my thesis, there is now the opportunity to try multiple and more specific neuronal

125

tracing strategies in future studies to further strengthen the current anatomical

frameworks for neural pathway of volitional control of breathing, such as: i) to

investigate the polysynaptic connectivity of the brainstem with higher brain regions

using retrograde polysynaptic viral tracing to identify the extended connectivity of the

primary respiratory control areas with the forebrain; ii) to inject two different retrograde

tracers in two brainstem respiratory control areas and verify if the tracer retrogradely

labels the same neurons in the forebrain; and iii) to use a conditional anterograde

tracer approach (similar approach to McGovern et al., 2015) that allows to differentiate

forebrain efferent pathways to the brainstem respiratory regions. Both approaches ii

and iii would be helpful to extend the present anatomical framework by investigating

whether different respiratory control areas in the brainstem (e.g., PAG and KFn)

receive projection neurons not only from the same forebrain region but also whether

a single forebrain neuron projects to different brainstem respiratory control areas.

Additionally, to identify the specific vocalisation forebrain-brainstem pathways, it would

be helpful to apply a specific Cre-dependent tracing approach that targets neuronal

population expressing FoxP2, a transcription factor linked to the development of

speech (FoxP2: a selective marker for KFn neurons; Gray, 2008), in Cre-driver

transgenic mice. Finally, investigating the ascending connectivity of the brainstem

respiratory network with the forebrain (similar approach to McGovern et al., 2012)

would also be crucial for future studies. For instance, it is known that respiratory activity

in the brainstem influences cognition and emotion (Heck et al., 2017; Varga and Heck,

2017; Del Negro et al., 2018; Tort et al., 2018; Kluger and Gross, 2020), but the

neuroanatomical organisation of ascending projections from the primary respiratory

control areas in the brainstem to forebrain regions that could contribute to modulation

126

and mediation of cognitive and limbic brain functions remains largely unknown, and

therefore warrants thorough investigation.

In conclusion, the present thesis presents several valuable discoveries to

advance our understanding of the neuroanatomical organisation of volitional control of

breathing. The new neuroanatomical framework of the present thesis might help to

pave future studies on behavioural adaptation of breathing and may provide better

understanding of respiratory disorders and impaired orofacial behaviours in a variety

of neurological and neurodegenerative diseases.

127

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Minerva Access is the Institutional Repository of The University of Melbourne

Author/s:Trevizan Bau, Pedro Henrique

Title:Neuroanatomy and neuropathophysiology of volitional control of breathing

Date:2021

Persistent Link:http://hdl.handle.net/11343/286070

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