UNIVERSITY of CALGARY the Role of the 5-HT1A Receptor in Normal

Home , Melanopsin

UNIVERSITY OF CALGARY

The role of the 5-HT1A receptor in normal and enhanced responses of the mouse

circadian clock to light

Victoria M. Smith

A THESIS

SUBMITTED TO THE FACULTY OF GRADUATE STUDIES

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE

DEGREE OF MASTER OF SCIENCE

DEPARTMENT OF PSYCHOLOGY

CALGARY, ALBERTA

SEPTEMBER, 2008

ISBN: 978-0-494-44633-1

Abstract

The serotonergic system modulates the effect of light on the mammalian circadian system, thus the role of the 5-HT1A receptor was examined in normal and enhanced

responses to light. 5-HT1A receptor knockout mice exhibit larger phase shifts to both short (15 min) and long (12 h) light pulses, as well as attenuated Fos and mPer1

expression in the retinorecipient cells of the SCN. The 5-HT1A receptor is crucial for the

potentiation of photic phase shifts observed using the 5-HT1A receptor mixed

agonist/antagonist NAN-190, with 5-HT1A receptor knockout mice exhibiting attenuated

phase advances. NAN-190 also attenuates the light-induced expression of Fos and the phosphorylation of CREB corresponding to potentiated photic phase shifts, with no effect on the expression of mPer1 or mPer2, nor the phosphorylation of ERK at the times

examined. Together, these result exemplify the important, yet complicated role that the 5-

HT1A receptor plays in the ability of light stimuli to alter the expression of circadian

rhythmicity.

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ACKNOWLEDGEMENTS

I would like to extend my sincerest gratitude to my supervisor and mentor Dr. Michael Antle, without whom this work would not have been possible. His encouragement and guidance have given me all the tools necessary to become a successful researcher. I would also like to thank Drs. Brian Bland, Penny Pexman, and Ben Rusak for serving on my defense committee. Finally, I would like to thank the members of the Chronobiology Lab (past and present) at the University of Calgary, especially those who participated in any portion of the work included in this thesis.

TABLE OF CONTENTS

APPROVAL PAGE…………………………………………………………………… ii

ABSTRACT……………………………………………………………………………iii

ACKNOWLEDGEMENTS………………………………………………………….. iv

TABLE OF CONTENTS………….………………………………………………….. v

LIST OF FIGURES……………….…………………………………………………viii

ABBREVIATIONS……………………………………………………………………ix

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

1.1 Circadian Rhythms: Overview……………………………………………... 2 1.1.1 Evolutionary Perspective .………………………………………... 2 1.1.2 Entrainment. ……………..………………………………………….4 1.2 Rhythmic Desynchronization……………………………………………….. 6 1.3 The Master Pacemaker..……..……………………………………………… 8 1.4 The Endogenous Circadian System………………………………... ……….9 1.4.1 Afferent Pathways to the SCN..…………………………… …….. .9 1.4.2 The Suprachiasmatic Nucleus.....………………………… ……...10 1.4.2.1 Regions of the SCN..………………………………. ……..10 1.4.2.2 Gene Expression in the SCN.………………………. …11 1.4.3 Efferent Pathways Out of the SCN.……………………………… 13 1.5 Photic Phase Shifting………..…………………………………………….. 15 1.5.1 Effects of Light Intensity and Duration……….……………… …..18 1.5.2 Mimicking and Altering the Effects of Light...………………... …19 1.5.3 Light-Induced Protein and Gene Expression………………….. ..20 1.5.3.1 Immediate-Early Genes...…..………………....…...…...21 1.5.3.2 Period Genes...... 23 1.5.3.3 Biochemical Pathways………………………………… 25 1.6 Serotonergic Modulation of Circadian Rhythmicity.…...... 27 1.6.1 The Role of Serotonin……………………………....………………27 1.6.2 Non-Photic Phase Shifts Mediated by the Serotonin System…....29 1.6.2.1 Serotonergic Mediation of Activity and Arousal.….. …29 1.6.2.2 Chronobiotic Mediation of Non-Photic Phase Shifts.. …30 1.6.3 Serotonergic Mediation of Photic Phase Shifts..………………..34 1.6.3.1 Affinity for Serotonergic Receptor Subtypes….. ……..35 1.6.3.2 Serotonin Agonists……………...………….….. ……..36 1.6.3.3 Serotonin Antagonists……………………...…... ……..38

1.6.3.4 Serotonin Mixed Agonists/Antagonists………... ……..38 1.6.3.5 Special Case of NAN-190………………….…... ……..40 1.6.4 Serotonergic Mediation of Light-Induced Gene Expression…… 43 1.6.5 Transgenic Mice…………………………..………………….. …… 46 1.6.6 Summary………………………………...... ……….…...…...... 48 1.7 Present Study: Objectives and Hypotheses ………………………………...49

2. Altered photic and non-photic phase shifts in serotonin1A receptor knockout mice………………………………………………………………………………….50

Abstract………………………………………………………………………...51 2.1 Introduction…………………………………………………… …………...53 2.2 Experimental Procedures...…………………………………… …………...57 2.2.1 Animals and Housing…...…………………………. …………….57 2.2.2 Activity Rhythms……...... ………………………….….…………58 2.2.3 Experiment 1 – Phase Shifts to Short Duration Light ….………...59 2.2.4 Experiment 2 – Phase Shifts to Long Duration Light……... ……. 60 2.2.5 Experiment 3 – Protein and Gene Expression Underlying Photic Phase Shifts..…………………….………………60 2.2.6 Experiment 4 – Non-photic Phase Shifts to the Serotonin Agonist 8-OH-DPAT…...……………………………….. 61 2.2.7 Immunocytochemistry..………………………………….………….. 62 2.2.8 In Situ Hybridization…………………………………..………… 63 2.2.9 Quantification of Fos and mPer1 Expression……...……………....64 2.2.10 Statistical Analysis.. ………………………………………………….65 2.3 Results ……………………………………………………………………66 2.3.1 Experiment 1 – Phase Shifts to Short Duration Light... …………66 2.3.2 Experiment 2 – Phase Shifts to Long Duration Light………….... 66 2.3.3 Experiment 3A – Fos Protein Expression Underling Photic Phase Shifts……………………. ……………………..….. 71 2.3.4 Experiment 3B – mPer1 Gene Expression Underlying Photic Phase Shift..…………..………………………………. 74 2.3.5 Experiment 4 – Non-photic Phase Shifts to the Serotonin Agonist 8-OH-DPAT..…………………………………… 74 2.4 Discussion……….……………………………………………………….... 79

3. Potentiation of photic phase shifts by the serotonergic mixed agonist/antagonist NAN-190: The role of the 5-HT1A receptor…………………………………….....87

Abstract...………………………………………………………………..…...... 88 3.1 Introduction.……………………………………...………………………...90 3.2 Experimental Procedures………………...…………………………………93 3.2.1 Animals and Housing…………………. …………………………93 3.2.2 Drugs and Reagents..………………...... …………96 3.2.3 ActivityRhythms….….…………………………………………… 96

3.2.4 Phase Shift Manipulations………..……………………………… 97 3.2.5 Statistical Analysis….…………………………………………… 98 3.3 Results…………………………………………………………………….98 3.3.1 Modulation of Phase Shifts to Bright Light Pulse….……………..98 3.3.2 Modulation of Phase Shifts to a Dim Light Pulse…...... ……… 101 3.3.3 Modulation of Phase Shifts to No Light Pulse…….……………..106 3.4 Discussion……………………………………………………………….109

4. Molecular underpinnings of photic phase shift potentiation with NAN-190…117

Abstract...………………………………………………………………..…....118 4.1 Introduction …………………………………...……………………….120 4.2 Experimental Procedures………………...………………………………..124 4.2.1 Animals and Housing…………………. ……………………….124 4.2.2 Drugs and Reagents…..………………...………………………126 4.2.3 Activity Rhythms…….…………………………………………..126 4.2.4 Protein and Gene Expression Underlying Photic Phase Shifts…127 4.2.5 Immunocytochemistry…………………………………………...128 4.2.5.1 Diaminobenzidine (DAB)…………………………….128 4.2.5.2 Fluorescent Triple-Label……………………………...129 4.2.6 In Situ Hybridization……………………………………………130 4.2.7 Quantification of Protein and Gene Expression………………..131 4.2.8 Statistical Analysis……………………………………………...132 4.3 Results…………………………………………………………………….133 4.3.1 Experiment 1: DAB-Labeled Fos Expression…………………..133 4.3.2 Experiment 2: Fluorescent Triple-Label Fos Expression.……...133 4.3.3 Experiment 3: mPer1 and mPer2 Gene Expression...…………..142 4.3.4 Experiment 4: Biochemical Pathway Components……………..142 4.3.4.1 P-ERK………………………………………………..142 4.3.4.2 P-CREB……………………………………………....151 4.4 Discussion………………………………………………………………...151

5. General Discussion………………………………………………………………..164

5.1 Summary of Major Findings…………………………………… ...………164 5.2 Overall Conclusions……………………………………………………... 165 5.3 Limitations and Future Directions……….……....……………………… 168 5.4 Clinical Importance………………...……………………………………..172

6. References………………………………………………………………………. ...174

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LIST OF FIGURES

Figure 1.1 Examples of a photic PRC for A) mice and B) hamsters….. ………….17

Figure 2.1 Actograms and mean phase shifts for short duration light……...……. 68 Figure 2.2 Actograms and mean phase shifts for long duration light.……...……. 70 Figure 2.3 Fos expression and mean Fos-IR cell counts….………………………73 Figure 2.4 mPer1 expression and mean mPer1 relative optical densitoes.…….....76 Figure 2.5 Actograms and mean phase shifts for 8-OH-DPAT.………………….78

Figure 3.1 Example of electrophoresis gel used to genotype animals………….... 95 Figure 3.2 Actograms for bright light pulse manipulations..…………………….100 Figure 3.3 Mean phase shifts with each drug treatment and a bright light pulse...103 Figure 3.4 Actograms for dim light pulse manipulations...... ………………..105 Figure 3.5 Mean phase shifts with each drug and a dim light pulse.……………108 Figure 3.6 Actograms for no light pulse manipulasitons………………………..111 Figure 3.7 Mean phase shifts with each drug alone……………………………..113

Figure 4.1 DAB-labeled Fos protein expression with each drug treatment……..135 Figure 4.2 Mean number of Fos-expressing cells with each drug treatment……137 Figure 4.3 Fluorescent triple-labeled Fos protein with each drug treatment……139 Figure 4.4 Mean number of Fos-IR cells with each drug treatment…………….141 Figure 4.5 mPer1 and mPer2 gene expression with each drug treatment………144 Figure 4.6 Mean ROD of mPer1 and mPer2 expression with each drug...……..146 Figure 4.7 P-ERK protein expression with each drug at three time points……..148 Figure 4.8 Mean ROD of P-ERK expression in the core and shell of the SCN...150 Figure 4.9 P-CREB protein expression with each drug at three time points…...153 Figure 4.10 Mean number of P-CREB-expressing cells in the core and shell…...155

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LIST OF ABBREVIATIONS

Measurement Terms: °C degrees Celsius kg kilogram lux unit of illuminance μm micrometer mg milligram mL milliliter mM millimolar M molar

Other Terms: 5-HT serotonin, 5-hydroxytryptamine 8-OH-DPAT (±)-8-hydroxy-2-(dipropylamino)tetralin hydrobromide ANOVA analysis of variance bp base pairs CalB calbindin cAMP cyclic adenosine monophospate CRE Ca2+/cAMP response element CREB cAMP response-element binding protein Cry cryptochrome gene Cry cryptochrome protein CT circadian time DD constant darkness DAB diaminobenzidine DIG digoxigenin DMSO dimethyl sulfoxide ERK extracellular signal-regulated kinase GABA gamma aminobutyric acid GHT geniculohypothalamic tract GRP gastrin-releasing peptide haPer1 hamster period1 gene Het heterozygous ICC immunocytochemistry IEG immediate-early gene IGL intergeniculate leaflet i.p. intraperitoneal -IR -immunoreactive KO knockout LD light/dark cycle LL constant light LP light pulse MAPK mitogen-activated protein kinase mPer1 mouse period1 gene

mPer2 mouse period2 gene mRNA messenger ribonucleic acid NAN-190 1-(2-methoxyphenyl)4-[4-(phthlaimido)butyl]piperazine hydrobromide NMDA N-methyl-D-aspartic acid NPY neuropeptide Y PACAP pituitary adenylate cyclase-activated polypeptide PBS phosphate buffered saline PBSx phosphate buffered saline with 0.1% Triton X-100 P-CREB phosphorylated cAMP response-element binding protein Per period gene Per period protein P-ERK phosphorylated extracellular signal-regulated kinase PRC phase response curve RHT retinohypothalamic tract ROD relative optical density SCN suprachiasmatic nucleus SEM standard error of the mean SP substance P tRNA transfer ribonucleic acid VIP vasoactive intestinal polypeptide VP vasopressin WL wheel lock WT wildtype

1. General Introduction

The chronic desynchronization between the endogenous circadian pacemaker and

the rhythmic external environment which is inherent to the 24-hour nature of society in the 21st century often leads to numerous health consequences. Shift work and the

phenomenon known as “jet lag,” which results from rapid travel across multiple time

zones, have become a normal part of everyday life. Currently, there are no practical

medicines or therapies commercially available which aid in the acceleration of the

resynchronization of the circadian system. Although light has the ability to reset the

mammalian circadian pacemaker, its effects are minimal in magnitude. Thus the

development of a chronobiotic which would potentiate these effects of light would be

extremely beneficial to thousands of individuals in all walks of life. Serotonin (5-HT) is

known to play a regulatory role in the circadian system, and has been shown to modulate

the effects of photic input to the circadian pacemaker. Thus, compounds such as NAN-

190, with both agonistic and antagonistic properties, which are capable of greatly

potentiating the effects of light on the circadian system could be a huge asset in today’s

fast-paced world. This study examined the role of the 5-HT1A receptor in the effects of

light on the circadian system, both on its own and in combination with the 5-HT1A mixed agonist/antagonist NAN-190, as well as the molecular underpinnings of the ability of

NAN-190 to greatly potentiate the effects of light.

1.1 Circadian Rhythms: Overview

1.1.1 Evolutionary Perspective

All organisms have basic needs which must be met in order to survive. They

require food, shelter, and access to a mate, in order to ensure their own survival as well as

the survival of the species. However, all organisms on Earth evolved in a rhythmic

environment, resulting from the geophysical properties of the solar system. The Earth

rotates around the sun, the moon rotates around the Earth, and the Earth rotates about its

own axis. Thus, temporal factors such as the changing of the seasons as well as the day- night cycle play a vital role in an organism’s ability to be successful in its particular niche. An organism’s ability to tailor its daily activities to the generally predictable rhythms in its surrounding environment is adaptive for maximizing its chances of survival and reproduction (Dunlap, 2004).

Every species has two options for the organization of its behaviour in a temporal

manner. Either an individual can follow exogenous timing cues, and directly respond to

environmental changes in determining when to perform certain activities, or it can rely on

an endogenous pacemaker which can be modulated by environmental stimuli to ensure it

remains properly synchronized to the external cycles. The anticipation of repetitive

temporal events allows an organism to optimally organize its behaviour and physiology,

such that everything occurs at the most advantageous time of day, and thus increasing genetic fitness (Dunlap, 2004).

Several types of rhythms can be observed in biological systems, including:

circannual rhythms (period of about one year), circalunar (period of about one lunar

cycle), infradian (period of more than a day), circadian (period of about one day) and

ultradian (period of less than a day; Refinetti, 2006). Circadian rhythms (circa =

approximately; dies = a day) can be found in species ranging from prokaryotes to

mammals, and constrain behaviour and physiology to species-specific times of the day

(Sharma, 2003). Although it is not explicitly known, it has been speculated that an

endogenous circadian rhythm may have developed as an evolutionary solution to the

problem of cellular processes which were sensitive to UV radiation. Thus some processes

could have been necessarily restricted to night-time when UV radiation was lowest,

leading to rhythmicity that was circadian in nature (Pittendrigh, 1993).

For centuries, it was believed that biological rhythms were solely the result of the

periodic nature of the external environment (Moore-Ede, 1982). Not until the middle of

the 20th century was it generally accepted by the research community that this

rhythmicity was in fact the product of an internal timekeeping mechanism, within the

organism itself (Edery, 2000). This fundamental paradigm shift led to an explosion of

scientific research investigating the biological rhythms of numerous species. Since that

time, extensive research has been done on the marked rhythmicity of many biological

processes in both plants and animals. The first experiment which tested this belief was

conducted by Jean Jacques d’Ortous de Mairan in 1729, where he observed that the leaves of a specific plant open and close at consistent intervals in conjunction with the

light/dark cycle, but persist regardless of the presence of those external time cues

(Moore-Ede, 1982). It seemed that this leaf behaviour was endogenously generated.

Subsequently, Augustine de Candolle showed that the rhythmic opening and closing of

leaves in the Mimosa pudica plant was not only endogenously generated, but also had a

period that was slightly shorter than the external light/dark cycle (LD) when in constant

conditions (Moore-Ede, 1982). This suggested to de Candolle that the plant was able to

‘freerun’ with a unique endogenous period when no longer synchronized to the light

cycle of the external environment (Moore-Ede, 1982).

1.1.2 Entrainment

Photic information received by the suprachiasmatic nucleus (SCN) from the

retinas is the primary cue for entraining, or synchronizing, an animal’s circadian rhythm

to the external light/dark cycle (Hirota and Fukada, 2004). The day-night cycle serves to

adapt mammals to their external environment such that the animal’s behaviour remains

tied to the rhythms in its environment, and the chances of survival are optimized with the ability to anticipate environmental changes (Cermakian and Sassone-Corsi, 2002; Gachon et al., 2004; Panda and Hogenesch, 2004). The circadian system is normally synchronized to the environmental light/dark cycle, and therefore has a period of approximately 24 hours (Moore-Ede, 1982; Reppert and Weaver, 2001). This synchronization allows organisms to optimally coordinate their physiological processes with the external world, and can predict daily changes in the environment (Holzberg and Albrecht, 2003; Antle and Silver, 2005). Yet, in the absence of external time cues, this endogenous rhythm

continues to oscillate with a period of about a day, although the exact period length is species-specific (Pittendrigh, 1976a; Moore-Ede, 1982; Roenneberg et al., 2003)

Since the period of the mammalian circadian rhythm is only approximately 24 hours, a correction must be made on a daily basis to keep the rhythm in synchrony with the length of the day. Gachon et al. (2004) use the example of a nocturnal animal being able to anticipate approximately when it is dusk. If the animal wakes up and the sun has not yet set, the subsequent day it will wake up a little later in an attempt to prevent being seen by a predator when it emerges from its burrow on future days. Similarly, the animal anticipates approximately when it will be dawn, such that if the animal stays up too long and risks exposing itself to diurnal predators it will attempt to wake up a little earlier on subsequent days to ensure it has returned to the burrow before the sun rises.

Only when the oscillation of a zeitgeber (“time-giver”) falls within a fairly narrow range of period lengths sufficiently close to 24 hours, called the limits of entrainment, can

entrainment result (Aschoff and Pohl, 1978). It is possible to experimentally entrain an animal to a light/dark cycle with a period within the species-specific circadian range, generally between 21 and 27 hours (Pittendrigh & Daan1976b). It does not matter what the actual time of ‘sunrise’ is in an animal’s environment, it is merely necessary that the length of the ‘day’ and ‘night’ be appropriate for the species.

Mammals also exhibit circadian rhythms under constant lighting conditions, either constant light (LL) or constant dark (DD). When an animal is not entrained to any light- dark cycle, it is said to be freerunning. In this situation, statements of ‘day’ and ‘night’ are changed to ‘subjective day’ and ‘subjective night.’ These terms simply make

reference to the portion of the normal activity cycle to which the animal is behaving most

similarly. Thus, in constant lighting conditions, ‘subjective day’ refers to when a

nocturnal animal is behaving as though it is daytime (generally sleeping/inactive) and

‘subjective night’ refers to when the animals is behaving as though it is nighttime

(generally active), regardless of what the actual external time-of-day is. The period of the

freerunning rhythm is no longer tied to the 24-hour light/dark cycle, and thus varies somewhat from 24 hours. It depends on the species whether an animal will tend to freerun with a period slightly less than 24 hours or slightly greater than 24 hours, and individuals within a species will not freerun with exactly the same period. Constant darkness is frequently used to determine the freerunning period of the activity cycle in

nocturnal rodents. Since it would be difficult to see any change in the circadian rhythm

for an animal entrained to a light/dark cycle, the majority of research involving

adjustments to the circadian rhythm is done under constant lighting conditions in order to

maximize the effects that can be observed. Light stimuli play an integral role in both the

entrainment of the circadian rhythm, and in adjustments that can be made to these

endogenous rhythms.

1.2 Rhythmic Desynchronization

Numerous health, safety and social problems arise when endogenous circadian

rhythms become desynchronized from the external environmental cycle. There are

several causes of desynchronization, including jet lag, shift work, and circadian rhythm

disorders, but regardless of the cause the results can be disastrous. Although the

percentage of individuals who are considered to be shift-workers (i.e. either working the

night shift, working on a rotating shift, or working extended shifts) is fairly small, some of the most important positions in society are subject to this type of work schedules

(Dunlap, 2004). Doctors, nurses, emergency service workers, pilots, and nuclear power plant operators, are just a few examples of the occupations in which individuals are often subjected to shift work schedules. The circadian disruptions inherent in these sorts of positions not only affect the circadian rhythms of the individual workers, but can also have disastrous effects on those people under their care.

The rapid desynchronization inherent to phenomena such as rotating shift work schedules and jet lag can lead to numerous negative consequences. A wide range of health problems, including increased risk of cardiovascular disease and cancer, are the more serious of the potential medical conditions which can result from chronic desynchronization between one’s internal circadian rhythm and the external environment

(Rajaratnam and Arendt, 2001; Knutsson, 2003; Dunlap, 2004; van Mark et al., 2006).

However, reduced concentration and increased reaction times are just a few of the adverse effects which can have disastrous effects on the innocent people around those experiencing the chronic desynchronization, potentially leading to accidents as small as traffic accidents and as major as the Exxon Valdez oil spill and the Chernobyl nuclear disaster (Dunlap, 2004). In order to minimize the effects of such desynchrony and develop potential treatments to accelerate resynchronization, it is necessary to more fully understand the mechanisms responsible for the synchronization of our endogenous circadian system to the rhythmic world.

1.3 The Master Pacemaker

Numerous behavioural and physiological processes in mammals, including activity cycles and the expression of certain genes, are governed by the daily oscillation of endogenously generated circadian rhythms (Reppert and Weaver, 2001). In 1972, two independent research groups made a ground-breaking discovery – the location of the endogenous mammalian circadian pacemaker (Moore and Eichler, 1972; Stephan and

Zucker, 1972). In the 36 years since that discovery, an enormous body of evidence has been accumulated confirming that the suprachiasmatic nucleus is indeed the location of the master circadian pacemaker in the mammalian brain (Weaver, 1998; Lee et al., 2003).

This circadian clock is seated in the anterior hypothalamus, directly dorsal to the optic chiasm, and consists of a group of approximately 20,000 neurons in paired nuclei

(Reppert and Weaver, 2001). This small bundle of neurons is responsible for managing and coordinating the timing of both physiology and behaviour. The exact mechanism of how these cells perform this immensely complex task of coordinating all biological rhythms in the body remains elusive to researchers.

Lesioning the SCN was shown to completely ablate the adrenal corticosterone rhythm, as well as the circadian drinking and locomotor rhythms of rats (Moore and

Eichler, 1972; Stephan and Zucker, 1972). However, only complete SCN lesions have the ability to cause an animal to become fully arrhythmic, and in fact it appears that only a small amount of SCN cells need remain intact, approximately 20%, in order for rhythmicity to be preserved (Rusak, 1977; Davis and Gorski, 1988). Specifically, the preservation of caudal pathways to the SCN appears to be more critical in the sparing of

entrainment and rhythmicity than pathways rostral to the SCN (Harrington et al., 1993).

Yet, transplants of fetal SCN tissue are able to restore rhythmicity to SCN-ablated animals (Sawaki et al., 1984; Lehman et al., 1987). The arrhythmic host receiving a fetal

SCN transplant not only becomes rhythmic once again, but instead of simply restoring the original period of the host, it takes on the specific circadian period of the donor

(Ralph et al., 1990).

1.4 The Endogenous Circadian System

1.4.1 Afferent Pathways to the SCN

Normal vision is not necessary for entrainment to a specific light cycle or to shift an animal’s circadian phase (Hirota and Fukada, 2004). The photoreceptor which serves to reset the circadian clock does not require functional rods or cones in the retina. Instead, the light information hitting a subset of intrinsically photosensitive retinal ganglion cells, containing melanopsin, projects to the SCN through the retinohypothalamic tract (RHT;

Berson, 2003; Hirota and Fukada, 2004). These projections release glutamate and pituitary adenylate cyclase-activated polypeptide (PACAP) onto retinorecipient cells in the SCN. This process leads to the remodeling of chromatin and the subsequent expression of clock genes (Pittendrigh, 1976b; Morse and Sassone-Corsi, 2002). Light information is also received by the SCN indirectly from the geniculohypothalamic tract

(GHT) via the intergeniculate leaflet (IGL; Morin, 1994; Harrington, 1997; Meijer and

Schwartz, 2003). A third major afferent connection to the SCN is a dense serotonergic

projection originating in the raphe nuclei, which predominately terminates in the

ventromedial region of the SCN (Meyer-Bernstein and Morin, 1996). In the hamster,

neurons from the median raphe project directly to the SCN, however projections from the

dorsal raphe connect to the SCN only indirectly through the IGL (Morin, 1999; Morin and Meyer-Bernstein, 1999).

1.4.2 The Suprachiasmatic Nucleus

1.4.2.1 Regions of the SCN

Some neurons in the SCN are intrinsically rhythmic in their gene and protein expression, exhibiting a multitude of different periods, but a daily signal from a subset of

SCN cells appears to synchronize the phases of those individual oscillators to the mean period of the group of neurons (Antle and Silver, 2005). However, not all neurons in the

SCN exhibit such endogenous rhythmicity (Aton and Herzog, 2005). The cells which do not express an overt rhythm receive input directly from the retinas, and then relay that light information to the rhythmic cells (Hirota and Fukada, 2004; Antle and Silver, 2005).

These rhythmic and non-rhythmic cells are located in distinct regions of the SCN – the rhythmic cells are located in what is called the dorsomedial ‘shell,’ whereas the light- sensitive retinorecipient cells are said to be located in the ventrolateral ‘core’ of the SCN

(Moore et al., 2002; Lee et al., 2003; Antle and Silver, 2005).

In the hamster, the SCN ‘core’ cells that receive direct retinal innervation are of four different phenotypes: gastrin-releasing peptide (GRP), vasoactive intestinal polypeptide (VIP), calbindin (CalB), and so-called ‘cap’ cells that are responsive to GRP

and contain phosphorylated extracellular signal-regulated kinase (P-ERK) at night (Antle and Silver, 2005). However, in the mouse, the cells of this ‘core’ region only express

GRP, with no further subdivision in cell sub-populations. This is also the region of the

SCN that predominantly receives extensive serotonergic projection from the midbrain raphe nuclei (Rea et al., 1994). In contrast, the SCN ‘shell’ cells are intrinsically rhythmic in their expression of Period (Per) and Cryptochrome (Cry) genes, and mainly express

vasopressin (VP; Antle and Silver, 2005).

1.4.2.2 Gene Expression in the SCN

Although only cells in the SCN shell are endogenously rhythmic in their

expression of clock genes, cells in the SCN core which receive retinal input express clock

genes immediately following exposure to light (Hamada et al., 2004). Core cells of the

SCN express immediate-early genes such as c-fos, as well as the clock genes Per1 and

Per2, in response to a light pulse during the subjective night, but do not rhythmically express these genes (Antle and Silver, 2005). In contrast, cells in the SCN shell

rhythmically express the clock genes Period and Cryptochrome through an

autoregulatory transcription-translation feedback loop (Holzberg and Albrecht, 2003).

Yet, these rhythmic shell cells do not all express Per genes in a common phase, and some

may actually be slave oscillators under the control of other pacemaker cells (Lee et al.,

2003; Hamada et al., 2004).

The mechanism of the negative autoregulatory transcription-translation feedback

loop of clock genes in the SCN shell has been described by many different review

articles, including Antle and Silver (2005), Reppert and Weaver (2001), and Okamura

(2004). Briefly, the loop begins with the dimerization of CLOCK and BMAL and the

subsequent binding of this complex to the promoter regions of the Per and Cry genes,

which activates the transcription process. As PER and CRY proteins accumulate in the

cytoplasm, they dimerize and move into the nucleus where they inhibit the activity of the

CLOCK-BMAL dimer. Thus, the PER-CRY dimer inhibits the further transcription of

the Per and Cry genes from which they are made. Once enough of the PER-CRY dimer has been degraded and removed from the nucleus, the process of gene transcription

resumes (Hastings and Herzog, 2004). Conversely, in the non-rhythmic retinorecipient

core of the SCN, Per genes are expressed following a light signal relayed to the neurons by the RHT (Antle and Silver, 2005). The light signal leads to the phosphorylation of cyclic adenosine monophosphate (cAMP) response-element binding protein (CREB), which then binds to the Ca2+/cAMP response element (CRE) in the promoter region of

the Period genes and induces transcription (Antle and Silver, 2005).

Light information must be relayed from the retinorecipient core region to the

endogenously rhythmic shell region in order to produce entrainment or photic phase

shifts in response to light signals (Antle and Silver, 2005). This communication between

the core SCN cells and the cells of the shell appears to be accomplished by multiple

neurotrasmitters and neuromodulators, including VIP, GRP, substance P (SP), and

gamma aminobutyric acid (GABA; Hamada et al., 2001; Antle and Silver, 2005). The

photic information relayed from the core cells then affects the rhythmic gene expression

in the shell cells of the SCN. The expression of clock genes has been shown to have a

very specific temporal and spatial organization (Hamada et al., 2004).

It has been demonstrated that Per1 gene expression first increases in the dorsomedial SCN, spreading ventrolaterally over the next several hours, subsequently

receding rapidly, first medially and then dorsally (Hamada et al., 2004). The expression of Per1 and Per2 extend across almost the entire SCN, spanning both the core and shell, except for a small area of GRP cells in the core in which Per expression is low and

arrhythmic (Hastings and Herzog, 2004). Based on real-time imaging of Per1-driven expression of a bioluminescent or fluorescent reporter, it was shown that the majority of

SCN cells express Per1 during the day, with peak expression occurring earlier in the

dorsal and lateral regions, and later in the ventral and medial regions (Hastings and

Herzog, 2004).

1.4.3 Efferent Pathways Out of the SCN

The SCN sends various efferent projections to targets located both within the

hypothalamus as well as other brain regions, with a projection pattern which appears to be predominantly ipsilateral (Swanson and Cowan, 1975; Berk and Finkelstein, 1981;

Watts and Swanson, 1987; Watts et al., 1987; Klein, 1991). One of the most pertinent efferent pathways consists of the SCN projections to the preoptic area, the most rostral portion of the hypothalamus, which is the region that controls many of the aspects of hypothalamic function which have circadian components, including fluid balance, reproduction, sleep, and thermoregulation (Klein, 1991). Another major efferent pathway involving many dorsocaudally directed fibers includes projections sent to the subparaventricular zone and the paraventricular nucleus of the hypothalamus. The

paraventricular nucleus is considered to be a major integrative center, concerned with

numerous processes relating to neuroendocrine, autonomic and behavioural functions

(Klein, 1991).

In terms of projections within the hypothalamus, the SCN has also been shown to send projections to the dorsomedial and ventromedial nuclei of the hypothalamus, which in turn have extensive connections to the hypothalamus, septum, amygdale, and brainstem. These areas have been implicated in several physiological functions including ingestive behaviour, rage, and female sexual behaviour (Klein, 1991). It also appears that there are a small number of fibers which may also project to the posterior hypothalamic area and the periaquaductal gray, as well as some efferent fibers which target the contralateral SCN. Although, there is little evidence as to whether they actually synapse with cells in the contralateral SCN or if they instead pass right through and continue on to more remote targets outside the SCN (Klein, 1991). Finally, the SCN sends projections through the retrochiasmatic area, which may serve as an alternate route by which circadian information can be relayed to neuroendocrine and autonomic circuits (Klein,

1991).

There are also a few efferent projections from the SCN which make their way to brain areas outside the hypothalamic area. Two of the most important of these targets of

SCN efferents include the rostral parts of the paraventricular and paratenial nuclei of the thalamus (Berk and Finkelstein, 1981; Watts and Swanson, 1987; Watts et al., 1987;

Klein, 1991). A much more minor projection of fibers from the SCN can be found targeting the ventral lateral septal nucleus, however this structure does receive a much

larger indirect input through the subparaventricular zone. Finally, a laterally directed

pathway has been observed which connects the SCN to the intergeniculate leaflet of the

lateral geniculate nucleus (Watts and Swanson, 1987; Watts et al., 1987). This pathway

may be involved in reciprocal connections regulating the activity of the geniculohypothalmic tract via the intergeniculate leaflet, a major source of neuropeptide

Y (NPY) innervation to the SCN which has been shown to play a role in photic modulation of circadian phase (Harrington et al., 1985).

1.5 Photic Phase Shifting

Light exposure around dawn and dusk, as well as inappropriate light during the night, can produce a shift in the time of an animal’s daily activity onset on subsequent days, leading to the animal either becoming active earlier or later than on previous days.

Such a shift in the phase of a circadian rhythm is termed a phase shift, and follows a very specific pattern called a photic phase response curve (PRC; Figure 1.1). Although there are subtle differences in exact timing between species, in nocturnal rodents this phase resetting by light is possible during the animal’s subjective night (when they are active),

but rarely during the subjective day (when they are inactive; Gillette and Mitchell, 2002).

During the early subjective night, brief exposure to light functions to delay behavioural

rhythms and causes the animal to have an activity onset that is later than expected based on previous onsets – i.e. a phase delay (Gillette and Mitchell, 2002). In contrast, a light stimulus during the late subjective night serves to phase advance an animal’s behavioural

Figure 1.1 Examples of a photic phase-response curve (PRC) for A) mice and B)

hamsters.

rhythms, such that activity onset on subsequent days will occur at an earlier time than expected (Gillette and Mitchell, 2002).

1.5.1 Effects of Light Intensity and Duration

Although timing of light exposure is of great importance, the specific properties of the light are also pertinent, especially the intensity and duration of the light. Although there has not been a large amount of data collected regarding the effects of intensity and duration on the magnitude of phase shift elicited, there is some research looking at the effects of these two aspects of light stimuli. For example, phase shifts induced by green light (λ = 503 nm) at circadian time (CT) 19 at various intensities and durations were examine by Nelson and Takahashi (1991). Regardless of stimulus duration, it was observed that no phase shifts could be elicited in hamsters at intensities below 0.2 lux, indicating that 0.2 lux is the threshold intensity for evoking phase shifts (Nelson and

Takahashi, 1991). Similar intensity thresholds have been found in hamsters, rats and mice as well (Foster et al., 1991; Bauer, 1992; Meijer et al., 1992; Sharma et al., 1999). These results also demonstrate the phenomenon of temporal summation, such that stimuli can be summed over time allowing for a trade-off between intensity and duration. For example, there are two ways to elicit a 60-minute phase shift – either by a short bright light pulse or by a long dim light pulse. Thus, in general, longer duration light exposure of a specific intensity induces larger phase shifts, and larger intensity light stimuli of a given duration also induce larger phase shifts.

Originally there appeared to be a ceiling effect to this temporal summation, such

that the peak phase shift appeared to be elicited following a light pulse somewhere

between 30 and 60 minutes in duration (Nelson and Takahashi, 1991; Daymude and

Refinetti, 1999). However, recently two independent studies have demonstrated that

much longer duration high intensity light exposure, of up to 12 hours in duration at ~1500

lux, can in fact increase the maximal phase shift to approximately 5 hours (Comas et al.,

2006; Antle et al., 2007). However, there may be a non-photic, activity-related, component to these phase shifts since blocking running-wheel activity following the transition back to dark at the end of the light pulse reduces the phase shift substantially

(Antle et al., 2007). Though for simplicity and efficiency, most studies examining the phase shifting effects of brief light stimuli tend to use light exposure of 5 to 30 minutes in duration at an intensity of 30-150 lux, which is generally considered to result in a sufficient activation to approximately saturate the circadian system (Foster et al., 1991;

Bauer, 1992; Meijer et al., 1992; Sharma et al., 1999).

1.5.2 Mimicking and Altering the Effects of Light

Exogenous application of some neurotransmitters or neuromodulators which are involved in the pathway responsible for the transduction of light information to the SCN are also able to induce phase shifts similar both in magnitude and direction to brief light exposure (Piggins et al., 1995; McArthur et al., 2000; Sterniczuk et al., 2008). GRP, N- methyl-D-aspartic acid (NMDA), and VIP are just a few examples of such photic-like chemicals (Antle and Silver, 2005; Sterniczuk et al., 2008). There are also many non-

photic stimuli which are able to modulate the effect that light has on the circadian phase.

For example, sleep deprivation for 8 hours during the subjective day has been shown to reduce the magnitude of light-induced phase delays (Challet et al., 2001). Sleep deprivation, by a slow rate of forced activity, for as little as 6 hours has even been shown to significantly attenuate the phase delay produced by an early night light pulse

(Mistlberger et al., 1997). Confinement to a novel running wheel late in the night cannot phase shift the circadian activity rhythm alone, but is capable of significantly attenuating the normal phase advance observed with a late night light pulse (Ralph and Mrosovsky,

1992; Mistlberger and Antle, 1998; Edelstein et al., 2003).

It has previously been demonstrated that there is a dramatic increase in serotonin release in the SCN during wheel-running activity (Dudley et al., 1998). Similarly, several studies have shown that the modulation of photic input to the circadian system induces an elevation of serotonin levels within the SCN (Asikainen et al., 1997; Grossman et al.,

2000). Thus, it appears that increased serotonin release in the SCN is able to inhibit light- induced phase shifts of the circadian activity rhythm (Pickard and Rea, 1997a).

Interestingly, pretreatment with the serotonergic antagonist metergoline is able to block the attenuation of late night photic phase advances, despite the inability of metergoline to significantly alter the photic phase shift itself (Mistlberger and Antle, 1998).

1.5.3 Light-Induced Protein and Gene Expression

Along with looking at the behavioural manifestations resulting from the modulation of circadian rhythms, it is also pertinent to investigate the changes in the

expression of specific genes within the SCN which coincide with these changes in

behaviour. Cells in the SCN shell are endogenously rhythmic in their expression of

several genes related to the functioning of the circadian clock, whereas cells in the SCN

core receiving retinal input express numerous genes immediately following exposure to

light (Hamada et al., 2004). Core cells of the SCN express immediate-early genes such as c-fos, as well as the clock genes Per1 and Per2, in response to a light pulse during the subjective night, but do not rhythmically express these genes (Antle and Silver, 2005). In contrast, cells in the SCN shell rhythmically express the clock genes Period and

Cryptochrome (Antle and Silver, 2005).

1.5.3.1 Immediate-Early Genes

Upwards of 20 genes in the mammalian system are rapidly and transiently induced following treatment with several different agents, including those which trigger mitogenesis, differentiation or membrane depolarization (Curran and Morgan, 1987).

They are considered to be a part of the signal-transcription coupling cascade which functions to link extracellular stimuli with long-term adaptive responses (Curran and

Morgan, 1987). They are proposed to function as third messengers, in that they mediate further changes in gene expression which convert short-term extracellular signals into long-lasting cellular changes (Guido et al., 1999a). Included in this unique class of genes termed immediate-early genes (IEGs), is the proto-oncogene c-fos. The two types of patterning in gene expression observed in the cells of the SCN, one dependent upon photic input during the subjective night and the other being spontaneously rhythmic

during the subjective day, can both be observed in the expression of the c-fos gene within

the SCN (Guido et al., 1999a).

These two expression patterns are generally observed in different cell sub-

populations of the SCN. The ventrolateral region of the SCN shows very little rhythmic

expression of c-fos, with only tiny peaks of expression in the early night and during the

day (Sumova and Illnerova, 2005). Instead, these ventral cells are the only SCN cells

which demonstrate light-induced expression of c-fos during the subjective night (Guido et al., 1999a). In contrast, the dorsomedial SCN cells are spontaneously rhythmic in their c-

fos expression, with a peak level of expression during the early day and the lowest levels of expression in the early night (Guido et al., 1999a; Sumova and Illnerova, 2005).

Circadian activity rhythms can be adjusted by aptly timed nocturnal light exposure, as previously discussed. Changes in the expression of several IEGs have been shown to coincide with such subjective night light exposure (Guido et al., 1999a; Guido et al., 1999b). It is thought that the light-induced expression of IEGs could be an early event in the response cascade which ultimately leads to light-induced phase alterations of the circadian pacemaker (Rea, 1992). However, the induction of the IEG c-fos is not sufficient, or even required to induce shifts in the circadian activity rhythm. This was discovered because mice with a null mutation at the c-fos gene locus are still able to respond to a phase-shifting light pulse (Honrado et al., 1996). Further, sensitivity to light exposure, in terms of induced gene expression, diminishes in the early subjective day despite robust spontaneous expression of c-fos (Guido et al., 1999b).

Light-induced c-fos expression appears to only be seen when the light exposure occurs at a time when behavioural phase shifts are generated (Rusak et al., 1990; Amir et al., 1998). This temporal specificity has been demonstrated by exposing animals to light in the early night and observing that the greatest concentration of Fos protein expression is found in the retinorecipient ventrolateral cells of the SCN (Amir et al., 1998). It has also been demonstrated that there is a unique spatial expression of c-fos, depending on whether the light is presented during the early or late subjective night (Rea, 1992). In the early subjective night, when light exposure results in a phase delay, Fos expression is mainly confined to the ventrolateral region of the SCN. In contrast, when light is presented late in the subjective night, and a phase advance of the rhythm is generally observed, Fos expression is found in both the ventrolateral as well as the dorsolateral cell sub-populations of the SCN (Rea, 1992).

1.5.3.2 Period Genes

The temporal and spatial expression of clock genes, discussed previously, is highly influenced by exposure to light during the subjective night (Yan and Silver, 2004).

Substantial light-induced expression of mPer1 in the core region of the SCN has been observed during the entire subjective night, whereas mPer2 expression is mostly seen in the SCN core after a light pulse in the early subjective night (Yan and Silver, 2002).

Quite a different pattern emerges in the shell of the SCN, where light-induced phase delays in the early subjective night cause mPer2 expression but very little mPer1 expression, and the opposite is true for phase-advancing light pulses in the late subjective night in which mPer1 is highly expressed and not mPer2 (Yan and Silver, 2002). After a

light pulse in the early subjective night, Per1 expression is mainly seen in the ventrolateral GRP cells of the SCN, whereas Per2 expression is much broader and is seen also in the VP cells (Dardente et al., 2002). Hamada et al. (2004) also found Per1 induction in the core region of the SCN (in the CalB-expressing cells) approximately one hour following a light pulse, and in the dorsomedial shell approximately 30 minutes later.

Thus, it appears that phase shifts occur only when light-induced Period gene expression spreads from the core to the shell of the SCN (Yan and Silver, 2002). More specifically, light-induced Per1 and Per2 gene expression in the SCN shell are associated with phase advances and phase delays respectively (Yan and Silver, 2002).

Further evidence of the spatial and temporal organization of Per gene expression in the SCN is seen when a light-pulse is given during the late subjective night, when

endogenous levels of mPer1 and mPer2 proteins are rising, and the resultant expression of mPer1 is further increased while mPer2 is unaffected (Yan and Silver, 2004). In

contrast, when a light pulse falls during the early subjective night, when endogenous levels of the two Per proteins are falling, expression of both mPer1 and mPer2 are

increased (Yan and Silver, 2004). These results seem to imply that there are likely

complex integrations between different cell types behind the differential expression of the

Per genes when induced by light, which appears to be a requirement for phase

adjustments to the circadian rhythm (Dardente et al., 2002).

During the subjective day, a very different pattern of expression emerges. Both

Per1 and Per2 are expressed similarly, restricted mainly to the region defined by the VP

cells of the SCN (Dardente et al., 2002). According to Dardente et al. (2002), it did not

appear that there was a role for light in the induction of Per gene expression during the

day. Yet, Challet et al. (2003) observed quite the opposite. It was found that even though

there was no phase-shifting effect of light exposure, there was still a general up-

regulation of Per1 and Per2 expression during the subjective day with a 30 minute light

pulse. Specifically, a more pronounced effect was observed with a light pulse

administered in the early subjective day. Also, it was found that Per2 expression was markedly increased with light exposure of up to 3 hours, whereas Per1 was not increased with extended light exposure (Challet et al., 2003).

1.5.3.3 Biochemical Pathways

Light-induced changes in the circadian system depend upon the photic information from the retina being relayed to the SCN cells and initiating the transcription of several pertinent genes mentioned above. SCN neurons contain second-messenger signaling pathways which couple the incoming photic stimuli with the transcription of immediate-early genes and subsequently Period genes within SCN cells (Butcher et al.,

2003). The mitogen-activated protein kinase (MAPK) pathway consists of three kinases, one of which is extracellular signal-regulated kinase (ERK; Butcher et al., 2003).

This MAPK cascade has been shown to be activated by light in a phase dependent manner in the SCN, such that it parallels the effects of light on the expression of

immediate-early genes (Obrietan et al., 1998). However, MAPK pathway activation occurs very rapidly, within minutes of photic stimulation, and before an increase in immediate-early gene mRNA expression is seen (Dziema et al., 2003). The disruption of this MAPK cascade has been shown to attenuate both the phase shifting effects of light

and photic-induced immediate-early gene expression (Butcher et al., 2002; Coogan and

Piggins, 2003; Dziema et al., 2003). The activation of the MAPK pathway by a light stimulus leads to the rapid activation of ERK, through its phosphorylation, which then dimerizes and moves into the nucleus of the cell (Butcher et al., 2003; Dziema et al.,

2003). This translocation of phosphorylated ERK into the nucleus is considered to be a key connection between the MAPK pathway and the activation of transcription of genes in the nucleus (Butcher et al., 2003). Specifically, P-ERK has been shown to interact directly with clock gene products of the transcription/translation feedback loop (Sanada et al., 2002). However, phase shifts can also be elicited when the levels of P-ERK are significantly decreased, as a result of a 3h sleep deprivation procedure during the mid- subjective day, also leading to a suppression of haPer1 mRNA (Antle et al., submitted).

Yet, administration of the drug U0126, which also prevents the phosphorylation of ERK, similar phase shifts were not induced (Antle et al., submitted).

In fact, is has also been determined that light-induced phase shifts are consistently accompanied by the phosphorylation of the transcription factor CREB protein within the

SCN (von Gall et al., 1998). Elevation of both Ca2+ and cAMP following light leads to the phosphorylation of CREB, which then binds to the cAMP response element of the promoter region several base pairs before the transcription start site of several circadian- related genes, including c-fos and Per1, thus leading to their transcription into mRNA

(Dunlap, 2004). Within minutes following light exposure, the phosphorylated form of

CREB (P-CREB) can be found in SCN cells, however this phosphorylation is phase- dependent, only occurring when light can induce phase shifts and immediate-early gene

expression (Ginty et al., 1993; Ding et al., 1997). Light has also been shown to trigger the colocalized expression of the activated forms of ERK 1/2 and the activated form of

CREB in cells of the SCN (Dziema et al., 2003).

In addition to the effects of photic stimulation only being observed during the subjective night, and not during the subjective day, levels of P-CREB have also been shown to be phase-dependent, with peak levels occurring during the mid- to late- subjective night (Obrietan et al., 1999). The pertinence of P-CREB to eliciting light- induced phase shifts and Per1 gene expression are highlighted when glutamate- as well as GRP-induced photic-like phase shifts are blocked following the application of an antisense oligonucleotide against Per1 and an antisense oligonucleotide decoy for the

CRE of the promoter region of the Per1 gene, thus preventing transcription of the gene

(Tischkau et al., 2003; Gamble et al., 2007).

1.6 Serotonergic Modulation of Circadian Rhythmicity

1.6.1 The Role of Serotonin

The neurotransmitter serotonin has been extensively researched in regards to its ability to modulate the activity of the SCN. The SCN receives dense serotonergic innervation arising from projections mainly from the median raphe nucleus (as in the hamster), but also partially from the dorsal raphe nucleus in some species (specifically in the rat; Pickard and Rea, 1997a). Both retrograde and anterograde tracing techniques have demonstrated that serotonergic neurons from the median raphe project to the SCN

(Meyer-Bernstein and Morin, 1996). Specifically, anterograde tracing of median raphe efferent neurons projecting to the SCN revealed synapses in a pattern similar to that of 5-

HT neuron fibers in the SCN, and serotonin-labeled fibers in the SCN demonstrate a retrograde tracing mainly to the median raphe (Jiang et al., 2000). These efferent projections from the median raphe form both axodendritic and axosomatic synapses with

SCN neurons receiving retinal input, and hence serotonergic innervation mainly resides in the core region of the SCN (Pickard and Rea, 1997a).

Serotonin is an inhibitory neurotransmitter that appears to modulate the effects of light on the SCN by dampening the excitatory effects light has on retinorecipient cells

(Meyer-Bernstein and Morin, 1996; Mistlberger, 2000). The daily profile of 5-HT release in the region of the SCN, in a light/dark cycle or constant darkness, indicates an abrupt extracellular increase in 5-HT which peaks at the light/dark transition or onset of subjective night, with a subsequent decrease during the night and into the following day

(Dudley et al., 1998). This rise in 5-HT levels corresponding to activity onset, at the beginning of the subjective night, lends support to the idea that the 5-HT output rhythm is not driven by light input. Serotonin has been shown to have numerous physiological effects on the SCN. A few of these actions include an inhibitory outward current at the postsynaptic membrane of some neurons, presynaptic inhibition of glutamate release from the RHT, reduced release of GABA from presynaptic terminals, and an excitatory inward current in some neurons (Jiang et al., 2000). Also, electrical stimulation of the median raphe and the application of 5-HT to the SCN have been shown to depress the firing rate of most SCN neurons (Pickard and Rea, 1997a).

Disruption of the serotonergic projections to the SCN, as well as pharmacological

manipulation of the synthesis or degradation of serotonin, has been extensively shown to alter various aspects of circadian rhythms in rodents (Rea et al., 1994). The removal of the afferent projections from the medial raphe to the SCN through the application of a neurotoxin has been shown to produce three main effects on the circadian system

(Pickard and Rea, 1997a). Re-entrainment to shifted light/dark cycles was accelerated,

larger light-induced phase shifts resulted from the administration of a 30-minute light pulse, and there was a significant increase of the freerunning period when in constant light. These three findings suggest that these 5-HT fibers play a role in the modulation of light input from the retina to the SCN. Thus, it is not surprising that increased endogenous serotonin in the SCN, stimulated by exogenous application of tryptophan, was able to attenuate the phase shifting effects of light (Glass et al., 1995).

1.6.2 Non-Photic Phase Shifts Mediated by the Serotonin System

1.6.2.1 Serotonergic Mediation of Activity and Arousal

Serotonin is thought to couple activity and arousal to the circadian system, and thus is also thought to participate in non-photic phase shifts of the circadian system, as well as activity-induced modulation of responses to light. Through the use of microdialysis to measure endogenous 5-HT, it was demonstrated that activity induced by novel running wheel access during the night suppressed 5-HT release, whereas the same induced activity during the day led to a stimulation of 5-HT release (Dudley et al., 1998).

Such novelty-induced running for 3h during the day has also been shown to cause phase

shifts, of approximately 3.5h, in the activity rhythm in hamsters (Bobrzynska et al.,

1996a). Although, this high amplitude activity does not seem to be strictly necessary, as

3h of daytime arousal, without induced activity, has been shown to also stimulate 5-HT release and induce large magnitude phase shifts (Antle and Mistlberger, 2000; Grossman et al., 2000).

However, high amplitude activity induced during the early or late night does not cause phase shifts in the activity rhythm (Mistlberger and Antle, 1998). Yet, novel wheel running-induced activity beginning before, and continuing through, a brief light pulse during the late subjective night significantly attenuates the normal phase shift induced by light at that circadian phase (Mistlberger and Antle, 1998). Induced activity combined with light exposure in the early subjective night, however, did not have any effect on the magnitude of phase shift normally observed with an early subjective night light pulse

(Mistlberger and Antle, 1998). This attenuation of the photic response by activity was found to be serotonin-dependent, such that pretreatment with the serotonin antagonist metergoline prevented the attenuation of the photic phase shifts by activity, but without affecting the levels of activity elicited by access to the novel running wheel (Mistlberger and Antle, 1998).

1.6.2.2 Chronobiotic Mediation of Non-Photic Phase Shifts

There is also a large body of evidence indicating a role for the serotonin system in modulating the circadian system and producing non-photic phase shifts through the use of pharmacological agents targeting serotonin receptors. For example, it appears that serotonin and its agonists can produce phase shifts during the subjective day, both in vivo

and in vitro (Smith et al., 2001). The systemic injection of 8-OH-DPAT has been shown

to result in phase advances during the mid-subjective day (Bobrzynska et al., 1996b;

Challet et al., 1998; Gardani and Biello, 2008). This non-photic phase shift induction by

8-OH-DPAT has even been observed in multiple types of lighting conditions, including a

light/dark cycle (LD), constant light, and constant darkness which are similar to those

induced by wheel-running (Shibata et al., 1992; Tominaga et al., 1992; Edgar et al., 1993;

Cutrera et al., 1994a; Bobrzynska et al., 1996b; Cutrera et al., 1996). Both the systemic

and intracerebro-ventricular injection of 5-HT agonists have been shown to result in phase advances during the mid-subjective day (Challet et al., 1998; Antle et al., 2003).

However, this non-photic effect is highly species-specific. For example, in both mice and hamsters, the 5-HT1,2 agonist quipazine was not able to induce phase shifts at a

range of circadian phases, whereas 8-OH-DPAT did induce phase shifts in hamsters but

only when delivered intraperitoneally and not with intra-SCN injections (Bobrzynska et

al., 1996b; Antle et al., 2003). Also, 8-OH-DPAT suppressed rhythmic daytime c-Fos in

hamsters, whereas there was no effect in mice (Antle et al., 2003). Similarly, the specific

effect can also depend on the exact agonist used along with which species is tested, with quipazine administered during the middle of the day serving to increase rhythmic Fos in mice but decrease it in hamsters (Antle et al., 2003).

In terms of the mechanism by which these chronobiotics produce their effects on the circadian system, 5-HT1A agonists may produce phase advances during the subjective

day by reducing the expression of Per1 and Per2 in the nocturnal rodent SCN (Horikawa et al., 2000). It was also postulated that the non-photic effects of 8-OH-DPAT may be the

result of binding to 5-HT7 receptors, with some possible involvement of 5-HT1A, as

opposed to the strict binding to 5-HT1A receptors to which this action was previously

attributed (Sprouse et al., 2004b). However, a 5-HT1A antagonist was shown to block 8-

OH-DPAT-induced phase shifts, further supporting the proposed role of the 5-HT1A

receptor (Sprouse et al., 2005). This is quite contrary to what this same research group

had found previously, where it appeared the primary action of 8-OH-DPAT was achieved

through binding to 5-HT7 receptors in hamsters (Sprouse et al., 2004b). Yet another study

by the same research group, still regarding the action of 8-OH-DPAT, showed that it was

actually the activation of 5-HT5A receptors which produced a shift in the circadian rhythm

(Sprouse et al., 2004a). Obviously, the effect of 8-OH-DPAT on the circadian system is

highly complex, and the mechanism of action has yet to be fully elucidated.

Thus, taking advantage of the generation of mice lacking 5-HT7 receptors allowed

more direct research into how the 5-HT agonist 8-OH-DPAT actually operates on the

circadian rhythm (Sprouse et al., 2005). It was observed that the in vitro application of 8-

OH-DPAT to SCN slices at mid-day induced phase advances in both normal mice and the

5-HT7 receptor knockout mice. Indicating that the lack of 5-HT7 receptors did not hinder

the effect of 8-OH-DPAT, and thus the effect was surmised to be the result of activity at

the 5-HT1A receptors. However, in vivo, the administration of 8-OH-DPAT systemically

to 5-HT7 receptor knockout mice produced no significant phase shifts in the mid-

subjective day, whereas phase shifts were induced in the wild type mice (Gardani and

Biello, 2008). Therefore, it appears that there is still a role for the 5-HT7 receptor in the

non-photic phase shifting abilities of 8-OH-DPAT when administered in the middle of

the day.

Not only does it appear that multiple serotonin receptors may be necessary for the

non-photic effects observed with 8-OH-DPAT, but it has also been shown that

serotonergic fibers which connect the raphe nuclei to the SCN are essential for the action

of 8-OH-DPAT (Schuhler et al., 1998). These projections from both the median and

dorsal raphe nuclei appear to be quite important to the ability of 5-HT and its agonists to

induce behavioural phase shifts. In fact, 8-OH-DPAT appears to act to alter the circadian system not by activity in the SCN itself, but instead via action in the raphe nuclei (Mintz et al., 1997). Further, stimulation of the dorsal and median raphe nuclei in hamsters has been found to elicit phase advances on its own (Meyer-Bernstein and Morin, 1999). Thus, it is obvious that the serotonergic projections from the raphe nuclei to the SCN are play some sort of role in eliciting phase shifts in the circadian system, and perhaps the involvement of 5-HT1A and 5-HT7 receptors in such phase shifts may be at least partly

through these raphe nuclei projections as well.

Along with the non-photic effects of serotonin agonists and the role of the raphe

nuclei on the circadian system, the effects that antagonists may have on their own as well

as on the effects induced by agonists must also be considered. As such, several serotonin

antagonists have been shown to block agonist-induced effects on the circadian system,

while having no effect on photic responsiveness when administered alone (Ying and

Rusak, 1997; Gannon, 2003). Pretreatment with three different specific 5-HT1A receptor antagonists (WAY-100635, NAN-190, and p-MPPF) were shown to block the effect of

the 5-HT agonist 8-OH-DPAT, while the weak partial agonist/antagonist BMY 7378 had

no blocking effect (Chen et al., 2002). Yet, due to its action as a mixed autoreceptor

agonist, systemic administration of BMY 7378 is able to suppress the excitatory effects

of 5-HT on the SCN (Mistlberger et al., 2000). The serotonin antagonist WAY-100635

was unable to induce any phase shifts when administered in isolation, but did induce

significant release of 5-HT in the SCN during the subjective day (Antle et al., 2000).

1.6.3 Serotonergic Mediation of Photic Phase Shifts

Currently, there are a total of 18 serotonin receptor subtypes recognized in the

brain, 6 of which are believed to be present in the SCN: 5-HT1A, 5-HT1B, 5-HT2A, 5-

HT2C, 5-HT5A, and 5-HT7 (Glennon, 1995; Smith et al., 2001; Niesler et al., 2007).

However, the specific contributions of each of these receptors in the functioning of the

SCN are still not well understood, and thus it is difficult to determine which serotonin

receptors are responsible for the mediation of photic effects on the circadian system. A

plethora of serotonin agonists and antagonists are available which can pharmacologically

manipulate serotonin activity by differentially activating receptors in the SCN. Each

individual agonist or antagonist has a unique receptor specificity and efficacy, which

results in varying effects on the circadian system following administration.

Much research has been done with serotonergic agonists and antagonists in order

to elucidate the role of serotonin in the circadian system. However, there are seemingly

conflicting results surrounding the effects of such pharmacological manipulations. The

agonistic or antagonistic effect of a substance on the circadian system is dependent upon

the mode of application, the species under consideration, and most importantly, the

location of the receptor within the brain at which the substance is an agonist or antagonist

– e.g. whether it has its action at a post-synaptic receptor or a pre-synaptic autoreceptor.

1.6.3.1 Affinity for Serotonergic Receptor Subtypes

One of the difficulties in determining exactly which receptor subtypes contribute

to, and mediate, the serotonergic effects observed on the mammalian circadian system, as

well as exactly how they accomplish these effects, lies in the lack of selective 5-HT

receptor ligands (Smith et al., 2001). The majority of serotonin receptor agonists and antagonists have an affinity for more than one receptor subtype, but to varying degrees

(Ying and Rusak, 1997). Some chronobiotics previously assumed to be highly selective,

to the point of being the prototypical choice for studies of a specific receptor subtype, have since been shown to bind to multiple receptor subtypes (Sprouse et al., 2004b).

An example of this is 8-OH-DPAT, which was originally thought to be a highly selective 5-HT1A receptor agonists, but is now known to be active at 5-HT7 receptors as

well (Sprouse et al., 2004b; Gardani and Biello, 2008). Other examples include the non-

selective 5-HT1,2 antagonist metergoline, the mixed 5-HT1A/1B agonist TFMPP, the mixed

5-HT2/7 receptor antagonist ritanserin, and the receptor antagonist Clozapine which is

primarily a dopamine antagonist but also has an affinity for 5-HT2A, 5-HT2C, 5-HT3, 5-

HT6 and 5-HT7 (Glass et al., 1995; Smith et al., 2001; Saifullah and Tomioka, 2003).

Similarly, the serotonin mixed agonists/antagonists BMY 7378, S 15535, and MDL

73005 EF are proposed to have their action mainly at the 5-HT1A receptors, but also have

affinities for other receptors as well (Gannon, 2003). There are also some serotonergic

chronobiotics which not only have affinities for multiple 5-HT receptor subtypes, but for

other receptor types as well. For example, although BMY7378 was thought to be highly

specific to the 5-HT1A receptors, it has also been shown to antagonize dopamine-D2

receptors (Millan et al., 1994). Another such drug is NAN-190 (Wesolowska et al.,

2002).

NAN-190 is generally considered to be a high affinity antagonist at postsynaptic

5-HT1A receptors, but it has also been shown to have an affinity for 5-HT2 and 5-HT7 as

well as dopamine-D2 and α1-adrenergic receptors (Glass et al., 1995; Rea et al., 1995;

Wesolowska et al., 2002; Saifullah and Tomioka, 2003). Plus, although NAN-190 is

mainly considered to be an antagonist at postsynaptic 5-HT1A receptors, and has potent

α1-adrenoceptor blocking properties (Claustre et al., 1991), it is also a partial agonist of

presynaptic 5-HT1A receptors and has been considered by some to be at least a partial

agonist at somatodendritic 5-HT1A autoreceptors (Portas et al., 2000; Wesolowska et al.,

2002). Therefore, it is obvious that much more research needs to be performed in order

to fully understand the effects that these different drugs are able to produce in the

mammalian circadian system.

1.6.3.2 Serotonin Agonists

In general, it has been shown that 5-HT agonists have the ability to modify the

response of the SCN to light exposure, and it has been reported that serotonin receptor

agonists may be able to mimic photic-like input to the SCN and independently induce

phase shifts of the circadian rhythm (Kohler et al., 1999; Kennaway et al., 2001; Smith et

al., 2001; Graff et al., 2005). Some researchers (Gannon, 2003) have made the assertion

that 5-HT and its agonists always inhibit photic responsiveness of the circadian system,

however there is an increasing body of evidence disagreeing with this statement.

For example, the serotonin1,2 agonist quipazine was shown to decrease the number

of cells expressing Fos-like immunoreactivity induced by a light pulse (Selim et al.,

1993). Similarly, the 5-HT1A agonist 8-OH-DPAT has been shown to inhibit both

behavioural phase shifts as well as c-fos expression in the SCN stimulated by light exposure during the latter part of the subjective night (Rea et al., 1994; Rea and Pickard,

2000). Although 8-OH-DPAT was not able to significantly alter the circadian phase when

administered alone in the late subjective night, it did dose-dependently inhibit light-

induced phase advances (Rea et al., 1994). It has also been shown to attenuate photic

phase shifts when administered systemically or locally (Rea and Pickard, 2000).

However, the effect of 8-OH-DPAT is species-dependent, with the attenuation of light- induced phase shifts observed in hamsters being completely absent in mice (Antle et al.,

2003). Yet, in rats, 8-OH-DPAT is actually able to mimic the immediate and long-term

effects of light on the melatonin rhythm, and reduced light-induced Fos expression to a

much smaller degree than seen in the hamster (Glass et al., 1994; Kennaway et al., 1996;

Recio et al., 1996).

Alternatively, and contrary to the assertion that serotonergic agonists always

inhibit light-responsiveness, the highly selective 5-HT1A agonist MKC-242 injected

systemically has been shown to potentiate phase advances and delays induced by a light

pulse (Moriya et al., 1998; Takahashi et al., 2002), exemplifying the necessity of looking

at exactly what type of receptor is being utilized by a drug. MKC-242 administered 30

minutes prior to a light pulse in the early subjective night was shown by Takahashi et al.

(2002) to prolong the light-induced expression of mPer1 and mPer2 in the mouse SCN.

This prolonged gene expression directly corresponded to the potentiation of photic phase delays. MKC-242 was also noted to have no effect on mPer1 or mPer2 expression in the

SCN without exposure to light. Further, mPer1 and mPer2 expression in the ventral portion of the SCN (likely the core area) was inducible by light, whereas their expression was not induced by light in the medial SCN (likely corresponding to the shell area) and instead demonstrated a strong rhythmic expression of the genes and proteins (Takahashi et al., 2002).

1.6.3.3 Serotonin Antagonists

It has been reported that 5-HT1A antagonists either reverse the action of agonists or produce the opposite effect (Ying and Rusak, 1997). However, it has also been shown that 5-HT1A antagonists are able to potentiate phase advances in the late subjective night

(Rea et al., 1995). For example, pre-treatment with the specific 5-HT1A antagonist WAY-

100635 has been shown to significantly potentiate a light-induced phase delay during the early subjective night. It was, however, ineffective at adjusting the magnitude of phase shifts during the late subjective night (Smart and Biello, 2001). Similarly, Rea et al.

(1995) found that the serotonin antagonist NAN-190, injected systemically, is also able to potentiate photic phase shifts by a substantial amount, when administered in both the early and late subjective night.

1.6.3.4 Serotonin Mixed Agonists/Antagonists

Although there are several other substances which act as serotonergic antagonists,

they also bind to some receptors agonistically, and are thus considered to be a class of drugs which is distinct from the strict serotonergic agonists and antagonists. Thus, along with the two classes of drugs which are considered to be only agonists or antagonists at serotonin receptors, there is a unique third class of drugs that act as mixed agonists/antagonists at serotonergic receptors (Gannon, 2003). This class of drugs is termed mixed agonists/antagonists due to their action as antagonists at presynaptic axoaxonic serotonin heteroreceptors, as well as agonists at postsynaptic 5-HT somatodendritic autoreceptors. The three drugs of this class used by Gannon (2003) were

BMY 7378, S 15535, and MDL 73005 EF. Both BMY 7378 and S 15535 administered during the late subjective night were able to potentiate photic phase shifts to a great degree, and MDL 73005 EF to a lesser degree (Rea et al., 1995; Gannon, 2003).

Considered to be highly efficacious, when it is administered in advance of a light pulse, BMY7378 was found to greatly potentiate late subjective night photic phase advances of the hamster circadian activity rhythm (Rea et al., 1995; Gannon, 2003).

Byku and Gannon (2000) found BMY7378 to potentiate phase advances by approximately 250-300% compared to control treatments. However, it is not able to significantly potentiate light-induced phase delays in the early subjective night (Gannon,

2003). More recently, S 15535 was shown to potentiate the effect of light nearly three- fold in the late subjective night (Gannon and Millan, 2006). It was also discovered that when BMY 7378 is administered during the mid- to late-subjective day it produced a phase advance in the circadian rhythm without any light exposure (Gannon, 2003).

BMY7378 has also been shown to decrease the amount of 5-HT released in the hamster SCN following a systemic injection of the drug (Dudley et al., 1998). This decreased 5-HT release coincides well with the suspected mechanism of action of this class of mixed serotonin agonist/antagonists, such that it could represent the outcome of an agonistic action at the median raphe to prevent serotonin release onto the SCN along with an antagonistic action at the RHT or SCN cells to block serotonergic inhibition at these location. Thus, this two-facetted decrease of inhibition to the SCN would result in an overall potentiated photic phase shifting effect of pre-treatment with this class of drugs.

1.6.3.5 Special Case of NAN-190

Included in the class of serotonin mixed agonists/antagnists is the drug 1-(2- methoxyphenyl)4-[4-(phthalimido)butyl]piperazine hydrobromide (NAN-190), originally thought to be strictly a 5-HT1A antagonist. However, the determination of the specific actions of NAN-190 is complicated by its marked lack of receptor specificity.

Specifically, NAN-190 has been shown to be a potent antagonist at postsynaptic 5-HT1A receptors, but it also has potent α1-adrenoceptor blocking properties (Claustre et al.,

1991). NAN-190 is also a partial agonist of presynaptic 5-HT1A receptors, and has been considered by some to be at least a partial agonist at somatodendritic 5-HT1A autoreceptors (Portas et al., 2000; Wesolowska et al., 2002). However, NAN-190 is also believed to bind to 5-HT7 receptors (Lovenberg et al., 1993). Therefore, it appears that

NAN-190 is a mixed agonist/antagonist at 5-HT1A receptors, but also has some affinity for numerous other receptors. More investigation is needed to conclusively determine the

receptor at which NAN-190 is most active, as pertaining to the mammalian circadian

system.

NAN-190 has been shown to be able to potentiate photic phase shifts by a substantial amount when administered in both the early and late subjective night (Rea et al., 1995). It is able to potentiate the effects of light in both dark-adapted and entrained

hamsters, it greatly decreases the number of days required to re-entrain to a 6h advance in

the light/dark cycle, and it even potentiates photic phase shifts when administered up to

6h after the onset of the light exposure (Kessler et al., 2008). However, the potentiating

effects of administering NAN-190 several hours after a phase-shifting light pulse have

yet to be replicated. NAN-190 has also been shown to block the effect of the 5-HT1A agonist 8-OH-DPAT (Claustre et al., 1991). It has been proposed that NAN-190 regulates part of the circadian system through its action to block the inhibitory effect of serotonin release (Rea et al., 1995). The agonistic action of NAN-190 at presynaptic somatodendritic 5-HT1A receptors also leads to decreased release of serotonin at the SCN,

as well as at the IGL (Recio et al., 1996). In addition, the ability of NAN-190 to block

the accumulation of the cyclic AMP stimulated by the 5-HT1A,1D agonist 5-CT has been shown to likely be the result of activation of 5-HT7 receptors (Lovenberg et al., 1993).

These results indicate that the modulation effects of NAN-190 on the photic

signaling involved in circadian rhythmicity may be a result of the presence of 5-HT7 receptors, despite the prevailing view that NAN-190 acts mostly through its activation of the 5-HT1A receptor (Lovenberg et al., 1993). Yet, part of the inhibitory effect of NAN-

190 on serotonergic transmission also appears to be due to its ability to block α1-

adrenoceptors (Claustre et al., 1991). Regardless of which receptor is mediating the effect, the blockade of serotonergic inhibitory signals to the SCN results in NAN-190 modulating the photic response of the circadian pacemaker and potentiating phase shifts to light stimuli (Rea et al., 1995; Takahashi et al., 2002).

Although the information regarding the receptor activation with NAN-190 has been somewhat inconclusive thus far, research into the cell populations in the SCN which are involved and the portions of the photic pathway affected in the photic potentiation observed with NAN-190 has been a bit more fruitful. In an attempt to better understand how NAN-190 exerts its effect on the circadian system, Sterniczuk et al. (2008) investigated the effects that NAN-190 has on different neurotransmitter systems which are known to be involved in the pathway of light information towards the SCN, and are released from distinct cell populations following light exposure. Administered without light exposure, these neurochemicals or their agonists are known to induce photic-like phase shifts, mimicking the effects of light on the circadian system. Sterniczuk et al.

(2008) have demonstrated that NAN-190 administered to hamsters can not only potentiate the effects of light, but is also able to potentiate phase shifts induced by the direct application of some of the peptides released along the pathway between the retina and the SCN.

Specifically, it was demonstrated that a systemic injection of NAN-190 significantly potentiates the phase shifts induced by NMDA application directly into the

SCN, both in the early and late subjective night (Sterniczuk et al., 2008). However,

NAN-190 was not shown to affect the Fos protein expression induced by the NMDA

treatment. Similarly, NAN-190 significantly potentiated phase advances in the late

subjective night following administration of GRP directly to the SCN. Yet, unexpectedly,

significantly smaller phase delays were observed when NAN-190 was combined with the

GRP treatment. This difference in phase delays induced in the early night was examined

further, such that it was shown to be accompanied by an increase in Fos protein

expression in a cell population called the ‘cap’ region of the SCN core, but no change

was observed in the VIP region of the core nor in the shell of the SCN. Interestingly, and

contrary to previous findings, Sterniczuk et al. (2008) also observed significant phase

advances, along with a trend towards larger phase delays, induced by the administration

of NAN-190 without any other pharmacological treatment. Thus, it is still not fully understood exactly how NAN-190 is able to induce such large magnitude phase delays in the circadian system of hamsters.

1.6.4 Serotonergic Mediation of Light-Induced Gene Expression

Administration of serotonin agonists, including 5-HT1A/7 receptor agonists, which

attenuate or block light-induced phase shifts of the rodent activity rhythm, have also been

shown to attenuate the expression of Fos in the hamster SCN (Amir et al., 1998). Both systemic and intra-SCN injections of the 5-HT agonist quipazine can reduce light- induced Fos expression in the SCN during the subjective night (Selim et al., 1993).

Similarly, two 5-HT1A receptor agonists, 8-OH-DPAT and buspirone, have been

demonstrated to inhibit light-induced Fos-like immunoreactivity when administered 30

minutes prior to a half-hour duration light pulse (Glass et al., 1994). This inhibition was

mainly observed in the ventrolateral core of the SCN, with little effect in the dorsal and lateral regions of the SCN. This inhibitory effect of 8-OH-DPAT could be completely abated by NAN-190 pretreatment, despite the inability of NAN-190 to induce a change in the Fos-like immunoreactivity when injected alone (Glass et al., 1994). However, this study was performed prior to the discovery that NAN-190 increases the effects of light on behavioural phase shifts, and thus they did not look at the effect of NAN-190 and light exposure without the 5-HT agonist pretreatment.

The selective 5-HT1A receptor agonist MKC-242 administered 30 minutes prior to a light pulse in the early subjective night was shown by Takahashi et al. (2002) to prolong the light-induced expression of mPer1 and mPer2 in the mouse SCN. This prolonged gene expression directly corresponded to the potentiation of photic phase delays. MKC-242 was also noted to have no effect on mPer1 or mPer2 expression in the

Although the 5-HT1A receptor agonist MKC-242 has also been shown to potentiate photic phase advances in the late subjective night, this action was antagonized by pre-treatment with the selective 5-HT1A receptor blocker WAY100635. However, despite the phase-shift potentiation ability of MKC-242, it has been shown to have no effect on the light-induced c-fos expression in the SCN when administered 30 minutes

prior to a light pulse (Moriya et al., 1998). One suggested explanation of this result

proposed by Moriya et al. (1998) was the MKC-242 could be acting downstream of the

light-induced c-fos expression seen in the SCN. Further, this receptor agonist could be

activating 5-HT1A autoreceptors in the midbrain raphe to reduce the serotonin activity in

the SCN, thus allowing for the potentiation of the effect of light (Moriya et al., 1998).

It has been previously shown that 5-HT1A antagonists either reverse the action of

agonists or produce the opposite effect (Ying and Rusak, 1997; Horikawa et al., 2000).

Plus, it is known that 5-HT1A antagonists are able to potentiate phase advances in the late

subjective night (Rea et al., 1995). Thus, it is possible that the administration of a 5-HT1A antagonist, such as NAN-190, along with a light pulse in the late subjective night could result in the increased expression of Per1 and Per2 in the SCN. This increase could then artificially induce the Per levels that would normally have been seen slightly earlier in the subjective night – inducing a phase advance.

Similarly, NAN-190 has been shown to attenuate the inhibitory effect of serotonin release, as stimulated by administration of tryptophan, on the c-fos expression normally induced by a light pulse (Glass et al., 1995). When tryptophan was administered 1 hour prior to light exposure, there was an approximately 77% reduction in the number of cells expressing Fos-like immunoreactivity compared to the light only condition. However, pretreatment with NAN-190 attenuated this suppressive effect of the tryptophan on the

Fos-like protein expression. As previously reported, NAN-190 administered alone had no significant effect on the amount of Fos-like immunoreactivity detected in the SCN (Glass et al., 1995). Yet, when NAN-190 was combined with a light stimulus, it was found to

affect Fos expression by partially blocking the expression normally induced in the

ventrolateral SCN by exposure to a light pulse (Recio et al., 1996).

1.6.5 Transgenic Mice

A major difficulty in determining exactly which receptor subtypes contribute to

and mediate serotonergic effects on the mammalian circadian system, as well as how they do so, lies in the lack of highly selective 5-HT receptor ligands (Smith et al., 2001). In order to better determine the location of action of nonselective chronobiotics, genetically

modified mice have been developed (Sprouse et al., 2005). These mice have been

engineered such that they are missing a specific receptor under consideration (Sprouse et

al., 2005). Parks and colleagues (1998) created a strain of knockout (KO) mice with an

inactivated gene for encoding the 5-HT1A receptor. In this case, the 5-HT1A receptor gene

was partially substituted with a neomycin gene segment, such that there is no longer a

start codon signaling to begin the translation of the gene into a protein (Parks et al.,

1998). The homozygous knockout mice then have two substituted alleles, and heterozygous animals have one normal receptor allele and one non-functional substituted

allele. Experimental animals can then be generated by the breeding of heterozygous

animals, such that both homozygous wildtype (WT) and homozygous knockout (KO)

animals can be generated in the same litter – this allows for litter-mate control animals to guard against the effects of treatment being due to some strain or litter effect instead of as

a result of the experimental manipulation. Therefore, this strain of mouse is highly useful

in order to better understand the action of serotonin agonists and antagonists at this

specific receptor subtype.

There is ample evidence suggesting that serotonergic modulation of photic

transmission into the SCN may be achieved at least partially through postsynaptic

mechanisms mediated by 5-HT1A or 5-HT7 receptors on SCN neurons as well as through

5-HT1B receptors located on retinal terminals (Pickard and Rea, 1997a). Interestingly, 5-

HT1B receptor knockout mice are known to have attenuated behavioural responses to

light, in the form of reduced light-induced phase shifts (Sollars et al., 2006b). Similarly,

5-HT7 receptor knockout mice also demonstrate attenuated phase shifts in activity rhythm

in response to light, although this effect is limited to late-subjective night light exposure

(Gardani and Biello, 2008). This result would not be expected, since the removal of a

receptor which would inhibit photic responsiveness would be predicted to result in an

increase in the magnitude of the photic response. Thus, it is important to examine the role

of individual receptors in the actual transmission of photic information to the SCN,

without the potential interactions with pharmacological agents which are generally used

to examine receptor contribution. However, receptor knockout technology must be

examined with some caution, as it is difficult to fully understand or predict the possible

compensatory mechanisms at work in a system lacking a receptor from birth which could

also affect photic responsiveness.

By comparing wild type mice to knockout mice with the inactivated receptor gene

on how they react to the administration of certain nonspecific receptor agonists and

antagonists, it may be possible to better understand the location of action producing the

effect observed on the circadian system (Sprouse et al., 2005). The non-selective 5-HT antagonist NAN-190 could be tested in this manner to determine the specific receptor at which it acts to produce the observed modulation of photic phase shifts in the mammalian circadian activity rhythm. It is known that NAN-190 does act as an antagonist at 5-HT1A receptors, but since it also binds at other receptors, including 5-HT7, it is difficult to

determine the receptor activation that is specifically producing the phase-shift

potentiation observed (Rea et al., 1995). By employing 5-HT1A receptor knockout mice, it

should be possible to determine if NAN-190 is producing its effect specifically through

the 5-HT1A receptor.

1.6.6 Summary

Through various different receptors, serotonin acts on the mammalian circadian

system to inhibit photic input to the SCN. Projections from the median raphe into the

SCN are the major source of this serotonergic input, acting to dampen the responsiveness of the SCN to photic input. Although effects are somewhat species-specific, serotonergic agonists generally attenuate photic responses during the subjective night, but can also potentially induce phase shifts on their own when administered during the subjective day.

In contrast, serotonin antagonists are able to either prevent or reverse the effects of

agonists, as well as potentiate the effects of light when delivered at the appropriate

phases. Several serotonin agonists and antagonists are also known to affect the expression

of certain genes and proteins involved in the molecular clockwork of the SCN. A unique

class of drugs termed mixed agonists/antagonists, which possess both agonistic and

antagonistic properties in different brain regions, have been shown to greatly potentiate

light-induced phase shifts of the circadian system. However, the complete

pharmacological mechanisms underlying the abilities of these drugs, including NAN-190,

remain to be fully elucidated. The precise mechanism of action is necessary before a

chronobiotic could be developed which is tailored to elicit the desired effects upon the

circadian system without any undesirable side-effects.

1.7 Present Study: Objectives and Hypotheses

Serotonin is known to play a major role in the regulation of circadian rhythmicity

and modulate the effects of photic input to the SCN. The 5-HT1A mixed

agonist/antagonist NAN-190 has been shown to greatly potentiate the phase-shifting

response of the circadian system to photic stimuli. However, it has still unknown exactly

how NAN-190 achieves this enhanced response to light, both in terms of the receptor to

which it binds and in terms of the effects it has on molecular components of the SCN

cells. The purpose of the present study is three-fold: 1) to better understand the role of the

5-HT1A receptor in the mouse circadian system, by characterizing the response of 5-HT1A receptor knockout mice to various photic and non-photic manipulations; 2) to better understand the contribution of the 5-HT1A receptor to the enhancement of behavioural

phase shifts with NAN-190, by comparing the phase shifts elicited in 5-HT1A knockout

mice compared to their heterozygous and wildtype counterparts; 3) to better understand

which clock-related genes and proteins demonstrate altered expression as a result of

NAN-190, and are thus involved in the behavioural potentiation effect observed.

2. Altered photic and non-photic phase shifts in serotonin1A receptor knockout mice

Victoria M. Smith1, Roxanne Sterniczuk1, Catherine I. Phillips1 and Michael C. Antle1,2,3

1 Department of Psychology, University of Calgary, 2500 University Drive NW, Calgary, AB, T2N 1N4, Canada. 2Hotchkiss Brain Institute and 3Pharmacology & Therapeutics, University of Calgary, 3330 Hospital Drive NW, Calgary, AB, T2N 1N4, Canada.

Running Head: Phase Shifting 5-HT1A Knockout Mice

Author Contributions:

Victoria Smith - Laboratory Research: 5-HT1A receptor knockout mouse colony generation and maintenance, animal genotyping, predictions for manipulation times, brief light pulse manipulations, long light pulse and wheel-lock manipulations, phase shift calculations, injections, perfusions, tissue collection, immunocytochemistry, in situ hybridization, tissue image capturing, cell counting and ROD analysis - Manuscript Preparation: Entered and organized data; ran statistical analyses, produced figures, composed methods and results Roxanne Sterniczuk: - Experiment 4 (8-OH-DPAT): Assisted with carrying out predictions for manipulation times, injections, and phase shift calculations Catherine Phillips: - Experiment 3A (Fos expression): Provided technical assistance with immunocytochemistry and counting cells Dr. Michael Antle: - Trained me on immunocytochemistry and in situ hybridization techniques - Provided laboratory, equipment and financial support necessary to carry out experiments - Manuscript Preparation: Composed introduction and discussion, provided feedback and editing of the manuscript

Modified version of:

Smith, VM, Sterniczuk, R, Phillips, CI, Antle, MC (Submitted). Altered photic and non-

photic phase shifts in serotonin1A receptor knockout mice. Neuroscience.

Abstract

The mammalian circadian clock located in the suprachiasmatic nucleus (SCN) is thought to be modulated by serotonin (5-HT). 5-HT is thought to inhibit photic phase shifts both by inhibiting the release of glutamate from retinal terminals, as well as by decreasing the responsiveness of retinorecipient cells in the SCN. Furthermore, there is also evidence that 5-HT may underlie, in part, non-photic phase shifts of the circadian system.

Understanding the mechanism by which 5-HT accomplishes these goals is complicated by the wide variety of 5-HT receptors found in the SCN, the heterogeneous organization of both the circadian clock and the location of 5-HT receptors, and by a lack of sufficiently selective pharmacological agents for the 5-HT receptors of interest.

Genetically modified animals engineered to lack a specific 5-HT receptor present an alternative avenue of investigation to understand how serotonin regulates the circadian system. Here we examine behavioral and molecular responses to both photic and non- photic stimuli in mice lacking the 5-HT1A receptor. When compared to wildtype controls,

these mice exhibit larger phase advances to a short late-night light pulse and larger delays

to long 12h light pulses that span the whole subjective night. Fos and mPer1 expression

in the retinorecipient SCN is significantly attenuated following late-night light pulses in

the 5-HT1A knockout animals. Finally, non-photic phase shifts to 8-OH-DPAT are lost in

the knockout animals, while attenuation of the phase shift to the long light pulse due to

rebound activity following a wheel lock is unaffected. These findings suggest that the 5-

HT1A receptor plays an inhibitory role on behavioral phase shifts, a facilitatory role on

light-induced gene expression, a necessary role in phase shifts to 8-OH-DPAT, and is not

necessary for activity-induced phase advances that oppose photic phase shifts to long light pulses.

Key Words: Circadian, Suprachiasmatic Nucleus, 8-OH-DPAT, Fos, Per1

2.1 Introduction

The mammalian circadian clock, located in the suprachiasmatic nucleus (SCN), regulates daily oscillations in physiology and behavior (Antle and Silver, 2005). These endogenously generated rhythms are entrained to the daily environmental cycles through exposure to light (Daan and Pittendrigh, 1976; Rusak and Zucker, 1979) as well as various non-photic cues such as activity (Mrosovsky, 1996), dark pulses (Boulos and

Rusak, 1982), and sleep deprivation (Antle and Mistlberger, 2000). The neurotransmitter serotonin (5-hydroxytryptamine, 5-HT) has been implicated in mediating and modulating both photic and non-photic phase shifts of the circadian system.

Photic information is relayed from the retina to the SCN by way of the glutamatergic retino-hypothalamic tract (RHT; Mintz and Albers, 1997; Mintz et al.,

1999). Serotonin innervation from the median raphe nucleus (Meyer-Bernstein and

Morin, 1996; Morin and Meyer-Bernstein, 1999) is thought to modulate the release of glutamate from the RHT (Rea and Pickard, 2000), as well as the postsynaptic response to glutamate by cells in the SCN (Ying and Rusak, 1994, 1997; Rea and Pickard, 2000).

These responses are thought to be mediated by 5-HT1B receptors on RHT terminals

(Pickard et al., 1996; Pickard and Rea, 1997b; Pickard et al., 1999; Rea and Pickard,

2000), as well as 5-HT1A, 1B, and 7 receptors on SCN cells (Ying and Rusak, 1994; Ying

and Rusak, 1997; Rea and Pickard, 2000; Belenky and Pickard, 2001). Blocking the

release and/or postsynaptic actions of 5-HT appears to enhance phase shifts both to light

(Rea et al., 1995; Smart and Biello, 2001; Gannon, 2003) and to treatments that mimic

light by activating the photic pathway downstream of glutamate release (Sterniczuk et al.,

2008).

In rats, agonists for the 5-HT2A/2C receptor produce photic-like phase shifts of

locomotor, temperature and melatonin rhythms (Kennaway et al., 1996; Kennaway and

Moyer, 1998; Kalkowski and Wollnik, 1999; Kohler et al., 1999) and induce expression

of the immediate early gene c-fos and the clock genes Per1 and Per2 in the SCN (Moyer

et al., 1997; Kennaway and Moyer, 1999; Kohler et al., 1999; Ferguson and Kennaway,

2000; Varcoe et al., 2003). These results suggest that, for this species, serotonin may

help to mediate photic phase shifts. Recent findings suggest that this effect may be

mediated by 5-HT3 receptors on RHT terminals (Kennaway and Moyer, 1999; Graff et

al., 2005; Graff et al., 2007). It has been hypothesized that activation of the 5-HT3 receptors on the RHT terminals stimulates or enhances glutamate release, thereby eliciting a photic response (Graff et al., 2007).

Serotonin has also been suggested to underlie non-photic phase shifts of the circadian clock. Serotonin release is associated with behavioral state (Jacobs and Fornal,

1995; Jacobs and Fornal, 1997; Jacobs and Fornal, 1999), with high levels of serotonin release being observed during locomotor activity. Serotonin release in the SCN is at its lowest during the day (Dudley et al., 1998), but output increases dramatically at this time if hamsters are kept awake (Grossman et al., 2000) or run in a wheel (Dudley et al.,

1998). Electrical stimulation of the median raphe elicits serotonin release at the SCN and produces phase shifts consistent with those elicited by non-photic stimuli (Meyer-

Bernstein and Morin, 1999; Glass et al., 2000). When the 5-HT1A/7 agonist 8-hydroxy-2-

(dipropylamino)-tetralin (8-OH-DPAT) is given systemically to hamsters (Tominaga et al., 1992; Antle et al., 2003) or in vitro to rat or mouse SCN slice cultures (Prosser, 2000;

Prosser, 2003; Sprouse et al., 2004b; Guscott et al., 2005; Sprouse et al., 2005), phase advances can be elicited during the daytime. Systemic application of 8-OH-DPAT was initially reported to have no phase shifting properties in mice (Antle et al., 2003), however more recent studies suggest that 8-OH-DPAT can phase shift the mouse circadian system when applied 6 hours before activity onset (Horikawa and Shibata,

2004; Gardani and Biello, 2008), a phase not examined in the original study.

Understanding how serotonin influences both photic and non-photic phase shifts has been complicated by a number of factors: the different species and techniques employed to investigate the system; the wide range of serotonin receptors, each with specific distributions in both the brain and the synapse; and a limited range of selective agonist and antagonists for the receptors of interest. Of particular interest to the serotonergic regulation of both photic and non-photic regulation of circadian rhythmicity is the 5-HT1A receptor. The 5-HT1A receptor serves not only as a postsynaptic receptor

(Kia et al., 1996; Riad et al., 2000), but also as a somatodendritic autoreceptor in the raphe nuclei (Verge et al., 1986; Sotelo et al., 1990), that decreases serotonin output when activated (Dudley et al., 1999). Thus systemic application of 5-HT1A agonists could conceivably activate postsynaptic receptors while simultaneously inhibiting serotonin output, thereby decreasing activation of all other 5-HT receptors. Conversely, systemic application of 5-HT1A antagonists could conceivably block all postsynaptic 5-

HT1A receptors, while simultaneously blocking the 5-HT1A autoreceptors, resulting in an

increase in 5-HT output, effectively increasing the activation of all other 5-HT receptors.

Furthermore, while early studies implicated the 5-HT7 receptor in regulation of non-

photic phase shifts to 8-OH-DPAT in vitro (Lovenberg et al., 1993; Prosser, 2000), recent

studies employing 5-HT7 knockout mice suggest that the 5-HT7 receptor is only partially

responsible for phase shifts to 8-OH-DPAT (Guscott et al., 2005; Sprouse et al., 2005).

In these studies the phase shifts were only fully blocked in 5-HT7 knockout mice when a

5-HT1A receptor antagonist was included. Given the problematic pharmacology involved

in studying serotonergic regulation of circadian rhythmicity, specific 5-HT receptor

knockout mice present a novel tool for understanding the role served by those particular

receptors. We examine here how circadian responses to both photic and non-photic

treatments are altered in 5-HT1A knockout mice.

This study examined knockout animals compared to their wildtype counterparts in

behavioural phase shifts elicited by a brief exposure to light in the early- and late-

subjective night. It is hypothesized that if the 5-HT1A receptor is involved in the

transmission of light information from the retina to the SCN, then the removal of one

serotonin receptor subtype will decrease the amount of inhibition delivered to the retinal

terminals, and there will be an increased responsiveness of the 5-HT1A receptor knockout

mice in comparison to the wildtype controls. Behavioural phase shifts were also

examined following long duration light exposure, with and without a subsequent

prevention of rebound activity. It was hypothesized that the removal of the 5-HT1A receptor would also increase the responsiveness of the knockout animals to longer duration light exposure; however prevention of the non-photic effect of rebound activity

would not necessarily be affected. Underlying protein and gene expression in the cells of

the SCN were also examined following a brief exposure to light, either in the early- and

late-subjective night (Fos protein) or only in the late subjective night (mPer1 gene). It

was predicted that the expression of Fos and mPer1 would be altered in the knockouts

compared to the wildtypes. Finally, non-photic phase shifts to the 5-HT1A/7 receptor

agonist 8-OH-DPAT were assessed, and if the 5-HT1A receptor activation is the source of the phase shift generation observed with this drug, as is predicted, then it is expected that the phase shift will be absent in the knockout animals.

2.2 Experimental Procedures

2.2.1 Animals and Housing

A total of 46 male mice were used, consisting of 13 C57BL/6J mice obtained from the University of Calgary Biological Sciences breeding colony (Calgary, AB,

Canada), as well as 24 5-HT1A receptor knockout (KO) mice and 9 5-HT1A receptor

wildtype (WT) mice. The knockout mice were generated from a breeding colony

established in our lab from mice originally developed by Dr. Thomas Shenk (Princeton

University, Princeton, NJ), provided by Dr. Miklos Toth (Cornell University Medical

College, New York, NY) and bred on the C57BL/6J background. For information regarding the generation of the 5-HT1A receptor KO mice see Parks et al. (1998). Animals

from our breeding colony were genotyped using a pair of KO primers (amplifying a 400

base-pair product that included a portion of the neomycin cassette that replaced the start

codon of the 5-HT1A receptor gene) as well as a pair of WT primers (amplifying a 238 base-pair product that included a portion of the 5-HT1A receptor gene replaced by the

neomycin cassette in the KO animals) to identify the presence or absence of each allele of

the 5-HT1A receptor gene (KO up primer: CTT TAC GGT ATC GCC GCT CCC GAT

TC; KO down primer: TGC AGG ATG GAC GAA GTG CAG CAC A; WT up primer:

AGT GCA GGC AGG CAT GGA TAT GTT; WT down primer: CCG ATG AGA TAG

TTG GCA ACA TTC TGA). Mice were individually housed in Nalgene Type L clear

polycarbonate cages (30.3 cm long × 20.6 cm wide × 26 cm high; Nalg Nunc

International, Rochester, NY), equipped with a stainless steel running wheel (diameter of

24.2 cm). Animals were housed in a temperature- (21 ± 1°C) and humidity-controlled

room, with ad libitum access to food and water. Cages were changed at least every 14

days (7-10 days prior to, and following a manipulation). All protocols were approved by

the Life and Environmental Sciences Animal Care Committee at the University of

Calgary, and adhered to the Canadian Council on Animal Care guidelines for the ethical

use of animals.

2.2.2 Activity Rhythms

Wheel-running activity was continuously monitored using magnetic switches

mounted on the running wheel, and connected to a computer running the Clocklab data

collection software package (Coulbourn Instruments, Allentown, PA, USA). Actograms

(graphical representation of wheel-running behavior) were generated and analyzed by

Clocklab analysis software. Mice were allowed to entrain to the light-dark (LD) cycle or

free-run in constant darkness (DD) for a minimum of 7 days prior to the start of

manipulations. Onset of running wheel activity for manipulation day was predicted using

a regression line fit to the activity onsets for the 7-10 days prior to the testing day.

2.2.3 Experiment 1 – Phase Shifts to Short Duration Light

Experiment 1 examined the effects of a brief exposure to dim light on behavioral phase shifts of the activity rhythm. Animals were housed in DD (0 lux) for the duration of this experiment. The light pulse was administered using a light-sealed box, which was located in the animal room, with a light bulb partially covered in aluminum foil such that the light intensity at cage level was measured as 40 lux. Mice (n = 12, 6 KO and 6 WT) were exposed to a 15-minute light pulse at two different phases in their circadian activity rhythm, either 4 h or 10 h after their predicted activity onset (i.e., either at circadian time

(CT) 16 or 22, where activity onset is designated to be CT12 by convention). Animals

received each light pulse treatment on separate occasions in a counterbalanced fashion.

Following the light pulses, actograms were analyzed using the automatic fitting function

included in the Clocklab software, creating one line of best fit for the activity onsets of

the 10 days prior to and including the manipulation day, and another line of best fit for

the 10 days following the light pulse (excluding the first 3 onsets to avoid transient

effects). The phase shift in the behavioral activity rhythm was calculated based on the

horizontal difference between the predicted onsets from the two regression lines on the

day following the light pulse.

2.2.4 Experiment 2 – Phase Shifts to Long Duration Light

In experiment 2, the effects of a long duration exposure to bright light, with and without access to a running wheel, on behavioral phase shifts of the activity rhythm were examined. A modified Aschoff type II design was used, such that animals were entrained to a 12h:12h LD cycle (at cage height the light intensity was ~1500 lux when the lights were on and 0 lux when the lights were off) prior to the manipulation and were subsequently released into DD following the end of the experimental light exposure.

Mice (n = 15; 9 KO and 6 WT) were exposed to two conditions, in counterbalanced fashion, beginning at the end of the normal lights-on period for manipulation day: 1) 12 h of light exposure (12h LP), and 2) 12 h of light exposure followed by the running wheel being locked for the first 12 h of darkness (12h LP+WL). Wheels were locked by manually inserting a stiff metal bar horizontally through the wheel rungs to prevent wheel rotation, which was fixed in place using lab tape so the animals were unable to push it out of the way. Wheels were manually locked and unlocked by the experimenter with the aid of night-vision goggles (BG15Alista, Richmond Hill, Ontario, Canada). Following the manipulation, animals were left to free run in DD for at least 10 days before being re- entrained to the LD cycle for the next manipulation, and phase shifts were calculated as stated above.

2.2.5 Experiment 3 – Protein and Gene Expression Underlying Photic Phase Shifts

Experiments 3a and 3b examined the effects of short duration, dim light exposure on the expression of Fos protein and mPer1 gene expression in the SCN, respectively.

Mice (3a: n = 30, 14 KO and 16 WT; 3b: n = 26, 14 KO and 12 WT) were housed in DD

for the duration of this experiment. Individuals from each genotype were randomly

assigned to a phase group (CT16 or CT22 for Fos; only CT22 for mPer1) and a treatment

group (light pulse or no light pulse at the assigned phase). Animals were killed 90 min

after their assigned phase (i.e., at either CT17.5 or CT23.5) with an intraperitoneal (i.p.) injection of Euthanyl (0.3mL). The head was wrapped in aluminum foil to prevent the retinas from being exposed to light prior to and during the perfusion. Animals were then perfused using approximately 50 mL of phosphate buffered saline (PBS), followed by 50 mL of buffered 4% paraformaldehyde. Brains were extracted and postfixed in 4% paraformaldehyde overnight, and subsequently cryoprotected in a 20% sucrose solution for 24 h. Brains were sliced through the anterior hypothalamus using a cryostat, with

alternate 35 μm sections through the SCN being collected for each of immunocytochemistry (ICC) and in situ hybridization.

2.2.6 Experiment 4 – Non-photic Phase Shifts to the Serotonin Agonist 8-OH-DPAT

In experiment 4, the effects of the serotonergic1A/7 agonist 8-OH-DPAT (5 mg/kg;

Sigma, St. Louis, MO) on behavioral phase shifts of the activity rhythm were examined.

Mice (n = 12, 6 KO and 6 WT) were allowed to free-run in DD for approximately 10

days prior to the first manipulation. The mice were then subjected to two manipulations

separated by 2 weeks presented in a counterbalanced fashion; 1) an i.p. injection of both

8-OH-DPAT and 2) vehicle control (sterile saline). Injections were delivered at CT6,

with the aid of night-vision goggles. Following each manipulation, actograms were

analyzed using the methods described above.

2.2.7 Immunocytochemistry

Sections were triple-labeled for Fos (goat anti-Fos; 1:20,000; Santa Cruz

Biotechnology, Santa Cruz, CA), vasopressin (VP; guinea-pig anti-VP; 1:5,000;

Peninsula Laboratories, San Carlos, CA), and bombesin/gastrin-releasing peptide (GRP;

rabbit anti-GRP; 1:5,000; ImmunoStar Inc., Hudson, WI). ICC staining was completed

using a previously described method from our lab (see Sterniczuk et al., 2008). Briefly,

brain sections were exposed to three 10-min washes in 0.1M PBS, followed by a 1 h

incubation in a blocking buffer (2% normal donkey serum, in 0.1M PBS and 0.1% Triton

X-100 (PBSx)), after which the tissue was incubated for 48 h at 4°C in the primary

antibodies diluted in blocking buffer. Tissue was then washed using six 10-min rinses in

0.1% PBSx, and left overnight in 0.1% PBSx. Finally, the brain sections were washed

with another six 10-min 0.1% PBSx rinses, incubated for 1 h in the following secondary antibodies: CY-2 donkey anti-rabbit, CY-3 donkey anti-goat, and CY-5 donkey anti-

guinea pig (each at 1:200; Jackson ImmunoResearch Laboratories Inc., West Grove, PA),

and exposed to three final 10-min washes in 0.1M PBS. Sections were then mounted on

gelatin-coated slides, air dried, dehydrated in a series of alcohol rinses, cleared with

xylenes, and coverslipped with Krystalon.

2.2.8 In Situ Hybridization

For mPer1 in situ hybridization, all instruments and trays were cleaned with

RNase Zap (Ambion, Austin, TX) and all solutions were prepared with RNase-free water.

Free floating tissue sections were processed using the probe and digoxigenin (DIG)

protocol of Yan and Silver (2002). First, tissue was treated with proteinase K (1 mg/mL,

0.1M Tris buffer pH 8.0; 50 mM EDTA; 10 min) at 37°C. This reaction was stopped by

adding 2 mL 4% paraformaldehyde. Following rinses in saline sodium citrate (300mM

NaCl, 30mM sodium citrate) the tissue was treated with 0.25% acetic anhydride in 0.1M

triethanolamine for 10 min. The sections were then incubated in 1.5 mL hybridization

buffer [50% formamide, 60mM sodium citrate, 600mM NaCl, 10% dextran sulphate, 1%

N-laurylsarcosine, 25 mg/mL tRNA, 1X Denhardt's, 0.25mg/ml salmon sperm DNA]

containing the DIG-labeled mPer1 (mPer1 plasmid courtesy of Dr. L. Yan, Michigan

State University, East Lansing, MI), antisense cRNA probes (0.1 µg/ml; for details see

Hamada et al., 2001) for 16 h at 60°C.

After two 30 minute high-stringency post-hybridization washes (50% formamide/saline sodium citrate at 60°C), sections were treated with RNase A. After more high-stringency washes, the tissue was then processed for immunodetection with a nucleic acid detection kit (Roche, Indianapolis, IN). The sections were incubated in 1.0% of blocking reagent in buffer 1 (100 mM Tris-HCl buffer, 150 mM NaCl, pH 7.5) for 1 h at room temperature. Sections were then incubated at 4°C in alkaline phosphatase- conjugated Dig antibodies diluted 1:5000 in buffer 1 for 3 days. On the following day, sections were washed in buffer 1 twice (15min each), and incubated in buffer 3 (100 mM

Tris-HCl buffer, pH 9.5, containing 100 mM NaCl and 50 mM MgCl2) for 5 min. They

were then incubated in a solution containing nitroblue tetrazolium salt (0.34 mg/mL) and

5-bromo-4-chrolo-3-indolyl phosphate toluidinium salt (0.18 mg/mL; Roche) for 16 h.

The colorimetric reaction was stopped by immersing the sections in buffer 4 (10 mM

Tris-HCl containing 1 mM EDTA, pH 8.0). The tissue was then mounted on gelatin

coated slides, air-dried, briefly dehydrated in an alcohol series, cleared with xylenes, and

coverslipped with Permount.

2.2.9 Quantification of Fos and mPer1 Expression

Images of the immunofluorescently-labeled tissue were captured via an Olympus

BX51 microscope equipped with filters for each of the three cyanine dyes. Bilateral

counts of Fos-immunoreactive (Fos-IR) cells were performed using computerized image

analysis (ImagePro Plus 5.1.2.59; Media Cybernetics, Inc., Bethesda, MD), and were

manually counted by two researchers who were blind to the treatment conditions. Cell

counting was completed at the mid-caudal level of the SCN, identified by the presence of

the GRP subnucleus. Separate cell counts were obtained for two regions of the SCN: shell (delineated by VP-IR) and core (delineated by GRP-IR). Images of the digoxigenin- labeled tissue were captured using the RGB color filter for the camera of the same microscope as used above. Bilateral counts of the relative optical density (ROD) of mPer1 expression were performed using computerized image analysis software (ImageJ

1.34s; National Institutes of Health, Bethesda, MD). ROD was calculated at the mid-

caudal level of the SCN, with SCN regions (core, shell) being delineated based on the general shape of each region identified previously while performing the Fos cell counts.

2.2.10 Statistical Analysis

For experiment 1, two-tailed independent-samples t-tests were conducted to determine if there were significant differences in the magnitude of the photic phase shifts between the two genotypes at each of the two phases. For experiment 2, paired t-tests were used to examine the effect of treatment, while independent samples t-tests were used to examine the effects of genotype. Experiments 3a and 3b were each analyzed using a mixed two-way analysis of variance (ANOVA), to determine if there were significant differences in the amount of protein or gene expression in each of the core and shell regions of the SCN between the two genotypes. Separate ANOVAs were used for each of the two light pulse conditions at each of the two phases examined in each part of experiment 3. A Pearson r correlation coefficient was also calculated to determine the magnitude of inter-rater reliability between cell counters for experiment 3a (r=0.973,

P<0.0001). For experiment 4, a mixed two-way ANOVA was used to analyze the magnitude of the phase shifts to the two treatment conditions between the two genotypes.

Statistical significance was set at P<0.05 for all tests, using SigmaStat Version 2.03

(Systat Software, Inc.; San Jose, CA). All significant ANOVAs were followed up by post hoc multiple comparisons using Student-Newman-Keuls tests. All means are reported as

± standard error of the mean in the figures.

2.3 Results

2.3.1 Experiment 1 – Phase Shifts to Short Duration Light

Exposure to a light pulse at CT22 induced significantly larger phase advances in wheel-running activity in the 5-HT1A receptor KO mice when compared to WT animals

(t(10)=4.75, P<.0001; Figure 2.1E). However, there was no difference found in the magnitude of the phase delays induced by a light pulse at CT16 between the KO and WT

mice (t(10)=0.43, P=0.67; Figure 2.1E). Figure 2.1A-D show representative actograms for

each genotype at each of the phases examined. While the freerunning period was usually shortened by a light pulse, by about 0.12h, there was no significant main effect of phase

(F(1,20)=1.762, p>0.05) or genotype (F(1,20)=.074, p>0.05), nor was there a significant interaction (F(1,20)=0.724, p>0.05) between phase and genotype on this phenomenon,

therefore any effect that period change had on phase shift estimates would not have

differed between groups.

2.3.2 Experiment 2 – Phase Shifts to Long Duration Light

Large phase shifts were observed to the 12 h extension of light (12h LP) in both

genotypes (Figure 2.2). Knockout animals exhibited significantly larger phase delays than did the WT animals (t(13)=2.17, p=0.048). Consistent with previous findings (Antle

et al., 2007), locking the wheels for the 12h following the LP (12h LP+WL) significantly

attenuated the phase shifts in both the WT (t(13)=3.10, p=0.026) and KO animals

Figure 2.1. (A-D) Representative actograms of running wheel activity for (A,C) a WT

and (B,D) a KO animal receiving a 15-minute light pulse, denoted by a circle (z),

at (A,B) CT16 or (C,D) CT22. Each horizontal line of an actogram represents a

single day, with each subsequent day being plotted directly below the previous

day. Daily activity is represented by black vertical marks in each row, with the

height of the vertical marks being proportional to the amount of activity. Dark

grey diagonal lines represent regression lines fit to activity onsets both before and

after exposure to a light pulse. (E) Mean phase shifts (± SEM) of WT and KO

animals treated with a light pulse at CT16 and CT22. A positive value indicates a

phase advance of the activity rhythm, while a negative value indicates a phase

delay. (**) indicates statistical significance between genotypes on a treatment to

be P<0.01.

Figure 2.2. (A,B) Representative actograms for (A) a WT and (B) a KO animal depicting

its response to each of two manipulations. Areas of no shading indicate times

when the lights were on, whereas light grey shading indicates darkness. The

hatched box during the second manipulation represents when the lights were off

and the running wheel was locked. Dark grey vertical and diagonal lines represent

regression lines fit to the activity onsets before and after a manipulation. The

treatments were: 12h of extended light followed by DD (12h LP), and 12h of

extended light followed by the wheel being locked for the first 12h of DD (12h

LP+WL). (C) Mean phase delays (± SEM) of WT and KO animals receiving a

12h light pulse and a 12h light pulse followed by a 12h wheel lock. (*) indicates

statistical significance difference between indicated groups (P < 0.05).

(t(13)=3.45, p=0.008). Again, the KO animals exhibited significantly larger phase delays

than did the WT animals in the 12h LP+WL treatment (t(13)=2.60, p=0.022).

2.3.3 Experiment 3A – Fos Protein Expression Underlying Photic Phase Shifts

A brief light pulse in the late subjective night (CT22) resulted in a significant

interaction between the two genotypes and the two regions within the SCN (F(1,7)=7.936,

P<0.05; Figure 2.3E). The majority of the Fos expression was located within the core region of the SCN in all animals. However, the KO animals demonstrated a significant attenuation in the amount of Fos which was expressed in the core when compared to their

WT counterparts, while no differences were found in the shell expression. Some Fos expression was observed the peri-SCN region in WT animals and this expression was conspicuously absent in the KO animals, although this phenomenon was not specifically analyzed as it fell outside the boundary of the SCN. Animals that were not exposed to a light pulse in the late subjective night, regardless of genotype, exhibited minimal Fos expression in either region of the SCN (Figure 2.3E). Representative examples of the Fos expression for each genotype in each light pulse condition in the late subjective night are presented in Figure 2.3A-D.

Similar results were observed following brief light exposure in the early subjective night (CT16), with reduced Fos expression in both the core and shell regions of the SCN in the KO mice compared to WT mice (Figure 2.3J). As with the CT22 light pulse, the vast majority of the Fos-expressing cells were located in the core of the SCN in all animals. However, the interaction between genotype and SCN region did not quite

Figure 2.3. Fos protein expression at CT23.5 (A-E) and CT17.5 (F-J). Representative

SCN tissue slices indicating Fos-IR cells (in green) in the late subjective night,

collected from (A) a WT and (B) a KO animal following no light exposure, as

well as (C) a WT and (D) a KO animal following treatment with a 15-minute light

pulse at CT22. (E) Mean number of Fos-expressing cells (±SEM) in the core and

shell regions of the SCN of WT and KO animals following each light treatment

condition in the late subjective night. Representative SCN tissue slices indicating

Fos-IR cells in the early subjective night, collected from (F) a WT and (G) a KO

animal following no light exposure, as well as (H) a WT and (I) a KO animal

following treatment with a 15-minute light pulse at CT16. (J) Mean number of

Fos-expressing cells (± SEM) in the core and shell regions of the SCN of WT and

KO animals following each light treatment condition in the early subjective night.

GRP-IR (red) and VP-IR (blue) were used to delineate SCN regions for the

purpose of cell counting. Scale bar = 200µm. (**) indicates a statistical

significance of P<0.01.

reach the level of significance (F(1,7)=4.933, P=0.062). Again, the peri-SCN region of WT

animals exhibited some Fos expression which was not observed in KO animals, although

this was not specifically analyzed as it fell outside the boundary of the SCN. Also similar to the late subjective night, when no light pulse was administered in the early subjective night, very few Fos-expressing cells were found in either region of the SCN in all animals

(Figure 2.3J). Representative examples of the Fos expression found in the early subjective night conditions across both genotypes can be found in Figure 2.3F-I.

2.3.4 Experiment 3B – mPer1 Gene Expression Underlying Photic Phase Shifts

Comparable with the results of the Fos expression, a significant attenuation in the amount of mPer1 gene expression was observed in KO animals relative to WT mice

(F(1,5)=10.525, P=0.023; Figure 2.4E) following a light pulse at CT22, with the majority of the expression again appearing in the core of the SCN (F(1,5)=21.413, P<0.01; Figure

2.4E). Samples of mPer1 gene expression observed in each light pulse condition for each

genotype can be found in Figure 2.4A-D.

2.3.5 Experiment 4 – Non-photic Phase Shifts to the Serotonin Agonist 8-OH-DPAT

The magnitude of phase shifts induced by the serotonin agonist 8-OH-DPAT and

its vehicle control (saline) at CT6, resulted in a significant interaction between genotype

and type of injection being observed (F(1,10)=11.591, P<0.01; Figure 2.5E). Specifically,

WT mice exhibited significantly larger phase advances when receiving a treatment of 8-

OH-DPAT compared to vehicle control (P<0.05). Alternatively, KO mice injected with

Figure 2.4. mPer1 gene expression at CT23.5 (A-E). Representative SCN tissue slices

indicating mPer1 gene expression in the late subjective night, collected from (A)

a WT and (B) a KO animal following no light exposure, as well as (C) a WT and

(D) a KO animal following treatment with a 15-minute light pulse at CT22. (E)

Mean relative optical density (ROD) of mPer1 expression (±SEM) in the core and

shell regions of the SCN of WT and KO animals following each light treatment

condition in the late subjective night. Scale bar = 200µm. (**) indicates a

statistical significance of P<0.01.

Figure 2.5. (A-D) Representative actograms of running wheel activity for (A,C) a WT

and (B,D) a KO animal receiving an i.p. injection of either (A,B) saline (control)

or (C,D) 8-OH-DPAT (5 mg/kg), denoted by a diamond (), at CT6. Dark grey

diagonal lines represent regression lines fit to activity onsets both before and after

exposure to a light pulse. (E) Mean phase shifts (± SEM) of WT and KO animals

treated with either saline or 8-OH-DPAT at CT6. (**) indicates statistical

significance between genotypes on a treatment to be P<0.01.

8-OH-DPAT demonstrated not just smaller phase advances, but actually reversed their

phase shift to a delay, which was significantly different from the small phase advance

observed with the injection of saline (P<0.05). The phase advances induced by 8-OH-

DPAT in the WT mice were also found to be significantly different from the delays

induced in the KO mice (P<0.001), whereas there was no difference in the phase

advances induced by saline between the two genotypes (P=0.919). Representative

actograms for each genotype receiving each injection can be found in Figure 2.5A-D.

2.4 Discussion

This is the first study to examine circadian responses to both photic and non-

photic stimuli in mice lacking the 5-HT1A receptor. Elimination of the 5-HT1A receptor

resulted in larger phase advances to short light pulses, larger delays in response to long

light pulses, decreased light-induced Fos and mPer1 expression in the SCN, and altered

responses to a non-photic stimulus, namely the administration of 8-OH-DPAT. While

compensatory mechanisms may cloud the interpretation of findings using any KO

models, our findings are largely consistent with the results from other serotonin receptor

KO models (Sollars et al., 2002; Sprouse et al., 2005; Sollars et al., 2006b; Sollars et al.,

2006a; Gardani and Biello, 2008) as well as the results from 5-HT1A agonist studies

(Tominaga et al., 1992; Ying and Rusak, 1994; Rea and Pickard, 2000; Smart and Biello,

2001; Antle et al., 2003) and lesions studies (Morin and Blanchard, 1991; Bradbury et al.,

1997).

Serotonin is thought to inhibit activation of both retinal terminals in the SCN (Rea and Pickard, 2000) and retinorecipient SCN neurons (Ying and Rusak, 1994), inhibiting the effects of light on the circadian clock. Consistent with this, phase advances to a 15 minute light pulse were enhanced in the 5-HT1A KO mice. Of the three 5-HT KO models

(5-HT1A, 5-HT1B, and 5-HT7), only the 5-HT1A KO mice used in the present study exhibited enhanced phase advances to late-night light pulses. The 5-HT1B KO mice have significantly smaller advances (Sollars et al., 2006a), while the 5-HT7 KO mice show phase delays rather than advances to light in the late night (Gardani and Biello, 2008).

Phase delays to short light pulses were not affected in our KO model.

When compared against their WT counterparts, neither the 5-HT1B KO nor the 5-

HT7 KO mouse exhibit significantly different phase delays to light, although the 5-HT1B

KO shows a non-significant attenuation at CT16 (Sollars et al., 2006a), while the 5-HT7

KO shows a non-significant potentiation at CT12 (Gardani and Biello, 2008). The lack of significant enhancement in the phase delay zone may represent a ceiling effect to short light pulses. Long light pulses have been shown to produce very large delays (Comas et al., 2006; Antle et al., 2007), and phase shifts to these long light pulses were potentiated in the 5-HT1A KO mice in the present study. Responses of other KO models to the long light pulse paradigm have yet to be examined.

Taken together, these data imply that only the 5-HT1A mouse is consistent with the position that serotonin inhibits light responses in the SCN, suggesting that serotonin may be acting through the 5-HT1A receptor to modulate behavioral responses to light.

The paradoxical response of the 5-HT1B KO mouse may in part be due to the

rd heterogeneity of receptor distribution in the SCN, in that only 1/3 of 5-HT1B receptors

are found on retinal terminals (Sollars et al., 2006a). As the retinally-derived 5-HT1B

receptors are thought to inhibit the transmission of photic signals to the SCN, selective

knockout of retina-specific 5-HT1B receptors may yield enhancement of photic phase shifts, in contrast to the global knockout of 5-HT1B studied to date (Sollars et al., 2006a).

It is rather puzzling that phase delays, but not advances, are potentiated when

hamsters are pretreated with the selective 5-HT1A antagonist WAY100635 (Smart and

Biello, 2001). The discrepancy between our findings with the 5-HT1A KO mouse and the

pharmacology study in hamsters may be explained by the fact that, while WAY100635

blocks postsynaptic 5-HT1A receptors, it also blocks 5-HT1A somatodendritic

autoreceptors on raphe cells, resulting in an increase in 5-HT release at the SCN (Dudley

et al., 1999; Antle et al., 2000) which would activate all other 5-HT receptors. Basal 5-

HT levels in the 5-HT1A KO mouse are not different from levels observed in WT mice

(Bortolozzi et al., 2004), suggesting that activation of other 5-HT receptors is no different

in the 5-HT1A KO than it is the WT mice.

The role of the 5-HT1A receptor in non-photic phase shifting is not clear. Despite

the numerous studies pointing towards involvement of 5-HT acting at the SCN to mediate

and mimic non-photic phase shifting (Lovenberg et al., 1993; Cutrera et al., 1994b; Penev

et al., 1995; Prosser, 2000; Prosser, 2002; Prosser, 2003; Guscott et al., 2005; Sprouse et al., 2005), there are numerous studies suggesting that this may not be the case. Direct

application of 8-OH-DPAT to the hamster SCN in vivo does not phase shift the circadian

system (Mintz et al., 1997; Antle et al., 2003). Furthermore, selective activation of

serotonergic cells produces a significant increase in 5-HT output at the SCN, but does not

phase shift the circadian clock (Antle et al., 2000). Lesioning 5-HTergic input to the

SCN or blocking 5-HT receptors in the SCN does not block non-photic phase shifts to

daytime exercise (Bobrzynska et al., 1996a; Antle et al., 1998; Meyer-Bernstein and

Morin, 1998). The discrepancy between the in vivo with comparable in vitro findings for

SCN application has been suggested to be caused by rapid regulation of 5-HT receptor

density in the in vitro model (Ehlen et al., 2001; Prosser et al., 2006).

Two studies examining the response of 5-HT7 KO mice to in vitro application of

8-OH-DPAT to the SCN (Guscott et al., 2005; Sprouse et al., 2005) suggest that 8-OH-

DPAT produces phase shifts through combined activation of both the 5-HT1A and 5-HT7 receptors. Another study demonstrated that phase shifts to 8-OH-DPAT at CT6 are lost in 5-HT7 KO mice (Gardani and Biello, 2008). Our finding demonstrated that 5-HT1A

KO mice also do not respond to 8-OH-DPAT. These data suggest that both the 5-HT1A

and 5-HT7 receptors are necessary for 8-OH-DPAT to elicit a phase shift. Because the 5-

HT1A and 5-HT7 receptors have opposite effects on cAMP (cyclic adenosine monophospate) production, it has been suggested that these receptors are found on different cell populations within the SCN to mediate the in vitro response to 8-OH-DPAT

(Sprouse et al., 2005). With systemic application of 8-OH-DPAT in vivo, it is even more

likely that the phase shifts to 8-OH-DPAT depend on the involvement of different 5-HT

receptors in different areas.

It has been suggested that 8-OH-DPAT may be acting in both the dorsal and

median raphe nuclei (Mintz et al., 1997) and may depend on the presence of 5-HTergic

fibers projecting from the raphe nuclei to the SCN (Schuhler et al., 1998). It is known

that 8-OH-DPAT acts in the raphe to decrease 5-HT output to the SCN (Dudley et al.,

1999) or at the IGL to regulate NPY release to the SCN (Challet et al., 1998). Further complicating the picture is the recent suggestion that 8-OH-DPAT may also be acting through the 5-HT5A receptor to produce phase shifts in the in vitro SCN slice model

(Sprouse et al., 2004a).

Despite the loss of the non-photic phase shift response to 8-OH-DPAT in the 5-

HT1A KO mice, it appears that these mice can still respond to non-photic stimuli. We

have previously shown that rebound activity following a wheel-lock after a 12h light

pulse attenuates the magnitude of the photic phase shift (Antle et al., 2007). In this study, we demonstrate that 5-HT1A KO mice also show attenuation of their shifts to long light

pulses when their wheels are locked for 12 hours following the light pulse. The fact that

the phase shifts to the 12h LP+WL condition is larger in the 5-HT1A KO mice than in the

WT mice may reflect the fact that 5-HT1A mice have larger photic phase delays but

identical non-photic phase advances when compared to WT controls. Take together with

the 8-OH-DPAT findings, this suggests that the network underlying non-photic phase

shifts is still intact in 5-HT1A KO mice, but that it can no longer be activated by 8-OH-

DPAT. This would be consistent with 8-OH-DPAT acting outside the SCN to produce

non-photic phase shifts in vivo. This would also be consistent with the studies that

suggest 5-HT activity at the SCN is not necessary for non-photic phase shifting

(Bobrzynska et al., 1996a; Antle et al., 1998; Meyer-Bernstein and Morin, 1998).

The observation that phase advances to light were significantly enhanced in the 5-

HT1A KO mice, yet light pulse-induced Fos and mPer1 expression in the SCN was

significantly decreased in the SCN core, was puzzling. Original reports on light pulse-

induced Fos expression suggested that the magnitude of the phase shift and the amount of

Fos expression were positively correlated (Kornhauser et al., 1990). However, a number

of reports have begun to challenge this conclusion (Recio et al., 1996; Sterniczuk et al.,

2008). The 5-HT1A partial agonist NAN-190 has been shown to dramatically increase

photic phase shifts (Rea et al., 1995), and yet pretreatment with NAN-190 decreases Fos expression in response to light (Recio et al., 1996) and NMDA microinjections to the

SCN (Sterniczuk et al., 2008).

Furthermore, NAN-190 significantly decreases phase shifts to gastrin-releasing peptide injections to the SCN, yet increases Fos expression to this treatment (Sterniczuk et al., 2008). A decrease in light pulse-induced Fos is also observed in mice lacking the

5-HT1B receptor, although these mice also show smaller behavioral phase shifts to light

pulses (Sollars et al., 2006a). While the retinorecipient SCN is the region expressing the

greatest amount of Fos and mPer1 following a light pulse, this region is not believed to

be the site of the cell autonomous circadian oscillators (Hamada et al., 2001; Karatsoreos

et al., 2004; Antle and Silver, 2005). The function of gene expression in light-responsive, non-rhythmic SCN cells is unknown. We also noted that Fos expression immediately outside the SCN appeared greater in WT animals, although this was not specifically quantified.

Interpretation of studies using genetically modified animals engineered to lack a particular gene must always be interpreted carefully, as unanticipated compensatory

mechanisms can lead to differences between the KO animal and its WT counterparts

beyond just the absence of a particular gene. Specifically, loss of one 5-HT receptor may lead to changes in the levels of expression or activity of other 5-HT receptors to compensate. Levels of 5-HT1B receptors in the 5-HT1A KO mouse have been suggested

to either be unaffected (Richer et al., 2002; Bortolozzi et al., 2004) or be supersensitive

(Boutrel et al., 2002). As mentioned, 5-HT levels in the brain are not different between

WT and 5-HT1A KO mice (Bortolozzi et al., 2004). Sleep/wake behavior in the 5-HT1A

KO mice is relatively normal with the exception of a significant increase in paradoxical sleep and no rebound paradoxical sleep following paradoxical sleep deprivation (Boutrel et al., 2002). However, 5-HT1A KO mice display an increased level of anxiety (Parks et

al., 1998), and mood and circadian rhythms are thought to interact (Wirz-Justice, 2006;

McClung, 2007). Stress and/or anxiety has been shown to affect phase shifts to non-

photic stimuli (Mistlberger et al., 2003) as well as re-entrainment to shifted LD cycles

(Mohawk and Lee, 2005). Therefore, it is possible that some of the observed effects in

the present study may be attributable to an increase in stress or anxiety, rather than the

absence of the 5HT1A receptor.

In conclusion, the present study supports a role for the 5-HT1A receptor in regulating behavioral phase shifts to light, as well as a role in mediating 8-OH-DPAT induced phase shifts of the circadian system. It is likely that non-photic phase shifts persist in these mice, but other models of non-photic entrainment should be examined in

these mice to further explore the role of the 5-HT1A receptor in this phenomenon. Overall, these findings contribute to our understanding of how serotonin regulates the circadian system. Given the tight interplay between circadian rhythms, sleep regulation and mood disorders (Wirz-Justice, 2006; McClung, 2007), care must be taken to ensure that treatments for disorders in one of these three systems does not produce detrimental effects on the other systems. Mice bearing selective KO’s of 5-HT receptors may complement pharmacological investigation in efforts to understand how serotonin regulates sleep, mood and circadian rhythmicity.

Acknowledgements

This work was supported by an NSERC discovery grant to MCA.

3. Potentiation of photic phase shifts by the serotonergic mixed agonist/antagonist

NAN-190: The role of the 5-HT1A receptor

Victoria M. Smith1 and Michael C. Antle1,2,3

Running Head: Phase Shift Potentiation and the 5-HT1A Receptor

Author Contributions:

Victoria Smith: - Laboratory Research: 5-HT1A receptor knockout mouse colony generation and maintenance, animal genotyping, predictions for manipulation times, injections, light pulse manipulations, phase shift calculations - Manuscript Preparation: Entered and organized data; ran statistical analyses, produced figures, composed manuscript Dr. Michael Antle: - Provided laboratory, equipment and financial support necessary to carry out experiments - Manuscript Preparation: Provided feedback and editing of the manuscript

Abstract

The mammalian circadian clock is located in the suprachiasmatic nucleus (SCN) in the

anterior hypothalamus, and generates daily oscillations in physiology and behaviour. The

circadian system is thought to be modulated by serotonin (5-HT), such that 5-HT is

presumed to inhibit photic phase shifts by dampening the excitatory signal sent to the

SCN following light exposure. Fully understanding the precise role played by 5-HT is

hampered by the variety of 5-HT receptors found in the SCN, and the lack of sufficiently

selective pharmacological agents for the 5-HT receptors of interest. The non-selective

drug NAN-190, is a serotonergic mixed agonist/antagonist which has been shown to

greatly potentiate photic phase shifts in hamsters. NAN-190 is presumed to bind to the 5-

HT1A receptor, but also has affinities for several other receptors, both serotonergic and

non-serotonergic. Genetically modified animals engineered to lack a specific 5-HT

receptor represent an alternative method by which to investigate the specific receptor

involvement in the induction of potentiated photic phase shifts. Here we examine the role

of the 5-HT1A receptor in behavioural responses to photic stimuli following pretreatment with NAN-190. When NAN-190 was administered to wildtype control mice, as well as mice heterozygous for the 5-HT1A receptor, significantly potentiated phase shifts to late

subjective night light pulses were observed. Conversely, mice lacking the 5-HT1A completely (knockouts) exhibited an attenuated behavioural response to NAN-190 pretreatment. NAN-190 administered without light exposure was, however, unable to induce any significant phase shifts in the activity rhythm of either wildtypes or knockouts. These findings suggest that the 5-HT1A receptor plays a critical role in the

potentiation effect observed with NAN-190, while one functional allele for the receptor appears to at least partially restore the ability of NAN-190 to potentiate the effects of light on the circadian system.

Key Words: Circadian, Suprachiasmatic Nucleus, 5-HT, Transgenic Mice

3.1 Introduction

Endogenously generated daily oscillations in physiology and behaviour are

evident throughout phylogeny, from prokaryotes to mammals (Sharma, 2003). Such

circadian oscillations in mammals are controlled by the circadian clock, located in the suprachiasmatic nucleus (SCN) of the anterior hypothalamus ( Moore and Eichler, 1972;

Stephan and Zucker, 1972), and are entrained to the daily environmental light/dark cycle

by photic input to the eye (Daan and Pittendrigh, 1976; Rusak and Zucker, 1979). Light

information is relayed from the intrinsically photosensitive retinal ganglion cells in the

retina, along the retinohypothalamic tract to the SCN, where it can alter the expression of

circadian rhythms (Berson, 2003; Hirota and Fukada, 2004). However, exposure at

specific times during the night can produce a shift in the onset of the activity rhythm,

such that the animal will become active a little earlier or later than on previous days,

depending on the exact timing of the light exposure (Gillette and Mitchell, 2002).

The SCN also receives dense serotonergic innervation from the median raphe

nucleus (Meyer-Bernstein and Morin, 1996). Serotonin (5-HT) is an inhibitory

neurotransmitter which has been shown to modulate the effects of light by dampening the

excitatory signals sent to the cells of the SCN as a result of light exposure (Meyer-

Bernstein and Morin, 1996; Mistlberger, 2000). Although there are believed to be 6

serotonin receptors present in the SCN (Smith et al., 2001), it is not fully understood the

precise role each one plays in serotonergic modulation of circadian rhythmicity. One

method for studying receptor contribution is through the use of drugs which either activate or block specific receptors. A class of compounds which are known to modulate

the effects of light on the circadian system is the set of drugs which act as serotonin mixed agonists/antagonists (Gannon, 2003).

These unique drugs are termed mixed agonists/antagonists due to their actions as antagonists at presynaptic axoaxonic heteroreceptors, as well as agonists at postsynaptic somatodendritic autoreceptors. When administered during the early or late subjective

night, these drugs are known to greatly potentiate photic phase shifts in the hamster, inducing shifts in the activity rhythm of up to 8h in magnitude (Rea et al., 1995; Gannon,

One specific example of this class of 5-HT mixed agonists/antagonists is the drug

1-(2-methoxyphenyl)4-[4-(phthalimido)butyl]piperazine hydrobromide (NAN-190).

Although NAN-190 is able to greatly potentiate photic phase shifts when administered in the early or late subjective night, and is effective when injected either before or after the exposure to light (Rea et al., 1995; Kessler et al., 2008), the receptor activation leading to this effect is still not well understood. Originally, NAN-190 was thought to be strictly a

5-HT1A antagonist, but more recently it has also been shown to have an affinity for

several other receptors as well, including 5-HT2, 5-HT7, dopamine D2, and α1-adrenergic

receptors (Claustre et al., 1991; Glass et al., 1995; Rea et al., 1995; Saifullah and

Tomioka, 2003). As a result of the lack of receptor specificity with NAN-190, more

investigation is necessary in order to determine exactly which receptors are involved in

the production of the potentiated photic phase shifts observed. One method by which to determine the location of action of non-selective drugs is through the use of mice genetically modified to lack a specific receptor subtype (Sprouse et al., 2005). Non- selective serotonin mixed agonists/antagonists are prime candidates for testing with transgenic mice missing the 5-HT1A receptor, allowing for the determination of the role of

the 5-HT1A receptor subtype in the photic phase shift potentiation observed with NAN-

190.

The present study sought to first determine if the potentiation effect previously

observed in hamsters was reproducible in mice. Subsequently, the contribution of 5-HT1A

receptor activation in the photic phase shift potentiation observed with the 5-HT1A mixed agonist/antagonist NAN-190 was examined. It is hypothesized that if activation of the 5-

HT1A receptor is the primary mechanism of action for NAN-190, then the potentiation of

photic phase shifts expected to be observed in the wildtypes will not be present in the

knockout mice. However, it is also predicted that heterozygous animals will demonstrate

a phase shift potentiation similar to that of the wildtypes, due to their possessing one

normal allele for the expression of at least some 5-HT1A receptors in the SCN. It is also

hypothesized that there will be a gradient of the magnitude of phase shifts induced by

different intensities of light in all genotypes, such that larger phase shifts will be observed

following bright light than dim light, and no phase shift will be observed when NAN-190

is administered alone.

3.2 Experimental Procedures

3.2.1 Animals and Housing

A total of 33 male mice were used, consisting of 13 5-HT1A receptor knockout

(KO) mice, 4 5-HT1A receptor heterozygous (Het) mice and 16 5-HT1A receptor wildtype

(WT) mice. The mice were generated from a breeding colony established in our lab from

mice originally developed by Dr. Thomas Shenk (Princeton University, Princeton, NJ),

provided by Dr. Miklos Toth (Cornell University Medical College, New York, NY) and

bred on the C57BL/6J background. For information regarding the generation of the 5-

HT1A receptor KO mice see Parks et al. (1998). Animals from our breeding colony were

genotyped using a pair of KO primers (amplifying a 400 base-pair product that included a

portion of the neomycin cassette that replaced the start codon of the 5-HT1A receptor

gene) as well as a pair of WT primers (amplifying a 238 base-pair product that included a

portion of the 5-HT1A receptor gene replaced by the neomycin cassette in the KO

animals) to identify the presence or absence of each allele of the 5-HT1A receptor gene

(KO up primer: CTT TAC GGT ATC GCC GCT CCC GAT TC; KO down primer: TGC

AGG ATG GAC GAA GTG CAG CAC A; WT up primer: AGT GCA GGC AGG CAT

GGA TAT GTT; WT down primer: CCG ATG AGA TAG TTG GCA ACA TTC TGA;

Figure 3.1).

Mice were individually housed in Nalgene Type L clear polycarbonate cages

(30.3 cm long × 20.6 cm wide × 26 cm high; Nalg Nunc International, Rochester, NY),

equipped with a stainless steel running wheel (diameter of 24.2 cm). Animals were

Figure 3.1. Example of an electrophoresis gel used to genotype the animals. The first

column indicates an animal with only wildtype alleles (wildtype mouse), the

second column indicates an animal with one wildtype allele and one knockout

allele (heterozygous mouse), and the third column indicates an animal with only

knockout alleles (knockout mouse).

housed in a temperature- (21 ± 1°C) and humidity-controlled room, with ad libitum access to food and water. Cages were changed at least every 14 days (7-10 days prior to, and following a manipulation). All protocols were approved by the Life and

Environmental Sciences Animal Care Committee at the University of Calgary, and adhered to the Canadian Council on Animal Care guidelines for the ethical use of animals.

3.2.2 Drugs and Reagents

The 5-HT1A receptor mixed agonist/antagonist NAN-190 was obtained from

Sigma-Aldrich Canada, Ltd. (Oakville, ON). NAN-190 was dissolved in 99% dimethyl sulfoxide (DMSO), also obtained from Sigma-Aldrich Canada, Ltd. Injections were administered intraperitoneally (i.p.), with the dose of NAN-190 being 2.5 mg/kg and vehicle control injections (99% DMSO) being a comparable volume based on weight.

The dose of NAN-190 was based upon the lowest dose which produced the largest effect in previous work performed by Rea et al. (1995). All injections were performed in constant darkness with the aid of night-vision goggles (BG15Alista, Richmond Hill, ON).

3.2.3 Activity Rhythms

Wheel-running activity was continuously monitored using magnetic switches mounted on the running wheel, and connected to a computer running the Clocklab data collection software package (Coulbourn Instruments, Allentown, PA, USA). Actograms

(graphical representation of wheel-running behavior) were generated and analyzed by

Clocklab analysis software. Mice were allowed to entrain to the light-dark (LD) cycle or free-run in constant darkness (DD) for a minimum of 7 days prior to the start of manipulations. Onset of running wheel activity for manipulation day was predicted using a regression line fit to the activity onsets for the 7-10 days prior to the testing day.

3.2.4 Phase Shift Manipulations

This experiment examined the effects of NAN-190 on a brief exposure to bright and dim light, as well as no light on behavioral phase shifts of the activity rhythm.

Animals were housed in DD (0 lux) for the duration of this experiment. The bright light pulse was administered by removing the animal’s cage from the housing room and leaving it on a counter in the main lab for the duration of the light pulse, such that the light intensity at cage level was ~1500 lux. The dim light pulse was administered using a light-sealed box with a light bulb partially covered in aluminum foil such that the light intensity at cage level was measured as 40 lux. A light meter was used to measure exact light intensities for all light pulses. Mice were exposed to a pretreatment of an i.p. injection of either NAN-190 or vehicle control, followed by a 15-minute light pulse at three different light intensities (1500 lux (KO: n = 3; Het: n = 3; WT: n = 4), 40 lux (KO: n = 12; Het: n = 4; WT: n = 11), or 0 lux (KO: n = 7; WT: n = 6)) in the late subjective night, 10 h after their predicted activity onset (i.e., circadian time (CT) 22, where activity onset is designated to be CT12 by convention). Animals received each injection treatment on separate occasions in a counterbalanced fashion. Following the manipulations, actograms were analyzed using the automatic fitting function included in the Clocklab

software, creating one line of best fit for the activity onsets of the 10 days prior to and including the manipulation day, and another line of best fit for the 10 days following the light pulse (excluding the first 3 onsets to avoid transient effects). The phase shift in the behavioral activity rhythm was calculated based on the horizontal difference between the predicted onsets from the two regression lines on the day following the manipulation.

3.2.5 Statistical Analysis

Each of the three light intensities was analyzed separately, using a mixed two-way analysis of variance (ANOVA), to determine if there was a significant difference in the magnitude of the photic phase shifts between the genotypes in each of the two drug conditions. For the bright light pulse (1500 lux) and dim light pulse conditions, all three genotypes were compared, whereas for the no light pulse condition, only WT and KO animals were compared. Statistical significance was set at P<0.05 for all tests, using

SigmaStat Version 2.03 (Systat Software, Inc.; San Jose, CA). All significant ANOVAs were followed up by Student-Newman-Keuls post hoc multiple comparisons. All means are reported as ± standard error of the mean in the figures.

3.3 Results

3.3.1 Modulation of Phase Shifts to a Bright Light Pulse

Representative actograms for each genotype receiving each of the treatments can be found in Figure 3.2A-F. Exposure to a bright intensity (1500 lux) light pulse at CT22

Figure 3.2. (A-F) Representative actograms of running wheel activity for (A-B) a

wildtype mouse, (C-D) a heterozygous mouse, and (E-F) a knockout mouse

receiving an i.p. injection of either (A,C,E) DMSO (control) or (B,D,F) NAN-190

(5mg/kg) at CT21, denoted by a diamond (), followed by a high intensity light

pulse (1500 lux) at CT22, denoted by a circle (z). Grey diagonal lines represent

regression lines fit to activity onsets both before and after exposure to a light

pulse.

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resulted in a significant interaction between the two drug treatments and the three genotypes (F(2,7)=18.194, P<0.01; Figure 3.3). The photic phase shift potentiation observed previously in hamsters was repeated in the wildtype mice, with phase shifts to

NAN-190 being significantly larger than those induced by DMSO pretreatment (P<.01).

However, the heterozygous animals did not demonstrate a similar potentiation with

NAN-190 compared to DMSO (P>0.05). The knockout mice also did not show a significant potentiation in the phase shift induced by NAN-190; in fact, they actually demonstrated a significant attenuation in the magnitude of phase shift induced by NAN-

190 compared to DMSO (P<0.01). There was a gradient in the magnitude of phase shifts elicited by DMSO pretreatment, such that KO animals demonstrated significantly larger phase shifts than either of the WT (P<0.01) or Het (P<0.05), however although Het animals did exhibit slightly smaller phase shifts than WT animals, this difference was not significant (P>0.05). There was also a gradient in the phase shifts observed with NAN-

190, with WT mice demonstrating phase shifts that were significantly larger than both the

Het (P<0.05) and KO (P<0.001) animals, but the phase shifts in Het animals were also significantly larger than those found in the KO animals (P<0.05).

3.3.2 Modulation of Phase Shifts to a Dim Light Pulse

Similar results were observed in the dim light pulse (40 lux) conditions as those discussed above. Representative actograms for each genotype in each drug group can be found in Figure 3.4A-F. As with the bright light exposure, exposure to a dim light pulse

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Figure 3.3. Mean phase shifts (± SEM) of wildtype, heterozygous and knockout animals

treated with either DMSO or NAN-190 at CT21, followed by a 1500 lux light

pulse. Bars not sharing a letter were significantly different, F(2,24)= 18.194,

P<0.01; Student-Newman-Keuls post hoc tests P<0.05.

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Figure 3.4. (A-F) Representative actograms of running wheel activity for (A-B) a

wildtype mouse, (C-D) a heterozygous mouse, and (E-F) a knockout mouse

receiving an i.p. injection of either (A,C,E) DMSO (control) or (B,D,F) NAN-190

(5mg/kg) at CT21, denoted by a diamond (), followed by a low intensity light

pulse (40 lux) at CT22, denoted by a circle (z). Grey diagonal lines represent

regression lines fit to activity onsets both before and after exposure to a light

pulse.

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at CT22 resulted in a significant interaction between the two drug groups and the three genotypes (F(2,24)=16.589, P<0.001; Figure 3.5). As with the bright light pulse condition, wildtype animals receiving a dim light pulse pretreated with NAN-190 demonstrated significantly larger magnitude phase shifts than following pretreatment with DMSO

(P<0.001). However, NAN-190 also elicited significantly larger phase shifts than did

DMSO in the heterozygous animals (P<0.05). Whereas, the knockout animals once again demonstrated significantly attenuated phase shifts to NAN-190 compared to DMSO

(P<0.05). A slightly different gradient of phase shifts emerged between the genotypes in the dim light pulse condition, than was seen in the bright light pulse condition, following

DMSO pretreatment. The KO animals again demonstrated the largest phase shifts to

DMSO, however the heterozygous animals had a magnitude of phase shift that was slightly less than the KOs but also slightly more than the WT animals. Although the phase shifts induced in the knockouts were significantly larger than those seen in the wildtypes (P<0.05), neither of these two genotypes were found to have phase shifts significantly different from those induced in the heterozygous animals (P>0.05 for both).

A slightly different pattern emerged following NAN-190 pretreatment, with both the wildtypes and heterozygotes exhibiting significantly larger phase shifts than knockouts

(P<0.001 and P<0.01, respectively), although there was no significant difference in the magnitude of phase shift elicited in the WT compared to Het animals (P>0.05).

3.3.3 Modulation of Phase Shifts to No Light Pulse

Representative actograms for the two genotypes in each drug condition, when

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Figure 3.5. Mean phase shifts (± SEM) of wildtype, heterozygous and knockout animals

treated with either DMSO or NAN-190 at CT21, followed by a 40 lux light pulse.

Bars not sharing a letter were significantly different, F(2,24)= 16.589, P<0.001;

Student-Newman-Keuls post hoc tests P<0.05.

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administered without any subsequent light pulse, can be found in Figure 3.6A-D. When either drug was injected alone at CT21, without any light exposure at CT22, no significant differences in phase shifts were found between the two genotypes

(F(1,11)=0.278, P>0.05) or between the two drug injections (F(1,11)=0.607, P>0.05). In fact, essentially no phase shifts were elicited at all by either NAN-190 or DMSO, in either wildtype or knockout animals (Figure 3.7).

3.4 Discussion

This was the first study to examine circadian responses to the 5-HT1A mixed agonist/antagonist NAN-190 in mice, and is therefore also the first study examining the effects of NAN-190 on the circadian system in mice lacking the 5-HT1A receptor. The first objective of this experiment was to reproduce in mice the phase shift potentiation effects of NAN-190 that have been well documented in hamsters (Rea et al., 1995). In wildtype mice, NAN-190 pretreatment did indeed result in a significant potentiation of late subjective night photic phase shifts. This effect was in part modulated by the intensity of the light pulse, with slightly greater phase shifts in the vehicle control condition as well as greater potentiation of the phase shifts with NAN-190, however it appears that the circadian system is only activated slightly more with the extremely high intensity of light in the bright light pulse condition compared to the dim light pulse condition. Overall, we have found that NAN-190 does have a similar ability to greatly potentiate photic phase shifts by up to 300% in mice, at least in the late subjective night,

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Figure 3.6. (A-D) Representative actograms of running wheel activity for (A-B) a

wildtype mouse, and (C-D) a knockout mouse receiving an i.p. injection of either

(A,C) DMSO (control) or (B,D) NAN-190 (5mg/kg) at CT21, denoted by a

diamond (), without a subsequent light pulse. Grey diagonal lines represent

regression lines fit to activity onsets both before and after exposure to a light

pulse.

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Figure 3.7. Mean phase shifts (± SEM) of wildtype and knockout animals treated with

either DMSO or NAN-190 at CT21, without a subsequent light pulse. All

comparisons were found to be non-significant.

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and a near maximal response can be observed with exposure to as little as 40 lux of light.

Plus, it was shown that this potentiation is not the result of some phase-shifting property

of the drug alone, but is rather a definite ability of NAN-190 to modulate the excitatory

effects of light information entering the SCN.

However, the complete elimination of the 5-HT1A receptor in the knockout

animals resulted in not only a loss of this potentiated response, as was predicted, but

actually led to a significant attenuation of the response. Had NAN-190 been solely

binding to the 5-HT1A receptor, pretreatment with NAN-190 in the knockout mice would have led to phase shifts no different in magnitude than when these animals were pretreated with an injection of the vehicle control. Therefore, NAN-190 is definitely also binding to some other receptor involved in the circadian system, which results in an attenuation of the phase shifting effects of light.

It is apparent though, that the potentiation effect observed with NAN-190 pretreatment is at least partially through the 5-HT1A receptor, due to the essentially

normal response observed in the heterozygous animals. Although these animals possess

only one normal allele for the 5-HT1A receptor, and are thus expected to exhibit a somewhat decreased level of expression of the receptor, they still exhibited a significant potentiation of the phase shift induced by a dim light pulse, which was no different than those induced in the mice with two normal alleles for the receptor. Despite the wildtype and heterozygous animals exhibiting magnitudes of phase shifts with NAN-190 that were significantly different from each other only when the circadian system was saturated with a bright intensity light pulse, but not with a dim intensity light pulse, this difference was

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less than half an hour in magnitude and the heterozygous animals still demonstrated a

potentiation of close to 200%. Thus, it does appear that the majority of the phase shift

potentiation observed with NAN-190 is indeed the result of binding to the 5-HT1A

receptor, but with some sort of mediating influence possibly resulting from binding to

some other receptor type as well.

One of the obvious receptor candidates for which NAN-190 has a high affinity

through which it could also be acting, and which is also found in the circadian system, is

the 5-HT7 receptor (Smith et al., 2001). It has previously been suggested that NAN-190 may function partially by also blocking the presumed activation of 5-HT7 receptors

(Lovenberg et al., 1993). However, it is unknown if the photic phase shift attenuation effect of NAN-190 unmasked in the 5-HT1A receptor knockout mice is due to the drug

also binding to the 5-HT7 receptor, or whether it is the result of binding to some other receptor. In order to more fully understand the specific roles other receptor subtypes may be playing in the effects on the circadian system following administration of NAN-190, it will be necessary to test the effects of NAN-190 in mice of other receptor knockout models, particularly the other receptors found in the SCN for which NAN-190 has a high binding affinity.

Another possible way to further examine the phenomenon of phase shift potentiation by serotonergic chronobiotics would be through the study of other drugs shown to have similar potentiation abilities to NAN-190. Compounds such as BMY

7378, S 15535, MDL 73005, and WAY-100635 have also been shown to potentiate photic phase shifts in the hamster circadian activity rhythm (Smart and Biello, 2001;

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Gannon, 2003). The precise serotonergic receptor binding profiles of these drugs is also not fully understood, although some of them are thought to be more specific in their binding affinities compared to NAN-190. Thus, the use of mice lacking the 5-HT1A receptor, as well as those lacking other serotonin receptors, could be extremely useful in further elucidating the serotonergic modulation of circadian rhythmicity inherent to this type of photic phase shift potentiation.

By understanding the precise pharmacology responsible for the modulation of the circadian system’s sensitivity to light stimuli which is observed with the class of drugs termed mixed agonists/antagonists, including NAN-190, it may be possible to eventually develop chronobiotics specifically targeted for the rapid resynchronization of circadian rhythms. Effectively managing or even preventing the chronic desynchronization between the circadian system and the external environment, such as that observed with shift work, jet lag, or circadian rhythm disorders, could play a large role in the elimination of the adverse health effects associated with this sort of desynchrony. Thus, it is of great importance in today’s 24-h society to more fully understand the pharmacology behind drugs such as NAN-190.

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4. Molecular underpinnings of photic phase shift potentiation with NAN-190

Victoria M. Smith1, Kimberly Hagel1 and Michael C. Antle1,2,3

Running Head: Molecular Underpinnings of Phase Shift Potentiation

Author Contributions:

Victoria Smith: - Laboratory Research: 5-HT1A receptor knockout mouse colony generation and maintenance, animal genotyping, predictions for manipulation times, light pulse manipulations, injections, perfusions, tissue collection, immunocytochemistry, in situ hybridization, tissue image capturing, cell counting and ROD analysis - Manuscript Preparation: Entered and organized data; ran statistical analyses, produced figures, composed manuscript Kimberly Hagel: - Experiment 1 and 2: Assisted with Fos (DAB and fluorescent triple-label) immunocytochemistry, tissue collection, tissue image capturing and cell counting (data included in her Honours Thesis) - Experiment 4: Provided technical assistance with immunocytochemistry for some animals in the 15- and 30-minute groups. Dr. Michael Antle: - Trained me on immunocytochemistry and in situ hybridization techniques - Provided laboratory, equipment and financial support necessary to carry out experiments - Manuscript Preparation: provided feedback and editing of the manuscript

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Abstract

Daily oscillations in physiology and behaviour are generated by the mammalian circadian pacemaker located in the suprachiasmatic nucleus (SCN). Serotonin (5-HT) is thought to modulate photic phase shifts by dampening the excitatory signals released from retinal terminals, along with inhibiting the responsiveness of the retinorecipient cells in the SCN.

Understanding the mechanism by which 5-HT accomplishes this inhibition is hampered by the wide variety of 5-HT receptors found in the SCN, as well as the lack of truly selective pharmacological agents for the 5-HT receptors of interest. The non-selective serotonergic mixed agonist/antagonist NAN-190 has been shown to greatly potentiate photic phase shifts in both mice and hamsters; however, the precise molecular mechanism by which this potentiation is accomplished is still largely unknown. Here we examine the molecular underpinnings, in terms of both gene expression and biochemical activation, of the behavioural phase shift potentiation observed with NAN-190. The protein product of the immediate-early gene c-fos induced 90 minutes following photic stimulation was found to be significantly attenuated following pretreatment with NAN-

190. Similarly, the levels of phosphorylated cyclic adenosine monophosphate response element-binding protein (P-CREB), which is involved in one of the biochemical pathways leading to gene transcription, were also attenuated by NAN-190 administration for up to 45 minutes following the light pulse. The majority of this modulated protein expression was observed in the retinorecipient cells of the core region of the SCN.

However, no effects were observed on the expression of the second-messenger signaling pathway component phosphorylated extracellular signal-regulated kinas (P-ERK), or on

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the expression of the circadian clock components Period1 and Period2 in the time course examined. These findings suggest that at least part of the mechanism by which NAN-190 potentiates photic phase shifts is through its modulation of the expression of certain genes induced in the retinorecipient core of the SCN, which are involved in coupling the light signal entering the SCN with the output of the circadian system. Yet, precisely how the expression of the molecular clock components, or ‘gears’ of the circadian clock, are being modulated in order to result in such potentiated phase shifts is still somewhat unclear.

Key Words: Circadian, Suprachiasmatic Nucleus, 5-HT, Fos, Period Genes, P-ERK,

P-CREB

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4.1 Introduction

Daily oscillations in physiology and behaviour are evident throughout phylogeny,

from prokaryotes to mammals (Sharma, 2003). In mammals, such endogenously generated circadian oscillations are controlled by the circadian clock located in the

suprachiasmatic nucleus (SCN) of the anterior hypothalamus (Moore and Eichler, 1972;

Stephan and Zucker, 1972). The circadian system is entrained to the daily environmental

light/dark cycle by photic input to the eye (Daan and Pittendrigh, 1976; Rusak and

Zucker, 1979), which is relayed from the intrinsically photosensitive retinal ganglion

cells in the retina, along the retinohypothalamic tract (RHT) to the retinorecipient cells of

the SCN (Berson, 2003; Hirota and Fukada, 2004). This signal is then able to alter the

expression of circadian rhythms in the endogenously rhythmic region of the SCN

(Berson, 2003; Hirota and Fukada, 2004). Light exposure at specific times during the

night can induce a shift in the onset of the circadian activity rhythm (Daan and

Pitendrigh, 1976). Depending on the precise timing of light exposure, the animal will

either become active a little earlier or a little later than on previous days (Gillette and

Mitchell, 2002).

Serotonin (5-HT) is an inhibitory neurotransmitter which has been shown to

modulate the effects of light by dampening the excitatory signals sent to the cells of the

SCN as a result of light exposure (Meyer-Bernstein and Morin, 1996; Mistlberger, 2000).

Although there are believed to be 6 serotonin receptors present in the SCN (Smith et al.,

2001), it is not fully understood the precise role each one plays in serotonergic

modulation of circadian rhythmicity. One method for studying receptor contribution is

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through the use of drugs which either activate or block specific receptors. A class of

compounds which are known to modulate the effects of light on the circadian system is

the set of drugs which act as serotonin mixed agonists/antagonists (Gannon, 2003).

night, these drugs are known to greatly potentiate photic phase shifts in the hamster, inducing shifts in the activity rhythm of up to 8h in magnitude (Rea et al., 1995; Gannon,

2003). The potentiation effects produced by this class of drug is presumed to be the result of the combination of an agonistic action at the median raphe to prevent serotonin release onto the SCN, along with an antagonistic action at the RHT terminals or SCN cells to block serotonergic inhibition at these locations. This decreased inhibition to the SCN is suspected to lead to increased excitation following photic input, thus allowing an overall potentiated phase shifting effect. One specific example of this class of 5-HT mixed agonists/antagonists is the drug 1-(2-methoxyphenyl)4-[4-(phthalimido)butyl]piperazine hydrobromide (NAN-190). Although NAN-190 is able to greatly potentiate photic phase shifts when administered in the early or late subjective night, and is effective when injected either before or after light exposure (Rea et al., 1995; Kessler et al., 2008), the changes to molecular components of the functioning of the circadian clock remain to be elucidated.

Along with inducing behavioural phase shifts, light exposure in the subjective night also results in the immediate expression of numerous genes in the cells located in

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the core of the SCN, including immediate-early genes (IEGs) such as c-fos, along with

the clock genes Per1 and Per2 (Guido et al., 1999a; Guido et al., 1999b; Hamada et al.,

2004). Although it is thought that the light-induced expression of IEGs could be an early

event in the response cascade which ultimately leads to photic phase alterations of the circadian pacemaker (Rea, 1992), the induction of the IEG c-fos is not sufficient, or even required to induce phase shifts in the circadian activity rhythm (Honrado et al., 1996).

Second-messenger signaling pathways, including the pathway leading to the phosphorylation of the extracellular signal-regulated kinase (ERK), serve to couple this incoming photic input to the transcription of IEGs and ultimately Period genes (Butcher et al., 2003). Similarly, the phosphorylated form of cAMP-response-element-binding protein (CREB), which is also induced following light exposure, subsequently binds to the cAMP response element (CRE) of the promoter region of several genes, including c- fos, Per1, and Per2, inducing transcription (Antle and Silver, 2005). These cascades are activated extremely rapidly by light, before an increase in IEG mRNA is even observed

(Dziema et al., 2003). The phase shifting effects of light are consistently accompanied by the colocalized expression of the phosphorylated forms of ERK (P-ERK) and CREB (P-

CREB; von Gall et al., 1998; Dziema et al., 2003).

As with shifts in the behavioural expression of circadian rhythms, the serotonergic system has also been shown to modulate the effects of photic input on the light-induced gene expression in the cells of the SCN. Both agonists and antagonists to the serotonin system which are known to alter circadian activity rhythms, have also been shown to modulate the light-induced gene expression in the SCN (Glass et al., 1994; Amir et al.,

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1998; Takahashi et al., 2002). Serotonergic agonists which attenuate light-induced phase shifts can also attenuate the subsequent Fos protein expression observed in the SCN

(Amir et al., 1998). Conversely, the selective 5-HT1A receptor agonist MKC-242, which

has the ability to potentiate photic phase advances and delays, has been found to

concomitantly prolong the light-induced expression of mPer1 and mPer2 in the mouse

SCN during the late subjective night (Moriya et al., 1998; Takahashi et al., 2002). The

serotonergic mixed agonist/antagonist NAN-190 which has also been shown to potentiate

photic phase shifts, leads to the partial blocking of the Fos expression normally induced

by a light pulse (Recio et al., 1996). However, this study did not examine the spatial

aspects of this reduced Fos expression within the SCN.

The molecular underpinnings of the photic phase shift potentiation observed with

serotonergic mixed agonists/antagonists such as NAN-190, both in terms of gene

expression and biochemical pathways leading to that gene expression, are still largely

unknown. It was originally proposed that mixed agonists/antagonists induce their

potentiation of light by blocking the inhibitory effects of serotonin at the retinal terminals

(Gannon, 2003). More recently though, it has been shown that NAN-190 affects both the

retinorecipient cells responsive to the glutamate agonist NMDA, as well as those cells in

the cap region which are responsive to gastrin-releasing peptide (GRP; Sterniczuk et al.,

2008). However, the effects of NAN-190 on the molecular components of the

retinorecipient cells of the SCN corresponding to potentiated photic phase shifts,

especially in terms of modulation of the clock genes which drive circadian rhythmicity,

remains to be fully elucidated.

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Thus the present study sought to determine the molecular underpinnings of the

photic phase shift potentiation observed with NAN-190 in mice, by examining the

regionally-specific expression of several different proteins and genes involved in the

transduction of the light signal into the phase shifts in the behavioural output of the

circadian clock. The proteins and genes examined include: Fos, the protein product of the

immediate-early gene c-fos; the clock genes mPer1 and mPer2; and the phosphorylated forms of the proteins ERK and CREB, which are involved in the biochemical cascades that regulate changes in clock gene expression. In general, it is hypothesized that all genes and proteins will be expressed to a greater degree in the ventrolateral core region of

the SCN compared to the dorsomedial shell. However, more specifically, it is

hypothesized that the number of cells expressing Fos protein following NAN-190

pretreatment will be decreased compared to a light pulse alone (Recio et al., 1996).

Conversely, it is hypothesized that pretreatment with NAN-190 will augment the

expression of mPer1 and mPer2, as well as p-ERK and p-CREB, such that there will

either be an increased number of cells responding or a prolonged activation of the cells.

4.2 Experimental Procedures

4.2.1 Animals and Housing

A total of 83 male mice were used, consisting of 74 C57BL/6J mice obtained

from the University of Calgary Biological Sciences breeding colony (Calgary, AB,

Canada), as well as 9 5-HT1A receptor wildtype (WT) mice. The 5-HT1A wildtype mice

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were generated from a breeding colony established in our lab from mice originally

developed by Dr. Thomas Shenk (Princeton University, Princeton, NJ), provided by Dr.

Miklos Toth (Cornell University Medical College, New York, NY) and bred on the

C57BL/6J background. For information regarding the generation of the 5-HT1A receptor

KO mouse colony see Parks et al. (1998). Animals from our breeding colony were

genotyped using a pair of KO primers (amplifying a 400 base-pair product that included a

portion of the neomycin cassette that replaced the start codon of the 5-HT1A receptor

gene) as well as a pair of WT primers (amplifying a 238 base-pair product that included a

portion of the 5-HT1A receptor gene replaced by the neomycin cassette in the KO

animals) to identify the presence or absence of each allele of the 5-HT1A receptor gene

(KO up primer: CTT TAC GGT ATC GCC GCT CCC GAT TC; KO down primer: TGC

AGG ATG GAC GAA GTG CAG CAC A; WT up primer: AGT GCA GGC AGG CAT

GGA TAT GTT; WT down primer: CCG ATG AGA TAG TTG GCA ACA TTC TGA).

Mice were individually housed in Nalgene Type L clear polycarbonate cages

(30.3 cm long × 20.6 cm wide × 26 cm high; Nalg Nunc International, Rochester, NY),

equipped with a stainless steel running wheel (diameter of 24.2 cm). Animals were

housed in a temperature- (21 ± 1°C) and humidity-controlled room, with ad libitum

access to food and water. Cages were changed at least every 14 days (7-10 days prior to,

and following a manipulation). All protocols were approved by the Life and

Environmental Sciences Animal Care Committee at the University of Calgary, and

adhered to the Canadian Council on Animal Care guidelines for the ethical use of

animals.

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4.2.2 Drugs and Reagents

The 5-HT1A receptor mixed agonist/antagonist NAN-190 was obtained from

Sigma-Aldrich Canada, Ltd. (Oakville, ON). NAN-190 was dissolved in 99% dimethyl

sulfoxide (DMSO), also obtained from Sigma-Aldrich Canada, Ltd. Injections were

administered intraperitoneally (i.p.), with the dose of NAN-190 being 2.5 mg/kg and

vehicle control injections (99% DMSO) being a comparable volume based on weight.

The dose of NAN-190 was based upon the lowest dose which produced the largest effect

in previous work performed by Rea et al. (1995). All injections were performed in

constant darkness with the aid of night-vision goggles (BG15Alista, Richmond Hill, ON).

4.2.3 Activity Rhythms

Wheel-running activity was continuously monitored using magnetic switches

mounted on the running wheel, and connected to a computer running the Clocklab data

collection software package (Coulbourn Instruments, Allentown, PA, USA). Actograms

(graphical representation of wheel-running behavior) were generated and analyzed by

Clocklab analysis software. Mice were allowed to entrain to the light-dark (LD) cycle or

free-run in constant darkness (DD) for a minimum of 7 days prior to the day of

manipulations. Onset of running wheel activity for manipulation day was based on the time of lights-on or predicted using a regression line fit to the activity onsets for the 7-10 days prior to the testing day.

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4.2.4 Protein and Gene Expression Underlying Photic Phase Shifts

This experiment examined the effects of short duration, dim light exposure on the expression of Fos protein (experiments 1 and 2), mPer1 and mPer2 gene expression

(experiment 3), and the expression of the phosphorylated forms of the ERK and CREB proteins (experiment 4) in the SCN. Mice (n = 30) were housed in LD (experiment 1) or

DD (experiments 2-4) for the duration of this experiment. Mice were exposed to a dim light pulse using a light-sealed box with a light bulb partially covered in aluminum foil such that the light intensity at cage level was measured as 40 lux. Mice were randomly assigned to a treatment group (NAN-190 or vehicle control), and only for experiment 4 they were also randomly assigned to a time group (15 min, 30 min, or 45 min following the start of the light pulse), being exposed to a pretreatment of an i.p. injection in the late subjective night, 9 h after predicted activity onset, followed by a 15-minute light pulse 10 h after predicted activity onset (i.e., circadian time (CT) 21 and 22, respectively, where activity onset is designated to be CT12 by convention).

For experiments 1-3, animals were euthanized, with an intraperitoneal (i.p.) injection of Euthanyl (0.3mL), 90 min after the start of the light pulse (i.e., at CT23.5).

Whereas, for experiment 4, animals were euthanized at one of three different durations following the start of the light pulse, 15 minutes, 30 minutes, or 45 minutes. The head was wrapped in aluminum foil to prevent the retinas from being exposed to light prior to and during the perfusion. Animals were then perfused using approximately 50 mL of phosphate buffered saline (PBS), followed by 50 mL of buffered 4% paraformaldehyde

(experiments 1-3), or perfused using only 50 mL of buffered 4% paraformaldehyde

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(experiment 4). Brains were extracted and postfixed in 4% paraformaldehyde overnight, and subsequently cryoprotected in a 20% sucrose solution for 24 h. Brains were sliced through the anterior hypothalamus using a cryostat, with alternate 35 μm sections through the SCN being collected for immunocytochemistry (ICC) or in situ hybridization.

4.2.5 Immunocytochemistry

4.2.5.1 Diaminobenzidine (DAB)

Diaminobenzidine ICC was completed using a previously described method from our lab (see Antle et al., submitted). Brain sections were exposed to a series of treatments at room temperature on a shaker tray unless otherwise indicated. Briefly, sections were rinsed for 30min in 0.5% H2O2 in PBS with 0.3% Triton X-1900 (PBSx) to inactivate endogenous peroxidase, followed by 3-10min PBSx rinses, a 90min incubation in 1% normal horse serum in PBSx (Fos) or 1% normal goat serum in PBSx (P-ERK and P-

CREB), and a 2-day incubation at 4°C in the primary antibody for Fos (goat anti-Fos;

1:20,000; Santa Cruz Biotechnology, Santa Cruz, CA), P-ERK (rabbit anti-Phospho- p44/42 MAP Kinase (Thr202/Tyr204); 1:600; Cell Signaling Technology, Inc., Danvers,

MA), or P-CREB (rabbit anti-Phospho CREB; 1:250; Cell Signaling Technology, Inc.,

Danvers, MA). Subsequently, sections were exposed to 3-10min PBSx rinses, a 1h incubation in the secondary antibody biotinylated horse anti-goat (for Fos; 1:200; Vector

Laboratories, Burlingame, CA) or biotinylated goat anti-rabbit (for P-ERK and P-CREB;

1:200; Vector Laboratories, Burlingame, CA), 3-10min PBSx rinses, a 1h incubation in

ABC (Vectastain Elite kit, Vector Laboratories, Burlingame, CA), 3-10min PBSx rinses.

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Slices were then developed for approximately 5min in 0.05% DAB and 0.02% NiCl in

25mL of 0.1M Tris buffer activated with 80μL of 30% H2O2. Finally, slices were

mounted on gelatin-coated slides, air-dried, dehydrated through an alcohol series, cleared

with xylenes, and coverslipped with Permount.

4.2.5.2 Fluorescent Triple-Label

Sections were triple-labeled for Fos (goat anti-Fos; 1:20,000; Santa Cruz

Biotechnology, Santa Cruz, CA), vasopressin (VP; guinea-pig anti-VP; 1:5,000;

Peninsula Laboratories, San Carlos, CA), and bombesin/gastrin-releasing peptide (GRP;

rabbit anti-GRP; 1:5,000; ImmunoStar Inc., Hudson, WI). ICC staining was completed

using a previously described method from our lab (see Sterniczuk et al., 2008). Briefly,

brain sections were exposed to three 10-min washes in 0.1M PBS, followed by a 1 h

incubation in a blocking buffer (2% normal donkey serum, in PBSx), after which the

tissue was incubated for 48 h at 4°C in the primary antibodies diluted in blocking buffer.

Tissue was then washed using six 10-min rinses in 0.1% PBSx, and left overnight in

0.1% PBSx. Finally, the brain sections were washed with another six 10-min 0.1% PBSx

rinses, incubated for 1 h in the following secondary antibodies: CY-2 donkey anti-rabbit,

CY-3 donkey anti-goat, and CY-5 donkey anti-guinea pig (each at 1:200; Jackson

ImmunoResearch Laboratories Inc., West Grove, PA), and exposed to three final 10-min washes in 0.1M PBS. Sections were then mounted on gelatin-coated slides, air dried, dehydrated in a series of alcohol rinses, cleared with xylenes, and coverslipped with

Krystalon.

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4.2.6 In Situ Hybridization

For mPer1and mPer2 in situ hybridization, all instruments and trays were cleaned with RNase Zap (Ambion, Austin, TX) and all solutions were prepared with RNase-free

water. Free floating tissue sections were processed using the probe and digoxigenin

(DIG) protocol of Yan and Silver (2002). First, tissue was treated with proteinase K (1 mg/mL, 0.1M Tris buffer pH 8.0; 50 mM EDTA; 10 min) at 37°C. This reaction was stopped by adding 2 mL 4% paraformaldehyde. Following rinses in saline sodium citrate

(300mM NaCl, 30mM sodium Citrate) the tissue was treated with 0.25% acetic anhydride in 0.1M triethanolamine for 10 min. The sections were then incubated in 1.5 mL hybridization buffer [50% formamide, 60mM sodium citrate, 600mM NaCl, 10% dextran sulphate, 1% N-laurylsarcosine, 25 mg/mL tRNA, 1X Denhardt's, 0.25mg/mL salmon sperm DNA] containing the DIG-labeled mPer1 or mPer2 (mPer1 and mPer2 plasmids courtesy of Dr. L. Yan, Michigan State University, East Lansing, MI) antisense cRNA probes (0.1 µg/mL; for details see Hamada et al., 2001) for 16 h at 60°C.

After two 30 minute high-stringency posthybridization washes (50% formamide/saline sodium citrate at 60°C), sections were treated with RNase A. After more high-stringency washes, the tissue was then processed for immunodetection with a nucleic acid detection kit (Roche, Indianapolis, IN). The sections were incubated in 1.0% of blocking reagent in buffer 1 (100 mM Tris-HCl buffer, 150 mM NaCl, pH 7.5) for 1 h at room temperature. Sections were then incubated at 4°C in alkaline phosphatase- conjugated Dig antibodies diluted 1:5000 in buffer 1 for 3 days. On the following day, sections were washed in buffer 1 twice (15min each), and incubated in buffer 3 (100 mM

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Tris-HCl buffer, pH 9.5, containing 100 mM NaCl and 50 mM MgCl2) for 5 min. They were then incubated in a solution containing nitroblue tetrazolium salt (0.34 mg/mL) and

5-bromo-4-chrolo-3-indolyl phosphate toluidinium salt (0.18 mg/mL; Roche) for 16 h.

The colorimetric reaction was stopped by immersing the sections in buffer 4 (10 mM

Tris-HCl containing 1 mM EDTA, pH 8.0). The tissue was then mounted on gelatin coated slides, air-dried, briefly dehydrated in an alcohol series, cleared with xylenes, and coverslipped with Permount.

4.2.7 Quantification of Protein and Gene Expression

Images of the immunofluorescently-labeled tissue were captured via an Olympus

BX51 microscope equipped with filters for each of the three cyanine dyes. Bilateral counts of Fos-immunoreactive (Fos-IR) cells were performed using computerized image analysis software (ImagePro Plus 5.1.2.59; Media Cybernetics, Inc., Bethesda, MD), and were manually counted by two researchers who were blind to the treatment conditions.

Cell counting was completed at the mid-caudal level of the SCN, identified by the presence of the GRP subnucleus. Separate cell counts were obtained for two regions of the SCN: shell (delineated by VP-IR) and core (delineated by GRP-IR). Images of the diaminobenzadine-labeled tissue were captured using the bright field setting, whereas images of the digoxigenin-labeled tissue were captured using the RGB color filter for the camera, of the same microscope. Bilateral counts of Fos- and P-CREB-labeled cells were performed using the same techniques as used above, whereas bilateral calculations of the relative optical density (ROD) of mPer1, mPer2, and P-ERK expression were performed

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using a different computerized image analysis software (ImageJ 1.34s; National Institutes

of Health, Bethesda, MD). Cell counts and ROD were calculated at the mid-caudal level of the SCN, with SCN regions (core, shell) being delineated based on the general shape of each region identified previously while performing the fluorescent triple-label Fos cell counts.

4.2.8 Statistical Analysis

The expression of Fos (experiments 1 and 2), mPer1 and mPer2 (experiment 3) were each analyzed using a mixed two-way analysis of variance (ANOVA), to determine if there was a significant difference in the expression induced in the core and shell regions of the SCN by a light pulse pretreated with each of the two drug conditions. The expression of P-ERK and P-CREB (experiment 4) were each analyzed using two two- way mixed ANOVAs, to determine if there was a significant difference in the expression induced at each of the three times following the start of the light pulse resulting from pretreatment with each of the two drug conditions, separately in the core and shell regions of the SCN. Statistical significance was set at P<0.05 for all tests, using

SigmaStat Version 2.03 (Systat Software, Inc.; San Jose, CA). All significant ANOVAs were followed up by a priori post hoc multiple comparisons. All means are reported as ± standard error of the mean in the figures.

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4.3 Results

4.3.1 Experiment 1: DAB-Labeled Fos Expression

Representative examples of the Fos expression for each drug treatment condition

are presented in Figure 4.1. A brief light pulse in the late subjective night (CT22) resulted

in an interaction between the two injection types and the two regions within the SCN

which approached significance using the DAB method of ICC staining (F(1,7)=3.604,

P=0.099; Figure 4.2). The majority of the Fos expression was located within the core region of the SCN in all animals. However, the animals pretreated with NAN-190

demonstrated a significant attenuation in the amount of Fos which was expressed in the

core when compared to animals pretreated with the vehicle control (P<0.01), while there

was no difference in the shell expression between the two treatment groups.

4.3.2 Experiment 2: Fluorescent Triple-Labeled Fos Expression

Representative examples of the Fos expression for each drug treatment condition

are presented in Figure 4.3. A brief light pulse in the late subjective night (CT22)

resulted in a significant interaction between the two drug pretreatments and the two

regions within the SCN using the fluorescence triple-label method of ICC staining

(F(1,10)=16.624, P<0.01; Figure 4.4). The majority of the Fos expression was again

located within the core region of the SCN in all animals. As with experiment 1, the

animals pretreated with NAN-190 demonstrated a significant attenuation in the number

of Fos-immunoreactive (Fos-IR) cells expressed in the core when compared to animals

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Figure 4.1. Representative SCN tissue slices indicating Fos-expressing cells at CT23.5

and stained using DAB immunocytochemistry following a brief light pulse in the

late subjective night (CT22), pretreated at CT21 with an i.p. injection of either (A)

vehicle control (DMSO) or (B) NAN-190. Scale bar = 200μm.

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Figure 4.2. Mean number of Fos-expressing cells (± SEM) in the core and shell regions

of the SCN of animals following a brief light pulse in the late subjective night

pretreated with either vehicle control (DMSO) or NAN-190. (**) indicates a

statistical significance of P<0.01.

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Figure 4.3. Representative examples of SCN tissue slices indicating Fos-IR cells (in

green) at CT23.5 and stained using fluorescent triple-label immunocytochemistry

following a brief light pulse in the late subjective night (CT22), pretreated at

CT21 with an i.p. injection of either (A) vehicle control (DMSO) or (B) NAN-

190. GRP-IR (red) and VP-IR (blue) were used to delineate the core and shell

regions of the SCN for the purpose of cell counting. Scale bar = 200μm.

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Figure 4.4. Mean number of Fos-IR cells (± SEM) in the core and shell regions of the

SCN of animals following a brief light pulse in the late subjective night pretreated

with either vehicle control (DMSO) or NAN-190. (***) indicates a statistical

significance of P<0.001.

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pretreated with the vehicle control (P<0.001), while there was a trend towards an attenuation in the shell expression between the two treatment groups as well (P=0.057).

4.3.3 Experiment 3: mPer1 and mPer2 Gene Expression

Representative examples of the mPer1 and mPer2 expression for each drug treatment condition are presented in Figure 4.5A-D. In contrast to the results of the Fos expression, no difference was observed in the amount of either mPer1 or mPer2 gene

expression in NAN-190 pretreated animals relative to DMSO pretreated mice

(F(1,11)=0.357, P>0.05 and F(1,11)=0.0214, P>0.05, respectively; Figure 4.6). Though, the

majority of gene expression was again observed in the core region of the SCN in all animals.

4.3.4 Experiment 4: Biochemical Pathway Components

4.3.4.1 P-ERK

Representative examples of the P-ERK protein expression for each drug treatment condition at each of the three times following the start of the light pulse in the late subjective night are presented in Figure 4.7. As with the results of the Period gene expression, no difference was found in the amount of P-ERK expression in either the core or the shell regions of the SCN in NAN-190 pretreated animals relative to DMSO pretreated mice (F(2,32)=0.171, P>0.05 and F(2,32)=0.547, P>0.05, respectively; Figure

4.8). There was also no discrete time course in P-ERK expression in either the core or the

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Figure 4.5. Representative examples of SCN tissue slices indicating (A,B) mPer1 and

(C,D) mPer2 cells at CT23.5 and stained using in situ hybridization following a

brief light pulse in the late subjective night (CT22), pretreated at CT21 with an

i.p. injection of either (A,C) vehicle control (DMSO) or (B, D) NAN-190. Scale

bar = 200μm.

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Figure 4.6. Mean relative optical density (ROD; ± SEM) of (A) mPer1 and (B) mPer2 in

the core and shell regions of the SCN of animals following a brief light pulse in

the late subjective night pretreated with either vehicle control (DMSO) or

NAN-190.

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Figure 4.7. Representative examples of SCN tissue slices indicating cells expressing

P-ERK at (A,B) CT22.25, (C,D) CT22.5, and (E,F) CT22.75 stained using DAB

immunocytochemistry following a brief light pulse in the late subjective night

(CT22), pretreated at CT21 with an i.p. injection of either (A,C,E) vehicle control

(DMSO) or (B, D,F) NAN-190. Scale bar = 200μm.

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Figure 4.8. Mean relative optical density (ROD; ± SEM) of P-ERK in the (A) core and

(B) shell regions of the SCN of animals euthanized 15, 30, or 45 minutes

following a brief light pulse in the late subjective night and pretreated with either

vehicle control (DMSO) or NAN-190.

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shell regions of the SCN across the three time points tested following the start of the light

pulse (F(2,32)=0.840, P>0.05 and F(2,32)=1.007, P>0.05, respectively).

4.3.4.2 P-CREB

Representative examples of the P-CREB protein expression for each drug treatment condition at each of the three times following the start of the light pulse in the late subjective night are presented in Figure 4.9. Similar to the results of the Fos expression discussed previously, there was a significant attenuation of P-CREB expression in the core region of the SCN in animals pretreated with NAN-190 compared to those administered DMSO prior to the light pulse, and this significant attenuation was observed across all three of the time points tested (15 min: P<0.01; 30 min: P<0.001; 45 min: P<0.01, Figure 4.10). However, a similar significant attenuation of P-CREB expression in the shell region of the SCN in animals pretreated with NAN-190 relative to

DMSO was only observed at the time point 30 minutes after the start of the light pulse

(P<0.01). A definite time-course in the expression of P-CREB was also observed in the

SCN core, with a significant increase in expression from the 15 minute to the 30 minute time point (P<0.05), followed by a subsequent non-significant decrease once again to the

45 minute time point (P>0.05). No such significant time course was observed in the shell region of the SCN.

4.4 Discussion

This was the first study to examine the molecular underpinnings of the phase shift potentiation induced by the 5-HT1A mixed agonist/antagonist NAN-190 in mice.

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Figure 4.9. Representative examples of SCN tissue slices indicating cells expressing

P-CREB at (A,B) CT22.25, (C,D) CT22.5, and (E,F) CT22.75 stained using DAB

immunocytochemistry following a brief light pulse in the late subjective night

(CT22), pretreated at CT21 with an i.p. injection of either (A,C,E) vehicle control

(DMSO) or (B, D,F) NAN-190. Scale bar = 200μm.

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Figure 4.10. Mean number of cells expressing P-CREB (± SEM) in the (A) core and (B)

shell regions of the SCN of animals euthanized 15, 30, or 45 minutes following a

brief light pulse in the late subjective night and pretreated with either vehicle

control (DMSO) or NAN-190. (*), (**) and (***) indicate a statistical

significance of P<0.05, P<0.01 and P<0.001, respectively.

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Although the modulation of Fos induced by a light pulse pretreated with NAN-190 was previously observed in hamsters, that study only looked at the expression of Fos in the entire SCN as a whole (Recio et al., 1996). Thus, the first objective of this experiment was to attempt to reproduce, in the mouse, the previous observation of a general attenuation of light-induced Fos expression following pretreatment with NAN-190.

However, the SCN is a heterogeneous structure, consisting of several different cell populations which form separate functional units, some of which receive retinal innervation while others are endogenously rhythmic in their clock gene expression (Antle and Silver, 2005). Therefore, the second objective of the present study was to further investigate the regional specificity, in the core and shell subdivisions within the mouse

SCN, of this attenuated Fos expression.

NAN-190 pretreatment in the mouse, at the same phase used for our behavioural studies (chapter 3), did indeed result in a significant attenuation of light-induced Fos expression in the late subjective night. This effect was observed using two different immunocytochemistry techniques, diaminobenzidine as well as fluorescent triple-label.

More specifically, the fluorescence staining allowed us to determine the regional nature of this decreased Fos expression, such that cells expressing GRP are found only in the core, whereas cells expressing VP are found only in the shell of the SCN (Antle and

Silver, 2005). Thus, by using these two neuropeptides which are expressed in distinct regions of the SCN, we were able to more precisely determine that the majority of the attenuation in light-induced Fos observed with NAN-190, is occurring in the cells of the retinorecipient core of the SCN.

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However, although this Fos protein expression is thought to be an early event in

the response cascade ultimately leading to photic phase alterations of the circadian system, it is not necessary or sufficient to produce phase shifts and is therefore considered to be more of an indication of cell activation (Rea, 1992; Honrado et al.,

1996). However, there are also several redundancies in the pathway linking photic

information to subsequent gene expression, and thus there could be a role for some other

immediate-early gene which behaves similarly to c-fos. Thus, it is still largely unknown

how the potentiation effects of NAN-190 affect the molecular components of the clock

itself. Plus, though a slight decrease in the amount of Fos induced by a light pulse in the

cells of the shell was observed following NAN-190 pretreatment, this difference was not

found to be significant, indicating that Fos expression cannot tell us the whole story as to

how NAN-190 modulates the photic signal sent from the retinorecipient cells to the

endogenously rhythmic cells allowing for the adjustment of circadian phase. For this

reason, we also investigated the expression of two of the clock genes involved in the

molecular workings of the circadian pacemaker, to see if they are specifically altered in

the phase shift potentiation of NAN-190.

Despite the clock genes Per1 and Per2 being known to be crucial to the so-called

‘ticking’ of the circadian clock, due to their involvement in the autoregulatory

transcription-translation feedback loop which drives the circadian cycle in the

endogenously rhythmic cells of SCN shell (Holzberg and Albrecht, 2003), these genes

are also expressed in cells of the SCN core in response to a light pulse during the

subjective night (Antle and Silver, 2005). As expected based on previous work by Yan

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and Silver (2002), mPer1 gene expression was induced in the core region following light

exposure, however the magnitude of this expression did not appear to be altered by NAN-

190 pretreatment. The negligible amount of mPer1 which was expressed in the cells of

the shell following vehicle treatment was surprising though, with previous findings

indicating a two-fold increase in mPer1 expression in the shell 90 minutes following a

light pulse (Yan and Silver, 2002). Therefore, the lack of effect observed with NAN-190

relative to vehicle treatment may not be properly representing the actual effect of the

drug, since we were unable to replicate previous findings in the control treatment.

Alternatively, the negligible amount of mPer2 expressed throughout the SCN with

vehicle pretreatment does coincide with previous work showing essentially no mPer2

expression 90 minutes after light exposure (Yan and Silver, 2002). This normal

expression was also not observed to be altered by the administration of NAN-190 prior to

the light exposure. Therefore, it appears that if NAN-190 administration induces

potentiated phase shifts by altering the expression of mPer1 and mPer2, this effect is not

observable 90 minutes following the start of the light exposure.

Yet, the 5-HT1A receptor agonist MKC-242, which is also able to potentiate

photic phase shifts in both mice and hamsters, was shown to also prolong the expression

of light-induced mPer1 and mPer2 expression in the mouse (Moriya et al., 1998;

Takahashi et al., 2002). MKC-242, administered at the same circadian phase at which it is

known to induce potentiated phase shifts, was able to increase the early subjective night

expression of both mPer1 and mPer2 at 3 hours following the light exposure, but had no

significant effect on either gene 1 hour after the light pulse (Takahashi et al., 2002).

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Perhaps NAN-190 induces potentiated phase shifts via a similar mechanism, which was

not observed due to our study only examining expression at a single time point instead of

the time course of the light-induced Period gene expression, and the possible effects that

NAN-190 may have on this expression over time. However, the effects of MKC-242 on

Period gene expression was only observed following a phase delaying early subjective

night light pulse, whereas we examined the effects of NAN-190 only in the late

subjective night. Thus, it is also possible that potentiated phase delays, but not phase

advances, are mediated by prolonged Period gene expression. Further investigation is

necessary to determine whether NAN-190 may also prolong the expression of the Period

genes, and whether there may be a phase-dependent component to this possible

prolonged expression.

It is also important to consider that changes in the light-induced Period genes may

not play any role at all in the potentiation induced by NAN-190, since its administration

up to 6 hours after light exposure still results in potentiated phase shifts (Kessler et al.,

2008). However, these results have not yet been replicated, but if they hold then it is still

pertinent to keep these results in mind when attempting to decipher exactly what NAN-

190 is doing on the molecular level to induce such large magnitude phase shifts. Perhaps the effect of NAN-190 on gene expression in the core region of the SCN is completely irrelevant, and we should instead be focusing on how it modulates the rhythmic expression of the Period genes, or other genes altogether.

Although we did not examine the time course of Period gene expression, we did examine the time course of P-ERK and P-CREB expression following pretreatment with

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NAN-190. Both of these substances are components of the biochemical cascades, or second-messenger signaling pathways, which couple incoming photic information with the subsequent expression of immediate-early genes, and eventually the Period genes as well (von Gall et al., 1998; Butcher et al., 2003; Dunlap, 2004). The phosphorylation, or activation, of these proteins occurs extremely rapidly following light exposure, but in a phase-dependent manner such that it can only occur at times when photic stimuli are able to induce phase shifts in the circadian activity rhythm (Ginty et al., 1993; Ding et al.,

1997; Obrietan et al., 1998). Indicating the pertinence of these biochemical cascades, the disruption of the ERK or CREB pathways has been shown to attenuate both the phase shifting effects of light as well as IEG expression (Butcher et al., 2002; Coogan and

Piggins, 2003; Dziema et al., 2003; Tischkau et al., 2003; Gamble et al., 2007). Thus, it is conceivable that NAN-190 could also modulate one or both of these pathways in its potentiation of photic phase shifts.

Although no effect was found with NAN-190 on the expression of P-ERK in either the core or shell, at any of the three times tested following the light exposure, there was also no time-dependent change in the vehicle-treated animals either. This indicates that although the expression of P-ERK was rapidly induced in animals in both drug groups, our time course did not extend long enough to observe the eventual decrease in light-induced P-ERK expression normally seen. Therefore, perhaps we would have found an effect with NAN-190, in the form of a prolonged expression for example, had we extended our time course further beyond the 45-minute time point used here. It is of course also possible that the mechanism employed by NAN-190 to induce potentiated

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photic phase shifts does not make use of the ERK pathway in order to alter the expression of the circadian clock.

Therefore, we also examined the parallel second-messenger signaling pathway involving the phosphorylation of CREB. Contrary to what was found with P-ERK expression, a significant effect was observed on the light-induced expression of P-CREB following NAN-190 pretreatment. The greatest effect was observed in the cells of the core, with a significant attenuation of the P-CREB expression, at all three time points following the light pulse, in animals administered with NAN-190 compared to vehicle control. This attenuation could be the source of the decrease in Fos which was also observed in the present study. As a result of light exposure, the CREB protein is phosphorylated, which then binds to the CRE promoter region on several light-induced genes (including c-fos, mPer1, and mPer2), allowing for the subsequent transcription of those genes into mRNA and perhaps their eventual translation into proteins (Dunlap,

2004). However, if the efficiency of this process is disrupted by decreased levels of P-

CREB, this would likely result in a decreased expression of those light-induced genes which possess a CRE promoter region.

There was also a significant attenuation of the light-induced P-CREB expression observed in the cells of the shell region of the SCN, however this was only true 30 minutes following the start of the light pulse. This appears to indicate that NAN-190 may achieve part of its phase shift potentiation via an attenuation of the basal levels of P-

CREB in the endogenously rhythmic cells of the dorsomedial cells of the SCN shell region. However, in order to determine whether NAN-190 may be modulating the basal

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levels of P-CREB in the rhythmic clock cells of the SCN, it would be necessary to

compare the present results to the expression induced by NAN-190 without any exposure

to a subsequent light stimulus.

In contrast to the lack of time course observed in the light-induced expression of

P-ERK, we did find a significant change in the general expression of P-CREB in the SCN

core across the three times examined. There was an increase in the levels of expression,

with a peak magnitude of expression observed 30 minutes following the start of the light

pulse, with a subsequent decline once again by the 45-minute time point. This coincides

quite well with the previously demonstrated time course of light-induced P-CREB, in

which it was observed that expression is induced rapidly, peaking somewhere between 15

and 30 minutes and tapering off over the next 90 minutes (Ding et al., 1997). However,

this general time course did not appear to be affected by NAN-190 pretreatment, which instead simply reduced P-CREB levels to a similar degree across all three time points.

Taken together, the present findings indicate that although it is still not fully

understood precisely how NAN-190 affects the molecular components of the cells of the

SCN in order to induce photic phase shift potentiation, it is clear that NAN-190 is

definitely altering the functioning of the retinorecipient cells of the SCN. This leads us to

the conclusion that the photic phase shift potentiation induced by NAN-190 administration cannot be solely the result of inhibition at the retinal terminals as has been previously suggested (Gannon, 2003). In fact, had the potentiation been the result of

NAN-190 blocking the inhibitory serotonergic innervation at retinal terminals, this should have resulted in a disinhibition and subsequent increased glutamate release. This

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would then be expected to lead to a greater activation of the second-messenger signaling pathways and thus greater c-fos and Period gene expression, which is contrary to what was found in the present study. Therefore, it seems quite evident that NAN-190 functions at least in part through its effects on retinorecipient cells in the core region of the SCN, although the exact mechanism by which this leads to the potentiated photic phase shifts observed in the circadian activity rhythm is yet to be fully elucidated.

By understanding the precise molecular mechanisms responsible for the modulation of the sensitivity of the circadian system to light stimuli, which is observed with the class of drugs termed mixed agonists/antagonists, including NAN-190, it may be possible to eventually develop chronobiotics more specifically targeted to the rapid resynchronization of circadian rhythms. Effectively managing or even preventing the chronic desynchronization between the circadian system and the external environment that is inherent to phenomena such as shift work, jet lag, and circadian rhythm disorders, could play a large role in the elimination of the adverse health effects associated with this

sort of desynchrony. Thus, it is of great importance in today’s 24-h society, to more fully

understand the molecular underpinnings involved with drugs such as NAN-190.

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5. General Discussion

5.1 Summary of Major Findings

The current study examined the role of the 5-HT1A receptor in both normal and

enhanced responses of the circadian clock to light stimuli. Behavioural phase shifting as

well as gene expression were examined in mice lacking the 5-HT1A receptor in order to

determine how the 5-HT1A receptor modulates both photic and non-photic stimuli in the mouse circadian system. Compared to wildtype controls, knockout mice exhibit larger phase advances to a short duration late-night light pulse and larger delays to long 12h light pulses spanning the whole subjective night. Fos protein and mPer1 gene expression in the retinorecipient cells of the SCN is significantly attenuated following late-night light pulses in knockout animals relative to their wildtype counterparts. Finally, non-photic phase shifts to 8-OH-DPAT are lost in the knockout animals, while the attenuation of the phase shift to a long light pulse due to the rebound activity which follows a wheel lock is unaffected.

The role of the 5-HT1A receptor in serotonergic potentiation of photic phase shifts,

using the serotonin mixed agonist/antagonist NAN-190, was also examined in mice

lacking the 5-HT1A receptor. Photic phase shifts were potentiated by NAN-190

pretreatment in both wildtype mice and mice heterozygous for the 5-HT1A receptor.

Conversely, knockouts exhibited an attenuated behavioural response to NAN-190.

Similar effects were observed between the three genotypes, regardless of the intensity of the light pulse. NAN-190 administered without subsequent light exposure was unable to

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induce any significant phase shifts in the activity rhythm of either wildtypes or

knockouts.

Finally, the molecular underpinnings of the behavioural phase shift potentiation

observed with NAN-190, in terms of both gene expression and biochemical activation,

were also examined. Both Fos (the protein product of the immediate-early gene c-fos)

expression and CREB (a component of one of the second-messenger signaling pathways

coupling photic input to clock gene expression) phosphorylation are attenuated by NAN-

190 for at least 90 and 45 minutes, following the light pulse, respectively. The majority of

this modulation is observed in the cells of the retinorecipient core of the SCN. However,

no effect was observed on either P-ERK (a component of another second-messenger

signaling pathway) or the circadian clock components Period1 and Period2 in the time

courses examined.

5.2 Overall Conclusions

Several different conclusions can be drawn from the present findings regarding

the role of the 5-HT1A receptor in normal and enhanced photic phase shifting. First, it

appears that the 5-HT1A receptor plays an inhibitory role on behavioural phase shifts, but

a facilitatory role on light-induced gene expression. In general, 5-HT is considered to

play an inhibitory role on the effects of light, and of the several 5-HT receptors found in

SCN, a postsynaptic mechanism involving the 5-HT1A receptor is proposed to be one way in which serotonin modulates photic information (Pickard and Rea, 1997a). Serotonergic agonists are able to reduce the firing rate in retinorecipient cells of the SCN (Miller and

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Fuller, 1990; Ying and Rusak, 1994), an effect which appeared to be mediate by a

receptor with properties similar to that of the 5-HT1A subtype (Ying and Rusak, 1994).

The inhibitory effects of serotonergic agonists which lead to attenuation of light-induced

phase shifts and inhibition of light-induced c-fos expression in the SCN also appear to

involve receptors which display pharmacological properties similar to 5-HT1A receptors

(Weber et al., 1998). The seemingly contradictory effects observed, such that NAN-190 behaves as an antagonist by potentiating photic phase shifts, but behaves more like an agonist by attenuating gene expression, have yet to be fully reconciled. It would be logical to expect that a greater response to light would lead directly to an increased activation of the light-induced biochemical pathways ultimately leading to increased gene expression.

One possible explanation is that these agonist-like and antagonist-like effects are mediated by 5-HT1A receptors in two different locations within the circadian system, with

an antagonistic action inducing potentiated phase shifts while an agonistic action

elsewhere leading to reduced gene expression. Yet it is also possible that the phase shift

attenuation which was uncovered in mice lacking the 5-HT1A receptor is an indication

that the binding of NAN-190 to some other receptor may also be mediating the attenuated

gene expression, especially since 5-HT1A receptor knockout mice show an increased responsiveness to light stimuli, through larger photic phase shifts, but also demonstrated

decreased light-induced Fos expression to a late night phase shifting light pulse. It is also

important to note that the majority of the responses observed in terms of gene expression

were found in the non-rhythmic core region of the SCN, and not in the rhythmic cells of

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the shell where the expression of clock genes must be ultimately adjusted in order to

induce a phase shift. More research is definitely necessary in order to determine exactly

what receptors are involved in each aspect of serotonergic modulation of photic phase

shifts, as well as the precise location of those receptors,.

Secondly, the 5-HT1A receptor appears not to be necessary for the activity- induced phase advances found to oppose phase delays to long duration light pulses. Yet it

does appear to play a necessary role in the non-photic phase shifts induced by 8-OH-

DPAT in the mid-subjective day. In vitro, 8-OH-DPAT has been found to induce phase

shifts in neuronal firing which were no different between wildtype and 5-HT7 receptor

knockout mice, and these phase shifts were significantly reduced in both wildtype and

knockout mice by the co-application of a highly selective 5-HT1A receptor antagonist,

indicating a major role for the 5-HT1A receptor in the effects of 8-OH-DPAT (Sprouse et

al., 2005). These findings are complicated by the recent studies also indicating a

necessary role for the 5-HT7 receptor in 8-OH-DPAT-induced non-photic phase shifts,

both in vitro and in vivo (Guscott et al., 2005; Gardani and Biello, 2008). The phase shifts

induced by 8-OH-DPAT administration in the mid-subjective day are sensitive to co-

application of 5-HT7 antagonists but not to 5-HT1A antagonists (Sprouse et al., 2004b).

Plus, in vivo, 5-HT7 receptor knockout mice show a significant attenuation of the non-

photic phase shifts induced by 8-OH-DPAT, demonstrating a necessary role for the 5-

HT7 receptor in these shifts as well.

Thirdly, the 5-HT1A receptor does also seem to be crucial to induce the photic

phase shift potentiation with NAN-190 in the late subjective night. Although one

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functional allele for the receptor appears to be sufficient to at least partially restore

essentially normal phase shift potentiation induced by NAN-190. Finally, NAN-190

appears to modulate the expression of genes and biochemical pathway components

linking photic input to clock gene expression in the retinorecipient cells of the SCN.

However, an attenuated activation of CREB at 30 minutes following the start of the light

pulse, as well as a trend towards decreased Fos expression 90 minutes after the light

pulse, in the cells of the SCN shell appears to indicate there may be a modulatory role for

NAN-190 in the endogenously rhythmic cells of the SCN as well as just in the

retinorecipient core. Although the present findings do not identify the exact mechanism by which NAN-190 induced potentiated photic phase shifts, they do suggest that the hypothesis put forth by Gannon (2003), that NAN-190 functions mainly by blocking the normal serotonergic inhibition at retinal terminals and thus allowing greater excitatory signals to reach the SCN and potentiating the phase shift response, cannot be the whole story. Our findings indicate that NAN-190 definitely appears to be modulating gene expression in some way directly in the retinorecipient cells of the SCN, and may also play a role in modulating the gene expression within the endogenously rhythmic cells of the SCN shell. Again, more research is still necessary to fully understand the mechanism by which NAN-190 potentiates photic phase shifts, the present findings do provide many possible avenues for investigating this phenomenon further.

5.3 Limitations and Future Directions

In order to examine the role of the 5-HT1A receptor, this study made use of

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animals genetically engineered to lack that receptor. Although this method can be

extremely useful in fully eliminating the effects of a specific receptor instead of simply

trying to selectively block a receptor subtype, there are issues involved in generating

enough animals to adequately examine a phenomenon of interest. The first major issue

involved the actual generation of knockout animals. Although our breeding colony was

fairly productive and litter sizes were relatively normal, very few knockouts were

generated overall. In fact, the ratio of each genotype that was generated by the colony was not at all conducive to the generation of a suitable number of knockout animals to

fully examine the role of this receptor in photic phase shifting. The colony also went

through cycles of productivity, as well as times when the colony was almost lost due to a

lack of productive breeding pairs, and the trait had to be back-crossed out to mice of the

same background on which the colony was originally generated. This is the main reason

for the relatively small sample sizes for some of the experiments using knockouts,

especially in the protein and gene expression components of chapter 2.

A second important limitation to this study involves the maintenance of the

transgenic colony and genotyping the animals. Since we were not using a previously

established colony, it was necessary to first determine an appropriate genotyping protocol

largely through the process of trial and error. However, once the protocol was working

accurately, there were also several instances when the protocol unexpectedly stopped

working. Thus, genotyping the animals in a timely manner, in order to start manipulations

as quickly as possible, was not always easily accomplished.

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Another limitation which should not be overlooked for the present study is

regarding the vehicle in which NAN-190 must be dissolved. Unlike other similar drugs

which can be easily dissolved in saline, such as BMY 7378, NAN-190 is insoluble in

water or saline and must instead be dissolved in DMSO. Although previous studies have

been able to use a 50% DMSO solution (Kessler et al., 2008), attempts were made to use

a combination of DMSO and saline, but we were unable to dissolve the powder form of

the drug in anything less than undiluted 99% DMSO. Thus, vehicle injections also had to

be 99% DMSO, which could potentially have led to undesirable side-effects, including

such effects as perivascular inflammation and intravascular thrombosis with repeated systemic injections, despite its relatively low toxicity in laboratory animals (Pope and

Oliver, 1966). It does not appear, however that the expression of circadian wheel-running activity was substantially affected despite these possible deleterious side-effects.

However, the use of water- or saline-soluble drugs might be a more suitable choice for future studies, especially since a drug such as BMY7378 has been shown to also greatly potentiate photic phase shifts in hamsters but is known to have a much higher affinity for and specificity to the 5-HT1A receptor (Gannon, 2003). However, pilot data from our lab

indicates that BMY 7378 may not be as effective in producing potentiated phase shifts in

mice (unpublished data) as has been shown previously in hamsters (Gannon, 2003).

One final limitation involves the specific time-courses examined for each gene or

protein included in this study. First, the unexpected lack of time-course effects observed

with the expression of P-ERK in the control injection condition does not allow us to

determine if NAN-190 may play a modulatory role in this expression. Therefore, it is

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possible that we may have found an effect of NAN-190 had we extended our time course to include times which would have shown the eventual decrease in the light-induced P-

ERK expression. With the current time course, we may have only examined times at which the circadian system was still saturated by the rapid induction of P-ERK, which would not allow us to discriminate between the normal photic induction and the effect of

NAN-190. Similarly, the single time point at which the expression of the clock genes mPer1 and mPer2 does not indicate whether NAN-190 may prolong the expression of these genes as is observed with the potentiating effects of MKC-242 (Takahashi et al.,

2002). Our results do not indicate that NAN-190 induces photic phase shifts without altering Period gene expression in some way; they merely indicate that the precise molecular mechanism by which NAN-190 modulates photic input was not uncovered by the specific gene expression measured and in the time courses examined. It would therefore be important to examine expression at more extended lengths of time following light exposure in order to determine whether the mechanism of action of NAN-190 may mirror the findings of MKC-242, or whether Period gene expression is even involved at all. Another avenue of investigation would also involve looking at several other immediate-early genes, such as fosB, Jun-B and Jun-D, or the clock genes cry1 and cry2 which could also play some role in modulating the potentiation of photic phase shifts with NAN-190.

A few other studies which would be extremely useful to extend our current findings include examining all of the aspects of the mechanism of action with NAN-190 studied in chapters 3 and 4 in the early subjective night as well. Plus, it would also be

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useful to determine the role of the 5-HT1A receptor in the gene expression modulation

found in chapter 4, through the use of mice missing the 5-HT1A receptor. Along with this, it would also be important to more fully examine the potential role of other serotonergic receptors for which NAN-190 has been shown to have an affinity. Since it has been proposed that at least part of the ability of NAN-190 to modulate photic signaling may be through binding at either 5-HT7 or α1-adrenergic receptors (Claustre et al., 1991;

Lovenberg et al., 1993), and our results from chapter 3 indicated that removing the 5-

HT1A receptor blocks the potentiating action of NAN-190 while uncovering an

attenuation effect through some other receptor activation, it is important to determine the

precise role of other receptors at which NAN-190 may be acting as well.

5.4 Clinical Importance

The development of a chronobiotic which could rapidly and extensively reset the

circadian system would be extremely useful in the today’s society. Numerous health

consequences are associated with those individuals who are consistently desynchronizing

their circadian system from the external environmental cycle through shift work

schedules or frequent intercontinental travel (Rajaratnam and Arendt, 2001; Knutsson,

2003; Dunlap, 2004; van Mark et al., 2006). Unfortunately it is difficult to avoid such

desynchronization with the 24h nature of our society, and the majority of the commonly

used means by individuals for resetting the circadian clock are only marginally effective

at best. Thus, the development of an oral chronobiotic which could alleviate or even

prevent the negative side effects of chronic desynchronization of circadian rhythms,

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would be invaluable for a large portion of the population. In order to accomplish this, it is critical to understand the pharmacological and molecular mechanisms underlying the action of drugs such as NAN-190.

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