Download The circadian visual system, 2005

BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
a v a i l a b l e a t w w w. s c i e n c e d i r e c t . c o m
w w w. e l s e v i e r. c o m / l o c a t e / b r a i n r e s r e v
Review
The circadian visual system, 2005
L.P. Morina,⁎, C.N. Allenb
a
Department of Psychiatry and Graduate Program in Neuroscience, Stony Brook University, Stony Brook, NY 11794, USA
Department of Physiology and Pharmacology and Centre for Research on Occupational and Environmental Toxicology,
Oregon Health and Science University, 3181 SW Sam Jackson Park Road, Portland, OR 97239, USA
b
A R T I C LE I N FO
AB S T R A C T
Article history:
The primary mammalian circadian clock resides in the suprachiasmatic nucleus (SCN), a
Accepted 9 August 2005
recipient of dense retinohypothalamic innervation. In its most basic form, the circadian
Available online 5 December 2005
rhythm system is part of the greater visual system. A secondary component of the circadian
visual system is the retinorecipient intergeniculate leaflet (IGL) which has connections to
Keywords:
many parts of the brain, including efferents converging on targets of the SCN. The IGL also
Circadian
provides a major input to the SCN, with a third major SCN afferent projection arriving from
Suprachiasmatic
the median raphe nucleus. The last decade has seen a blossoming of research into the
SCN
anatomy and function of the visual, geniculohypothalamic and midbrain serotonergic
Intergeniculate leaflet
systems modulating circadian rhythmicity in a variety of species. There has also been a
IGL
substantial and simultaneous elaboration of knowledge about the intrinsic structure of the
SCN. Many of the developments have been driven by molecular biological investigation of
the circadian clock and the molecular tools are enabling novel understanding of regional
function within the SCN. The present discussion is an extension of the material covered by
the 1994 review, “The Circadian Visual System.”
© 2005 Elsevier B.V. All rights reserved.
⁎ Corresponding author. Fax: +1 631 444 7534.
E-mail address: [email protected] (L.P. Morin).
0165-0173/$ – see front matter © 2005 Elsevier B.V. All rights reserved.
doi:10.1016/j.brainresrev.2005.08.003
2
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
Abbreviations:
5-HT, serotonin
CALB, calbindin
CALR, calretinin
CCK, cholecystokinin
ENK, enkephalin
GABA, gamma amino butyric acid
GRP, gastrin releasing hormone
IGL, intergeniculate leaflet
NPY, neuropeptide Y
NT, neurotensin
PACAP, pituitary adenylate
cyclase-activating polypeptide
SCN, suprachiasmatic nucleus
SP, substance P
SS, somatostatin
VIP, vasoactive intestinal polypeptide
VP, vasopressin
Contents
1.
2.
3.
4.
5.
6.
Introduction and caveats . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Psychophysics of the circadian rhythm system . . . . . . . . . . . . . . . . .
Organization of the suprachiasmatic nucleus . . . . . . . . . . . . . . . . .
3.1. Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.2. Distribution of cell phenotypes . . . . . . . . . . . . . . . . . . . . .
3.3. Intra-SCN connections . . . . . . . . . . . . . . . . . . . . . . . . . .
Retinal photoreceptors and projections . . . . . . . . . . . . . . . . . . . .
4.1. Melanopsin ganglion cell characteristics . . . . . . . . . . . . . . . .
4.2. Retinal ganglion cell projections and bifurcation . . . . . . . . . . .
4.3. Classical and ganglion cell photoreceptor contribution to regulation
light reflex and melatonin suppression . . . . . . . . . . . . . . . .
4.3.1. Circadian rhythms . . . . . . . . . . . . . . . . . . . . . . .
4.3.2. Masking, pupillary light reflex and melatonin suppression. . .
Neuromodulators of the retinohypothalamic tract . . . . . . . . . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
of circadian
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
rhythms, masking,
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. . . . . . . . . . .
. .
. .
. .
. .
. .
. .
. .
. .
. .
the
. .
. .
. .
. .
. . . . 3
. . . . 4
. . . . 5
. . . . 5
. . . . 5
. . . . 8
. . . . 8
. . . . 8
. . . . 9
pupillary
. . . 11
. . . 11
. . . 11
. . . 12
5.1. Glutamate . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.2. N-Acetylaspartyl glutamate . . . . . . . . . . . . . . . . . . .
5.3. Glutamate receptors . . . . . . . . . . . . . . . . . . . . . . .
5.4. Nitric oxide and RHT signal transduction . . . . . . . . . . .
5.5. PACAP. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.6. Substance P . . . . . . . . . . . . . . . . . . . . . . . . . . . .
SCN structure–function relationships . . . . . . . . . . . . . . . . .
6.1. Function of cellular neuromodulators in the SCN . . . . . . .
6.1.1. GABA . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.1.2. VIP and VIP/PACAP receptors. . . . . . . . . . . . . .
6.1.3. Calbindin and the central subnucleus of the hamster
6.1.4. Bombesin and gastrin releasing peptide. . . . . . . .
6.1.5. Neuromedin U and neuromedin S . . . . . . . . . . .
6.1.6. Vasopressin . . . . . . . . . . . . . . . . . . . . . . .
6.1.7. Diurnality–nocturnality . . . . . . . . . . . . . . . . .
6.2. Acetylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.3. Neurophysiology of the SCN action potential rhythm . . . . .
6.4. Inter-SCN differences in gene expression . . . . . . . . . . .
6.5. Regional expression of Fos . . . . . . . . . . . . . . . . . . .
6.5.1. Hamster . . . . . . . . . . . . . . . . . . . . . . . . .
6.5.2. Rat . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6. Regional clock gene expression . . . . . . . . . . . . . . . . .
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
SCN
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
. . .
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
12
12
13
13
14
15
17
18
18
18
19
21
21
21
22
22
23
25
25
25
25
26
3
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
6.7.
7.
6.8.
SCN
7.1.
6.6.1. Mouse. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6.2. Rat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6.3. Hamster . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Stimulus induction of gene expression . . . . . . . . . . . . . . . . . .
6.7.1. Photic stimuli . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.7.2. Nonphotic stimuli . . . . . . . . . . . . . . . . . . . . . . . . .
6.7.3. Inconsistencies between phase response and gene expression
Polysialic acid and SCN function . . . . . . . . . . . . . . . . . . . . .
connectivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Humoral factors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
7.2. Efferent anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
7.3. Afferent anatomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.
Intergeniculate leaflet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.1. Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.2. Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.3. Anatomy. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.3.1. Geniculohypothalamic tract. . . . . . . . . . . . . . . . . . . . . .
8.3.2. Connectivity and relationship to sleep, visuomotor and vestibular
8.4. Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.4.1. Neurophysiology . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.4.2. Nonphotic regulation of rhythmicity . . . . . . . . . . . . . . . . .
8.4.3. Photic regulation of rhythmicity . . . . . . . . . . . . . . . . . . .
9.
Serotonin and midbrain raphe contribution to circadian rhythm regulation . . .
9.1. Effects of serotonin-specific neurotoxins . . . . . . . . . . . . . . . . . . .
9.2. Serotonin receptor function . . . . . . . . . . . . . . . . . . . . . . . . . .
9.3. Regulation and consequences of serotonin release . . . . . . . . . . . . .
9.4. Serotonergic potentiation of light-induced phase shifts . . . . . . . . . . .
10. Failure to re-entrain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
11. Masking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
12. The future . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1.
Introduction and caveats
There have been a large number of new developments with
respect to knowledge about the anatomy and physiology of
circadian rhythm regulation since publication of our 1994
review, “The Circadian Visual System” (Morin, 1994). As is
nearly always the case with such projects, many of the
developments were well underway, and in some instances
completed, prior to the actual publication of that review. Most
notably, much of the presentation in the 1994 review
concerning the serotonergic system was incomplete or
wrong and understanding of the issues changed substantially
soon after publication of the review. Other issues emerged as
unanticipated major developments. One of these has been the
discovery of the photopigment, melanopsin, in retinal ganglion cells and the subsequent establishment of those ganglion
cells as a special class of photoreceptors.
The steady growth of molecular biological contributions
to the understanding of circadian clock mechanisms in
invertebrates has been translated into parallel evaluation of
clock mechanisms in mammals. This development has
created new avenues for architectural analysis of the
suprachiasmatic nucleus (SCN) function according to the
regional distributions of cell phenotypes comprising the
circadian clock.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
26
27
27
28
28
28
29
30
31
31
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
systems
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
. . . . .
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
31
32
34
34
34
34
34
34
35
35
36
37
38
38
38
39
40
41
42
43
43
43
The purpose of the current review is to continue the
theme of the original, but to do so with minimal redundancy. The focus of the current presentation will be on
necessary modifications to the perspective presented 10
years ago, and to the substantial additions to the general
body of knowledge relating to the circadian visual system.
The title of the 1994 review, “The Circadian Visual System,”
remains of some import. We believe it is worth emphasizing
that, taken as a whole, the research topic under consideration is “circadian,” is “visual” and certainly is a “system.”
That is an aspect of the research topic of importance that
should not be underestimated.
We do not intend to cover all facets of circadian rhythm
research and we also expect to have inadvertently overlooked
research worthy of inclusion here. To the extent that has
happened, we apologize. On a technical note, when specific
cell types are mentioned as containing a particular neuromodulator, the comment refers to the fact that it has been
demonstrated by immunohistochemical means.
The work that is particularly ignored concerns the molecular biology of the SCN or other tissues. There will be no effort
to review the burgeoning literature concerning the molecular
mechanisms of the central circadian clock or of molecular
oscillations in peripheral structures. Rather, we will touch on
this literature when it can be used to amplify understanding of
4
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
the system properties governing circadian rhythm regulation.
One issue at the forefront of the list of mysteries presently
confronting that understanding is the spatial distribution of
SCN neurons exhibiting clock-like behavior. The general
question is, “Are all SCN cells pacemakers?” While some
experimental work suggests that the answer is no, the topic is
an area of intense research and is discussed in some detail
below.
A second question is, “Do all pacemaker cells in the SCN
oscillate with the same phase?” Evaluation of a variety of
molecular oscillations in SCN cells has given the impression
that the molecular behavior of a whole SCN reflects the
behavior of the individual cells. While this may be true on
average, many individual SCN neurons do not express
rhythmicity in phase with the average of the whole (Quintero
et al., 2000). This development in the analytical approach to
pacemaker activity among groups of cells emphasizes the
importance of molecular activity within individual cells and
points to the need to evaluate spatial relationships and
connectivity among oscillatory neurons within the SCN.
When microelectrode arrays are used for long-term
recordings, the action potential firing rhythm periods of
individual of SCN neurons show a greater diversity in
dispersed cultures compared to organotypic cultures. These
data suggest that interneuronal communication is required to
fine tune the period of the clock (Nakamura et al., 2002; Herzog
et al., 1998; Liu et al., 1997a; Albus et al., 2002). The final
behavioral period is an average of these individual neuronal
periods (Liu et al., 1997a). The demonstration of phase
differences among SCN neurons is a variant on the larger
theme indicating differences in phase between the two SCN in
a single animal (De la Iglesia et al., 2000). Thus, from the point
of view of this review, utility of the molecular analysis of
cellular clocks serves to highlight the need to understand the
system properties intrinsic to the SCN, as well as its regulation
of efferent response to afferent information, whether that
information be visual or some other sensory modality.
2.
Psychophysics of the circadian rhythm system
The Pickard et al. (1987) IGL lesion paper was one of the first to
employ nonsaturating light stimuli to elicit circadian rhythm
phase responses. The method eliminated ceiling effects
caused by saturating light, providing a more accurate view of
psychophysical sensitivity to light, rather than a simple
statement that animals respond (or not) to the stimulus.
Subsequently, the psychophysics of circadian rhythm phase
response to light presented at CT19 has been explored in
greater detail. The hamster (Nelson and Takahashi, 1991, 1999;
Zhang et al., 1996), mouse (van den Pol et al., 1998) and gerbil
(Dkhissi-Benyahya et al., 2000) circadian systems estimate the
importance of an acute photic stimulus by effectively “counting” photons. It may be more appropriate to refer to this
activity as “temporal integration” which is a broader term. The
process is demonstrated by the fact that there is a clear
relationship between the stimulus duration and irradiance
(photons/cm2) with respect to the magnitude of the phase
shift. That is, dim, long duration stimuli have the same effect
as brief but bright stimuli, if the irradiances (i.e., total photons
or Joules/cm2) are equivalent. Most impressively, the photic
intensity information in a series of light pulses appears to be
summed across the series (Nelson and Takahashi, 1991), as
long as the individual pulses are of sufficient duration (i.e.,
minutes).
The integrative capacity of the circadian system appears to
be time-limited to the extent that, under test conditions
explored to date, if a photic stimulus is sufficient to produce a
maximal phase shift, more photons at the same time or up to 2
h later have no additional effect on phase shift magnitude
(Nelson and Takahashi, 1999). In such cases, the circadian
system is deemed to be “saturated” with respect to its ability to
show a further phase response to light.
An unexpected result (van den Pol et al., 1998), demonstrated the important point that a form of “temporal
integration” may occur even when the light stimulus is
subthreshold. For example, in the mouse or hamster, a brief
(2–3 ms), bright (1013 photons/cm2/s) flash will not elicit a
phase shift (Nelson and Takahashi, 1991; van den Pol et al.,
1998), but a series of flashes totaling 120 ms will yield a normal
phase response (van den Pol et al., 1998). Thus, some form of
summation has occurred to elicit the phase shifts. It is not yet
clear, however, that there is a systematic relationship between
the number of flashes and the magnitude of the phase shift.
Evidence from studies employing a series of 5 to 45 light
flashes, each only 10 μs long, suggests otherwise (Arvanitogiannis and Amir, 1999). Each such series effectively induces
FOS-IR in the SCN and yields 75 min phase shifts when
delivered to rats at CT13. Thus, the fact of a light-induced
phase shift may be separable from the process of
“integration.”
The entire visual system has been intact in all studies to
date that involve “temporal integration.” Therefore, there is
presently no information regarding the extent to which the
integration mechanism includes or excludes the retina, SCN,
IGL, visual midbrain, the serotonergic or other systems.
Unilateral enucleation eliminates 50% of the retinal photoreceptors and their associated RHT projections, but had no
effect on hamster re-entrainment rate (Stephan et al., 1982).
On the other hand, the same publication showed that both
advance and delay shifts of rats were retarded following
unilateral enucleation. D. Earnest (unpublished work cited by
Pickard and Turek, 1983) failed to find an effect of unilateral
enucleation on hamster circadian period in LL. Similarly,
unilateral enucleation appears to have no effect on lightinduced FOS in rat (Beaulé et al., 2001a) or hamster SCN
(Muscat and Morin, unpublished data), or in SCN of hamster
with one eye occluded, except in response to high irradiance
or long duration light (Muscat and Morin, in press). In each of
these investigations, saturating stimuli were employed and
the experimental methods not sufficiently sensitive to determine whether there is an effect of unilateral enucleation on
rate of re-entrainment or any other measure of circadian
rhythm regulation (but see Tang et al., 2002 for a broad set of
results suggesting a large effect of unilateral enucleation).
Unilateral enucleation is also reported to shorten the
circadian period of rats in LL, while a unilateral SCN lesion
has no similar effect (Donaldson and Stephan, 1982). Given
that the IGL regulates 50% of the hamster circadian period
response to LL (Pickard et al., 1987; Morin and Pace, 2002) (no
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
equivalent studies have been done in the rat), the effect of
enucleation could be attributable exclusively to loss of the
retinal input to the IGL.
Light induction of FOS-IR in the gerbil SCN (DkhissiBenyahya et al., 2000) has been interpreted as obeying the
temporal integration rule found in the hamster for phase
shifts and melatonin suppression. However, over the same
irradiance range, the effect of light on FOS in ganglion cells of
the gerbil retina appears to be all-or-none, rather than
additive. Similar results have been found for light-induced
FOS in hamster IGL cells (Muscat and Morin, unpublished data)
and imply that neither the ganglion cell nor IGL neuron
properties are altered by light in a manner consistent with
“photon counting” responses of the circadian rhythm system.
3.
Organization of the suprachiasmatic nucleus
3.1.
Nomenclature
An accepted anatomical nomenclature affords the opportunity to standardize the description of brain structures. This has
not yet happened with respect to the SCN intrinsic anatomy.
Historically, there have been two approaches to SCN organization, one based on the rat model and the other based on the
hamster. The rat SCN has been commonly divided into
dorsomedial and ventrolateral divisions. Although these are
geographic designations, the conceptualization is based on
the fact that, in this species, the vasoactive intestinal
polypeptide (VIP) neurons plus most of the geniculohypothalamic tract (GHT) and retinal afferents occupy the latter, while
the former is the location of most vasopressin (VP) neurons.
The hamster SCN has a caudocentral region containing
neurons identified by several different peptides and this has
been referred to as the “central subnucleus” which partially
overlaps with the region containing VIP neurons. In this
species, there has been no anatomical attempt to designate a
“ventrolateral” SCN division, although some investigators
have applied rat-derived descriptors to the hamster. As
indicated in Section 3.2, identification of a dorsomedial area
based upon the location of VP-immunoreactive neurons is of
dubious value, even in the rat.
The nomenclature has become further confused with
references to a “core and shell” SCN organization. For
example, in a 1996 review, one of us (LPM) used the terms as
an analogy to facilitate understanding of the relationship
between the hamster central subnucleus and its surround
(Miller et al., 1996). However, another contributor to that
article employed the same terms in a different review, also
published in 1996, as labels of regions more or less equivalent
to the previously used dorsomedial and ventrolateral divisions in the rat SCN (Moore, 1996). Given the reasonable
likelihood that a common SCN organizational plan will
emerge and be applicable to rodents, and that apparent
structural differences between rat and hamster SCN will be
reconciled, we believe it is premature to adopt the “core and
shell” terminology based upon their application to anatomical
description in one species or another. More important, and as
discussed below, we are of the opinion that the “core and
shell” terminology (A) is not supported by the data in any
5
species; (B) is not consistent with the apparent anatomical
differences between species; (C) has been employed with
different meanings in different species, as well as within the
same species; (D) unduly simplifies a structure which has a
still-emerging level of organizational complexity and (E) if
accepted in an unquestioning manner, will have a direct
negative influence on experimental design, while augmenting
the likelihood of erroneous and incomplete interpretation of
new and available data.
Another reason for caution in using the core/shell
terminology is that they are based primarily on the localization of the neuronal cell body. SCN neurons have a soma that
is approximately 10–15 μm in diameter and generally contain
1–3 dendrites. These dendrites extend for a distance of about
250 μm. Given the small size of the SCN (300–400 μm wide and
400–500 μm tall), a dendrite has the ability to project across
much of the SCN (van den Pol, 1980, 1991; Silver and Brand,
1979; Jiang et al., 1997). This implies, for example, that a cell
body in the dorsal part of the SCN could send a dendrite
ventrally into the region where the densest retinal terminals
are located. Indeed, individual dendrites from the dorsal SCN
have been shown to extend into the ventral SCN and further
into the optic chiasm (Silver and Brand, 1979). Rejection of
“core and shell” nomenclature specifically does not mean
that there is no specialized organization within the SCN.
However, we believe the research goal should be to determine
anatomical reality with a view toward consistent and
constructive use of a nomenclature that is sufficiently fluid
to permit revisions reflecting the evolution of knowledge
about the SCN.
3.2.
Distribution of cell phenotypes
Essential to the understanding of circadian rhythm system
function is knowledge about the intrinsic organization of the
SCN, site of the predominant circadian clock in mammals. It
has been commonly stated that all SCN neurons appear to
contain GABA (Moore and Speh, 1993; Morin and Blanchard,
2001; Abrahamson and Moore, 2001), although an immunohistochemical, electron microscopic study of the rat SCN
suggests that only 40–70% contain GABA (Castel and Morris,
2000). There is a dense GABAergic plexus throughout the
nucleus, but a clear spatial organization of the SCN is
evident if other cellular characteristics are evaluated. The
two most obvious are the distribution of cell phenotypes and
the distribution of cells projecting to non-SCN targets. The
former can be ascertained by immunohistochemical means,
while the latter relies largely upon retrograde tracer
injections.
A cogent argument has been made supporting the view
that the SCN consists of two major divisions, a dorsomedial
area related to the presence of VP neurons and a ventral area
related to the presence of VIP cells, dense retinal afferents and
neuropeptide Y-containing terminals of the GHT (Moore et al.,
2002). However, the evidence seems to suggest the presence of
more than two subdivisions within the SCN, with some more
expressly related to cell phenotypes than others. For example,
the rat SCN has VP cells distributed in a crescent extending
from a ventral location, medially, dorsally and laterally,
almost encircling a region that contains the VIP neurons.
6
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
These two areas are distinct from, but appear to overlap with,
a predominantly dorsolateral region containing calretinin
(CALR) and enkephalin (ENK) neurons (Moore et al., 2002).
Encompassed by the region of VIP cells is a smaller domain of
neurotensin neurons.
In the mouse, the regional situation resembles that of the
rat to the extent that there appear to be multiple SCN areas
characterized by one or more particular cell phenotypes. VP
cells are scattered across the rostral SCN, are located
predominantly in the typical dorsomedial aspect of the central
SCN and are again scattered through the entire caudal SCN.
Some cells are also present in the ventral and ventrolateral
SCN. ENK cells are grouped as a collection in the dorsal SCN,
while VIP neurons are generally localized to the ventral part of
the nucleus. GRP cells occupy a more central region, slightly
dorsal to the VIP neurons and ventral to the ENK cells
(Abrahamson and Moore, 2001; Silver et al., 1999). A green
fluorescent protein reporter which minimizes interference
from GRP immunoreactivity in cell processes, indicates that
these neurons occupy a large percentage of the central mouse
SCN and that the GRP region extends dorsolaterally to the
perimeter of the nucleus (Karatsoreos et al., 2004).
Interpretation of regional distinctiveness based on cell
phenotype may be limited by the experimental methods. For
example, recent data suggest that discrete VP and VIP regions
may be more apparent than real. Immunohistochemistry for
VP peptide generally reveals dense staining in the dorsomedial hamster SCN where there is a mix of cells and processes in
which cells are often difficult to identify (e.g., Miller et al., 1996;
Kalsbeek et al., 1993). In marked contrast, in situ hybridization
methods detecting VP mRNA enable clear visualization of
individual neurons. This procedure reveals that, at certain
circadian times, the gene is active over a much wider portion
of the SCN (Silver et al., 1999; Hamada et al., 2001), including
areas that do not contain neurons clearly identifiable by VP
peptide content. In fact, at certain circadian times, virtually no
VP mRNA at all is detected in the SCN (CT20), whereas at CT8,
the VP gene appears to be active in the majority of anterior
SCN neurons (Hamada et al., 2004a). In situ hybridization
methods also demonstrate a much larger distribution of VP
mRNA in the rat SCN than expected from comparable results
based on immunohistochemistry (Dardente et al., 2002). The
method-related differences are also apparent when evaluating rat VIP. These results plus those for VP mRNA or peptide
indicate clearly different, but extensively overlapping, distributions of VP and VIP cells in the rat SCN. The data have
implications for all allegations of discreet VP or VIP regions
and to any specific functions related to those cells or the areas
in which they are purportedly found.
Despite the foregoing caution, analysis of the hamster SCN
indicates that it shares with other rodents the apparent
separation of a VP region from a VIP region (Fig. 1) because
there is no question that the most persistently evident
dorsomedial part of the VP region does not overlap with the
VIP region (Hamada et al., 2004a; Card and Moore, 1984). The
hamster SCN also has a dorsolateral region generally devoid of
a presently known specific cell phenotype. The hamster SCN
also contains a feature initially thought to be unique to this
species, namely a small, fairly central region identifiable by its
distinct sets of neurons. The region was originally identified as
Fig. 1 – Schematic organization of the hamster SCN at a level
through the central subnucleus identified by SP, CALB, CALR
and GRP immunoreactive cells. (A) Approximate locations of
cell types. VP, SS and CCK are found in a generally
dorsomedial location, although this may vary according to
time of day (indicated by shading). CCK cells are also evident
ventrolaterally (black dots); (B) terminal fields of the three
major afferent pathways. PACAP and glutamate from retinal
projections occupy the entire SCN, to a varying degree (see
Part C of this figure). 5-HT terminals from the median raphe
nucleus fill essentially the entire nucleus, but only sparsely in
the central and dorsolateral regions. NT, NPY, ENK and GABA
arrive from the IGL, although the GABA terminals from the IGL
have not been explicitly identified (see text) and (C)
identifiable zones of retinal projections. Black = region of
dense retinal innervation, predominantly from the
contralateral retina. Cross-hatched = region of modest
innervation predominantly from the ipsilateral retina. Shaded
gray = region of lesser, approximately equal, innervation from
each retina. This diagram is a guide, not a rule. The lines do
not represent absolute boundaries.
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
the location of substance P (SP) cells (Morin et al., 1992)
visualized with the use of colchicine. However, the same
region contains numerous cells synthesizing calbindin (CALB)
(Silver et al., 1996a) which is more easily visualized. The region
also contains neurons immunoreactive to gastrin releasing
peptide (GRP) (Miller et al., 1996) and calretinin (CalR; Silver et
al., 1996a; Blanchard and Morin, unpublished data). Furthermore, this caudocentral SCN subnucleus overlaps to some
degree with the VIP region (Aioun et al., 1998).
The complexity of the caudocentral region of the hamster
SCN is borne out by colocalization studies. About 8% (Silver et
al., 1996a) or 65% (LeSauter et al., 2002), 37 and 33% of CalB cells
have colocalized SP, GRP and VIP, respectively. About 80–90%
of SP cells and 43% of GRP cells also contain CalB and 60% of
VIP neurons located within the region of the CalB neurons
actually containing CalB (LeSauter et al., 2002). GRP has also
been observed colocalized with VIP (Aioun et al., 1998).
Colocalization of CalB and cholecystokinin (CCK) does not
occur (LeSauter et al., 2002). CalR has not been examined for
the possibility of colocalization in the SCN, nor have most
other combinations of antigens.
The region containing VP neurons is matched reasonably
well by the locations of CCK cells. Nevertheless, to the limited
extent that the issue has been examined, there is no
colocalization of VP and CCK in the hamster (Blanchard and
Morin, unpublished data). In the most ventrolateral part of the
nucleus, CCK cells are evident at a location containing VP
fibers (LeSauter et al., 2002).
The caudocentral SCN subnucleus may be special with
respect to circadian rhythm regulation. This issue is considered separately below. From the comparative anatomical
perspective, it is useful to question the extent to which other
species have a clearly defined region of likely homology. The
answer is not clear. The only other species known to date with
SP cells at a similar location is the Djungarian hamster (Reuss
and Bürger, 1994). The grass rat has CALB-IR neurons clustered
in the SCN (Mahoney et al., 2000), apparently more centrally
located than those in the hamster in which the subnucleus is
located in the caudal half of the SCN. In rats, CALB-IR neurons
are more prevalent in the ventral SCN, but are not as tightly
clustered as in the hamster (Mahoney et al., 2000; Arvanitogiannis et al., 2000). In mice, CALB neurons are present within
the general central SCN area, the surrounding region and
generally throughout the entire nucleus (Abrahamson and
Moore, 2001). However, the mouse SCN does have a clearly
defined central region notable because of the absence of a
specific set of identifiable neurons (LeSauter et al., 2003). This
region corresponds most closely to the location of neurotensin
and GRP neuron clusters (Abrahamson and Moore, 2001). A
region with reasonably clear homology to the hamster central
subnucleus has been described in the golden mantled squirrel,
but is recognized by a distinct cluster of ENK, rather than SP,
cells (Smale et al., 1991).
The rat SCN is more compressed along the dorsoventral
axis than it is in the mouse or hamster. A cell grouping
obviously homologous to those in the hamster caudocentral
subnucleus has not been described in the rat. However, an
argument can be made that cells containing GRP occupy a
similar position. The GRP cells, in particular, meet the criteria
of being tightly distributed, more or less centrally located
7
within the SCN, with a distribution not identical to another
known phenotype (Dardente et al., 2002 but see Moore et al.,
2002 for a different perspective). The group of rat GRP cells
appears to be smaller than the group of VIP cells and
positioned within the distribution of those cells. A small
number of NT neurons has been described in the rat SCN in a
distribution that overlaps with the lateral collection of VIP
cells (Moore et al., 2002). Another candidate rat SCN homolog
of the caudocentral hamster SCN subnucleus consists of the
region occupied by CalR cells. These are present in a location
distinctly different from the other cell phenotypes in this
species, overlapping the dorsomedial VP cells to some extent,
but largely occupying a separate region in the dorsal part of
the nucleus (Moore et al., 2002).
As has been noted previously (Morin, 1994), there are substantial species differences in SCN structure. However, there are
at least two organizational characteristics which many species
seemtoshare.ThesearetherespectivegenerallocationsoftheVP
and VIP cell groups. The former tends to occupy a dorsomedial
SCN position, while the latter group sits centrally in the ventral
part of the nucleus (Fig. 1). This spatial characteristic has been
observedintheSCNofrat(Moore,1983),hamster(CardandMoore,
1984), grass rat (Smale and Boverhof, 1999), golden mantled
groundsquirrel(Smaleetal.,1991),Europeangroundsquirrel(Hut
et al., 2002), Octodon degus (Goel et al., 1999), blind mole rat
(Negroni et al., 1997), monkey (Moore, 1993) and human (Moore,
1993).However,theminkandmuskshrewarestrikingexceptions
to this rule as these species have no VP neurons present at the
location normally identified as the SCN (Martinet et al., 1995;
Tokunaga et al., 1992).
Cassone et al. (1988) evaluated comparative SCN organization by analyzing retinal projections to the SCN and the
relative distribution of various peptidergic neurons. They
distinguish “visual” and “nonvisual” regions of the SCN, with
the latter receiving a much reduced density of retinal
projections compared to the former. The “visual” region
usually contains VIP cells, although it is noteworthy that
the specific location of this region varies. In the eutherian
mammals examined (house mouse, guinea pig, cat, pig), the
“visual” region resides in the ventral or ventrolateral SCN.
Among marsupials, there are no VIP cells evident in the SCN
of the kowari or stripe-faced dunnart, while in the bandicoot,
VIP cells are codistributed medially and dorsomedially with
the VP neurons. The retinal projection of the Virginia
opossum and gray short-tail opossum distributes ventrally
and medially in the caudal half of the SCN (Cassone et al.,
1988). In these two species, VIP and VP neurons are
codistributed through the retinorecipient region and medially
in more rostral SCN. Thus, in general, VIP neurons tend to be
associated with the densely retinorecipient region, whereas
the VP neurons are also located there in some species, but not
in others, including most of the eutherian mammals.
Whether these differences in peptidergic cell distribution
reflect the functional organization of the SCN remains to be
demonstrated. It is also worth noting that the laterodorsal
SCN of most mammals is a “specialized” region remarkable
because of the general absence of non-GABA cell phenotypes
(Abrahamson and Moore, 2001; Moore et al., 2002; Silver et al.,
1999; Card and Moore, 1984; Smale and Boverhof, 1999; Smale
et al., 1991; Goel et al., 1999; Cassone et al., 1988).
8
3.3.
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
Intra-SCN connections
Studies of intra-SCN connections fall into two basic categories.
One involves filling individual cells and mapping processes
within the nucleus. The second involves the use of tract
tracers applied iontophoretically or by small volume injection.
The first procedure is limited by the fact that relatively few
cells can be examined, usually only one per SCN. A successful
application of the cell filling technique has demonstrated that
dendritic processes of CALB neurons in the hamster SCN have
a predominantly dorsomedial orientation (Jobst et al., 2004).
This suggests that CALB neurons receive input from dorsomedially located neurons, many (but not all) of which contain VP
or CCK (LeSauter et al., 2002).
The tract tracer application method has the advantage of
labeling more cells, but the disadvantage of usually spilling
label into regions adjacent to the intended target. With that
caveat in mind, small iontophoretic injections of biotinylated
dextran amine (BDA) into the SCN have provided a good initial
appraisal of connectivity within the nucleus. Confocal microscopic analysis of the injected brains shows that most, but not
all, of the cell phenotypes are connected to CALB neurons (Fig.
2), the focus of this particular study (LeSauter et al., 2002).
Particularly noteworthy is the absence of efferent or afferent
connections between CALB and VP neurons. Interconnections
among other pairs of neuron phenotypes have not been
examined.
Use of the Bartha pseudorabies virus as a tract tracer has
provided a novel perspective on intrinsic SCN connectivity. For
example, an injection into the dorsomedial hypothalamus
retrogradely labels, by 48 h postinjection, many neurons in the
dorsal third of the ipsilateral rat SCN (Leak et al., 1999). About
20–24 h later, labeled cells are present throughout much, if not
all, of the SCN. An injection of virus into the subparaventricular zone initially labels cells largely in the ventral two
thirds of the rat SCN, but by 54 h, labeled cells densely
populate the whole nucleus. The data suggest that cells in the
dorsal SCN receive projections from cells in the ventral SCN
and vice versa (Leak et al., 1999). It has not been determined
whether all SCN cells become labeled with virus after injection
into a target of SCN efferents. Comparable studies do not exist
for other species.
4.
In nonmammalian species, the existence of photoreceptors
not specialized for classical visual function has long been
recognized (Gaston and Menaker, 1968; Menaker, 1968; Menaker and Keatts, 1968; Menaker and Underwood, 1976; Enaker et
al., 1970; Underwood and Menaker, 1970). These photoreceptors regulate photoperiodism and entrainment of circadian
rhythms in nonmammals. For the same purposes, mammals
require photoreceptors in the eye (Foster et al., 2003; Yamazaki
et al., 1999). A critical study by Foster et al. (1991) demonstrated
that rd/rd mice, sustaining the near-total loss of rods and cones
along with their respective photopigments, retain substantially normal phase responses to light. In addition, the rd/rd mice
perform similarly to controls even after 767 days, although
there are no electroretinogram responses after 210 days of age
(Provencio et al., 1994). The implication of these studies was
that a novel retinal photoreceptor was sufficient for lightinduced phase shifts in mammals. In 1998, Provencio et al.
(1998a) identified the molecule, melanopsin, and described its
presence in Xenopus brain, eye and photosensitive skin
melanophores. The human and mouse melanopsin genes
were then cloned and demonstrated in cells of the mouse and
monkey retinal ganglion cell layer (Provencio et al., 2000).
4.1.
Fig. 2 – Connectivity of cell types within the hamster SCN.
CCK = cholecystokinin; GRP = gastrin releasing peptide; VIP =
vasoactive intestinal polypeptide; VP = vasopressin. Arrows
indicate the direction the cell types project. Numbers indicate
the percentage of cells with afferent contacts. Dashed arrows
indicate SCN-afferent projections containing serotonin (5-HT)
and neuropeptide Y (NPY) from the median raphe nucleus
(Mn) and intergeniculate leaflet (IGL), respectively. Data
drawn from Table 1 in LeSauter et al. (2002).
Retinal photoreceptors and projections
Melanopsin ganglion cell characteristics
The results of Provencio and colleagues were rapidly adapted
to specifically identify photoreceptive retinal ganglion cells
(Berson et al., 2002) with additional work demonstrating that
melanopsin behaves as a photopigment capable of activating a
G-protein (Newman et al., 2003; Melyan et al., 2005; Qiu et al.,
2005; Panda et al., 2005; Isoldi et al., 2005). Although there is
debate concerning the shape of the action spectrum, the best
indications are that the melanopsin action spectrum peaks at
about 480 nm.
Several studies have been designed to evaluate light
sensitivity of photoreceptive cells contributing to the retinohypothalamic tract (RHT). The results clearly demonstrated
that a special class of ganglion cells projecting to the
suprachiasmatic nucleus (SCN) are both photoreceptive in
the absence of known synaptic input (Berson et al., 2002) and
contain melanopsin (Hattar et al., 2002; Gooley et al., 2001). In
the hamster retina, there are approximately 1600 melanopsincontaining cells (Morin et al., 2003), comprising an estimated
1.5% of all ganglion cells (Tiao and Blakemore, 1976; Hsiao et
al., 1984; Rhoades et al., 1979). In the rat, the estimate is 2.5%
(about 2455 melanopsin cells) and, in the mouse, 1% (about 730
cells) (Hattar et al., 2002). These cells provide the retina with
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
what has been described as a “photoreceptive net” provided by
overlapping, large arbors of beaded dendrites (Berson et al.,
2002; Provencio et al., 2002). Apparently, all portions of the
melanopsin-containing cells are photoreceptive (Berson et al.,
2002). Melanopsin of retinal origin has been described in the
SCN of rats (Beaulé et al., 2003b), although it has not been
observed in the SCN of mouse or hamster (Provencio,
unpublished data; Blanchard and Morin, unpublished data).
In the hamster, the melanopsin cells are spread homogeneously throughout the retina, as they also are in the cat
(Semo et al., 2005). The mouse may have a higher density of
melanopsin in the superior retina (Hattar et al., 2002), a feature
that has been reported in the rat (Hattar et al., 2002; Hannibal
et al., 2002) and the diurnal species, the Nile grass rat
(Blanchard, Novak and Morin, unpublished data).
Nearly all the melanopsin-containing cells are present in
the ganglion cell layer of the retina. However, there is a small
percentage of the cells found as “displaced” to the inner
nuclear layer (Berson et al., 2002). In either case, the dendritic
arbor is most extensive along the border between the inner
nuclear and inner plexiform layers. The predominant location
of the dendrites is in the OFF sublayer of the inner plexiform
layer. The frequency histogram of soma diameters of mouse
melanopsin ganglion shows a mean of about 16.3 μm (Hattar et
al., 2002). In the cat, SCN-projecting ganglion cell diameter is
similar to the rat (17.2 μm) with dendrites ramifying primarily
in the inner plexiform layer (Pu, 1999). The size distribution is
not symmetrical, but skewed toward larger diameters for both
species.
Photoreceptive ganglion cells were first described following
intra-SCN injection of a retrograde tracer. This permitted
identification of ganglion cells contributing to the RHT and
could be combined with recording from those cells in the
absence of input from other retinal neurons (Berson et al.,
2002). With or without the presence of drugs blocking synaptic
input, the cells are slow to respond to light stimuli with
saturating light pulses requiring hundreds of milliseconds, and
near threshold stimuli requiring about 60 s, to response onset.
Latency to maximal depolarization is reduced as stimulus
irradiance increases. Subsequently, the depolarization slowly
decays to a plateau that more or less endures for the duration
of the stimulus, with the plateau proportional to the stimulus
energy. The cells are wavelength-sensitive with a peak
response at about 484 nm, similar to the results from action
spectra for rodent circadian rhythm phase response (Takahashi et al., 1984; Provencio and Foster, 1995). For a fixed
wavelength stimulus, peak depolarization increases linearly
until reaching a saturating irradiance. Spike frequency appears
related to the extent of depolarization. Subsequent to stimulus
offset, repolarization is very slow, taking minutes after stimuli
of high irradiance. Spike discharges are maintained until
polarity returns near baseline. The membrane depolarization
is produced by an inward current that is activated by the light
pulse (Warren et al., 2003). The current activates slowly
reaching a peak several seconds after the application of a
light pulse. The time course of the membrane depolarization
and the inward current are similar. The signal transduction
pathways coupling melanopsin to the activation of the inward
current are not known. The current was unaffected by
substitution of extracellular Na+ and shows both inward and
9
outward rectification. These data are consistent with the
current being mediated by channels of the trp family (transient
receptor potential) of channels (Warren et al., 2003). Properties
of SCN-projecting ganglion cells (identified by retrograde
tracer) recorded from a cat eyecup preparation in which there
was no elimination of rod/cone contribution indicate that they
exhibit sustained responses of the “on” or “on–off” center
variety; have a spectral sensitivity peak at about 500 nm and
response is optimal with motionless or slow-moving stimuli
(Pu, 2000).
4.2.
Retinal ganglion cell projections and bifurcation
Retinal projections to the SCN were definitively described in
1972 (Moore and Lenn, 1972; Hendrickson et al., 1972) and
focused scientific attention squarely on that nucleus as the
probable site of the neural circadian clock. Evaluation of the
RHT has continued as methods have improved and more
species are evaluated. The extent to which the retina projects
to each SCN is highly variable across species. The diurnal
grass rat has bilateral retinal innervation of the SCN, although
a preponderance arrives from the contralateral retina (Smale
and Boverhof, 1999). This differs from what is seen in the
diurnal golden mantle ground squirrel in which RHT innervation is exclusively from the contralateral retina (Smale et
al., 1991). A recent report shows modest ipsilateral SCN
innervation by the retina in the diurnal California ground
squirrel (Major et al., 2003). In contrast, following analysis of
darkfield images, the hamster RHT was described as bilaterality symmetrical (Johnson et al., 1988a). The issue of laterality
and pervasiveness of the retinal projection in the hamster
SCN has now been evaluated with confocal microscopy
(Muscat et al., 2003). The data provide several novel observations. First, nearly the entire SCN is innervated by the RHT,
consistent with earlier HRP results (Pickard and Silverman,
1981). Second, despite the extent of innervation, it is not
homogenous within the SCN. And third, the axons from one
retina project to most or all of the SCN bilaterally. The central
and dorsal SCN receive dense innervation predominantly
from the contralateral retina, while the ventromedial SCN
receives moderate innervation largely from the ipsilateral
retina. Thus, innervation of the SCN by one retina is generally
pervasive, but with clear regional specificity. Of particular
interest is the presence of the densest retinal innervation both
to the region of the CALB-containing central subnucleus of the
SCN and to a region slightly more dorsal. The heavily
innervated dorsal area corresponds to a zone in which
phosphorylated extracellular signal-regulated kinase (pERK)
is found (Lee et al., 2003), a characteristic that is apparently
important to pERK activity.
Other species also appear to have overlapping, but preferred SCN regions of innervation by retinal projections. This is
apparent in the rat, but not the mouse (Shivers and Muscat,
2004). One apparent difference between the RHT terminal
fields of these species and that of the hamster is the
substantially absent innervation of the dorsomedial SCN that
largely corresponds to the area in which most VP cells are
present.
The retina projects to the SCN via the RHT, a major direct
projection, and through a secondary, indirect pathway
10
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
through the IGL and the GHT. In addition, there are
numerous retinal projections to disparate brain regions
many of which have potential, reciprocal connections to
the SCN through the IGL (Morin and Blanchard, 1998). One
such is a direct retinal projection to the dorsal raphe
nucleus, with an initial description in cats given in 1978
(Foote et al., 1978). Interest was further heightened with the
demonstration, based on both anterograde and retrograde
tracing data, of a similar pathway in rat (Shen and Semba,
1994). This pathway has also been observed in gerbil (Fite et
al., 1999, 2003) and O. degus (Fite and Janusonis, 2001). The
pathway has not been found in hamster, although it has
been sought several times (Morin and Blanchard, unpublished data; Fite, unpublished data).
Bifurcation of ganglion cell axons and subsequent projection of these cells to the SCN and IGL (Pickard, 1985) appears to
be a common characteristic of the circadian visual system.
The presence of ganglion cells that bifurcate to innervate both
the SCN and IGL has been recently confirmed, with the
additional information that similar bifurcation is present for
individual cells projecting to both the SCN and superior
colliculus or OPT (Morin et al., 2003). Injection of two
retrograde tracers, one into each SCN, also demonstrates
that at least some ganglion cells bifurcate and send at least
one axon branch to each SCN. Further, at least some of these
cells contain melanopsin, as do some that project to the OPT.
In contrast, individual ganglion cells apparently do not
bifurcate and project bilaterally to the IGL, OPT or the SC.
Other retinorecipient regions have not been simultaneously
examined for both bifurcation and melanopsin cell afferents.
Nor has the question of individual ganglion cells projecting to
more than two visual targets been examined. The prospect is
likely that individual ganglion cells have multiple axonal
processes (beyond two) that would allow a rather small
number of photoreceptors to commonly influence a broad
range of visual functions.
A novel contribution to the analysis of retinal projections
to the SCN comes from two studies by Abe and Rusak (1992).
In the initial paper, hamsters were electrically stimulated in
the IGL with the result that FOS protein synthesis was
induced in the centrodorsal SCN. The preliminary interpretation was that GHT activation was causal to FOS expression
(Abe et al., 1992). However, a subsequent study (Treep et al.,
1995) showed that FOS expression could not be stimulated in
bilaterally enucleated animals. This result suggested an
alternate interpretation, namely, electrical stimulation may
have elicited antidromically activated retinal ganglion cells
which produced orthodromic activation of the SCN via the
collaterals from bifurcating axons in the RHT (Morin et al.,
2003; Pickard, 1985). The resulting pattern of FOS protein
expression would uniquely represent the distribution of
retinal projections that bifurcate and project to both the
IGL and SCN. The extent to which such bifurcating projections originate from ganglion cells that do or do not contain
melanopsin is unknown.
Photoreceptive ganglion cells contributing to the RHT
contain melanopsin (Hattar et al., 2002; Gooley et al., 2001;
Morin et al., 2003; Hannibal et al., 2002; Gooley and Saper,
2003). To date, however, there has not been a comprehensive
description of melanopsin ganglion cell projections to other
brain areas. The initial report of melanopsin cell targets
showed X-gal staining for β-galactosidase activity in a
transgenic mouse strain with tau-lacZ targeted into the
melanopsin gene locus (Hattar et al., 2002). This allowed
visualization of β-galactosidase fused to a protein anterogradely transportable to axon terminals. The blue reaction
product emphatically revealed projections to the SCN, IGL
and olivary pretectal nucleus (OPT). Subsequently, retrograde
tracing studies have confirmed melanopsin-containing ganglion cells projecting to these retinorecipient targets, as well
as to the ventral lateral preoptic area, and the subparaventricular zone of the hypothalamus (Morin et al., 2003; Gooley
and Saper, 2003; Sollars et al., 2003). Melanopsin cells in the
hamster (Morin et al., 2003), but not the rat (Gooley and
Saper, 2003), have been reported to project to the superior
colliculus. The retrograde studies confirm the projections
from melanopsin cells to initially defined major visual
targets, but suggest that not all the ganglion cells projecting
to these areas contain melanopsin. Estimates are that 10–
20% of ganglion cells projecting to the SCN do not contain
melanopsin (Morin et al., 2003; Gooley and Saper, 2003;
Sollars et al., 2003).
The suggestion that melanopsin cells and their projections create a “nonimage forming” visual system has been
recently put under pressure with the observation that at
least some of these cells project to the monkey dorsal lateral
geniculate nucleus (Peterson et al., 2003; Dacey et al., 2005).
These cells appear to provide both irradiance and color
information to the dorsal lateral geniculate nucleus, a site
involving in the processing of feature-related information,
i.e., images. Melanopsin cell projections to dorsal lateral
geniculate nucleus have not been found in the rat (Gooley
and Saper, 2003). As indicated above, the melanopsin cells
are known to project to several other retinorecipient nuclei
and bifurcation may be a common feature of axons
projecting through the RHT to the SCN. Therefore, it is likely
that the melanopsin cells projecting to the dorsal lateral
geniculate nucleus also bifurcate and innervate many, if not
all, other subcortical visual regions.
An unexpected development has arisen from a thorough
evaluation of an engineered version of the Bartha strain
pseudorabies virus (PRV152), as a retrograde transsynaptic
tracer (Pickard et al., 2002). Previously, viral tracing unequivocally demonstrated a subset of retinal ganglion cells following injection into the contralateral eye. These cells
presumably gave rise to the RHT (Moore et al., 1995;
Provencio et al., 1998b; Hannibal et al., 2001a). However, a
new interpretation of these results is based on the fact that
PRV152 is exclusively a retrograde tracer that makes its way
to the SCN multisynaptically through the nucleus of
Edinger–Westphal in the midbrain oculomotor complex
(Pickard et al., 2002). Cells in the SCN, IGL and OPT are
infected, as are several ganglion cell types in the contralateral retina. Only a subset of these contribute to the RHT.
Viral transsynaptic tracing is a powerful method and has the
added advantage of revealing second order retinal neurons,
bipolar and amacrine cells, that synapse with the primary
ganglion cells (Belenky et al., 2003). This observation has
significant implications for the manner in which light
regulates circadian rhythm phase.
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
4.3.
Classical and ganglion cell photoreceptor contribution
to regulation of circadian rhythms, masking, the pupillary
light reflex and melatonin suppression
4.3.1.
Circadian rhythms
Three classes of photoreceptive cell types reside in the
mammalian retina. The current view is that while all effects
of light on the circadian visual system are accounted for by
the three types, no single photoreceptor is necessary for
entrainment. The rd/rd mouse strain suffers postnatal
degeneration of rods and cones. These individuals have
essentially normal phase response to light (Foster et al.,
1991; Provencio et al., 1994; Freedman et al., 1999) and have
an action spectrum for phase shifts that peaks at 480 nm
(Yoshimura and Ebihara, 1996; Hattar et al., 2003) (but see
Provencio and Foster, 1995). Recently, entrainment stability
at threshold levels of illumination was assessed (Mrosovsky,
2003) and, with this simple procedure, only 10% of rd/rd
mice were able to remain entrained to a 12:12 LD
photoperiod when the daytime light intensity was 15
stops (b2 lx), compared to about 80% of wild-type mice.
This result supports the view (Yoshimura et al., 1994) that
there is a circadian system response to light that is
exclusively rod and/or cone-sensitive. Rods may regulate
hamster entrainment under very dim lighting conditions
(Gorman et al., 2005). Neurophysiological methods also
support the view that both rods and cones (primarily 510
nm green and secondarily 375 nm near-ultraviolet) provide
photic information to the SCN (Aggelopoulos and Meissl,
2000).
A flurry of investigations followed the discovery of
melanopsin photoreceptive retinal ganglion cells. Melanopsin gene knockout (KO) mice were made by two different
methods, yet the effects on light-induced rhythm responses
were very similar (Panda et al., 2002; Ruby et al., 2002a). The
animals entrain and free-run properly in normal light–dark
photoperiod and constant dark conditions, respectively.
However, phase response to light is attenuated in the KO
animals. With a 480 nm stimulus, phase shift magnitude
was only about half that of wild-type mice at each of three,
nonsaturating irradiance levels (Panda et al., 2002). A
saturating, white light pulse also yielded a diminished
phase shift from the KO animals, although it was about
two thirds normal (Ruby et al., 2002a). A second defect in
response to light by the melanopsin KO animals was the
failure of constant light (LL) to elicit a properly lengthened
circadian period. In both strains, LL (white light) effectively
increased the period, but in melanopsin KO animals, the
change was only about 55–65% of controls (Panda et al., 2002;
Ruby et al., 2002a). It is worth noting that similar effects on
light-induced circadian period lengthening can be achieved
by IGL lesions (Morin and Pace, 2002), possibly because such
lesions would destroy the functional effectiveness of melanopsin cell projections to the IGL (Hattar et al., 2002). Light
administered at CT16 was able to induce c-fos mRNA in SCN
cells of melanopsin KO animals, although there has been no
test of the normality of this expression (Ruby et al., 2002a).
An important outcome of the melanopsin KO studies is the
implication that both classical photoreceptors and intrinsically photoreceptive, melanopsin-containing retinal ganglion
11
cells contribute, approximately equally, to circadian rhythm
regulation by light.
Subsequently, two sets of investigators, again using
somewhat different methods, addressed the issue of photoreceptor types and circadian rhythm regulation using animals
defective in both melanopsin production and classical photoreceptor function. Again, the results are substantially the
same: animals lacking melanopsin but bearing the rd/rd
genotype (Panda et al., 2003) or undergoing rod and cone
degeneration plus having defective phototransduction in
cones (Hattar et al., 2003) free-run under normal light–dark
photoperiod conditions. Lack of entrainment was not altered
by light intensity nor was circadian period under LL different
from the circadian period under DD. In addition, light was
unable to inhibit nocturnal melatonin synthesis (Panda et al.,
2003). It should be noted that the defects in rhythm response
to light are not the result of loss of ganglion cells that normally
synthesize melanopsin or their projections (Hattar et al., 2003).
4.3.2.
Masking, pupillary light reflex and melatonin suppression
Other accessory visual functions are also modified by the
absence of melanopsin. Although light-induced negative
masking is reduced, particularly at lower light intensities, in
rd/rd mice, the melanopsin KO alone yields masking that is
about 30% greater than in the rd/rd mice (Panda et al., 2003).
When the melanopsin KO and rd/rd traits are combined, the
mice do not exhibit negative masking to light (Hattar et al.,
2003; Panda et al., 2003). Melanopsin does not seem to be
involved in the positive masking that occurs in response to
dim light exposure (Mrosovsky and Hattar, 2003), but apparently contributes to negative masking in the presence of light
≥10 lx (∼6.7 μW/cm2).
Pupillary constriction in response to photic stimuli is much
reduced in rd/rd mice and the response has a longer latency. At
high irradiances, pupillary response equals that in wild-type
mice, but the response of rd/rd mice is markedly attenuated at
intermediate and lower irradiances (Hattar et al., 2003; Panda
et al., 2003; Lucas et al., 2001). Unlike masking and phase
response to light, the pupillary light reflex is relatively
unmodified in the absence of melanopsin. Only at irradiances
N10 × 1012 photons/cm2/s is there a modest sensitivity deficit.
The full effect of light on the pupillary light reflex can be
accounted for by the additivity of rod/cone and melanopsin
photoreceptor contributions (Panda et al., 2003; Lucas et al.,
2003), an observation supported by the absence of any
pupillary light reflex in melanopsin KO animals lacking rod
and cone function (Hattar et al., 2003; Panda et al., 2003). A
single opsin/vitamin A-based photopigment with a peak
sensitivity around 480 nm appears to drive the pupillary
light reflex in rodless, coneless mice (Hattar et al., 2003; Lucas
et al., 2001).
A contribution of the pupil to circadian rhythm regulation
has not been studied and is seldom considered as a factor
regulating circadian rhythm response to light. However, it is
useful to note (a) that both classical and melanopsin photoreceptors contribute to both responses (Hattar et al., 2003;
Panda et al., 2003; (b) that, within limits, phase response
magnitude is proportional to the total number of photons
assessed by the circadian visual system across the duration of
light exposure (Nelson and Takahashi, 1991; (c) that the pupil
12
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
area can vary by a factor of 16 (Hattar et al., 2003; Lucas et al.,
2003) and (d) that the pupil acts to gate exposure of the
circadian visual system to the very stimulus type to which it
optimally detects and responds.
Mice which cannot synthesize melanopsin or which bear
the rd/rd mutation show, to the extent tested, normal lightinduced suppression of pineal melatonin (Lucas and Foster,
1999; Lucas et al., 1999). The two photoreceptor classes appear
redundant to the extent that loss of both eliminates melatonin
suppression by light (Panda et al., 2003).
5.
tract
Neuromodulators of the retinohypothalamic
5.1.
Glutamate
It is generally believed, based on abundant indirect evidence,
that glutamate is the primary neurotransmitter of the
retinohypothalamic tract. This topic (up to 1996) has been
extensively reviewed (Ebling, 1996; Hannibal, 2002a) and will
not be considered in depth here. However, despite the
numerous studies of function involving use of glutamate
receptor agonists and antagonists, there is much less certainty
regarding its actual presence in the RHT. Most convincing in
this regard has been the electron microscopic demonstration
that glutamate immunoreactivity occurs in cholera-HRP
identified terminals of retinal projections to the mouse and
rat SCN (De Vries et al., 1993; Castel et al., 1993). Electrical
stimulation of the optic nerve induces SCN release of [3H]glutamate or [3H]-aspartate (but not [3H]-GABA) loaded onto
rat slice preparations (Liou et al., 1986). However, direct
stimulation of the SCN yields even greater release of the two
amino acids, suggesting their presence in nonretinal terminals and SCN cells, in addition to RHT terminals. Pharmacological studies suggest that aspartate may be the most potent
excitatory neurotransmitter in the rat RHT (Shibata et al.,
1986). Two laboratories have evaluated the uptake and
retrograde transport of [3H]-aspartate injected into the rat
SCN. While the two investigations agreed with respect to
nonretinal projections to the SCN, one identified retinal
ganglion cells contributing to the RHT (Devries et al., 1995)
and the other did not (Moga and Moore, 1996). More recently,
glutamate has been immunohistochemically identified in the
RHT colocalized with PACAP which may identify most, if not
all, retinal projections to the SCN (Hannibal et al., 2000).
Vesicular glutamate transporters, VGluT1 and VGluT2,
account for nearly all the glutamate transporters in the brain
and are generally abundant in retinorecipient nuclei
(Fujiyama et al., 2003; Land et al., 2004), for example, in the
dorsal lateral and ventral lateral geniculate nuclei. However,
they are not abundant in either the IGL (where their presence
is weak, at best) or the SCN. The two transporters are weakly
evident in the SCN, but not at the identical locations (Fujiyama
et al., 2003). Unilateral enucleation has little, if any, effect on
transporter visibility in the SCN, although bilateral enucleation may do so.
One emergent difficulty resulting from the studies of
glutamate function has been the general inability to obtain a
light-type phase response curve (PRC) to direct SCN applica-
tion of excitatory amino acid. The initial study using
glutamate generated a PRC more of the NPY-type (Meijer et
al., 1988), while aspartate had no effect (De Vries and Meijer,
1991). Subsequently, NMDA was shown to elicit phase delays
when administered at CT13.5 and advances at CT19, consistent with a light-type PRC (Mintz and Albers, 1997) and a full,
light-type PRC (although the subjective night is longer and
maximal delays equal maximal advances) to direct SCN
application of NMDA, have been demonstrated (Mintz et al.,
1999). The effects of NMDA on circadian rhythm phase were
blocked by NMDA receptor antagonists. In vivo glutamate
yields a PRC that differs from that for NMDA for reasons that
are not clear.
Injection of NMDA onto the SCN during the early subjective
night elicits light-like phase delays which cannot be blocked
by simultaneous tetrodotoxin (TTX) application (Gamble et al.,
2003). NMDA treatment during the subjective day has no direct
effect on rhythm phase (Mintz et al., 1999), but it can inhibit
phase advances induced by the GABAA agonist, muscimol
(Gamble et al., 2003; Mintz et al., 2002). In this respect, NMDA
during the subjective day mimics the effect of light (Mintz et
al., 2002). However, the action of NMDA during the day is
blocked by TTX treatment suggesting that sodium-dependent
action potentials are necessary for effects of light on rhythm
phase during the subjective day, but not during the subjective
night.
A corollary to the issue of glutamate in the RHT concerns
the biochemical pathway that is likely necessary for processing the neurotransmitter in the SCN. This pathway involves
the de novo synthesis of glutamate in neurons and glia, plus
the recycling of extracellular glutamate through glia back to
neurons (Hertz, 2004). Extracellular glutamate concentration
in the peri-SCN region is greatest during the hours of darkness
(Rea et al., 1993a; Glass et al., 1993). Glial fibrillary acidic acid
concentration in SCN astrocytes is lowest after lights off (Glass
and Chen, 1999).
5.2.
N-Acetylaspartyl glutamate
A significant question that has not been resolved is whether
glutamate is the sole amino acid transmitter in the RHT
contributing to circadian rhythm regulation. N-Acetylaspartyl
glutamate (NAAG) is also present in the RHT (Moffett et al.,
1990, 1991; Tieman et al., 1991), although at the synaptic cleft,
NAAG is converted to its constituent amino acids and each is
available for postsynaptic activity (Baslow, 2000; Neale et al.,
2000; Coyle, 1997). However, unlike NAAG, neither aspartate
nor glutamate is released from retinal ganglion cell terminals
in a calcium-dependent manner (the RHT has not been
specifically tested; see Tieman and Tieman, 1996 for a
review). Whether this constitutes the source of photically
induced glutamate release or the transmitter is independently released by RHT terminals remains to be discovered. The
most recently available information (Moffett, 2003) suggests
the presence of NAAG in a portion, but not all, of the rat RHT
(ventrolateral part of the SCN, i.e., not the whole retinorecipient region). Analysis of prospective NAAG function is
complicated by the presence of numerous NAAG-IR neurons
in the SCN with particularly dense expression in dorsomedial
cells (Moffett, 2003).
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
5.3.
Glutamate receptors
One presumption, given that the RHT is likely to use glutamate
as a transmitter and that glutamate agonists or antagonists
alter circadian clock function, is that there are glutamate
receptors in the SCN. The most comprehensive study of
glutamate receptor localization in the SCN has been performed with hamsters (Stamp et al., 1997) and demonstrates
that the GluR1 receptor is most widely distributed in the SCN
with greatest density in a region corresponding approximately
to that which receives dense innervation from the contralateral retina (Muscat et al., 2003). Both the GluR2/3 and GluR4
receptors are distributed primarily in the dorsomedial SCN,
although there are also substantial GluR4 receptors in the
central and ventral part of the nucleus.
One important consideration regarding glutamate receptors is the fact that they can also be abundant on astrocytes
(Porter and McCarthy, 1997), and may indirectly modulate
neuronal activity by altering glial response to neurotransmitters (Haak et al., 1997). Astrocytes may also be a source of
glutamate that can activate neuronal glutamate receptors (Liu
et al., 2004). Astrocytes may be part of a route yielding
glutamate receptor-activated release of astrocytic endothelial
nitric oxide synthase that can act in parallel with glutamateactivated neuronal nitric oxide synthase through which light
can induce NO release in the SCN thereby altering circadian
rhythm phase response (Caillol et al., 2000).
5.4.
Nitric oxide and RHT signal transduction
Early articulation of the prospect that nitric oxide might
contribute to circadian rhythm regulation was provided by
Amir (1992) who tested whether the SCN mediated the
increased heart rate response to acute photic stimulation.
Infusion of L-NG-nitro-arginine-methyl ester (L-NAME), a
competitive inhibitor of nitric oxide synthase, over the SCN
greatly attenuated the heart rate increase. This attenuation
could be overcome by simultaneous infusion of excess Larginine, the natural substrate for the enzyme. Similar effects
on the heart rate response were also obtained with a
glutamate NMDA-type receptor antagonist and it was suggested (Amir, 1992) that nitric oxide mediates the effects of
light on the SCN.
Subsequently, Ding et al. (1994) published an extensive
series of experiments probing the contribution of nitric oxide
to photic signal transduction in the SCN. As a precursor to the
work, the investigators evaluated phase response of the SCN
neuronal firing rhythm to acute perfusion with glutamate
applied to a rat brain slice preparation. This yielded a very tidy
glutamate-induced PRC (Ding et al., 1994) that closely mimicked the normal rat light PRC (Summer et al., 1984), with
glutamate yielding concentration-dependent phase shifts
across a range of 10−6 to 10−2 M glutamate. The phasic effects
of glutamate were mimicked by in vitro and in vivo application
of NMDA (Ding et al., 1994; Colwell and Menaker, 1992; Colwell
et al., 1990, 1991; Takeuchi et al., 1991), a specific agonist for
one type of glutamate receptor.
Using in vitro activation of the NMDA glutamate receptor
sequelae as the indicator of a functional photic signaling
pathway, the photic effects were mimicked by administering
13
nitric oxide generators to the slice (Ding et al., 1994). Sodium
nitroprusside, S-nitroso-N-acetyl-penicillamine and hydroxylamine application yielded equivalent, glutamate-like phase
advances and delays during the subjective night, but no phase
change during the subjective day. Further, the glutamateinduced phase shifts could be blocked by any of several types
of competitive substrates for nitric oxide synthase. The
procedures were extended to the in vivo situation in which
L-NAME was able to completely block hamster phase advances
induced by light. This blockade was reversed by increasing the
availability of L-arginine substrate (Ding et al., 1994). L-NAME
was also able to attenuate hamster phase delays (Watanabe et
al., 1995). The data suggest that light acting through the RHT
releases glutamate in the SCN, activating the NMDA receptors
on SCN neurons. This increases intracellular calcium concentration, followed by nitric oxide synthase stimulation and
nitric oxide production (Ding et al., 1994). Nitric oxide may act
as a short distance neuromodulator in the photic signal
transduction pathway. The effects of melatonin and serotonin
on rat circadian rhythm phase may also require nitric oxide
synthesis activation in the SCN (Starkey, 1996).
The foregoing basic principles have been linked to lightinduced phosphorylation of the cAMP response element
binding protein (CREB) in SCN neurons (Ginty et al., 1993).
The appearance of phosphorylated CREB (pCREB) is the
earliest known postsynaptic biochemical marker of photic
signal transduction in the SCN. In vivo light-induced pCREB
immunoreactivity in rat SCN cells attained maximal expression within 10 min following a 10 min 150 lx light pulse at CT19
and this could be mimicked by glutamate application to a slice
preparation (Ding et al., 1997). The effects of glutamate on
pCREB immunoreactivity were blocked by an NMDA receptor
blocker (APV) or a nitric oxide synthase inhibitor (L-NAME). As
yet, the steps between postsynaptic glutamate action and
induction of CREB phosphorylation or nitric oxide synthesis/
release have not been determined. It is important to note that
light- or glutamate-induced pCREB and nitric oxide activity are
dependent on circadian clock phase indicating that the
responsiveness of postsynaptic targets of the RHT is limited
at certain times by feedback from the circadian clock. Whether
this means the postsynaptic cells are intrinsically “clock” cells
or they receive rhythmic inhibitory feedback from elsewhere
is not known.
Nitric oxide synthase activity in hamsters is rhythmically
related to the presence of an alternating light–dark cycle. Light
administered at any time of day elevated enzyme activity and
the day–night enzyme rhythm did not persist in DD (Ferreyra
et al., 1998; Agostino et al., 2004; Chen et al., 1997). This is
consistent with the view that the consequences of nitric oxide
release, but not the release itself, are regulated by the
circadian clock.
There is no complete consistency in the literature
concerning nitric oxide function in circadian rhythm regulation. Light-induced FOS protein, thought to be an event
downstream of CREB phosphorylation (Kornhauser et al.,
1990), has been blocked in rat SCN, but not IGL, by peripheral
injection of L-NAME (Amir and Edelstein, 1997). In contrast, the
same procedure in hamsters has blocked light-induced
activity rhythm shifts, but not light-induced FOS in the SCN
(Weber et al., 1995). This may indicate a species difference.
14
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
However, in the hamster, it is further recognition of the fact
that FOS protein expression is not a necessary component of
phase response to light (Colwell et al., 1993a,b).
The anatomical evidence in support of the SCN as a site for
nitric oxide modulation of circadian rhythmicity is not
particularly encouraging. The earliest descriptions of potential
nitric oxide synthesizing substrate based on NADPH-diaphorase staining in the circadian rhythm system revealed no cells
in the rat SCN (Vincent and Kimura, 1992) and sparse cells in
the Siberian hamster SCN (Decker and Reuss, 1994). Occasional neurons immunoreactive for the neuronal form of nitric
oxide synthase have also been observed in rat SCN, and some
of these contain VIP (Reuss et al., 1995). More recently, a dense
plexus of processes immunoreactive for the neuronal form of
nitric oxide synthase has been identified surrounding a few
immunoreactive cells in ventral rat SCN (Chen et al., 1997), but
this has not been seen by other investigators (Caillol et al.,
2000; Wang and Morris, 1996). The most convincing anatomical study indicates no cells in the SCN of rat or mouse
identifiable by NADPH-diaphorase staining, but in both
species, immunoreactivity for neuronal nitric oxide synthase
revealed distinct cell populations within the SCN. In the
mouse, the labeled neurons are evenly distributed through the
SCN with about one tenth of the SCN neurons labeled. In
contrast, all labeled rat neurons are grouped laterally in the
ventral SCN among immunoreactive neuronal processes.
Neither labeled cells nor processes were evident in the
dorsomedial SCN (Wang and Morris, 1996). However, under
special slice conditions, no nitric oxide synthase immunoreactivity was observed in the rat SCN although the ventral
lateral part of the nucleus is densely labeled with cyclic GMP
immunoreactivity. NADPH-diaphorase is not always a useful
indicator of nitric oxide synthase (see Wang and Morris, 1996
for discussion). A subsequent report using similar methods,
but different antisera, identified no cells containing neuronal
nitric oxide synthase in the hamster SCN and a few in the
ventral rat SCN. Nonneuronal cells (probably astrocytes) and
processes immunoreactive for endothelial nitric oxide
synthase were observed in the SCN of both species (Caillol et
al., 2000). Despite the biochemical manipulations indicating
that nitric oxide synthase activity is necessary for phase
response to light or glutamate, mutant mice lacking the gene
coding for endothelial (Kriegsfeld et al., 2001) or neuronal
(Kriegsfeld et al., 1999) nitric oxide synthase apparently
entrain normally to the light–dark cycle and light induces
normal FOS expression in the SCN. As suggested by Caillol et
al. (2000), it is possible that there are parallel neuronal and
glial pathways by which photic information gains access to
the circadian clock. If this is the case, then the only effective
knockout might require deleting the genes coding for both
endothelial and neuronal nitric oxide synthase.
5.5.
PACAP
Pituitary adenylate cyclase activating polypeptide (PACAP) did
not become a molecule of interest to the circadian rhythm
field until 1997 when Hannibal et al. (1997) described its
presence in retinal ganglion cells plus a projection of these
cells to retinorecipient regions of the SCN and IGL. The same
paper demonstrated that, in a concentration-dependent
fashion, PACAP administered at CT6 onto a rat SCN slice
preparation could induce 3.5 h phase advances (tests at CT14
or 19 yielded no phase change). VIP, which can act on
receptors similar to those for PACAP, also yielded phase
advances, but the concentration sensitivity and resulting
phase response were much lower (Hannibal et al., 1997). The
argument was offered that PACAP constitutes a neuromodulator in the RHT which, along with glutamate, normally acts to
modify circadian rhythm phase in response to retinal
photoreception. Subsequently, large, concentration-dependent, PACAP-induced (CT6) phase advances were observed in
hamster SCN slices (Harrington et al., 1999). However,
substantial phase delays were also observed if 1 nM PACAP
was administered at CT14, but lesser or greater concentrations
had progressively diminished effects. In addition, small phase
delays or advances were obtained in vivo following infusion of
1 nM PACAP onto the hamster SCN at CT14 and CT18,
respectively (Harrington et al., 1999). Thus, PACAP in vivo
mimicked the effect of light in terms of the direction of the
elicited phase shift (tests during the subjective day were not
performed).
Replication of the foregoing PACAP effects has proven
elusive. Additional in vivo hamster tests have usually, but
not always, yielded phase delays following administration of
various PACAP concentrations at CT14. Only phase delays
were obtained at CT18, and small phase delays (no
advances) were observed with one PACAP concentration
given at CT6 (Piggins et al., 2001a). Recently, a third
investigation of hamster in vivo effects obtained small
phase delays following a moderately high concentration of
PACAP administered at CT14 (Bergstrom et al., 2003). As is
obvious, there are numerous inconsistencies in the available
data which require reconciliation before a function of PACAP
can be accepted.
The Hannibal lab has developed a monoclonal PACAP
antibody (Hannibal et al., 1995) that has enabled production of
a large amount of information regarding the anatomy of
structures containing this peptide (see Hannibal, 2002a for a
review). PACAP is present in the RHT (Hannibal and Fahrenkrug, 2004; Hannibal et al., 1997, 2002; Bergstrom et al., 2003)
where it colocalizes with glutamate (Hannibal et al., 2000).
PACAP-ir neurons in the rat SCN have also been reported
(Piggins et al., 1996), although there may have been crossreactivity with VIP. In addition, PACAP appears to be found in
axons and terminals in most of the retinorecipient regions of
the rat brain (Hannibal and Fahrenkrug, 2004) including the
IGL, ventral lateral preoptic area and subparaventricular
hypothalamus. Two exceptions may be the commissural
pretectal area and the dorsal lateral geniculate nucleus,
although that information is difficult to discern. PACAP is
widespread in brain (Hannibal, 2002b) and it is not always
clear whether the PACAP observed is actually present in
retinal projections or is from other sources.
PACAP immunoreactivity, in addition to identifying projections to the SCN, identifies a small population of retinal
ganglion cells in the rat most of which are found in the
superior retina (Hannibal et al., 2002). Most, if not all, of these
cells also contain melanopsin and glutamate. Hannibal et al.
(2002) has suggested that all ganglion cells projecting to the
SCN contain both PACAP and melanopsin. However, the issue
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
has not been directly evaluated using retrograde tracers
injected into the SCN. The retrograde tracer procedure has
shown that 10–20% of RHT neurons do not contain melanopsin (Morin et al., 2003; Gooley and Saper, 2003; Sollars et al.,
2003). The apparently close relationship between PACAP and
melanopsin raises the possibility that PACAP released by
melanopsin cell electrical activity may generate prolonged
postsynaptic activity in the SCN that endures beyond the rapid
release effects of glutamate.
The suggestion that all melanopsin-containing ganglion
cells contain PACAP also implies a major role for that peptide
in normal photic regulation of circadian rhythms, assuming it
acts as a neuromodulator. In the context of light and
glutamate effects on circadian rhythm phase, in vivo application of PACAP antibody or receptor antagonists to the SCN
attenuates the phase delaying or advancing effects of light
(Bergstrom et al., 2003; Chen et al., 1999). Ordinarily, glutamate
applied to an SCN slice preparation mimics the effect of light
on the phase of the neuron firing rate rhythm with phase
delays and advances induced during the early and late
subjective night, respectively (Ding et al., 1994; Chen et al.,
1999). When glutamate and PACAP are coinfused onto the slice
preparation, phase delays are enhanced and phase advances
are abolished. In contrast, the PAC1 receptor antagonist,
PACAP6-38, enhanced glutamate-induced phase advances
while blocking phase delays (also blocked by PACAP antibody
infusion). The implication is that glutamate cannot induce
phase delays without cofunctioning PACAP, whereas glutamate is better able to induce phase advances in the absence of
functional PACAP (Chen et al., 1999).
The role of PACAP in circadian rhythm regulation has been
recently addressed through the use of PACAP KO mice (Colwell
et al., 2004; Kawaguchi et al., 2003). The results are mixed, both
within and between studies. Absence of the gene coding for
PACAP does not modify rate to re-entrain to an advanced or
delayed LD12:12 photoperiod, phase response to dark pulses at
CT6, the circadian period in LL or the negative masking effect
of light on wheel running (Colwell et al., 2004). Nevertheless,
the same study showed that, in the PACAP KO mice, both
advance and delay phase shifts to normally saturating or
nonsaturating light stimuli administered at CT16 or 23 are
greatly reduced, the circadian period is shorter in DD and the
phase angle of entrainment is earlier. The second study
(Kawaguchi et al., 2003) failed to find a difference in circadian
period during DD or an effect of the KO on phase delays to light
at CT15, although the phase advances to light at CT21 were
about half of the expectation. To make matters more
confusing, wild-type mice had greater light-induced FOS
protein in the SCN at CT15 than did PACAP KO mice, but not
at the time at which light yielded reduced phase advances
(CT21). At the present time, it is difficult to reconcile the
observation (Colwell et al., 2004) of (a) a large effect of the gene
KO on phase response to light pulses with the absence of an
effect on rate of re-entrainment; (b) the presence of reduced
FOS at a time at which the same stimulus yields a normal
rhythm phase shift; (c) the presence of a reduced phase shift at
time at which the same stimulus yields normal FOS expression and (d) a failure of the two studies (Colwell et al., 2004;
Kawaguchi et al., 2003) to find equally deficient phase delays
in PACAP KO mice.
5.6.
15
Substance P
Confusion also abounds with respect to the presence of SP in
the RHT. In contrast to PACAP, there is more positive evidence
regarding function of SP than there is for its anatomical
presence in the RHT. SP has been suggested as a peptide native
to retinal ganglion cells projecting to the SCN (Takatsuji et al.,
1991; Mikkelsen and Larsen, 1993) and the distribution of SP
fibers and terminals in the rat SCN is similar to that for retinal
input (Piggins et al., 2001b). Other attempts to confirm a SP
projection have not succeeded (Otori et al., 1993; Hartwich et
al., 1994), and the presence of a SP terminal plexus in the SCN
varies substantially across species (Piggins et al., 2001b). A
fairly comprehensive analysis of the rat RHT has demonstrated PACAP and anterograde tracer in the SCN terminal plexus
of the RHT, but apparently there is no colocalization with SP
(Hannibal and Fahrenkrug, 2002). Similarly, SP does not
colocalize with PACAP in the rat retina. According to one
report, most retinal SP cells are present in the inner nuclear
layer and outer part of the inner plexiform layer. The few SP
cells in the rat ganglion cell layer differ in size from the PACAP
neurons and are considered to be “displaced amacrine cells”
(Hannibal and Fahrenkrug, 2002). This contradicts an earlier
report demonstrating SP-containing ganglion cells identified
by retrograde label as projecting to the SCN (Takatsuji et al.,
1991). However, the results of Hannibal and Fahrenkrug (2002)
(Negroni et al., 1997) are consistent with the presence of SP in
the SCN surviving bilateral enucleation (Otori et al., 1993). A
similar conclusion has been reached by other investigators
following bilateral enucleation of either rat or hamster
(Hartwich et al., 1994).
The SP receptor, neurokinin 1 (NK1), is generally absent
from the ventral SCN of both rat and hamster (Mick et al., 1994,
1995), with low levels of NK1 receptor present in neurons
which are found along the dorsolateral margin of the rat SCN.
Another report (Takatsuji et al., 1995) also indicates a general
absence of the receptor from the ventral rat SCN, but
demonstrates its presence on a few cells in the most ventral
part of the nucleus and in the underlying optic tract. Most evident receptor is on neurons and dendrites scattered through
the dorsal and lateral part of the nucleus. Dendrites of the
labeled neurons are described as extending ventrally and, in
enucleated animals, NK1 is found in dendrites contacting
degenerating retinal projections via asymmetric axo-dendritic
synapses (Takatsuji et al., 1995). This suggests that SP-containing retinal fibers terminate on cellular elements in the SCN
with NK1 as a receptor. A detailed evaluation of NK1 receptor
distribution in the rat, mouse, golden and Siberian hamsters
generally confirms the above observations as holding across
species (Piggins et al., 2001b). The major point of difference is
the absence of receptor from the ventral SCN in all species
except the rat. In all species, the SCN is surrounded by a clear
margin of dense NK1 receptor and there is a distinct cluster of
cells identifiable by NK1 immunoreactivity just inside extending to just outside the dorsolateral border of the SCN.
SP is present in fibers and terminals of the rat, mouse,
golden and Siberian hamster IGL (Morin et al., 1992; Piggins et
al., 2001b), although the origin of this input is not known. The
NK1 receptor is densely distributed throughout the entire IGL
of the same species (Piggins et al., 2001b).
16
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
With respect to function, SP yielded increased 2-deoxyglucose uptake in the rat SCN during the subjective night, but not
during the day. It also produced a light-like PRC in the action
potential rhythm (Shibata et al., 1992; Hamada et al., 1999; Kim
et al., 2001). Both effects were blocked by the SP receptor
antagonists. Cycloheximide also blocked SP-induced phase
shifts in the slice preparation (Shibata et al., 1994). About 43%
of SCN neurons were excited by application of SP to rat brain
slices (11% inhibited), but there was no night/day difference
(Shirakawa and Moore, 1994). Similar results have been found
in the hamster SCN (Piggins et al., 1995a). In both species, SP
modulated neuronal response to glutamate (Shirakawa and
Moore, 1994; Piggins et al., 1995a). SP also induced glutamate
release in rat SCN slices (Hamada et al., 1999). In optic nerve
stimulated slices, an SP antagonist reduced excitatory postsynaptic currents in rat SCN neurons. The effect on NMDA and
non-NMDA receptor-mediated responses was similar (Kim et
al., 1999). However, SP-induced phase delays could be blocked
by NMDA or non-NMDA receptor antagonists, whereas SP
antagonists were unable to block glutamate-induced phase
delays (Kim et al., 2001). The latter results are interpreted as
indicating that SP works upstream of glutamate.
Intracranial SP application to the hamster SCN in the early
subjective night elicited small phase delays. The effects were
not dose-dependent in the pico/nanomolar range and were not
seen at other circadian times (Piggins and Rusak, 1997). The use
of SP receptor agonists or antagonists has yielded more
consistent and sometimes larger effects. Light-induced FOS
was significantly reduced by pretreatment with the nonspecific SP antagonist, spantide. The reduction was concentration-specific and varied according to SCN region. Greater
reduction occurred in the dorsocaudal SCN than in the
ventrocaudal SCN and in the rostral SCN than in the caudal
SCN (Abe et al., 1996). The authors compare the effects of
spantide to those of NMDA and non-NMDA antagonists with
respect to the distribution of light-induced FOS and concluded
that the glutamate receptor antagonists were involved in light
responsiveness of the ventrocaudal SCN, whereas SP
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
contributed to light responsiveness of dorsocaudal SCN
neurons. Intraperitoneal injection of an NK1 receptor antagonist had no effect on hamster rhythms in DD, but in LL, phase
advances were obtained following treatment at CT8 (Challet et
al., 1998a, 2001). These were not associated with increased
wheel running at the time of injection. NK1 antagonists had no
effect on light-induced phase delays at any of three intensities
(Challet et al., 1998a, 2001). However, at CT19, they attenuated
light-induced phase advances by about 30–45%, if administered prior to the light pulse. The antagonists were unable to
block the effect of light on phase response if administered after
the light pulse (Challet et al., 2001). The effect of spantide on
rhythm regulation in hamsters has apparently not been
investigated.
6.
SCN structure–function relationships
The structure–function relationship that appears to be emerging is one of both time and space. That is to say, the phases of
intrinsic oscillatory events vary according to SCN region.
Attempts to refine the designation, “region,” according to cell
phenotype are progressing, but must be considered as fluid. As
is discussed below, the actual number of “regions” in the SCN
is highly debatable (2, 3 or even 4 for the rat; Figs. 3A–E), even
within a single species, as is the correspondence of a nominal
region to a particular cell type or organizational attribute.
Perhaps the dominant question pertaining to function of
SCN neurons concerns the extent to which each is an
oscillator. The prevailing view, until fairly recently, has been
that the SCN is composed of many intrinsically oscillatory
neurons which interactively synchronize to yield a monophasic rhythm (Herzog et al., 1998; Liu et al., 1997a). Gillette
17
et al. (1993) provided evidence, using extracellular recording
methods, that separated dorsomedial and ventrolateral parts
of the rat SCN oscillated more or less equally in vitro,
although amplitude of the rhythm from the ventrolateral
region was reduced. This early attempt to distinguish
regions of functional significance in which one SCN area
differs from another with respect to oscillatory spike
discharge provided a negative result, but presaged a topic
that has evolved along with the availability of molecular
biological tools. More recently, more refined methods have
allowed greater detail to be gleaned from individual SCN
slices (Schaap et al., 2001, 2003). Subtle differences in phase
and waveform are apparent both between the bilateral SCN
and across regions within a single rat SCN.
Whereas most attempts to evaluate rhythmicity of SCN
cells have been done with reference to location or phenotype,
Saeb-Parsy and Dyball (2003) evaluated cells based on their
targets of projection. Cells projecting to the arcuate and/or the
supraoptic nucleus had two peaks in firing rate, one in the
early night and the other in the late night, in contrast with
cells projecting elsewhere.
As described below (Section 6.1.3), there is accumulating
evidence that at least some cells may not oscillate. However,
attempts to identify individual SCN pacemaker cells, while
successful in doing so, have not been able to provide
information regarding the percentage of SCN cells that are
intrinsically oscillatory or their particular cell characteristics
(Herzog et al., 1997, 1998; Liu and Reppert, 2000, Liu et al.,
1997a; Welsh et al., 1995; Aujard et al., 2001). Three generalities
arise from these studies (Liu and Reppert, 2000). One is that the
behavior of oscillatory cells evinces a wider range of intrinsic
circadian periods if the cells are isolated from one another
than if they are coupled. The second is that GABA is able to
Fig. 3 – Suggested organization of the SCN varies according to species, rostrocaudal location within the SCN, time of day,
laboratory, criteria for region designation and the evolution of ideas. RAT: (A1 is rostral to A2) Structure using the “core” and
“shell” nomenclature (Leak and Moore, 2001). Note fairly substantial differences in position that depend on the level within the
SCN. (B) Geographical structure using “dorsomedial” (DM) and “ventrolateral” (VL) designations. Drawn from histology in
Nagano et al. (2003). The area VL′ set off by the dashed line has been added to indicate a portion of the SCN that behaved
differently from either the DM or VL, but was not acknowledged by the authors. (C1, C2) Divisional structure as suggested from
the variation in density of FOS-IR cells (dots). Drawn from the histology in (Beaulé et al., 2001b). Note that the smaller number of
FOS-IR cells within the middle of the 3 divisions of (C2) results from light exposure. The dashed line in panels C1 has been added
to demarcate a lateral region that did not appear to vary with respect to FOS induction. (D) Three divisions (dorsomedial
periventricular, dorsomedial central, ventrolateral) derived from the expression of Per1 (Yan and Okamura, 2002). Dashed line
has been added to set off a region that varies minimally with respect to Per1 expression. Compare to panels C1 and C2. (E) Three
divisions (dorsomedial, ventrolateral main, ventrolateral medial) derived from the expression of VIP-IR and Per1. The VLmed
receives sparse or no retinal innervation (Kawamoto et al., 2003). MOUSE: (F) Expression of Cry1-IR (dots;
F1—ZT23, F2—ZT14-light induced) and Per1 (dots; F3—ZT23, F4—ZT14-light induced). Drawn from histology in Field et al.
(2000). The circular central area was drawn to encompass most of the Cry1-IR cells and held constant in panels F2–F4. Note that
light-induced Per1 tends to be absent from the region of rhythmic Cry1 expression (F1—ZT23), but has substantial overlap with
the area showing rhythmic Per1 expression (F3—ZT23). (G) Regions indicated by antiphase rhythmic expression of Per2. Drawn
from histology in (King et al., 2003). The central area in panel G1 was drawn to encompass most of the Per2-IR cell nuclei (dots;
CT0) and retained in panels G2 which shows the widespread expression of Per2 in nuclei (dots) at CT12. (H) Compartments
indicated by the location of VP-IR cells and fibers or GRP cells as indicated by a green fluorescent protein signal. Drawn from
Karatsoreos et al. (2004). (I) Solid lines diagram the mouse SCN “core” and “shell” according to one formulation (Yan and Silver,
2004). The dashed line indicates the border of the “core” as presented in another formulation (Abrahamson and Moore, 2001).
HAMSTER: (J) SCN and CALB-identifiable central subnucleus (drawn from Hamada et al., 2001). (K) Cells expressing VP mRNA
(dots) in the rostral SCN at (K1) CT20 and (K2) CT4 or in the caudal SCN at (K3) CT20 and (K4) CT4 (drawn from histology of
Hamada et al., 2004b), outlines included, with the enclosed ventrolateral area designating the region containing CALB neurons
in this study).
18
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
shift the phase of individual, noninteracting SCN neurons in a
phase-dependent manner. Third, the phases of oscillations
among a collection of independent cells are synchronized by
repeated bath application of GABA at the same time every day.
There is no information concerning the cell phenotype
whether from hamsters (Herzog et al., 1998), rats (Welsh et
al., 1995) or mice (Liu and Reppert, 2000; Liu et al., 1997a).
The ability to evaluate the simultaneous rhythmic behavior of multiple individual cells has provided a new perspective on the oscillatory organization of the SCN. In mouse SCN
with a green fluorescent protein (GFP) reporter of Per1 gene
activity (Kuhlman et al., 2000), most neurons oscillate in
synchronized fashion, but with numerous distinct phases
(Quintero et al., 2003). An estimated 11% (LD) and 26% (DD) of
the cells evaluated were judged as nonrhythmic. Unexpectedly, cells tend to be grouped, having one of three preferred
phases that differ if the animals have been entrained or freerunning. In SCN obtained from mice in LD, 78% of neurons
had peak GFP expression at ZT5, ZT8 or ZT11 (most cells
peaked at ZT8). With a history of DD, most cells peak at CT12
and 92% had peaks in the CT12, CT15 or CT18 phase groups.
The half-maximum phase plot indicates CT12 for the DDhoused animals and ZT8 for the LD housed animals. Several
valuable observations are evident in this study (Quintero et
al., 2003): (1) not all SCN neurons are rhythmic; (2) different
rhythmic neurons have different phases; (3) there are three
preferred phases of rhythmic SCN neurons, a characteristic
previously observed using multiunit in vivo electrophysiological methods (Meijer et al., 1997); (4) the preferred phase of
rhythmic neurons varies according to photoperiod history.
The results support the conclusion that, assuming clock
genes actually determine the pacemaker properties of SCN
neurons, other factors control the phase relationship between the locomotor rhythm and circadian clock phase. This
is demonstrated by the fact that the preferred phase of gene
expression is about 4 h earlier in LD housed mice compared
to DD housed mice when assessed relative to activity onset
(Quintero et al., 2003).
An exception to the above evidence concerning the timing
of oscillations in SCN neuronal activity has been observed in
the hamster. Whereas there is a neuronal activity rhythm
peak recorded approximately once every 24 h if the SCN tissue
is cut in the coronal plane, there are two distinct peaks if the
SCN is cut in the horizontal plane (Jagota et al., 2000). The
phases of the two peaks are modulated by photoperiod history
(Mrugala et al., 2000). This phenomenon appears to apply only
to hamsters, and has not been seen in mouse or rat (Burgoon
et al., 2004). These results support the view that there are
functionally distinct regions within the SCN; different regions
oscillate differently; and the “standard” perspective (here, the
coronal view) is not always the fully correct and adequate
perspective.
6.1.
Function of cellular neuromodulators in the SCN
6.1.1.
GABA
As indicated above, GABA is the predominant neurotransmitter in cells of the rat, mouse and hamster SCN. It is, therefore,
not a surprise that manipulation of GABA neurotransmission
in the SCN alters circadian rhythm function. One potential
complication to the interpretation of the functional analyses
of GABA in the SCN is the presence of a GABAergic afferent
projection from the IGL (Morin and Blanchard, 2001).
The basic observation is that peri-SCN administration of
the GABAA agonist, muscimol, substantially reduces the
magnitude of light-induced phase shifts, while similar application of the GABAA antagonist, bicuculline, substantially
increases phase shift magnitude (Gillespie et al., 1996). These
results are inconsistent with prior data obtained using
systemic drug injections (Ralph and Menaker, 1987, 1989).
Nevertheless, the effects of intracranial (i.e., SCN) drug
application appear to be repeatable and consistent, although
the effect of bicuculline may depend on circadian time
(Gamble et al., 2003; Mintz et al., 2002; Gillespie et al., 1997).
Similarly, bicuculline augments and muscimol suppresses
light-induced Fos protein in the hamster SCN (Gillespie et al.,
1999).
The GABAB receptor may also be involved in modulation of
light-induced phase shifts. Peri-SCN injections of a GABAB
receptor agonist, baclofen, block light-induced phase delays
and advances, but the antagonist, CGP-35348, has no effect
(Gillespie et al., 1997). Baclofen also blocks glutamate-induced
phase delays and this action is not affected by TTX (Mintz et
al., 2002). The GABAB agonist also effectively blocks lightinduced Fos at CT13.5 and CT19 (Gillespie et al., 1999),
although an antagonist has no effect. The effect of GABAB
receptor activation is the result of an inhibition of glutamate
release from the RHT terminals (Jiang et al., 1995).
The GABAA agonist, muscimol, is also able to induce phase
advances when administered to DD-housed hamsters during
the middle subjective day (Novak et al., 2004; Smith et al., 1989)
and this effect is inhibited by a concomitant light pulse (Mintz
et al., 2002).
6.1.2.
VIP and VIP/PACAP receptors
VIP neurons are clustered in the ventral SCN of most
mammalian species and use VIP as a neuromodulator to
communicate with other neurons in the SCN through activity
at the PAC1 or VPAC2 receptor, both of which are found in the
SCN (Cagampang et al., 1998a,b). A complicating factor in the
analysis of receptor function in the context of the circadian
system is the fact that both receptors bind VIP and PACAP with
approximately equal affinity. PACAP-containing terminals are
found in projections from the retina, as well as from other,
unidentified brain regions (Hannibal and Fahrenkrug, 2004;
Hannibal et al., 1997). VIP peptide has been implicated in the
regulation of circadian rhythm phase and period, although
conflicting observations have been reported (Gillespie et al.,
1996; Piggins et al., 1995b; Albers et al., 1991). More recent work
has focused on function of the PAC1 and VPAC2 receptors with
respect to circadian rhythm regulation and the results have
been quite provocative.
Calcium imaging methods suggest that the effects of
PACAP on SCN neurons are elicited through the PAC1 receptor
because of the absence of response to equimolar VIP
application (Kopp et al., 1999). The PAC1 knockout mouse
tends to have a shorter circadian period in DD than wild-type
and light-induced phase shifts are less after stimulation at
certain circadian times compared to the wild type (Hannibal et
al., 2001b). Fos protein and Per1/2 gene induction were also
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
much less in the knockout animals following a light pulse
during the early subjective night, although they were not
different during the late subjective night.
The effects of PAC1 receptor knockout are small compared
to the consequences of losing a functional VPAC2 receptor.
The latter appears to be an essential component of the
circadian rhythm system, at least in mice. Its absence in the
Vipr2 KO mouse is associated with nearly complete loss of the
circadian locomotor rhythm assessed in constant dim red
light (Cutler et al., 2003). Similarly, rhythms in SCN gene
expression (Per1, Per2, Cry1, VP were lost, although a vastly
attenuated, but statistically significant, rhythm in BMAL1
expression was evident (Harmar et al., 2002; Cutler et al., 2002).
In the Vipr2 KO mice, there was also very little light-induced
FOS protein compared to what is normal in the wild type. In
the SCN slice preparation, the ensemble circadian rhythm in
neuronal firing rate is lost in the Vipr2 KO mouse (Cutler et al.,
2003). An additional feature of this study was the demonstration that a specific VPAC2 receptor antagonist had the same
effect on the ensemble neuronal firing rhythm of wild-type
animals as the gene defect does on the Vipr2 KO animal. In
contrast to the gene knockout model, a transgenic mouse
overexpressing the VPAC2 receptor has a much longer
circadian period and appears to be more sensitive to light as
indicated by faster re-entrainment to an advance of the light–
dark photoperiod (Shen et al., 2000).
At the level of individual SCN cells, there is also an absence
of rhythmic gene expression in Vipr2 KO mice (Harmar et al.,
2002). An explanation for the function of the VPAC2 receptor
within the context of circadian clock structure and intercellular pacemaker regulation has not been determined, although the suggestion has been made that rhythmic VIP
serves to maintain coupling between oscillating neurons in
the SCN (Harmar et al., 2002). This explanation does not
account for the fact that about 15% of the congenitally
anophthalmic ZRDCT-An mouse strain are arrhythmic despite
the presence of VIP neurons in the SCN (Laemle and
Ottenweller, 2001). Thus far, there is a demonstrable requirement for the presence of the VPAC2 receptor in order for mice
to display circadian rhythmicity, but no demonstration that
VIP itself is explicitly necessary or greatly involved in rhythm
generation or regulation.
Nearly half the mouse SCN neurons immunoreactive for VP
also have the VPAC2 receptor as do about 31% of the VIP and
CalR neurons (Kallo et al., 2004). In general, there is a high
degree of regional correspondence between the distribution of
mouse neurophysin (marker for VP) and for PHI-containing
(marker for VIP) neurons with the distribution of cells
containing VPAC2 receptor (King et al., 2003), although the
degree of colocalization was not established in this paper. The
regional correspondence is not perfect as a central zone in the
mouse SCN is devoid of VIP or VP neurons, but contains a few
cells with VPAC2 receptors.
The specific importance of VIP for circadian rhythms has
been emphasized in mice (Colwell et al., 2003; Aton et al.,
2005). VIP neurons project widely from the SCN to various
targets (Kalsbeek et al., 1993), but also appear to have intraSCN connections (LeSauter et al., 2002; Daikoku et al., 1992;
van den Pol and Gorcs, 1986). Two thirds of cells in the
mouse SCN have spontaneous inhibitory postsynaptic
19
potentials that are enhanced by VIP. These cells are
distributed throughout the SCN with no discernible regionrelated pattern (Itri and Colwell, 2003). VIP infused into the
hamster peri-SCN region or onto the rat SCN slice during the
early subjective night induces phase delays, while infusions
during the late subjective night yield phase advances, with a
PRC similar to that for light (Piggins et al., 1995b; Albers et
al., 1991; Reed et al., 2002) (although there is controversy
concerning whether or not PHI and GRP are necessary for
obtaining phase shifts). In mice lacking genes coding for PHI
and VIP, a variety of rhythm-related measures were altered.
In particular, the circadian period was shorter by about an
hour, phase response to light was greatly reduced or
eliminated, there was more variability in activity onset
under LD and cohesive rhythms were generally not maintained under DD (Colwell et al., 2003). Mice lacking only the
gene coding for VIP had similar characteristics. Most notably,
about two thirds of the animals had running rhythms in DD
that expressed multiple noncircadian components. Importantly, the behavior of VIP KO mice was strikingly similar to
mice lacking the gene (Vipr2) coding for the VPAC2 receptor
(Aton et al., 2005). The importance of VIP to normal
rhythmicity in the VIP KO mice has been established by
reinstating SCN rhythmicity and synchrony among SCN
neurons by periodic application of VIP agonist to a tissue
culture preparation, but this has no effect on VPAC2 KO mice
(Aton et al., 2005).
6.1.3.
SCN
Calbindin and the central subnucleus of the hamster
There has been emphasis on the central subnucleus of the
SCN and the role its constituent CALB cells may play in
circadian rhythm regulation. A subset of SCN neurons
expresses the Ca2+ binding protein CALB (Arvanitogiannis et
al., 2000; LeSauter and Silver, 1999). CALB contains six highaffinity Ca2+ binding domains and contributes to intracellular
Ca2+ signaling and buffering (Leathers et al., 1990; Roberts,
1994). As described above, the CALB neurons are clustered
within the caudocentral SCN. This region contains about 250
CALB neurons and is approximately 300 μm long, 180 μm high
and 130 μm wide (Silver et al., 1996a).
Partial lesions of the SCN that completely destroyed the
CALB region yielded animals with arrhythmic locomotor
activity. In contrast, if the lesions spared a portion of one or
both CALB regions, behavioral rhythmicity was maintained
(LeSauter et al., 1996). The circadian locomotor rhythm is
restored after implantation of fetal SCN tissue into the third
ventricle of previously SCN-lesioned adult hamsters (Ralph et
al., 1990). Restoration of rhythmicity requires the presence of
CALB neurons within the transplanted grafts and the strength
of the restored locomotor rhythm is correlated with the
number of surviving CALB neurons (LeSauter and Silver,
1999). Small lesions of the SCN that damage the CALB region
bilaterally eliminate a number of behavioral and physiological
rhythms. If the CALB region on either side was partially or
completely spared, the rhythms remained intact (Kriegsfeld et
al., 2004). These data suggest that the region of the SCN
containing these CALB positive neurons is important in the
generation of circadian rhythms. An important question to be
answered is whether it is the CALB neurons (or other neurons
20
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
in this region) or whether it is the network connections
between the cells in this region that are critical for the
generation of circadian rhythms.
The central subnucleus receives dense input from the
retina via the retinohypothalamic tract and the CALB neurons
are directly contacted by retinal projections (Muscat et al.,
2003; Bryant et al., 2000). A light pulse during the subjective
night increased FOS expression in the majority of CALB
neurons of the hamster SCN. These data led to the development of a model in which the CALB neurons serve as a “gate”
for photic input to the SCN (Antle et al., 2003a). This model
must be reconciled with anatomical data demonstrating that
the RHT projects to all regions of the hamster SCN and the fact
that the densest retinal terminal field extends well beyond the
CALB subnucleus (Muscat et al., 2003). In contrast to results in
the hamster, CALB and FOS colocalization are rare in the grass
rat (Mahoney et al., 2000). Destruction of CALB neurons in the
rat SCN does not alter light-induced FOS expression, freerunning rhythms or photic entrainment in that species
(Beaulé and Amir, 2002).
Neurons in the hamster central subnucleus do not show a
rhythmic expression of messenger RNAs for Per1 and Per2
(Hamada et al., 2001, 2004a). However, light can induce the
expression of both these clock genes in the central subnucleus. This has led to the conclusion that the central
subnucleus is a nonrhythmic region of the SCN, while the
dorsomedial region corresponding to the general location of
VP neurons is rhythmic because the cells in this area express
Per1 and Per2 in a rhythmic manner. However, it may not be
possible to determine the rhythmicity of individual neurons
with in situ hybridization techniques if only a portion of the
neurons in a region are rhythmic. Further, electrophysiological evidence suggests that at least some neurons in the
central subnucleus fire action potentials in circadian manner.
The mean spontaneous firing rate (SFR) of non-CALB neurons
in the central subnucleus has a peak at ZT6 and a nadir at ZT19
(Jobst and Allen, 2002). Whether the rhythmicity of these nonCALB neurons is driven by endogenous clocks located in
neurons of the central subnucleus or whether it is due a
network phenomena of interconnections between rhythmic
and nonrhythmic neurons is an important area for future
research. In contrast, CALB neurons had a mean SFR of less
than 1 Hz and showed no circadian pattern of action potential
firing (Jobst and Allen, 2002). It is noteworthy, but of unknown
significance, that some CALB neurons contact a neighboring
CALB or non-CALB cell via gap junctions (Jobst et al., 2004). In
the mouse, increased firing frequency is correlated with
increased expression of Per1 (Kuhlman et al., 2003). This
suggests that CALB neurons are a functionally distinct
subpopulation within the SCN that do not contain a rhythmic
circadian clock. The data raise the intriguing possibility that
the interactions between rhythmic and nonrhythmic neurons
are required to generate circadian locomotor rhythms. There
is also the untested possibility that the GRP, SP and CALR
neurons that occupy the central subnucleus along with the
CALB cells are likewise nonrhythmic.
CALB expression is dynamic depending on the genotype,
developmental age and the lighting conditions. Tau hamsters
have significantly more CALB expressing neurons than wildtype hamsters (LeSauter et al., 1999). In mice, there is
significantly more CALB expression in young animals that
gradually decrease as the animals approach adulthood (Ikeda
et al., 2003).
CALB expression is inversely correlated with the duration
of the light cycle (Menet et al., 2003). Hamsters maintained on
short duration photoperiods had significantly more CALB
expression than those maintained on long photoperiods
(Menet et al., 2003). Hamsters reared in DD or short photoperiod conditions had more CALB than controls (Tournier et al.,
2003). Similarly, rats maintained in LL conditions had fewer
CALB-containing neurons then in rats maintained in DD
(Arvanitogiannis et al., 2000). Neonatal monosodium glutamate (MSG) exposure increased the number of CALB-expressing SCN neurons and prevented the disruption of circadian
rhythms produced by LL in rats (Arvanitogiannis et al., 2000).
The parallel effects of MSG on CALB and LL changes on
rhythms suggest that CALB may be important in the regulation of response to LL.
CALB appears to play a key role in the photic responses in
hamsters. CALB is present in the cell nucleus and cytoplasm
of central subnucleus SCN neurons during the day, but is
absent from the cell nucleus during the night (Hamada et al.,
2003). Reduction of CALB expression by antisense oligonucleotides blocked light-induced phase shifts during the
night, but light could produce shifts of both behavioral and
Per rhythms during the day (Hamada et al., 2003). Interestingly, these effects occurred despite the fact that the CALB
mRNA was only reduced 38% and the CALB protein only
reduced 28% (Hamada et al., 2003). Also interesting was the
rapidity of the effect, which occurred only 4 h after the
antisense injections. Therefore, while the CALB neurons do
not have rhythmic clock gene expression or show rhythms
in action potentials, CALB expression in the nucleus may be
controlled in a circadian manner. Thus, these cells appear to
be rhythmic according to at least one criterion, but not by
others. The mechanisms controlling this cellular regulation
are unknown. They may involve rhythmic feedback from the
dorsomedial “rhythmic” SCN or may result from the
interaction between rhythmic and nonrhythmic neurons in
the central subnucleus.
The central subnucleus is heterogeneous and CALB cells
constitute a minority of the neurons (LeSauter et al., 2002;
Jobst and Allen, 2002). A better understanding of mechanisms
underlying intercellular communication between non-CALB
and CALB neurons and how they respond to synaptic input
will be important to understand how the SCN generates a
functional circadian output. Double-label methods have
demonstrated that 94% of CALB neurons also contained
GABA (Jobst et al., 2004). Of note, most of the cellular profiles
in the central subnucleus contain GABA, although many GABA
neurons in the central subnucleus did not contain CALB. In
addition to CALB and GABA colocalization in the central
subnucleus, glutamate decarboxylase 67 (GAD67), one of two
related isoforms of the synthetic enzyme for GABA, was
detected in CALB cell bodies. Immunoreactivity for the GABAA
receptor α2 and β2/3 subunits was prominent in the SCN (Jobst
et al., 2004). Confocal microscopic analysis demonstrated that
β2/3 and α2 did not colocalize with CALB. Labeling for the β2/3
subunits was diffusely distributed around CALB neurons,
whereas labeling for the α2 subunit appeared as discrete
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
puncta. The data indicate that α2 and β2/3 subunits contribute
to the composition of GABAA receptors in the CALB region. A
limitation of this study is that the GABAA receptor subunits
were not localized to either postsynaptic or presynaptic
regions. GAD65, an isoform of GAD exclusively located within
synaptic terminals (Erlander et al., 1991), is expressed in a
punctate pattern throughout the SCN, including the central
subnucleus (Jobst et al., 2004).
6.1.4.
Bombesin and gastrin releasing peptide
Bombesin contains 14 amino acids, GRP has 27 and they share
a 7 amino acid sequence homology. There is a very small
literature concerning the effects of bombesin on SCN function.
Immunoreactive cells are abundant in the Djungarian hamster (Reuss, 1991), although cross-reactivity with GRP is a
potential problem (Panula et al., 1984). Bombesin increased the
firing rate of about 50–60% of golden hamster neurons studied,
while inhibiting the rate in half of the remainder (Piggins and
Rusak, 1993; Piggins et al., 1994). Neuromedin B, another in the
class of bombesin-like peptides, activated about 40% of SCN
neurons and inhibited very few (Piggins et al., 1994). The
responses of individual cells to application of several different
bombesin-like peptides were similar in more than 80% of the
tests. Responses to the peptides were attenuated by pretreatment with antagonists to the GRP-preferring receptor which is
present in the SCN (Ladenheim et al., 1992, 1993). More cells
were activated by GRP and neuromedin B at ZT12-16 than at
ZT4-8 (Piggins et al., 1994). Bombesin has not been similarly
tested.
GRP was originally reported to induce light-like phase
shifts in hamster only if infused onto the SCN along with VIP
and PHI (Albers et al., 1991), but may be equivalently
functional if applied alone (Piggins et al., 1995b). The action
potential rhythm in rat or hamster slice preparations
exhibited large, light-like phase responses to GRP and
these effects could be blocked by bombesin 2 receptor
antagonists. Bombesin 2 and bombesin 1 receptors were
identified in SCN, although there is no evidence that
bombesin 1 contributes to rhythm regulation (McArthur et
al., 2000). GRP injected into the hamster third ventricle at CT
13 induces sizable phase delays in association with induction of Per1, Per2 and FOS (Antle et al., 2005). Interestingly,
the gene expression occurs almost exclusively in the densely
retinorecipient region in the dorsal SCN, just above the
central subnucleus identified by CALB cell presence (Muscat
et al., 2003). This is the same area in which pERK is
expressed, dependent upon an intact RHT, during the
subjective night (Lee et al., 2003). And, phosphorylation of
ERK is apparently necessary for phase response to GRP
because if phosphorylation is pharmacologically blocked, the
phase shifts are also blocked (Antle et al., 2005). Although
spontaneously rhythmic ERK phosphorylation does not
occur in the absence of the RHT, the effect of enucleation
on phase response to GRP has not been examined.
In the mouse, GRP infused into the peri-SCN region at
CT16 elicited small phase delays and induced Per and Fos
gene expression (Aida et al., 2002). GRP receptor was also
found throughout most of the SCN, although density was
clearly greatest in the dorsocaudal part of the nucleus. Mice
lacking the GRP receptor had normal phase delays in
21
response to a 15 min 30 lx light pulse at CT16, but unlike
wild-type animals, the delays were not greater if the light was
300 lx. Small (approximately 20%) decrements were also seen
in Per2 (but not Per1) and Fos response to the 300 lx light (Aida
et al., 2002).
In ovariectomized, estrogen-primed rats, pituitary prolactin is released in a daily surge timed by the SCN (Mai and Pan,
1993, 1995; Freeman et al., 2000). Properly timed bombesin
infusion into the peri-SCN region blocks the prolactin surge
(Mai and Pan, 1995). The implication is that bombesin neurons
participate in the efferent control of prolactin release timing.
This raises the general issue of whether individual SCN cell
phenotypes are concerned with the control of single or
multiple efferent systems.
6.1.5.
Neuromedin U and neuromedin S
Neuromedin U (NMU) is a 24 amino acid peptide that shares a 7
amino acid sequence homology with the somewhat larger
neuromedin S (NMS) (Mori et al., 2005). NMU and its receptors,
NMU-R1 and NMU-R2, are both present in the rat and mouse
SCN (Mori et al., 2005; Graham et al., 2003, 2005; Nakahara et
al., 2004). NMU and NMU-R1 have high amplitude rhythmic
gene expression that persists in DD, whereas rhythmic
expression of the NMU-R2 gene is lower amplitude (Graham
et al., 2003; Nakahara et al., 2004). NMU is present in some, but
not all, VP neurons and some, but not all, NMU-containing
neurons also have VP; it apparently does not colocalize in VIP
neurons (Graham et al., 2003). The distribution of NMU in rat is
most similar to the distribution of VIP neurons, but is present
throughout much of the SCN. NMU-R1 and R2 receptor
distributions are similar to that for the peptide. Phase
advances of about an hour followed ventricular injection of
NMU at CT0 into rats; injections at CT6 yielded phase delays of
about 0.6 h; and from CT8 to CT23, there was no phase
response. FOS protein expression was widely induced by
intraventricular NMU (Graham et al., 2003). Mice lacking the
gene coding for NMU have rhythmic feeding behavior, but the
phase angle of entrainment may be altered (Hanada et al.,
2004).
NMS is distributed in the rat SCN in a pattern similar to VIP
and there was rhythmic expression peaking at ZT11 (Mori et
al., 2005). The rhythm was lost in DD and a light pulse acutely
reduced expression. Intraventricular injection of NMS at CT23
induced 1 h phase delays and, at CT6, about 2 h phase
advances with a dead zone between CT9 and CT22. FOS
expression in dorsal SCN cells was also induced by NMS. The
PRC for NMS was similar to that for NPY (Albers and Ferris,
1984; Albers et al., 1984), whereas that for NMU appeared to be
somewhat different. The general shape of the PRC for NMS
was very much light-like, but was shifted such that phase
delays began at CT24 with a cross-over point at CT3 and the
phase advance region ended at approximately CT8-9 (Nakahara et al., 2004).
6.1.6.
Vasopressin
The Brattleboro rat is deficient in VP, yet has circadian
rhythms (Ingram et al., 1998). It has been concluded that VP
is not necessary for circadian rhythm expression. In fact,
transplanted SCN from Brattleboro rats will restore normal
circadian rhythmicity in arrhythmic hosts (Boer et al., 1999).
22
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
Although VP is not a necessity for rhythm expression in
Brattleboro rats, transplanted SCN from normal animals can
restore rhythmicity in paraventricular hypothalamic neurons
and this rhythmicity is likely to be the result of VP stimulation
(Tousson and Meissl, 2004). The issue is rendered more
complicated by the demonstration that rhythmicity of VP
cells evaluated in vitro shows advances and delays according
to circadian time of VIP stimulation. Interestingly, the same
study failed to observe a phase shifting effect of glutamate
which would be expected to act either directly on the VP cells
or on the VIP cells projecting to the VP cells (Watanabe et al.,
2000).
6.1.7.
Diurnality–nocturnality
In the grass rat, a diurnal rodent, the above effect of GABA
agonists administered during the subjective day may occur at
phases opposite to those effective in the nocturnal hamster
(Novak and Albers, 2004a). In particular, muscimol injected in
the peri-SCN region at this time induces phase delays rather
than phase advances and these effects are not blocked by TTX.
Opposite to the effect in nocturnal species, phase shifts
caused by peri-SCN muscimol are not inhibited by simultaneous light exposure (Novak and Albers, 2004b). However,
nocturnal and diurnal species appear to respond similarly
with respect to the ability of GABAA and GABAB agonists to
inhibit phase response to light (Novak and Albers, 2004b;
Novak et al., 2004). Assuming that the action of injected GABA
agonists is occurring within the SCN, then the antiphase GABA
activity in diurnal vs. nocturnal animals is the first indication
that the SCN contributes to this species-specific behavioral
trait.
In a subparaventricular region contiguous to the dorsal
SCN, the lower subparaventricular zone, the grass rat has
higher expression of FOS during the night than during the
day, an effect not observed in rats (Smale et al., 2001;
Schwartz et al., 2004; Nunez et al., 1999). Moreover, the day–
night pattern of expression is roughly opposite that which
occurs in the SCN of either the Norway lab rat or grass rat.
The suggestion has been made that the neuron firing rate
rhythm can become inverted in the peri-SCN region for
nocturnal (Inouye and Kawamura, 1979), but not for diurnal
(Sato and Kawamura, 1984), species and may contribute to
the manifestation of nocturnal or diurnal activity traits
(Smale et al., 2003).
6.2.
Acetylcholine
Cholinergic innervation of the SCN has been of interest for
many years and there are numerous reports of cholinergic
receptors on SCN cells and modification of circadian rhythm
function by cholinergic agonists or antagonists (see Morin,
1994 for a review). Several anatomical reports have described
choline acetyltransferase (ChAT) immunoreactive processes
in the SCN, although the histology has not been robust and the
demonstrable fibers have been located in somewhat different
places, depending upon the particular study (van den Pol and
Tsujimoto, 1985; Bina et al., 1993; Ichikawa and Hirata, 1986;
Kiss and Halasz, 1996). Cholinergic fibers have also been
shown to make contacts directly on SCN neuron (Kiss and
Halasz, 1996). Perhaps the most convincing evidence of
cholinergic input to the SCN is the demonstration that,
following an intra-SCN injection of retrograde tracer, labeled
cells containing ChAT are found in several brain nuclei
including the lateral dorsal and pedunculopontine tegmentum, areas concerned with sleep regulation (Bina et al., 1993).
Retrogradely labeled cholinergic cells were also found in the
basal forebrain. Excitotoxic lesions of the nucleus basalis
magnocellularis destroyed about 75% of cholinergic cells in
this location yielding a decrease in cholinergic processes in
the SCN (Madeira et al., 2004). Bilateral nucleus basalis lesions
also diminished the numbers of countable VP and VIP cells in
the SCN by about 30% and reduced volume and mRNA levels of
VP and VIP cells by about 34%.
The immunotoxin, 192 IgG-saporin, destroys cells bearing
the low affinity nerve growth factor receptor, p75NTR, which
are generally cholinergic (Leanza et al., 1995). Intraventricular infusion of the immunotoxin abolished p75NTR in the rat
SCN without affecting VIP neurons (Moga, 1998). However,
intrahypothalamic immunotoxin injection into the rat greatly reduced numbers of CALB neurons (Beaulé and Amir,
2002). The circadian period length was related to the extent
of cell loss. Typically, the period was much shorter in
lesioned rats and about a third of them became arrhythmic.
Animals sustaining immunotoxin lesions of cholinergic cells
also had about a 50% reduction in light-induced FOS cell
counts. In another study (Erhardt et al., 2004), 192 IgGsaporin was injected into the ventricle or directly into the
SCN, but neither procedure yielded deficits in CALB or VIP
neurons and changes in circadian period or general rhythmicity were not observed. Nevertheless, the immunotoxin
generally reduced the ability of light to produce phase delays
or advances. The rhythm-related effect of 192 IgG-saporin
treatment may be related to disruption of effective retinal
projections to the SCN. Unilateral enucleation reduces
optical density of p75NTR in the contralateral SCN by about
50% (Bina et al., 1997).
As reviewed (Morin, 1994; O'Hara et al., 1998), the cholinergic agonist, carbachol, has generally yielded light-like phase
responses. However, the topic is not without controversy. For
example, peri-SCN carbachol induced robust phase delays
when injected at CT14 and advances at CT22, but also
produced large phase advances in about half the animals
receiving the drug at CT6 (Bina and Rusak, 1996). The phase
shifts during the subjective night are consistent with a lightlike action, but the shifts during the day are not. The responses
were blocked by pretreatment with atropine, but not by
mecamylamine, suggesting that a muscarinic receptor mediates the cholinergic effects. The functional presence of
muscarinic receptors in the SCN region is supported by
ample additional data, particularly from SCN slice preparation
studies (Liu and Gillette, 1996; Liu et al., 1997b; Artinian et al.,
2001) and there is evidence supporting a rhythm function
specifically for the M1 receptor (Gillette et al., 2001).
Despite the foregoing, the nicotinic receptor may also
mediate the effects of acetylcholine on circadian rhythmicity.
The nicotinic antagonist, mecamylamine, blocked the phase
shifting effects of light at CT19 and greatly attenuated lightinduced FOS in the dorsomedial hamster SCN with little
inhibition more ventrally (Zhang et al., 1993). In the rat slice
preparation, the PRC to nicotine was phase-independent with
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
advances occurring throughout the circadian day and the
phase shifts could be blocked by mecamylamine (Trachsel et
al., 1995). In vivo, nicotine administered subcutaneously at
CT16 elicited significant phase delays and induced FOS in the
SCN and mecamylamine greatly attenuated FOS expression
(Ferguson et al., 1999). A preliminary description of rat
response to nicotine indicated that phase shifts were unreliable following systemic administration, but were light-like
following nicotine intraventricular infusion (O'Hara et al.,
1998).
There are also reports that a strain of rats bred for
cholinergic supersensitivity has a shorter circadian period
than controls, an altered phase angle of entrainment and
reduced FOS induction in SCN neurons (Shiromani et al., 1988;
Ferguson and Kennaway, 1999).
6.3.
Neurophysiology of the SCN action potential rhythm
Action potential firing by SCN neurons is regulated as a
circadian rhythm. Higher frequencies are observed during the
subjective day than during the subjective night with the mean
population firing frequency peaking near CT6 (Gillette, 1991).
During the subjective day, action potential firing frequencies
range from less than 1.0 Hz to 14 Hz, compared to the
subjective night when action potentials fire with frequencies
of 0.5 to 5.0 Hz. The presence of fast firing neurons during the
day is responsible for the increased mean population firing
frequency observed in SCN slices (Gillette, 1991). This circadian rhythm of action potential firing is a property of individual
neurons because the firing difference is maintained when SCN
neurons are grown in dispersed cell cultures (Welsh et al.,
1995; Herzog et al., 1997; Abe et al., 2000; Honma et al., 1998a;
Shirakawa et al., 2000).
A current hypothesis proposes that the molecular circadian
clock sends signals to regulate the activity of ion channels
responsible for setting the membrane potential and action
potential firing rate. This hypothesis predicts that disruption
of clock gene function would alter the action potential firing of
SCN neurons. Mutations of clock genes alter the period of
action potential firing rhythms. Microelectrode array recordings of slice cultures from Clock homozygous mice demonstrated neurons with a period near 27 h that was significantly
longer than neurons of wild-type or Clock heterozygous mice
(period near 24 h) (Nakamura et al., 2002). In a similar study, the
period of Clock heterozygous mice was longer than that of the
wild type mice (24.5 vs. 23.3 h), while the Clock homozygous
mice were arrhythmic (Herzog et al., 1998). In the tau mutant
hamster, which has a shortened circadian period, the circadian
rhythm of action potential firing is also shortened to approximately 20 h (Liu et al., 1997a). SCN neurons from mice with
mutations of the cryptochrome 1 or cryptochrome 2 genes are
arrhythmic (Albus et al., 2002). Also consistent with this model
are the observations of McMahon and colleagues who used a
transgenic mouse expressing a green fluorescent protein
driven by the Per1 promoter to demonstrate that the frequency
of action potential firing was linearly correlated with Per1
transcription (Quintero et al., 2003).
SCN neurons express a wide repertoire of ion channels and
regulation of the activity of these ion channels by signals from
the molecular clock would lead to regulation of action
23
potential firing. The presence of a diffusible messenger is
supported by the observation that spontaneous firing of SCN
neurons “washes out” during the dialysis of the intracellular
milieu that occurs during whole-cell recording (Schaap et al.,
1999). Currently, the signaling pathways connecting the
molecular clock in the nucleus to changes in membrane
potential are unknown. However, significant progress has
been made toward describing the ion channels responsible for
the regulation of spontaneous firing.
Many membrane properties of SCN neurons show a diurnal
or circadian variation. SCN neurons have input resistances in
the range of 1–3 GΩ when measured with whole cell recording
techniques. The input resistance of SCN neurons is higher
during the day compared to the night in slices maintained in
either LD or DD (De Jeu et al., 1998; Kuhlman and McMahon,
2004). The resting membrane potential of SCN neurons has
been reported to be more depolarized during the day than
during the night (De Jeu et al., 1998; Kuhlman and McMahon,
2004; Pennartz et al., 2002). These differences in membrane
potential are maintained in rats maintained in DD consistent
with their regulation by the circadian clock (Kuhlman and
McMahon, 2004). However, a significant day–night difference
in the membrane potential has not always been observed
(Schaap et al., 1999; Kim and Dudek, 1993; Teshima et al.,
2003). Difficulties in estimating the “resting” membrane
potential of spontaneously firing neurons and dialysis of
intracellular contents may contribute to these different
results. The resting membrane potential has been proposed
to be set, in part, by a Ba2+-sensitive, fast-activating, outwardly
rectifying K+n current (De Jeu et al., 2002). Interestingly, this
current did not show a circadian rhythm in activity, although
given the high input resistance of SCN neurons, a small
change in this current could significantly change the resting
membrane potential. The membrane potential of some SCN
neurons has a low frequency (2–7 Hz) oscillation that is
mediated by L-type Ca2+ channels and is observed when action
potentials are blocked by TTX (Jiang et al., 1997; De Jeu et al.,
1998; Pennartz et al., 2002; Jackson et al., 2004).
Action potentials in SCN neurons are characterized by a
rapid upstroke of membrane depolarization (∼+25 mV) followed by membrane potential repolarization. An afterhyperpolarization (AHP; ∼−80 mV), a membrane potential that
hyperpolarizes below the resting membrane potential and
which is dependent on extracellular Ca2+, follows the action
potential upstroke (Cloues and Sather, 2003; Thomson, 1984;
Thomson and West, 1990). The rapid upstroke of the action
potential is driven by current flowing through voltage-gated
Na+ channels and is completely blocked by TTX (Thomson and
West, 1990; Huang, 1993; Wheal and Thomson, 1984).
During the interval between individual action potentials
(interspike interval), the membrane potential shows a steady
depolarization that reaches spike threshold (Kononenko et al.,
2004; Pennartz et al., 1997). This interspike interval can be
modulated by synaptic input, particularly inhibitory postsynaptic potentials generated by GABAergic input (Pennartz et al.,
1998; Kononenko and Dudek, 2004). These “depolarizing
ramps” are sensitive to the Na+ channel blocker, TTX, and to
riluzole, a compound used to treat amyotrophic lateral
sclerosis, that blocks a persistent Na+ current. Voltage-gated
Na+ channels rapidly activate and inactivate during a
24
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
membrane depolarization. However, many neurons have a
small Na+ current that shows very slow or no inactivation (for
review see Crill, 1996). This “slowly-inactivating” or “persistent” Na+ current (INa,S or INa,P) is TTX-sensitive and may
represent current flowing through a noninactivating class of
voltage-gated Na+ channels or a modal variation of the
normally transient Na+ current. In SCN neurons, a slowly
inactivating Na+ current (in I = 50–250 ms at − 45 mV) is blocked
by TTX, and riluzole similar to the INa,P. The INa,S may represent
a third modal state of voltage-gated Na+ channels (Jackson et
al., 2004).
It has been hypothesized that the INa,S is primarily
responsible for driving the membrane depolarization during
the interspike interval (Kononenko and Dudek, 2004; De Jeu
and Pennartz, 2002). Jackson et al. (2004) recorded from acutely
isolated dorsomedial SCN neurons and used action potential
waveforms as voltage signals under voltage clamp to determine the currents flowing during the interspike interval. The
ionic currents flowing during the action potential waveform
are the same as those that flow during the action potential
firing. The majority of the current during the interspike
interval was carried by TTX-sensitive Na+ current. A nimodipine-sensitive Ca2+ current was also present during the during
the interspike interval. However, the amplitude and charge
carried by the Ca2+ current are only 26% of that carried by the
Na+ current (Jackson et al., 2004). Further, the Ca2+ current
turns on later in the interspike interval than the Na+ current
and is not significant until the membrane potential
approaches threshold. The authors conclude that the slowly
inactivating Na+ current is primarily responsible for setting the
action potential frequency while contribution of voltage-gated
Ca2+ currents to the action potential firing frequency is small
(Jackson et al., 2004).
Other candidate ion currents have been proposed to
contribute to setting the action potential frequency (Kim and
Dudek, 1993; Pennartz et al., 1997). The H-current (IH), a
nonselective cation channel activated by membrane depolarization has been hypothesized as contributing a depolarizing
current, but a direct test of this hypothesis did not yield any
evidence for an IH contribution (Akasu et al., 1993; De Jeu and
Pennartz, 1997). The A-current (IA), a fast-activating, rapidly
inactivating K+ current present in all SCN neurons, may
contribute to setting the duration of the interspike interval
(Huang, 1993; Bouskila and Dudek, 1995). The low-threshold Ttype Ca2+ current has also been proposed as a contributor to the
setting of action potential frequency (Akasu et al., 1993). No
direct evidence for the IH or IT has been presented.
The AHP that follows membrane potential repolarization
also contributes to setting action potential firing frequency.
The AHP duration correlates with the spontaneous firing rate,
the slower the AHP decay, the slower the firing rate (Cloues and
Sather, 2003). If the AHP is blocked, then the rate of firing of SCN
neurons increases. In SCN neurons, the AHP is dependent on
Ca2+ entering the cell via predominately L- and R-type voltagegated Ca2+ channels that activate both large-conductance
(BKCa) and small-conductance (SKCa) Ca2+-activated-K+ channels. Blockade of apamin-sensitive SKCa channels (only SK2
subunits in SCN) has been reported to produce an increase or
no effect on spontaneous action potential frequency. The
difference probably reflects a diversity of SKCa expression in
SCN neurons. Block of iberiotoxin-sensitive BKCa did not alter
firing frequency. An apamin- and iberiotoxin-insensitive KCa
current was diurnally modulated and contributes to the spike
frequency (Cloues and Sather, 2003). The AHP is dependent on
both small conductance (SKCa) and large conductance (BKCa)
Ca2+-activated-K+ channels.
Abundant experimental evidence demonstrates that Ca2+
plays an important role in the activity of SCN neurons. First,
removal of extracellular Ca2+ abolishes the circadian pattern of
action potential firing (Shibata et al., 1984). Second, some SCN
neurons have a regular pattern of action potential firing
characterized by small deviations in the interspike interval
(Thomson, 1984). Reduction of the extracellular Ca2+ concentration disrupts this regular firing pattern, whereas an increase
potentiates the action potential spike amplitude and the rate of
membrane repolarization (Thomson and West, 1990). Removal
of Ca2+ or block of Ca2+ channels with high concentrations of
extracellular Mg2+ reduces the spike amplitude, rate of
membrane potential repolarization and the amplitude of the
AHP (Thomson and West, 1990). Third, there is a circadian
rhythm of intracellular Ca2+ levels with higher levels during
the subjective day than during the subjective night (Ikeda et al.,
2003; Colwell, 2000). The Ca2+ rhythm is not blocked by TTX
although action potentials are completely eliminated. Ryanodine, which depletes intracellular Ca2+ stores, significantly
blocked both the intracellular Ca2+ rhythm and disrupted the
action potential firing (Ikeda et al., 2003).
GABA is the most prominent neurotransmitter in the SCN
(Moore and Speh, 1993; Morin and Blanchard, 2001) and there
are abundant inhibitory postsynaptic potentials that can be
recorded in SCN neurons (Jiang et al., 1997; Kim and Dudek,
1992). Activation of GABAA receptors is traditionally thought to
produce cellular inhibition due to activation of a chloride
conductance and membrane hyperpolarization or the shunting of excitatory currents. However, in the SCN, descriptions of
the actions of GABA on the SCN neuronal activity have been
complicated and controversial. The majority of recordings had
shown GABA to be an inhibitory neurotransmitter during the
day (Bos and Mirmiran, 1993; Gribkoff et al., 1999; Liou and
Albers, 1991; Liou et al., 1990). However, other investigators
described the activation of GABAA receptors to be primarily
excitatory, producing an increase in firing activity or membrane depolarization during the day reducing action potential
firing and producing a membrane hyperpolarization during
the night (Wagner et al., 1997, 2001). This diurnal rhythmicity
was proposed to be caused by a daily change in the
intracellular chloride concentration (Wagner et al., 1997,
2001). A high daytime intracellular chloride concentration
was proposed to account for the observations. An excitatory
GABA response would be observed when the equilibrium
potential of Cl− (ECl) is less negative than the resting
membrane potential. Contrary to those results, a different
group of investigators reported that GABAA receptor activation
produces only inhibitory effects during the day and excitatory
effects in a subpopulation of SCN neurons during the night (De
Jeu and Pennartz, 2002). The reversal potential during the
night was more depolarized than during the day consistent
with an increased intracellular chloride concentration. The
specific reasons for these dramatically different observations
remain unknown. Technical issues such as style of whole cell
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
recording, duration of GABA application and the GABA
concentration may all contribute to the different observations.
6.4.
Inter-SCN differences in gene expression
One of the more intriguing developments during the last 10
years has been the observation that the two SCN can
oscillate out of phase (De la Iglesia et al., 2000). In male
hamsters with split rhythms caused by LL exposure, the
putative circadian clock gene, Per1, is maximally expressed
in one SCN while another clock gene, BMAL1, is simultaneously expressed maximally in the other SCN (normally,
Per1 and BMAL1 are expressed about 180° out of phase and
maximal expression cannot be seen simultaneously; Honma
et al., 1998b; Shearman et al., 2000). High Per1 expression
normally occurs during the middle subjective day, a
characteristic that is also true in SCN from hamsters with
split rhythms. The genes coding for VP and FOS proteins are
also expressed rhythmically in normal animals at the same
phase as Per1. In the animals with split rhythms, activity of
these genes remains temporally synchronized with that for
Per1. Despite the continuous presence of light, photically
induced Per or FOS expression does not appear to occur
during the subjective night (De la Iglesia et al., 2000). This
observation is curious because it implies that a photic action
normally occurring in DD-housed animals in is being
blocked, at least in the case of animals with split rhythms.
Apparently, LL-housed animals with unsplit rhythms have
not been tested to determine if there is light-induced gene
expression during the subjective night. It would also seem
possible, although perhaps unlikely, that the rhythmic gene
expression observed in SCN of animals with split rhythms is
induced by a temporally gated action of light rather than
being an expression of intrinsic cellular rhythmicity.
The observation that the two SCN may oscillate in
antiphase has raised the possibility that each SCN may
have specific control over the timing of individual physiological events. This has been examined by evaluating the
control of the gonadotropin, luteinizing hormone (LH), in
animals with split running rhythms. Ordinarily, estrogentreated, ovariectomized female hamsters have a single daily
LH peak timed by a circadian clock (Alleva et al., 1971;
Bittman and Goldman, 1979), but in animals with split
rhythms, there are two such LH peaks in antiphase (Swann
and Turek, 1985). The circadian clock action occurs through
regulation of pituitary LH release by septal-preoptic neurons
synthesizing gonadotropin releasing hormone (GNRH) (De la
Iglesia et al., 1995). Activation of GNRH neurons, as indicated
by FOS expression, is maximal at the time of LH release
(Urbanski et al., 1991; Doan and Urbanski, 1994). When
estrogen-treated, ovariectomized hamsters have split
rhythms in LL, there is daily FOS expression in GNRH cells
predominantly (about 5:1) localized to the side of the
forebrain in which SCN FOS expression is also dominant
(about 5.5:1) (De la Iglesia et al., 2003). The implications of
this form of lateralized circadian organization for regulatory
physiology are far-reaching and there are preliminary data
supporting the view that each SCN controls the ipsilateral
expression of rhythmicity in peripheral organs (Yamazaki et
al., 2000).
6.5.
Regional expression of Fos
6.5.1.
Hamster
25
An early observation (Rea, 1992) was that light did not
homogeneously induce FOS expression throughout the hamster SCN. Rather, the protein was heavily induced in the
ventral half of the nucleus both at CT13 and CT18, but there
was also large induction of FOS in the dorsal half at CT18. This
study lacked no-light controls, but the result has been
corroborated (Chambille et al., 1993; Guido et al., 1999). FOS
induction by light is similar in the dorsal and middle thirds of
the hamster SCN at both ZT15 and ZT22 (no response and
large response, respectively) and in the ventral third (very
large responses at both times) (Guido et al., 1999). Also, the FOS
response to light is greater in the ventral third of the SCN than
in the other two thirds at both ZT15 and ZT22. Much of the
ventral third of the SCN, in which dense, light-induced FOS
occurs, corresponds to the central subnucleus that can be
identified by the presence of CALB neurons and dense retinal
innervation. One point of the Guido et al. (1999) results is that
FOS induction by light occurs throughout the SCN, not just in
the central subnucleus.
The effects of GABA agonists on facilitation or inhibition of
light-induced FOS are proportionally equivalent in rostral,
middle and caudal thirds of the SCN (Gillespie et al., 1999).
When FOS counts were made through an entire SCN section at
the midcaudal level, there was no response at all to a 1 min
light exposure at ZT15, whereas the same stimulus at ZT18, 21
or 24 yielded a near maximal result. A longer exposure to light
at ZT15 greatly increased the FOS response, but never to the
level seen at the other times. A complementary analysis
(Chambille et al., 1993) has additionally demonstrated that
while light at CT14 induces a small increase in FOS in the
rostral 3/4 of the SCN, there is substantial FOS induction in this
area when light is administered at CT19. The above-cited
papers demonstrate a region by time interaction with respect to
gene induction by light stimulation, with region being defined
by rostrocaudal or dorsoventral level. Further, the Guido et al.
(1999) work indicates that the sensitivity of SCN neurons to
photic input varies according to time during the subjective
night. Implicit, but not studied, is the likelihood of region by
time interaction with respect to sensitivity to light. The
observations of region by time interaction with respect to
light-induced FOS expression have not had an impact on more
recent developments focused on photic regulation of putative
clock genes.
6.5.2.
Rat
Fos gene expression spontaneously oscillates in the SCN with
most of the amplitude accounted for by the expression in the
dorsomedial region (Schwartz et al., 1994; Koibuchi et al.,
1992). However, a low amplitude oscillation is present in the
ventrolateral region (Sumova et al., 1998). Regional FOS
oscillation persists in vitro (Prosser et al., 1994; Geusz et al.,
1997). Multiple Fos-family genes are light-induced in the
ventrolateral region of the rat SCN. These include c-fos, fra-2
and fosB. It has been suggested that each gene is a candidate
mediator of light effects on SCN neurons and regulation of
circadian rhythm phase that may enable reasonably normal
rhythm function in the absence of normal gene activity
26
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
(Schwartz et al., 2000; Honrado et al., 1996). A double label
study has shown that only a small percentage of VP cells
contain spontaneously expressed FOS and that a similarly
small percentage of cells expressing FOS also contain VP
(Sumova et al., 2000). The daily profile of spontaneously
expressed FOS is altered according to the length of the
entraining photoperiod, a phenomenon with implications for
photoperiodism (see Sumova et al., 2004 for a review).
Analysis of light-induced FOS expression in various SCN
regions has been rendered more complex by the observation
that light can inhibit it (Beaulé et al., 2001b). In the rat, light
induces large increases in FOS expression in neurons
throughout the ventral SCN. As indicated above, the dorsomedial region does not show light-induced gene expression, but
does have constitutive FOS that varies according to circadian
time. A particularly novel result in this study is the demonstration of an actual drop in the number of cells expressing
FOS in an apparently specialized sector of the dorsomedial
SCN (Fig. 3C1, 2) following light administered at CT2 or CT14,
or in entrained animals, at ZT1 or ZT2 (Beaulé et al., 2001b).
6.6.
Regional clock gene expression
6.6.1.
Mouse
The advent of digoxigenin procedures for in situ hybridization has permitted analysis of light-induced clock genes at
the cellular level. As indicated by these procedures, light
applied at CT16 induced Per1 mRNA in the ventral, but not
the dorsomedial, mouse SCN (Shigeyoshi et al., 1997). In
contrast, the dorsomedial region exhibited a circadian
oscillation of Per1 expression with a peak near CT4 and
similar rhythmicity in Per1 expression was present throughout the mouse SCN. A two component model of SCN
operation was proposed in which the SCN is organized into
dorsomedial “type A” cells and ventral “type B” cells
(Shigeyoshi et al., 1997). The latter are strongly influenced
by light, while the former are light-independent. Similar
results have been obtained in the rat (Yan et al., 1999).
In the Per1-GFP mouse, the extent of regionality appears to
depend upon the lighting history. In LD housed animals,
approximately equal numbers of cells rhythmic for Per1 were
found in the medial and lateral halves of the SCN (Quintero et
al., 2003). In contrast, in DD housed animals, about 66% of the
medial neurons were rhythmic compared to 34% in the lateral
SCN. There were also differences in the dorsoventral distributions of rhythmic cells, with twice as many found in the
ventral SCN of LD housed animals compared to similar
percentages in the two halves of DD housed mice. As
described above, the Per1-GFP mice have rhythmic neurons
with peak activity at one of three preferred phases. In DD
housed animals, the vast majority of cells peaking at CT18 are
present in the medial SCN, whereas the medial–lateral
difference becomes progressively less at CT15 and CT12,
respectively. A similar relationship between peak phase and
laterality is evident in LD housed mice, but is not as
pronounced (Quintero et al., 2003). In organotypic slice
cultures of SCN from mice carrying the Per1promoter-driven
luciferase reporter gene, most cells identified by the reporter
show oscillatory gene expression and these cells are widely
distributed throughout the SCN (Yamaguchi et al., 2003).
In the organotypic slice preparation, luminescence rhythm
peaks of Per1 expression in cells of the dorsal mouse SCN have
a broad distribution of phases, but most are maximal in
advance of those in middle and ventral thirds of the nucleus
(Quintero et al., 2003). The cells rhythmically expressing Per1 in
the dorsal third are unable to collectively sustain a cohesive
rhythm, unlike those more ventrally located. It is noteworthy,
however, that each dorsal cell continues a seemingly normal
oscillation uncoupled from the others in the same dorsal
region. This suggests that the more ventral oscillatory cells are
imparting a signal to those in the dorsal SCN, thereby
controlling phase of rhythmic cells in that region. In addition,
the investigators note that results from the horizontal slice
preparation indicate the dorsal neurons are not necessary to
maintain either rhythmicity of the ventral cells or their
oscillatory synchrony (Yamaguchi et al., 2003). The presence
of preferred phases by regionally specific collections of
oscillatory neurons (Quintero et al., 2003; Yamaguchi et al.,
2003) is consistent with the view described for hamsters
(Hamada et al., 2004a) (discussed below), although the
interpretation may be substantially different. On the one
hand, discussion of mouse SCN tends to emphasize the
presence of many oscillatory neurons collectively synchronized, but with different phases of peak Per1 gene expression.
On the other, the hamster work describes the “slow spread of
gene expression” from one area to another. The appearance of
“spread” may be the consequence of what has been described
in mice (Yamaguchi et al., 2003), namely, the presence of
oscillatory neurons with grouped or a gradient of phases. The
mouse data have an implication for the direction of causation.
The dorsal oscillatory neurons cannot maintain synchrony
when separated from those more ventrally located. At least in
the mouse, this strongly suggests that the direction of
causality with respect to phase angle and synchrony among
oscillators is from the more ventral rhythmic cells to the dorsal
rhythmic cells.
Other studies of the mouse SCN note a central region (Fig.
3F3) with greatly reduced rhythmic Per1 and Per2 expression
(Karatsoreos et al., 2004; LeSauter et al., 2003; King et al., 2003;
Reddy et al., 2002). This area corresponds closely to the
distribution of GRP neurons (note: the GRP cells were identified
in a CALB-GFP reporter mouse in which, for some unknown
reason, about 90% of the GFP cells in the SCN also contained
GRP, and for an equally unknown reason, no cells in the SCN
known to contain CALB were identified by GFP (Karatsoreos et
al., 2004). However, the most central part of the region
containing putative GRP cells has distinct expression of both
Per1 and Per2 at ZT2 or CT2, times when rhythmic expression is
minimal elsewhere in the SCN (King et al., 2003). Previously, a
similar sized, fairly discrete collection of neurons in the
centrodorsal (not dorsomedial) mouse SCN was described as
expressing Per1 at ZT0 or CT0 (Hastings et al., 1999).
With respect to light-induced clock gene expression in the
mouse, a light pulse at CT13-14 has been reported to cause
Per1 expression throughout the central and dorsolateral SCN
in a region complementary to the medial area in which
rhythmic Per1 is expressed (Karatsoreos et al., 2004). There is a
high degree of correspondence between light-induced Per1
and the GRP cell distribution. The same appears true for FOS
expression. The region of light-induced Per1 and GRP cells is
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
completely filled with retinal projections which tend to be
much sparser in the medial part of the SCN in which rhythmic
Per1 or Per2 expression predominate. However, another report
indicates that light-induced Per1 is found in the ventral third
of the SCN corresponding largely to the area containing PHI
neurons (King et al., 2003). This is consistent with observations
that VIP neurons containing Per1 jump from 31% to 59%
following a light pulse at ZT21 (Kuhlman et al., 2003). In
contrast, Per1 is expressed in about 35% of VP neurons at this
time, but is not elevated in response to light. VIP and VP cells
expressing Per1 comprise about 51% of the total neurons
spontaneously expressing that gene at ZT10, indicating that
there is no specific correspondence between cell phenotype
and rhythmic clock gene activity. The information also
indicates that rhythmic gene expression is not solely the
province of dorsomedial neurons, as the VIP cells are
consistently present in the ventral part of the SCN.
Light-induced Per2 protein in the SCN is described (King
et al., 2003) as being expressed with breadth and magnitude
equivalent to that which occurs at its maximal rhythmic
expression. One specific characteristic of the rhythmic
expression is its relative absence from the GRP cell region
(Karatsoreos et al., 2004; LeSauter et al., 2003; King et al.,
2003) (Figs. 3G1, 2; H). Cells identifiable as containing the
VPAC2 receptor had both rhythmic and light-induced
expression of Per2 (King et al., 2003). There appear to be
some differences in the results of two comparable mouse
studies (Karatsoreos et al., 2004; King et al., 2003), but it is
difficult to determine whether or not they are real or simply
the result of substantially different methods.
6.6.2.
Rat
A comprehensive analysis of Per1 and Per2 gene expression in
the entire rat SCN has produced a modification of the
dorsomedial/ventrolateral SCN divisional model to include
three functionally distinct regions (Yan and Okamura, 2002).
The single dorsomedial region has been divided into periventricular and central parts (Fig. 3D). The former overlaps a fairly
thin, dorsomedial portion of the SCN identifiable by VP
neurons. The central portion overlaps the ventrolateral part
of the VP cell group as well as the dorsal part of the VIP cell
distribution. The third, ventrolateral, portion of the SCN
generally corresponds to the bottom 80% of the region
containing VIP cells. The three regions are reported to be
distinct with respect to their expression of Per1. For example,
the peak expression in the periventricular division is at CT4,
but at CT8 in the central division. Overall, the profiles of Per1
expression are different in the three regions. A second
distinguishing feature is the Per1 response to light which, for
the periventricular and central divisions, is nonexistent at
CT16, while extremely robust for the ventrolateral division. In
contrast to Per1, rhythmic expression of Per2 is similar in the
periventricular and the central divisions, thereby indicating a
different SCN divisional structure.
An analysis of Per2 protein has shown rhythmic expression
in dorsomedial and ventrolateral rat SCN, but no induction in
either location after a light pulse at CT13 (Beaule et al., 2003a).
It has been conceptually convenient to consider the rat SCN
as having two parts, the dorsomedial and ventrolateral
“divisions.” Moreover, it has also been convenient to view
27
the ventrolateral division as the sole SCN region with retinal
innervation and VIP neurons, while the dorsomedial division
has no retinal innervation and contains VP neurons. However,
convenience must give way to a more complex reality as
suggested by the presence of two functionally distinct parts of
the dorsomedial rat SCN (Yan and Okamura, 2002). Similarly,
when the ventrolateral region containing VIP neurons has
been examined closely, it also can be subdivided into
differently functioning units. VIP cells identify an elliptical
area occupying the ventral half of the rat SCN. However, cells
in the medial quarter of this region do not express lightinduced Per1 mRNA nor, as demonstrated in adjacent
sections, does this area receive much, if any, retinal innervation (Kawamoto et al., 2003). Thus, the nominal ventrolateral
rat SCN containing VIP cells is divisible into a lateral part in
which Per1 is induced by light and a medial part in which the
cells do not have this response (Fig. 3E).
The foregoing results are of significance because they
demonstrate, at least for the rat, that (a) cells throughout the
entire SCN oscillate, although the magnitude is greater in some
regions than in others; (b) there are regional distinctions
between the rhythmic expression of Per1 and Per2; (c) there are
at least four functionally distinct regions within the SCN; (d)
the SCN appears to have a compartment in which cells rapidly
express Per genes in response to light, but this does not occur in
the other three compartments and (e) there is a large, but
clearly imperfect, anatomical correspondence between known
cell phenotypes and the three regions. In addition, it is
noteworthy that the ventrolateral division, unlike the others,
does not extend the entire length of the SCN (Yan and
Okamura, 2002).
One obvious burden imposed on investigators by the
increased number of SCN subdivisions is the need to minimize
confusion by using a common nomenclature characterized by
fully understandable definitions. It is difficult to compare
studies when, in one study, a region may be referred to as the
“VP” area, a second study refers to the same area as the
“dorsomedial region,” while a third study emphasizes a
difference between the “medial dorsomedial” and the “lateral
dorsomedial.” The difficulty is compounded by the fact that,
because the information is so new, the laboratories making
most of the anatomical observations have yet to achieve a
standard nomenclature across publications.
6.6.3.
Hamster
A functional analysis of SCN divisions has also been made in
the hamster, although without the same level of detail. A twocompartment description has been demonstrated, with one
unresponsive to light and the other highly responsive to light
(Hamada et al., 2001). The shape of the hamster SCN lends
itself to a descriptive terminology more closely related to
peptide content of cells than to the geographic descriptors of
dorsomedial vs. ventrolateral. Thus, there is more emphasis
on the central subnucleus and the fact that cells in this region
behave differently than the more dorsomedial cells. As in the
rat, there is a pronounced endogenous rhythm in Per1 and
Per2 gene activity among neurons closely associated with the
distribution of VP cells in the hamster SCN. As a generality, the
distribution of these cells wraps around the medial half of the
collection of CALB neurons in the central subnucleus. Cells in
28
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
the CALB-containing region express the Per1 and Per2 genes
robustly in response to light. It is also the case that neurons in
the CALB cell region do not have endogenous oscillations in
Per1, Per2 or Fos gene expression unlike their more medial
counterparts (Hamada et al., 2001). It is not known to what
extent the Per1 or Per2 gene expression occurs rhythmically in
VP neurons or is light-induced in CALB neurons.
Further analysis of the hamster SCN has demonstrated
that while the most robust Per gene response to light occurs in
cells in and near the CALB cell region, a similar phenomenon
occurs among cells in the dorsomedial SCN, the region of VP
neurons (Hamada et al., 2004a). The time course of lightinduced Per1 gene expression in cells of this region is
significantly longer than in the CALB region, suggesting an
altogether different pathway for induction. Nevertheless, the
result is consistent with the smaller number of retinal
projections to the dorsomedial region than to the central
CALB cell region in the hamster (Muscat et al., 2003). It is also
the case that the VP gene itself not only has an intrinsic
oscillatory expression in SCN neurons, it is also light-inducible
in a slow onset manner similar to that for the Per1 gene in cells
of the VP region (Hamada et al., 2004a).
Rhythmic expression of VP mRNA in the hamster SCN
appears to have at least two characteristics: there is always
expression in the dorsomedial region (minimal at CT20), and is
greatly increased in that region at CT8, while also appearing
throughout a much larger part of the entire SCN (Hamada et
al., 2004a). Per gene expression has similar rhythmic expression characteristics, but Per1 and Per2 mRNAs are virtually
nonexistent at CT20. The investigators suggest that rhythmic
gene expression is initiated among a small number of
dorsomedial cells and spreads slowly through more caudal
and ventrolateral parts of the SCN.
There is clear intra-SCN organization to the temporal and
spatial rhythmic expression of the Per and VP genes. However,
in an oscillatory system where there is no beginning and no
end, it is not clear that an origin of the gene expression rhythm
can be defined. More important is the issue of whether cells in
one part of the SCN transmit neural or humoral rhythmic
information to other cells that is requisite for the expression
or phasing of rhythmicity in those cells. Related to this issue is
the question of cellular “gradients” in gene expression within
the SCN (Hamada et al., 2004a; Yan and Okamura, 2002). A
“gradient” implies gradual spatial change, a view contrary to
observations of rhythmic gene expression occurring within
specific sectors of the SCN (see Fig. 3 in Yan and Okamura,
2002). The observations of temporal and spatial rhythmic organization within the SCN are likely to be of great importance
with respect to understanding the movement of information
within the SCN itself as well as to the understanding of
function attributable to the various sectors of the SCN.
To our knowledge, there has not been an investigation of
light-induced clock gene expression comparable to the comprehensive study of FOS by Guido et al. (1999). However,
possibly unlike the situation in the rat (Yan and Okamura,
2002), a CT19 light pulse first induces Per1 expression in an area
of the hamster SCN encompassing the central subnucleus, but
this is followed, with time, by induction of gene expression in
cells throughout most of the dorsomedial region as indicated
by the location of VP neurons (Hamada et al., 2004a). Whether
there is a “periventricular” part of the hamster SCN, as there is
for the rat (Yan and Okamura, 2002), remains to be determined.
6.7.
Stimulus induction of gene expression
6.7.1.
Photic stimuli
Rea (1989) showed photic induction of FOS protein in SCN
neurons in 1989. During the past decade, there have been
many other investigations directed at the expression of a
variety of genes thought to be components of the circadian
clock. Substantial information exists which shows rapid
induction of Per1 and Per2, but not Cry1, Cry2 or Per3, gene
expression following a light pulse (Shigeyoshi et al., 1997; Best
et al., 1999; Field et al., 2000; Zylka et al., 1998; Shearman et al.,
1997). Brief (15 min) light pulses at ZT14 yielded Per1 and Per2
gene expression increases that were 26 and 30%, respectively,
greater than controls when measurements were made 1
h later. However, the changes in mRNA levels occurred
without a noticeable increase in protein which is not evident
until many hours later (Field et al., 2000). These results
emphasize the investigators' point that there is not a reliable
relationship between the phase shift magnitude and the level
of light-induced Per gene expression.
In a novel experiment, Paul et al. (2003) tested whether
glutamate receptors mediate light-induced suppression of
nocturnal pineal melatonin as well as light-induced expression of Per1 mRNA. As expected, both nocturnal environmental light and the glutamate receptor agonist, NMDA, applied to
the SCN suppressed pineal melatonin and elevated Per1
mRNA. Joint application of AMPA or NMDA receptor antagonists with the NMDA blocked the effect of light on Per1 mRNA
expression, but in the same animals, the antagonists were
unable to prevent the light-induced reduction in melatonin.
This study demonstrates the existence of two different pathways in the SCN. Both are light-responsive, with one presumably related to the regulation of circadian rhythm phase
response and the other contributing to photic inhibition of
nocturnal pineal melatonin. The study also raises an issue that
should be of concern to all rhythm investigators. That is,
whether the changes observed in the SCN are directly related
to the phenomenon under investigation or simply correlates.
This issue has been extended to evaluation of photic induction
of the clock genes, Per1 and Per2 (Paul et al., 2004). The results
indicate that both light and NMDA induced the expected gene
expression in the SCN and also suppressed pineal melatonin.
Blockade of sodium-dependent action potentials with TTX
blocked both light-induced gene expression and melatonin
suppression consequent to either light or NMDA. In contrast,
induction of Per1 and Per2 by NMDA was not blocked by TTX
(Paul et al., 2004). The results of the above studies are
consistent with the view that there are two distinct mechanisms regulating light-induced gene expression and lightsuppression of pineal melatonin.
6.7.2.
Nonphotic stimuli
Whereas light during the subjective night induces Per gene
activity in the SCN and causes rhythm phase shifts, it has
little, if any, effect on these two measures during the
subjective day. In contrast, phasically active nonphotic
stimuli administered during the subjective day elicit rhythm
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
shifts when applied during the subjective day which is
associated with decreased Per gene activity in the SCN.
Several classes of nonphotic stimuli have this effect,
including benzodiazepine treatment (Yokota et al., 2000),
novel wheel activity (Maywood and Mrosovsky, 2001),
injection-related arousal (Maywood et al., 1999), peri-SCN
infusion of NPY (Maywood et al., 2002; Fukuhara et al., 2001)
and serotonin receptor agonists (Horikawa et al., 2000).
Antisense oligonucleotide to the Per1 gene administered
directly into the peri-SCN region during the subjective day
suppresses Per1 mRNA expression by about 60% (Hamada et
al., 2004b). This treatment at CT6 induces a phase advance
that averages about 60 min, a result consistent with the view
that enhanced Per1 expression, in this case caused by a
nonphotic stimulus, induces phase shifts.
A proposed unifying hypothesis suggests that elevated
Per gene expression and Per protein synthesis is causal to
phase shifts induced either by light or by nonphotic stimuli
(Maywood and Mrosovsky, 2001). Moreover, the hypothesis
makes an unconventional specific prediction that light
should be able to modify circadian clock function even
during the “dead zone” of the PRC. The rationale is that,
during the normal dead zone, light cannot alter phase
because Per expression is already high and further expression cannot be elicited by light during the subjective
daytime. However, light can be shown to alter SCN activity
during the subjective day because light exposure at that time
prevents the drop in Per gene activity normally induced by
nonphotic stimuli (Maywood and Mrosovsky, 2001) or direct
NPY infusion into the peri-SCN region (Maywood et al., 2002).
The converse of this procedure is also demonstrable with
nonphotic stimuli presented during the subjective night able
to attenuate light-induced phase shifts (Mistlberger and
Antle, 1998; Ralph and Mrosovsky, 1992).
According to the Maywood and Mrosovsky (2001) unifying
hypothesis, nonphotic stimulus should achieve its attenuating
effects on light during the subjective night by reducing the
extent of light-induced Per gene activity. However, direct tests
of this proposition have yielded mixed results. In the first
instance, benzodiazepine treatment produced an approximately 50% reduction in light-induced (5 lx, 15 min, CT20)
phase advances, but had no effect on phase delays (Yokota et
al., 2000). The effect on phase advances was associated with an
approximately 40–50% drop in Per1 or Per2 mRNA (the
corresponding test for phase delays was not performed). In
contrast, access to a novel running wheel reduced lightinduced phase delays without altering light-induced phase
advances, but the novel wheel activity had no effect on lightinduced Per1 mRNA (Christian and Harrington, 2002). In a
similar investigation, access to a novel wheel for 1 h after a
light pulse resulted in virtual abolition of phase advances in
response to less intense illumination and substantial attenuation of response to more intense illumination. However, as in
the above study, there was no reduction in either Per1 mRNA or
FOS protein associated with the reduced phase response to
light (Edelstein et al., 2003a). As noted by the authors, Per1
appears to be insufficient for phase shifts in locomotor
rhythms and that, “The relationship between light and
nonphotic stimuli is more complex than has been previously
suggested (Maywood and Mrosovsky, 2001)” (see Edelstein et
29
al., 2003a for a discussion that considers a variety of issues
related to the above inconsistencies).
6.7.3. Inconsistencies between phase response and gene
expression
Light-induced Fos gene activity is closely correlated with the
ability of light to entrain circadian rhythms, but there is a
general consensus that FOS is not an essential part of either
the clock or the input pathway mediating entrainment
(Colwell et al., 1993a,b,c). Moreover, FOS protein induction in
response to light is usually, but not always, predictive of
phase response. Quipazine injection at CT8 is significantly
reduced FOS expression in hamster SCN, but there was no
effect on phase shifts at this time and in mice, this drug had
no effect on phase response, but increased FOS expression
during the subjective day (Antle et al., 2003b). In mice
entrained to a skeleton photoperiod of 1 h light pulses
presented to create a 1:14:1:8 LDLD photoperiod, some
animals ran during the long, and others during the short,
interval of dark. The light manipulations revealed that only
the light pulse occurring near activity onset (ZT12) appeared
to influence locomotor rhythm phase. Irrespective of the
effect on entrainment, among animals running during the
long dark interval, light pulses at the beginning or the end of
that interval equally induced Fos mRNA expression. Moreover, if animals ran during the short dark interval, Fos mRNA
expression was induced by both pulses, but was much greater
after the pulse at ZT21 (Schwartz et al., 1996). Thus, in this
paradigm, there is a clear disparity between the entraining
capability of light and the magnitude of Fos gene expression
in response to light. Molecular absence of Fos does not
prevent entrainment of mice (Honrado et al., 1996). In vivo
blockade of Fos availability is associated with an inability of
light to induce phase shifts in rats (Wollnik et al., 1995).
However, JunB gene activity was simultaneously blocked in
this study and the negative effects on light-regulated phase
control may have been the consequence of the suppression of
the activity of both genes, rather than only Fos (see Schwartz
et al., 2000 for a discussion).
Edelstein et al. (2003a) suggest that their data from
nonphotic stimulus conditions are consistent with the view
that several different mechanisms contribute to the regulation
of circadian phase depending on the nature of the stimulus
involved. Most importantly, they conclude that phase shifts
can occur without necessarily modulating Per gene expression.
This is contrary to the general view that phase shifts are
dependent upon Per gene activity, although not at all
inconsistent with the available literature. It is also the case
that Per1 and Per2 expression is not the same in the SCN of
animals showing rhythm splitting in LL as it is in animals
showing a form of induced splitting in DD referred to as
“behavioral decoupling” (De la Iglesia et al., 2000; Edelstein et
al., 2003b). In the latter case, “splitting” consisting of a daily
bout of induced locomotion is followed by a resting interval
during which Per gene expression is increased throughout
most of the SCN. However, when the elevated gene expression
occurred during the rest phase following bouts of spontaneous
locomotion, Per gene expression was also elevated, but only in
the dorsomedial part of the nucleus (Edelstein et al., 2003b).
Also, despite having several characteristics similar to
30
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
spontaneous splitting in LL conditions, the DD-housed behaviorally decoupled animals had no indications of bilateral
asymmetry in Per gene expression in the two SCN, unlike the
situation in animals showing LL-induced splitting (cf., De la
Iglesia et al., 2000).
There are several forms of dissociation between the
regulation of gene expression and the phasic effects of
light on circadian rhythms. Perhaps the simplest is the
temporary loss of synchrony between gene expression
rhythms during clock resynchronization to a shifted photoperiod. The re-entrainment rate of certain gene rhythms in
mouse is faster than that of others and depends upon the
direction of the phase shift. (Reddy et al., 2002). The rhythm
of mPer1/2 expression in the mouse phase advances rapidly
in response to light, while re-entrainment lags for mCry1. In
contrast, the rhythms of mCry1 and mPer1 or mPer2 reentrain at the same rate during a phase delay. These gene
expression results match corresponding behavioral data
showing slower advances and faster delays (Reddy et al.,
2002). A variation on the theme of temporary desynchronization among gene expression rhythms is the demonstration
that they (Per1, Per2, Cry1) shift rapidly in the ventrolateral
rat SCN, but more slowly in the dorsomedial region (Nagano
et al., 2003).
A second form of dissociation has been demonstrated in
mice lacking the PAC1 receptor (Hannibal et al., 2001b). A light
pulse that induces a 100 min phase delay in wild-type mice
elicits a larger phase delay in the mutant mice. However, the
mutant mice have a vastly attenuated Fos or Per gene
induction response to light. A substantially different experiment has yielded the observation that imposition of a 6
h photoperiod advance rapidly induces a large phase advance
in the Per1-luc bioluminescence rhythm relative to unshifted
controls. This phase differential persists for at least 6 days in
DD. In contrast, such advances did not occur in in vivo SCN
electrophysiological or behavioral rhythms (Vansteensel et al.,
2003a).
A third form of dissociation addresses the question of
whether particular genes, or change in their expression, are
always necessary for rhythm expression. One example is the
gene, Clock. Mice homozygous for a mutant form of Clock
rapidly which when absent yields mice that rapidly become
arrhythmic in DD (Vitaterna et al., 1994). More recently,
however, a similar test procedure has been applied to Clock
mutant mice, but with an LL background to the rhythm
expression. In this circumstance, many of the mice exhibit
clear circadian rhythms despite the absence of the Clock
gene (Spoelstra et al., 2002). Other investigators have
suggested that running may be an inadequate variable to
assess in order to determine whether or not the Clock
mouse in DD is rhythmic, and present data supporting the
view that such mice are indeed fundamentally rhythmic and
entrainable, but with a much longer period than normal
(Kennaway et al., 2003).
In quite a different situation with the mixed in vivo/in vitro
preparation, novel wheel access for 3 hduring the subjective
day rather quickly reduced Per1 expression in the SCN as
might be expected during the midsubjective daytime. However, under the conditions of the experiment, there was no
associated shift in the phase of the SCN firing rate rhythm in
animals so treated (Yannielli et al., 2002). On the other hand, if
an additional 2 h of wheel access was permitted (with or
without additional running), a the firing rate rhythm advanced
about 2.4 h. Brains taken 5 h, rather than 3 h, after the initial
access to wheels had reduced Per2 expression, but Per1 had
returned to normal. One conclusion is that clock resetting in
response to activity-related stimulation may require a 3–5
h poststimulus latency (Yannielli et al., 2002; Antle and
Mistlberger, 2000), whereas other nonphotic stimuli or light
may not require the same amount of time (Mead et al., 1992;
Sumova and Illnerova, 1996).
6.8.
Polysialic acid and SCN function
There are a variety of possible neurobiological roles that may
be played by the molecule, polysialic acid (PSA) and its
carrier, the neural cell adhesion molecule (NCAM), which
form PSA-NCAM (particularly during development and in the
context of neural plasticity; see Rutishauser and Landmesser, 1996). PSA-NCAM is distributed along plasma
membranes of appositions between neurons, at synapses
and between glia (Shen et al., 1997, 1999). It is of interest that
the SCN of Siberian (Glass et al., 1994) and golden hamsters
(Fedorkova et al., 2002), mice (Shen et al., 1997) and rats
(Prosser et al., 2003) contains concentrations of PSA that
fluctuates rhythmically. In LD, PSA concentration drops
precipitously and by more than 90% during the dark hours.
Although the shape of the curve describing PSA content in
the SCN is substantially different during DD, clear rhythmicity persists (Glass et al., 2003a). Moreover, light greatly boosts
transient PSA expression in the SCN (but not in hippocampus). NCAM rhythmicity is similar in the SCN, although its
rhythm in DD is much less robust than that for PSA (Glass et
al., 2003a).
Mice lacking the NCAM-180 isoform that carries PSA have
a shorter than normal circadian period, a longer activity
phase and become arrhythmic within 3 weeks of DD (Shen et
al., 1997). The mutant mice also show light-induced phase
delays smaller than control mice (Glass et al., 2000a). The
role of PSA itself has been investigated using an enzyme,
endoneuraminidase (endoN), to remove PSA. Animals treated with endoN have shorter circadian periods in DD,
although they do not appear to become arrhythmic (Shen
et al., 1997). Similar to the mutant animals, the endoNtreated mice have diminished phase delays in response to
light; they also have reduced light-induced FOS expression
(Glass et al., 2000a).
In contrast to the diminished effects of light in mice
sustaining endoN removal of PSA, hamsters so treated had
augmented phase response to nonphotic stimuli (61%
increase to 3 h arousal; 120% increase to 8-OH-DPAT
injection) applied at ZT6. In the PSA-depleted rat SCN
slice preparation, direct application of 8-OH-DPAT elicited a
41% greater phase advance (Fedorkova et al., 2002). Also in
the slice preparation (Prosser et al., 2003), PSA levels
remain rhythmic. Glutamate treatment of the slice induced
a 100% increase of PSA in the SCN, but this was blocked by
endoN pretreatment. Treatment with endoN also blocked
the phase shifting effects of glutamate or electrical
stimulation of the slice. The strong implication of this
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
research is that retinal input to the SCN requires PSA to be
functional.
A further analysis of the molecular requirements form
normal mouse rhythmicity was performed using three strains
deficient, to varying degrees, in PSA and NCAM. Animals
lacking both PSA and all forms of NCAM become arrhythmic in
DD and tended to run so much when lights are on that
entrained rhythms are often difficult to discern (Shen et al.,
2001). A similar strain, but not lacking NCAM isoform 120 has
normal entrainment under LD, but is arrhythmic in DD,
suggesting that NCAM-180 and/or NCAM-140 are required to
avoid arrhythmicity in PSA deficient mice. The conclusion
from these studies is that some combination of PSA and NCAM
is necessary for normal rhythmicity and entrainment. The
mechanisms through which this action is achieved are
presently unknown.
7.
SCN connectivity
7.1.
Humoral factors
One of the more surprising results during the last decade of
research on SCN function has been the demonstration that
rhythmic events can be controlled, in all likelihood, by
nonneural linkage to the circadian clock. The availability of
perinatal SCN transplant methods enabled this conclusion.
The procedure developed with hamsters by Ralph et al. (1990)
used embryonic SCN to restore locomotor rhythms in lesioned,
arrhythmic hosts, but SCN efferent connections were restored
to an uncertain degree (Canbeyli et al., 1991; Lehman et al.,
1987). Subsequently, transplantation of viable, membrane
encapsulated SCN restored locomotor rhythmicity without
any apparent neural connections to adjacent hypothalamus
(Silver et al., 1996b). In contrast, rat to hamster or mouse to
hamster SCN transplants restore rhythmicity in association
with substantial growth of donor SCN projections into host
hypothalamus, preoptic area and septum (Sollars and Pickard,
1993; Sollars et al., 1995; Lehman et al., 1998). It appears to be
the case that behavioral rhythms, but not endocrine rhythms,
are restored by transplantation of embryonic SCN (MeyerBernstein et al., 1999; LeSauter and Silver, 1998).
Despite declarations that the SCN contains the timing
apparatus governing transplanted circadian periodicity, there
are other possible explanations of the phenomenon and there
may be species differences in the extent to which period is the
exclusive property of the SCN (Sollars et al., 1995). It is also the
case that the specific location of the transplant within the host
brain is relatively unimportant as long as the site is in or near
the third ventricle (such as adjacent lateral ventricle) (LeSauter et al., 1996; Lehman et al., 1987; Aguilar-Roblero et al., 1994).
In addition, transplants consisting of dissociated SCN cells in
the anterior hypothalamic parenchyma or medial hypothalamus also restore rhythmicity (Silver et al., 1990). Precision of
activity onset reinstated is related to the transplant distance
caudal or dorsal (but not rostral) to the SCN. Based on these
data, the suggestion has been made that the target of a
neurohumor conveying period information from the SCN is
located near or slightly rostral to the SCN (LeSauter et al.,
1997).
7.2.
31
Efferent anatomy
Watts et al. (1987) identified six general regions to which the
rat SCN projects. Subsequent studies using the anterograde
tracer, Phaseolus vulgaris leucoagglutinin, yielded generally
similar pathways in the hamster brain for which there is (1)
an anterior projection to the ventral lateral septum, bed
nucleus of the stria terminalis and anterior paraventricular
thalamus; (2) a periventricular hypothalamic projection to
the medial preoptic region, subparaventricular zone, the
paraventricular, ventromedial and dorsomedial nuclei and
the premammillary area and (3) a posterior projection to the
posterior paraventricular thalamus, precommissural nucleus
and olivary pretectal nucleus (Kalsbeek et al., 1993; Morin et
al., 1994). In the hamster, it is likely that the cells projecting
to the olivary pretectal nucleus are actually located in the
peri-SCN, as is indicated by retrograde tracer injections into
the pretectum (Morin and Blanchard, 1998). Such cells are
also seen following retrograde tracer injections into the IGL,
ventrolateral geniculate, posterior limitans nucleus, commissural pretectal nucleus and medial pretectal nucleus
(Morin and Blanchard, 1998; Morin et al., 1992; Vidal and
Morin, 2005). In particular, cells projecting to the IGL from
the SCN region do not reside within the SCN proper, but
rather around its dorsal perimeter with a substantial
probability that their dendrites extend ventrally into the
nucleus. Watts identified an IGL-efferent projection in the
SCN of rats and the cells of origin can be seen within the
SCN in this species (Watts and Swanson, 1987). The hamster
also differs from the rat in that it has a distinct caudal
projection that ascends to innervate posterior paraventricular thalamus (a separate projection innervates the anterior
paraventricular thalamus in both rats and hamsters) (Morin
et al., 1994). Given that retrograde tracer studies identify
cells around, but not in, the SCN for some of the putative
SCN targets (IGL, VLG, pretectum), it is distinctly possible
that others (such as the posterior amygdala or posterior
paraventricular thalamus) might also receive afferents from
peri-SCN cells.
Lesion methods have been used in conjunction with
immunohistochemistry to identify SCN projections having
specific peptide content (Kalsbeek et al., 1993). In the
hamster, VP cells project to the medial preoptic, parvicellular
paraventricular and dorsomedial hypothalamic nuclei and
the anterior paraventricular thalamus. Cells containing VIP
project to the anterior paraventricular thalamus, parvicellular paraventricular hypothalamus, subparaventricular area
and the dorsomedial hypothalamus. SCN efferent projections
containing GRP were identified in the subparaventricular
area, dorsomedial hypothalamus and supraoptic nucleus.
The topography of projection neurons in mouse SCN has
been inferred (Abrahamson and Moore, 2001) from the
presence of VP or VIP immunoreactive fibers in many of
the areas identified as receiving SCN projections in other
species.
Analysis of SCN projection neurons by retrograde tracer
methods provides at least a preliminary indication of a
structure–function relationship. Successful retrograde procedures indicate both targets of SCN efferent projections and the
location of neurons providing those efferents. Retrograde
32
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
tracing methods have provided much of the recent information regarding the putative “core” and “shell” organization of
the SCN (Moore et al., 2002). In this regard, there are two
references that need to be compared. The 1987 study of Watts
and Swanson (1987) reached conclusions different from those
of Leak and Moore (2001), although the two investigations
were equally broad and used basically similar methods. As a
result, it is not possible to determine which interpretation is
correct. On the one hand, Watts and Swanson (1987) emphasized the result that, “...cells in dorsal as well as ventral parts
of the SCh project to each of the terminal fields examined.” (p.
230, abstract). Three cases showing labeled cells following
retrograde tracer injection into the paraventricular hypothalamus were illustrated. In Case E46, the labeled cells were
predominant in the dorsomedial SCN with few in the ventral
part of the nucleus. In Case E47, there were many more cells
labeled and the great preponderance was found in the ventral
half of the SCN. In Case PVH33, fewer labeled cells were
present, and the greatest density of such cells was in the
ventrolateral quadrant of the SCN. On the other hand, a
predominance of retrogradely labeled neurons in the dorsomedial SCN compared to the ventral part following paraventricular nucleus tracer injection has been reported (Leak
and Moore, 2001; Vrang et al., 1995a). The observation of
injection-related variability by Watts and Swanson is consistent with an equivalent observation by Leak and Moore who
noted opposite patterns of dorsomedial:ventrolateral SCN cell
labeling that depended upon the precise location of tracer
injection in the subparaventricular zone (medial injection = 5.6:1; lateral injection 1:5.6). In the hamster, tracer
injection into the paraventricular hypothalamus yields a
“pattern of retrogradely labeled cells in the SCN […] uniform
across cases […] scattered throughout the subdivisions of the
SCN, and no tendency for backfilled cells to be located in the
dorsomedial SCN was observed” (Munch et al., 2002; p. 53), i.e.,
there is an absence of topographic organization in the species
in which it might be most expected. The Watts and Swanson
study remains the most substantial examination of the extent
to which individual SCN cells containing a particular neuropeptide project to various targets. In general, tracer injection
into a target nucleus retrogradely labels SCN neurons containing VP and VIP, although the great majority do not contain
either peptide.
In the mouse, SCN cells projecting to the subparaventricular area have been shown to express FOS protein in
response to night time light exposure (De la Iglesia and
Schwartz, 2002). In the hamster, SCN neurons projecting to the
paraventricular hypothalamus have also been shown to
express FOS in response to light. The percentage of the cells
expressing FOS is much greater at ZT19 than at ZT14 (Munch
et al., 2002). It was also determined that some of the same cells
contained VP, VIP or GRP. Of these, cells with the combined
traits of projecting to the paraventricular nucleus, expressing
FOS in response to light and containing VIP were most
common and the combination in which the cells contained
VP was rare. Direct SCN projections to gonadotrophin releasing hormone cells of the basal forebrain (De la Iglesia et al.,
1995; Van der Beek et al., 1997) and to the rat paraventricular
hypothalamic nucleus and corticotropin releasing hormone
cells, in particular, have been reported (Vrang et al., 1995a,b).
SCN neurons have also been reported to project to, and make
likely contact with, both hypocretin (orexin) and melanin
concentrating hormone neurons in the rat lateral hypothalamus (Abrahamson et al., 2001).
Transneuronal viral tracers have been utilized to determine polysynaptic routes from the SCN. Following virus
injection into the rat locus coeruleus, cells are retrogradely
labeled in the dorsomedial hypothalamic nucleus, a known
target of SCN neurons. The transneuronal tracer also labels
SCN cells, but this does not occur following lesions of the
dorsomedial hypothalamus. Furthermore, the circadian
rhythm in firing rate of locus coeruleus cells is abolished
after by the same lesions, suggesting that the SCN control of
rhythmic neuronal activity in the locus coeruleus is via a
two synapse circuit with a first order synapse in the
dorsomedial hypothalamic nucleus (Aston-Jones et al.,
2001). Injection of virus into the hamster medial vestibular
nucleus also labels a few neurons in the SCN. This is a
polysynaptic pathway that probably includes the locus
coeruleus which projects to the medial vestibular nucleus
(Horowitz et al., 2005).
Viral tracers injected into fat pads (brown or white
adipose tissue) identified neurons in Siberian hamster and
rat SCN (Bamshad et al., 1998, 1999). The actual number of
retrogradely labeled SCN neurons is small, and an estimated
2% of these also contain VP (Shi and Bartness, 2001).
Injections placed in the adrenal medulla (Ueyama et al.,
1999), pancreas (Buijs et al., 2001) and other structures (see
Ueyama et al., 1999; Bartness et al., 2001 for reviews) label
cells in the SCN. The general theme of these studies is that
the SCN regulates sympathetic outflow, a feature evident in
the viral tract tracing of pineal innervation (Larsen, 1999;
Larsen et al., 1998; Teclemariam-Mesbah et al., 1999). However, it appears that there is also SCN control of parasympathetic output (Ueyama et al., 1999; Kalsbeek et al., 2000).
The combination of the two types of peripheral innervation
is consistent with the presence of a circuitous polysynaptic
route from the SCN through the midbrain to the eye
identified by virus injection into that organ (Pickard et al.,
2002; Smeraski et al., 2004).
There is a large amount of convergence by SCN-efferent
projections onto brain regions receiving IGL input (Fig. 4). In
addition, many of the same areas are also innervated by
retinal and olfactory system projections (Johnson et al., 1988a;
Morin and Blanchard, 1999; Morin et al., 1994; Cooper et al.,
1994; Ling et al., 1998).
7.3.
Afferent anatomy
The SCN-afferent projections in the hamster were initially
described in a fairly comprehensive fashion by Pickard (1982)
in the early 1980s. There have been two additional such
investigations, and in another species, the rat (Moga and
Moore, 1997). In this study, a combination of retro- and
anterograde tracing studies demonstrated additional projections to the SCN arising from infralimbic cortex, lateral
septum, substantia innominata, ventral subiculum, subfornical organ, ventromedial hypothalamus, arcuate nucleus,
posterior hypothalamus, pretectal nuclei, median raphe and
IGL, but not in several of the brainstem regions identified by
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
33
Fig. 4 – Convergence of SCN (solid lines) and IGL (dashed lines) projections onto diencephalic and basal forebrain regions
(black areas) in the hamster. Gray areas receive innervation only from the IGL. Only the premammillary area (white) receives
input solely from the SCN. Thickness of the lines indicates approximate density of the projection. Note that the hamster
SCN does not project to the IGL, although peri-SCN cells do so. See text regarding further convergence of visual and olfactory
input to many of these areas. Drawn after Morin et al. (1994) and data from Morin and Blanchard (1999).
Bina et al. (1993), as containing cells projecting to the SCN. The
observations of Moga and Moore (1997) in the rat are
substantially similar, but broader than, those of the older,
more limited hamster study (Pickard, 1982). As part of a larger
analysis, Krout et al. (2002) evaluated monosynaptic inputs to
the rat SCN identified by CTB labeling. These investigators
observed about 40 regions sending direct projections to the
SCN, largely in agreement with the results of Moga and Moore
(1997). However, Krout et al. (2002) also observed a more
widespread distribution of labeled cells in areas such as the
locus coeruleus and divisions of the parabrachial complex,
among others, but did not see such cells in the substantia
innominata and only a few or none in the olivary pretectal
nucleus. In all studies involving tracer injection into the SCN,
the most likely cause of label disparity across injections is the
extent of label spread outside the nucleus, uptake by
processes in close proximity to the nucleus or uptake by cell
processes extending significant distances to enter the nucleus
(Morin and Blanchard, 1998; Morin et al., 1992, 1994; Gerfen
and Sawchenko, 1984).
The above-mentioned report (Krout et al., 2002) described
about 40 brain regions with cells monosynaptically projecting to the SCN. Injection of an exclusively retrograde
(Pickard et al., 2002), transneuronal virus label into the SCN
expanded the number of brain nuclei with cells projecting to
the rat SCN to about 100–140 regions (variation may relate to
label spread outside the SCN) (Krout et al., 2002). The regions
with likely polysynaptic projections to the SCN include
numerous additional olfactory structures, divisions of the
cortex, hippocampus, nuclei of the amygdala, thalamus,
hypothalamus, as well as nuclei in the midbrain and
medulla from which there are very few reported direct SCN
projections. While this study is important, it should be noted
that any uptake by terminals in the peri-SCN region is likely
to identify regions not connected to the SCN. A similar
thorough examination of polysynaptic projections to the
SCN has not been systematically completed in other species.
However, some information is available for the hamster and
shows both similarities and significant dissimilarities with
the rat data. In particular, there appears to be little or
nothing in the way of a polysynaptic projection from the
hamster tectum and pretectum to the SCN, yet more
oculomotor midbrain structures contain labeled cells than
in the rat SCN (Horowitz et al., 2004). A polysynaptic
projection from the medial vestibular nucleus to the SCN
has been reported in the hamster.
Focused studies have documented SCN-afferent projections from the pretectal complex, with cells identified in the
olivary, medial and posterior pretectal nuclei (rat) (Mikkelsen
and Vrang, 1994) and from the median raphe nucleus (MeyerBernstein and Morin, 1996; Hay-Schmidt et al., 2003). Cholinergic cells in the substantia innominata, preoptic nucleus,
diagonal band of Broca, medial septum, pedunculopontine,
laterodorsal tegmental and parabigeminal nuclei have also
been reported to project to the SCN in the rat (Bina et al., 1993).
As noted above, there is disagreement concerning which, if
any, these nuclei send projections to the SCN (Moga and
Moore, 1997; Krout et al., 2002).
34
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
8.
Intergeniculate leaflet
8.1.
Nomenclature
The IGL has been, and continues to be, confused with the
ventral lateral geniculate nucleus. Focus on the term, “leaflet,”
as a descriptor of the IGL has drawn attention away from the
actual boundaries of the nucleus and, until fairly recently, has
not had a clearly identifiable homolog in nonrodent species. It
is now established that the medial division of the ventral
lateral geniculate nucleus is the cat homolog of the IGL
(Nakamura and Itoh, 2004; Van der Gucht et al., 2003; Pu and
Pickard, 1996). In the monkey, the homologous structure is
known as the pregeniculate nucleus (Van der Gucht et al.,
2003; Moore, 1989). However, there is uncertainty regarding
the neuromodulator content of the GHT across primate
species (Chevassus and Cooper, 1998).
In the hamster, the definition of the IGL has gradually
evolved to include all regions of the lateral geniculate complex
in which NPY neurons and cells projecting to the SCN may be
found. As indicated from the developmental data described
below, NPY cells found in medial areas near the acoustic
radiation and ventral to the leaflet portion of the IGL have not
completed full migration (Botchkina and Morin, 1995). The
region of the laggard migratory NPY cells also contains
neurons projecting to the SCN which are not found elsewhere
in the lateral geniculate complex outside of the IGL (Morin and
Blanchard, 1995; Morin et al., 1992). There is also a
corresponding group of enkephalin-containing cells, especially in the medioventral part of the IGL (Morin and Blanchard,
1995). This description of the hamster IGL is generally
consistent with that for the rat (Moore and Card, 1994),
although there are slight species differences with respect to
its cross-sectional appearance at various rostrocaudal levels.
8.2.
Development
IGL development has been described for the hamster (Botchkina and Morin, 1995). The IGL is probably best considered as
part of the so-called ventral thalamus with a developmental
origin different from the dorsal lateral geniculate nucleus
(Altman and Bayer, 1979a,b). The hamster IGL develops out of
the reticular neuroepithelium with the first NPY cells evident
on embryonic day 9–10. These are evident against a background of radial glial cells extending dorsolaterally. NPY
neurons apparently migrate along a specialized set of radial
glial processes reaching their final destination in what will
become the IGL on E14. The radial glial cell bodies dislocate
from the ventricular germinal zone on E15, and transform into
the astrocytes that become evident in the IGL across postnatal
days 5–10. Some NPY neurons do not achieve complete
migration and remain relatively displaced in what is now
considered ventromedial IGL. Fiber outgrowth from NPYcontaining IGL neurons is evident by P0, and by P3, they
penetrate the SCN. The adult-like NPY terminal plexus in the
SCN is achieved by approximately P10.
The P3 arrival in the SCN of the NPY component of the GHT
approximately coincides with the end of SCN synaptogenesis
(Speh and Moore, 1990), arrival of serotonergic innervation
(Botchkina and Morin, 1993) and precedes the initial retinal
innervation of the SCN by about 2 days (Speh and Moore, 1993).
The arrival of retinal projections in the lateral geniculate
region occurs on E13, but there is a delay in terminal
arborization until about E15.5 (Jhaveri et al., 1991). The timing
of retinal innervation of the IGL is not known.
8.3.
Anatomy
8.3.1.
Geniculohypothalamic tract
The NPY neurons were the first IGL cell phenotype shown to
project to the SCN (Card and Moore, 1982; Moore et al., 1984).
The initial description of IGL projections to the contralateral
nucleus in the rat indicated that many of the cells contained
enkephalin, but none of this phenotype projected to the SCN
(Card and Moore, 1989). There are substantial species differences with respect to IGL organization and in the hamster,
only a very small percentage of NPY or enkephalin neurons
project to the contralateral IGL (Morin and Blanchard, 1995). In
both species, GABA neurons are present in the IGL and project
to the SCN (Morin and Blanchard, 2001; Moore and Card, 1994).
In the rat, there is a fourth, unidentified, cell type (Moore and
Card, 1994). In the hamster, a fourth class consists of
neurotensin neurons (Morin and Blanchard, 2001). It is useful
to recognize, as is further indicated below, that while ample
numbers of the IGL neurons also project to the pretectum, only
a small percentage contain any of the various neuromodulators thus far evaluated, implying existence of at least one
other cell type in hamster.
NPY cells can be used to identify the IGL as they occur
nowhere else in the lateral geniculate complex (Morin and
Blanchard, 2001; Moore and Card, 1994). In the hamster,
neurotensin, which is almost completely colocalized with
NPY, can likewise identify the nucleus, albeit with somewhat
greater difficulty because it is present in only about half the
NPY neurons. There is colocalization ranging from 7% (percent
of GABA cells containing enkephalin) to 98% (percent of
neurotensin cells containing NPY) of all identified peptides
with each other in hamster IGL neurons (Morin and Blanchard,
2001).
8.3.2. Connectivity and relationship to sleep, visuomotor and
vestibular systems
Harrington (1997) has reviewed the substantial literature
concerning the ventral part of the lateral geniculate complex. This review was titled, with a good deal of foresight,
“The ventral lateral geniculate nucleus and intergeniculate
leaflet: interrelated structures in the visual and circadian
systems.” Most of the literature cited in that review was
unaware that an “intergeniculate leaflet” existed, let alone
the possibility that it might have functions different from
those of the ventral lateral geniculate. At the time of the
review, it was established that the IGL contributes to the
photic regulation of circadian period and entrainment, and
that IGL lesions could block the ability of certain nonphotic
stimuli to induce phase responses normally consistent with
an NPY-type PRC (Janik and Mrosovsky, 1994; Wickland and
Turek, 1994; see Morin, 1994; Harrington, 1997 for reviews).
The prediction in Harrington's review, evident in the title,
that the IGL has functions beyond those of circadian rhythm
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
regulation has not yet been directly established, but the
groundwork is now in place for doing so.
The most important anatomical feature of the IGL appears
to be the enormity of its connections with the rest of the brain
(cf., Fig. 4, which only includes a portion of the ascending
projections). Not only are they widespread, but they are
generally bilateral and often reciprocal. The magnitude of
IGL connectivity was initially suggested by an investigation of
hypothalamic-IGL connectivity in the gerbil (Mikkelsen, 1990).
This study, and others in hamster (Morin and Blanchard, 1999)
and rat (Horvath, 1998; Moore et al., 2000) revealed a major
efferent target extending from the lateral optic chiasm
medially through the lateroanterior hypothalamus to the
midline, including the SCN, and dorsally to include the
subparaventricular zone and adjacent perifornical anterior
hypothalamus. In addition, IGL projections extend to several
divisions of the bed nucleus of the stria terminalis and variety
of basal forebrain areas extending laterally as far as the
piriform cortex (Morin and Blanchard, 1999). Projections to
nuclei of the midline thalamus are also fairly extensive (Morin
and Blanchard, 1999; Moore et al., 2000). Many of the same
regions also send projections back to the IGL (Morin and
Blanchard, 1999; Vrang et al., 2003).
Comparison of regions receiving retinal projections
reveals convergence with IGL efferents. Four distinct routes
providing retinal innervation of rostral diencephalon have
been described (Johnson et al., 1988a; Morin and Blanchard,
1999): (a) to the SCN and anterior hypothalamus; (b) to the
ventral lateral hypothalamic area; (c) rostrally to the ventral
preoptic area and (d) to the bed nucleus of the stria
terminalis (Cooper et al., 1994) (earlier identified as projecting to the anterodorsal thalamic nucleus; Johnson et al.,
1988a). With very few exceptions, IGL projections innervate
the retinorecipient regions, although the extent of IGL
projections is substantially greater than those from the
retina (Morin and Blanchard, 1999). In a similar vein, areas
receiving IGL input are also very likely to receive projections
from the SCN (Morin et al., 1994) and from the olfactory
system (Cooper et al., 1994).
IGL connections with the midbrain and brainstem are just
as extensive as those for the forebrain. The concept of a
“subcortical visual shell” has been invoked to facilitate
discussion of connectivity of the IGL (Morin and Blanchard,
1998). The subcortical visual shell consists of 12 contiguous
retinorecipient nuclei of the thalamus and midbrain. All but
those projecting to cortex (lateral posterior and dorsal lateral
geniculate nuclei) receive both ipsi- and contralateral projections from the IGL. In addition, most of the regions also supply
reciprocal input to the IGL bilaterally (Morin and Blanchard,
1998), although ipsilateral connections are more robust. The
functional importance of IGL connections with nuclei of the
subcortical visual shell has not yet been tested. However, the
presence of bilaterally symmetrical connectivity strongly
suggests involvement of the IGL in the regulation of visuomotor function (for a review, see Morin and Blanchard, 1998;
Horowitz et al., 2004).
The further suggestion has been made that the IGL may
contribute to two discrete classes of events, one concerning
visuomotor, and the other circadian rhythm, regulation
(Horowitz et al., 2004, 2005; Morin and Blanchard, 2005). The
35
IGL has at least two classes of neurons identified by their
projections to the SCN, one being so connected and the other
not. Those IGL cells that contribute GHT input to the SCN
appear not to receive any input from the pretectum or superior
colliculus despite abundant IGL-afferent projections from
those areas (Morin and Blanchard, 1998; Horowitz et al.,
2004). This further suggests that photic input to the pretectum
and tectum does not influence circadian rhythmicity by
passing through the IGL and GHT. This inference is supported
by lesion studies indicating that loss of the IGL, but not visual
midbrain areas afferent to the IGL, modifies circadian rhythm
response to light (Morin and Pace, 2002).
There have been recent reports describing brainstem
origins of IGL-afferent connections (Horowitz et al., 2004;
Vrang et al., 2003). Two of these, a serotonergic input from
the dorsal raphe and a noradrenergic projection from the
locus coeruleus, have been previously documented (MeyerBernstein and Morin, 1996; Kromer and Moore, 1980). Other
regions sending projections to the IGL include several
nuclei of the oculomotor complex, the caudal cuneiform,
pedunculopontine and laterodorsal tegmental nuclei (Horowitz et al., 2004; Vrang et al., 2003). In addition, projections
from the pararubral nucleus (which also is retinorecipient)
and vestibular nuclei to the IGL have been described
(Horowitz et al., 2004). Several of these regions, including
the medial vestibular nucleus, locus coeruleus, divisions of
the dorsal tegmental and pontine tegmental nuclei and the
dorsal raphe, are also recipients of projections from the
hypocretin/orexin neurons in the lateral hypothalamic area
(Horowitz et al., 2005; Mintz et al., 2001). These regions are
also thought to be involved in the modulation of sleep and
arousal processes (Saper et al., 2001). Most also receive
projections directly from the IGL (however, the locus coeruleus and vestibular nuclei do not) (Morin and Blanchard,
2005). Thus, it seems likely that the IGL contributes to the
regulation of sleep and arousal in addition to its role in
circadian rhythm regulation and its hypothesized role as a
mediator of visuomotor function.
8.4.
Function
8.4.1.
Neurophysiology
A novel observation has been made concerning the firing rate
of IGL neurons. In a series of paper, Lewandowski and
colleagues have described a slow oscillation in the rat IGL
that is not evident in adjacent thalamic structures. Maximal
firing rate is approximately 20 Hz occurring as a burst that lasts
about 70 s with the overall oscillation of firing having a period
of about 106 s (Lewandowski et al., 2000). Unlike the firing
pattern observed in the dorsal lateral geniculate, there is no
correlation between IGL cell firing and the pattern of firing in
the visual cortex. A lesion of one IGL does not disrupt
rhythmicity in the other (Lewandowski et al., 2002). The origin
of the oscillatory activity is unknown, but is not exclusively
intrinsic to the IGL as it fails to persist in a slice The function of
this burst oscillation is not known, nor is it even known to be
related to circadian rhythm regulation. However, it is present
only in the light with bursting suppressed by dark (Bina et al.,
1993). The bursting pattern can be obliterated and rendered a
tonic discharge pattern by bicuculline or picrotoxin applied
36
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
intraventricularly (Blasiak and Lewandowski, 2004a). In addition, the IGL burst discharge frequency is increased about 100%
by lesions of the dorsal raphe nucleus (Blasiak and Lewandowski, 2003), although serotonin agonists injected intraperitoneally unexpectedly induce an increase in discharge
frequency as well. The implications of the slow oscillatory
bursting activity are not clear and may be related more to
suspected visuomotor functions of the IGL.
Intriguingly, the ultradian rhythm in IGL neuron burst
discharge is very similar to what has been described as a burst
pattern in SCN neurons (Miller and Fuller, 1992). About 20% of
the cells recorded in and near the SCN burst at a rate of about
10 Hz with an average period of about 143 s. It may be that the
IGL burst rhythm is derived from the SCN region because the
IGL rhythm is lost in a slice preparation (Blasiak and
Lewandowski, 2004b).
8.4.2.
Nonphotic regulation of rhythmicity
One major question concerning the influence of nonphotic
stimuli on circadian rhythm regulation is whether it is
mediated by activity (i.e., muscle/joint movement) or a
correlate of activity. Dark pulses, triazolam and novel wheel
locomotion each elicit large phase advances when administered at CT6 (see Morin, 1994; Mrosovsky, 1996) and each of
these is usually associated with increased wheel running. At
least two experiments have now demonstrated that different
parts of the brain mediate the phase shifts elicited by
different stimuli. In the clearest case (Marchant and Morin,
1999), the benzodiazepine, triazolam, failed to elicit phase
shifts at any circadian time in hamsters sustaining neurotoxic
lesions of the deep superior colliculus. Lesions of the
pretectum were at least partially effective in eliminating
such responses and, because of the lesion method, it has not
been possible to fully distinguish between collicular and
pretectal effects. However, lesions confined to the superficial
layers of the superior colliculus had no effect on phase
response to triazolam.
In contrast to the ability of midbrain lesions to block phase
shift responses to triazolam, the same lesions had no effect on
phase shifts induced by prolonged running in a novel wheel.
Thus, the neural substrates for triazolam and novel wheelinduced phase shifts are different, although each appears to
require an intact GHT as a final common path conveying
information to the SCN (Janik and Mrosovsky, 1994; Wickland
and Turek, 1994; Johnson et al., 1988b; Marchant et al., 1997).
These results support previous observations that chlordiazepoxide (a benzodiazepine) and fentanyl (an opiate mu opioid
receptor agonist), neither of which enhances locomotion in
hamsters, nevertheless induce equivalent phase shifts (Biello
and Mrosovsky, 1993; Maywood et al., 1997; Meijer et al., 2000)
to those induced by triazolam.
The second experiment differentiating response to triazolam from response to novel wheel involved serotoninspecific neurotoxic lesions of the median or dorsal raphe
nuclei. Loss of serotonin neurons from the median (but not
the dorsal) raphe eliminated phase shifts in response to
triazolam treatment (Meyer-Bernstein and Morin, 1998).
Novel wheel access elicited equivalent phase shifts regardless of dorsal raphe, median raphe or control lesions. Thus,
it is clear that while a final common path from the IGL to
SCN mediates the phase shift response to two distinct
stimuli, there are different anatomical substrates subserving
those stimuli. Whether the ability of DD to elicit phase shift
responses corresponding to the NPY-type PRC depends upon
locomotion has not been determined.
NPY, the first neuromodulator identified in the IGL (Card et
al., 1983), has also received the greatest amount of attention
with respect to IGL and GHT function. Stimulation of the IGL
has been thought to elicit phase shifts because of NPY release
from GHT terminals in the SCN (see Morin, 1994). This concept
has been reaffirmed by antagonism of NPY activity in vivo by
infusion of NPY antiserum onto the SCN which blocked phase
shifts in response to novel wheel running (Biello et al., 1994).
As indicated above, cells contributing to the hamster GHT
also contain GABA, neurotensin and enkephalin. Neurotensin
appears not to have been evaluated with respect to its ability
to modify circadian rhythmicity in vivo. However, a CT6
application of neurotensin to the rat SCN in a slice preparation
generally reduced neuron firing (Coogan et al., 2001), while
inducing large phase advances of the firing rhythm (MeyerSpasche et al., 2002).
A small literature exists concerning the function of
enkephalin in the hamster GHT. The delta opioid receptor
visualized by immunohistochemistry is densely distributed
on cells in the SCN, particularly in the ventral and medial parts
of the nucleus (Byku et al., 2000). A few cells in the IGL also are
identifiable from the delta opioid receptor labeling. The
observed distribution of receptor label in the SCN is consistent
with the distribution of enkephalin terminals (Morin et al.,
1992). Delta opioid agonists administered at CT8 or CT10 (but
not at other times) will induce moderate phase advances that
are not related to the amount of activity also induced by the
drugs (Byku and Gannon, 2000a,b). Agonists of kappa or mu
opioid receptors apparently do not elicit phase shifts, although
other reports indicate that fentanyl, a mu opioid agonist, can
generate an NPY-type PRC (Meijer et al., 2000; Vansteensel et
al., 2003b). The delta (but not mu or kappa) opioid agonists are
also able to prevent light-induced phase shifts when administered in advance of a light pulse at CT19 (Tierno et al., 2002).
This result is consistent with a presynaptic inhibitory action
on RHT terminals, as suggested by the receptor localization in
the SCN (Byku et al., 2000). However, electrophysiological
evidence suggests that the primary site of action of the
effective delta opioid agonists might be outside the SCN
(Cutler et al., 1999).
The ability of delta opioid agonists to block light-induced
phase shifts is similar to the effect of NPY. In vivo infusion of
NPY onto the SCN greatly attenuates light-induced phase
advances and delays (Lall and Biello, 2002, 2003a,b; Weber and
Rea, 1997). In contrast, infusion of NPY antiserum onto the
SCN augments phase advances to light during the late
subjective night (Biello, 1995). Wheel running, which is
thought to release NPY into the SCN through the GHT (Biello
et al., 1994), appears to attenuate light-induced phase
advances, but not delays (Mistlberger and Antle, 1998; Ralph
and Mrosovsky, 1992). Inhibition of phase response to light is
observable in the slice preparation if the light pulse is
administered in vivo followed by removal of the brain and
acute treatment of the SCN with NPY (Yannielli and Harrington, 2000). Similarly, NMDA stimulation of the SCN slice yields
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
large phase shifts that are blocked by coapplication of NPY
(Yannielli and Harrington, 2001; Yannielli et al., 2004) and the
NPY-blocked phase response to NMDA can be restored by
simultaneous treatment with an NPY Y5 (but not Y1) receptor
antagonist. NPY induces phase advances when applied by
itself to the SCN slice during the subjective day, but no
observable phase shift during the subjective night (Harrington
and Schak, 2000; Biello et al., 1997b). In vivo studies using
intracranial injections showed that a Y5 receptor antagonist
enhances light-induced phase shifts. If the locomotor rhythm
phase response to light is blocked by intracranial NPY, the
shifts can be restored to normal by coinjection of NPY and the
Y5 receptor antagonist (Lall and Biello, 2003a; Yannielli et al.,
2004). Additional in vivo studies also demonstrated probable
involvement of the Y5 receptor in the NPY suppression of
light-induced phase shifts (Lall and Biello, 2003a) and lend
credibility to the view that light and NPY (consequent to
locomotor activity) normally act together to regulate phase
shift magnitude (Lall and Biello, 2002).
The Mistlberger lab has added a new twist to the topic of
nonphotic control of circadian rhythm phase. Initially, there
were two significant observations. One was that 24 h forced
treadmill activity blocked light-induced phase delays, whereas simple treadmill activity simultaneous with the light pulse
did not alter phase. The second observation yielded the
converse result, namely, a shorter interval on the treadmill
(6 h) blocked phase response to light (Mistlberger et al., 1997).
Studies were then performed, using the Aschoff type II
procedure, to elaborate on the ability of “sleep deprivation” to
induce rhythm phase shifts. In these experiments, sleep
deprivation was accomplished using gentle handling (not
confounded by simultaneous locomotion) beginning at CT6. A
3 h procedure elicited a significant phase advance shift and a 1
h procedure yielded an even greater (about 150 min) shift
(Antle and Mistlberger, 2000). The shifts appear to occur in
about two thirds of the animals and it was observed that the
fewer the number of interventions needed to keep an animal
awake, the larger the phase shift to the sleep deprivation
procedure. Additionally, exposure to light (10 lx) during a 3
h gentle handling interval effectively eliminated phase
advances. This observation suggests that the phase shifts
consequent to “gentle handling” are not caused by the absence
of sleep, but rather by a correlate. Such a correlate could be
“stress” generated by the sleep deprivation procedure.
In a comprehensive series of experiments (Mistlberger et
al., 2003), several of the stress-related issues consequent to
sleep deprivation were clarified. In particular, stress as
reflected by elevated cortisol levels, is not associated with
phase advance shifts, nor are shifts elicited by restraint which
also elevates cortisol (Mistlberger et al., 2003). Nor is sleep
deprivation caused by the “pedestal over water” method
associated with phase shifts. In contrast, mere access to a 1
m2 open field did cause phase shifts similar in magnitude to
those resulting from gentle handling. Phase shift magnitude
was related to the amount of forward locomotion and those
animals classified as “shifters” engaged in considerably more
locomotion than the nonshifters. The conclusion of these
studies is that some level of locomotion is indeed necessary to
achieve phase advance shifts (Mistlberger et al., 2003).
Observations that such shifts occur following the simple
37
intraperitoneal injection of saline (Mead et al., 1992; Hastings
et al., 1992) may reflect an action of this stimulus complex
through an alternate route to the circadian clock not requiring
locomotion for phase shifts (cf., benzodiazepine; Marchant
and Morin, 1999; Biello and Mrosovsky, 1993), or the response
to saline may in fact include a locomotor component not
measured in those studies.
One variation on the theme of nonphotic stimuli capable of
inducing phase advances has involved use of “dark pulses”
administered at CT6. The dark pulse is associated with
increased locomotion and consequent phase shifts in hamsters housed in LL (Boulos and Morin, 1985; Boulos and Rusak,
1982). Phase advance responses can be greatly increased (up to
7 h) if the dark pulses coincide with the “gentle handling”
paradigm for sleep deprivation (Mistlberger et al., 2002).
Although the reference to dark pulses as a nonphotic stimulus
implies that it is of the same class as benzodiazepines, novel
wheel, gentle handling and saline injection, the mechanism of
action of dark pulses is not necessarily the same. In particular,
IGL lesions block the ability of triazolam (Johnson et al., 1988b),
novel wheel (Janik and Mrosovsky, 1994; Wickland and Turek,
1994) and saline injection (Maywood et al., 1997) (gentle
handling has not been tested) to elicit phase advances, but
they apparently do not block such responses to dark pulses
(Harrington and Rusak, 1986).
The effect of restraint has also become more ambiguous
than expected when considered in the context of dark pulses.
Restraint during the midsubjective day has no effect on
rhythm phase (Van Reeth et al., 1991; Rosenwasser and
Dwyer, 2002). However, during the late subjective day, brief
or 6 h restraint will induce phase delays. At both circadian
times, restraint blocks the ability of a dark pulse to induce a
phase advance (Rosenwasser and Dwyer, 2002; Van Reeth and
Turek, 1989; Reebs et al., 1989). Thus, on the one hand, phase
advances during the midsubjective day are blocked by
unknown mechanism invoked by restraint, but one that is
apparently not directly related to the circadian clock; on the
other hand, blockade of the dark-induced phase advance at
dusk may be the simple consequence of a restraint-induced
phase shift of equal magnitude but opposite direction (Rosenwasser and Dwyer, 2002; Dwyer and Rosenwasser, 2000).
8.4.3.
Photic regulation of rhythmicity
One major function of the IGL is modulation of the circadian
period during LL conditions (Pickard et al., 1987; Harrington
and Rusak, 1986). However, the visual system projects to a
variety of midbrain regions that connect with the IGL leaving
the possibility that photic information effecting the circadian
clock acts indirectly through, rather than directly in, the IGL.
This perspective has been enhanced by the demonstration that
visual midbrain structures are involved in circadian rhythm
response to a benzodiazepine and that phase shift magnitude
to a novel wheel stimulus is modulated by light (Marchant and
Morin, 1999). IGL lesions reduce the circadian period lengthening effect of LL by about 50%, but do not alter the normal
period in DD (Morin and Pace, 2002). This result suggests the
IGL mediates the tonic effects of light. In contrast, large lesions
of the pretectum and tectum that eliminate rhythm response
to benzodiazepine treatment (Marchant and Morin, 1999) have
no effect on circadian period in LL (Morin and Pace, 2002).
38
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
In mouse, IGL lesions lengthen the circadian period of DDhoused mice, but do not modify the period lengthening effect
of LL (Pickard, 1994; Lewandowski and Usarek, 2002). In the rat,
IGL lesions do not alter the circadian period in DD nor do they
prevent LL-induced circadian rhythm disruption (Edelstein
and Amir, 1999a). Period in LL before rhythm disruption was
not assessed. Under skeleton photoperiod conditions (1 h of
light from ZT0-1 and ZT11-12, rats with IGL lesions generally
fail to entrain indicating the necessity of the IGL under these
conditions; Edelstein and Amir, 1999b). Equivalent studies
have not been done in other species.
Just as NPY is able to attenuate light-induced phase shifts,
effects of NPY on phase are also inhibited by light. Novel
wheel-induced phase advances are greatly attenuated by a
subsequent light pulse. Similarly, NPY-induced phase advances are curtailed by a light pulse (Biello and Mrosovsky, 1995).
A glutamate agonist (NMDA), injected onto the SCN to mimic
the effect of photic input, also greatly reduces the phase
advance effects of NPY administered in the midsubjective day
(Gamble et al., 2004). At CT20, NMDA injection onto the SCN
induces phase advances (Mintz et al., 1999), but when given
simultaneously with NPY, small phase delays occur (Gamble
et al., 2004). This result, in conjunction with the observation
that the effect occurs in the presence of TTX, suggests that
NPY is acting postsynaptically to induce phase shifts, rather as
a presynaptic inhibitor of RHT function.
Glutamate receptor antagonists, known to modify SCN and
rhythm response to light (Abe et al., 1992; Rea et al., 1993b;
Vindlacheruvu et al., 1992), fail to alter light-induced FOS
expression in the rat IGL (Edelstein and Amir, 1998). This is
consistent with the likelihood that there appears to be little or
no glutamatergic innervation of the IGL (Fujiyama et al., 2003).
9.
Serotonin and midbrain raphe contribution
to circadian rhythm regulation
The 1994 review (Morin, 1994) fell victim to the typical problem
of being partially out of date by the time it was published. No
topic was more affected by this than serotonin and its effect
on circadian rhythmicity. In the year of publication, research
on the topic began to blossom and there have been several
major reviews published (Morin, 1999; Rea and Pickard, 2000a;
Mistlberger et al., 2000; Yannielli and Harrington, 2004).
One of the more important changes in perspective
concerning the role of serotonin has been a change of focus
away from the dorsal raphe nucleus, previously thought to be
the source of SCN innervation by serotonin fibers (Smale et al.,
1990; Azmitia and Segal, 1978). Anatomical studies, using three
separate methods (anterograde and retrograde tract tracing of
raphe projections, and lesion studies designed to destroy
raphe projections), have unequivocally demonstrated that the
median raphe sends mixed serotonergic and nonserotonergic
projections to the SCN, and the dorsal raphe sends mixed
serotonergic and nonserotonergic projections to the IGL (Moga
and Moore, 1997; Meyer-Bernstein and Morin, 1996; HaySchmidt et al., 2003). This simple anatomy is complicated by
the presence of reciprocal serotonergic and nonserotonergic
connections between dorsal and median raphe nuclei (Tischler
and Morin, 2003).
9.1.
Effects of serotonin-specific neurotoxins
The sense of the literature is that, in hamsters, serotonin has
an inhibitory effect on rhythm response to light which, in
animals lacking median raphe serotonin neurons, is evident
as a larger amplitude PRC, greater prevalence of, and shorter
latency to, light-induced rhythm splitting and longer circadian
periods under LL (Meyer-Bernstein and Morin, 1996; Morin and
Blanchard, 1991; Bradbury et al., 1997). No effects on rhythmicity are seen following destruction of only the dorsal raphe
serotonergic cells. Most recently, evaluation of irradiance
response curves in animals with neurotoxic lesions of median
raphe serotonin neurons has shown similar sensitivities of
lesioned and unlesioned controls to light, but a greater phase
response by lesioned animals to high intensity (probably
saturating) light (Muscat et al., 2005).
The drug of abuse, MDMA (‘ecstasy’), is rapidly toxic to
serotonin terminals (Battaglia et al., 1991). Treatment of rats
with this drug for several days reduces phase advances of the
rhythm in SCN firing rate by about 50% in response to 8-OHDPAT applied to a slice at ZT6 (Biello and Dafters, 2001). The
effects of MDMA on rhythmicity have been extended to the
evaluation of its effects on hamsters in vivo. After drug
injections of MDMA, phase advances to 8-OH-DPAT administered at CT8 were blocked. Light-induced phase delays were
convincingly reduced about 50% (Colbron et al., 2002),
although the effect is not consistent with the literature
showing that serotonin loss increases phase response to
light. Damage to the serotonin terminals varies according to
brain region, with some areas being completely unaffected by
the drug (Battaglia et al., 1991). In the above MDMA studies, it
is unclear as to which part of, or to what extent, the serotonin
system has been damaged.
9.2.
Serotonin receptor function
A major development in the study of serotonergic effects on
circadian rhythm regulation has been the increased focus on
particular receptor types and their function. One critical
development has been the recognition that 8-OH-DPAT, long
thought to a 5HT1A-specific agonist and frequently used
because of that specificity (see Morin, 1999), is actually specific
to the 5HT1A, 5HT5A and 5HT7 receptors (Lovenberg et al.,
1993; Sprouse et al., 2004a; and reviewed by Gannon, 2001).
There are now reports indicating that the SCN contains the
5HT1A, 5HT1B, 5HT2A, 5HT2C, 5HT5A and 5HT7 receptor types
(Sprouse et al., 2004a,b; Belenky and Pickard, 2001; Duncan et
al., 1999, 2000; Prosser et al., 1993; Oliver et al., 2000; Moyer and
Kennaway, 1999). A useful review of the evidence favoring the
presence and function of 5HT7 receptors in the SCN was
published in 2001 (Gannon, 2001), but needs to be interpreted
in the context of subsequent information.
The 5HT5A receptor has most recently been appreciated as
a serotonin receptor type contributing to circadian rhythm
regulation. It is distributed in the dorsal and median raphe, as
well as in the SCN and IGL (Duncan et al., 2000). Thus, the
5HT1A, 5HT5A and 5HT7 receptors are found in similar places
and respond to similar agonists/antagonists thereby increasing the difficulty of discerning receptor-specific function. One
suggestion is that 8-OH-DPAT injection into the dorsal raphe
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
induces phase shifts via 5HT7 (but not 5HT5A) receptor
activation (Duncan et al., 2004), whereas systemic injection
of the drug induces phase shifts by activating the 5HT1A
receptor (Tominaga et al., 1992). The phase response of
hamsters to dorsal raphe application of serotonin 5HT1A/5A/
7 agonists diminishes with age (Duncan et al., 2004; Penev et
al., 1995), and this may be the result of specific loss of available
5HT7 receptors (Duncan et al., 1999).
In the rat, the 5HT2C receptor may contribute to the
regulation of circadian rhythm phase. The receptor type is
present in the SCN (Moyer and Kennaway, 1999). Agonists of
the 5HT2C receptor are able to induce phase delays in rats and
induction of FOS in SCN neurons at CT18, but not at CT6
(Kennaway and Moyer, 1998), and generally mimic the effects
of light. The suggestion has been made (Kennaway and Moyer,
1998) that previous observations of quipazine-induced phase
shifts in rat (Prosser et al., 1993; Kennaway and Moyer, 1998)
may have been the result of that drug's affinity for the 5HT2
class of receptors. Kennaway and Moyer (1998) suggest that the
literature is ambiguous with respect to the certainty that the
rat SCN contains 5HT1A and 5HT7 receptors. Further, they
suggest the possibility that light may act through the known
retina-dorsal raphe projection in the rat (not evident in the
hamster; see Section 4.2) and it is the modulation of the 5HT2C
receptors on this pathway that results in light-like phase
response and FOS induction (Kennaway and Moyer, 1998).
Consistent with this view is the observation that reduction of
serotonin in rats by pCPA injection attenuates light-induced
FOS (Moyer and Kennaway, 2000), suggesting that normal
serotonin innervation is necessary for light-induced FOS.
Agonist activation of 5HT2A/C receptors induces substantially
(but not completely) light-like FOS expression in the rat SCN
(Kennaway et al., 2001). The investigators suggest that, in the
rat, but probably not in the mouse or hamster, there are two
pathways by which light can modulate SCN rhythmicity. One
of these is the RHT, while the other involves the retinal
projection to the dorsal raphe which, it is hypothesized, sends
a direct serotonergic projection to the SCN that effects
glutamate release at that location (Kennaway et al., 2001). A
dorsal raphe projection to the SCN is probably absent in the rat
(Moga and Moore, 1997), as it is in hamster, but there is
abundant dorsal raphe innervation of adjacent hypothalamus
(Moga and Moore, 1997; Vertes, 1991) and peri-SCN neurons
modulated by serotonin from the dorsal raphe may provide
glutamate signaling to the SCN. Involvement of the 5HT2C
receptor in rat rhythm regulation is further supported by the
observation that agonists specific for this type of receptor
induce expression of FOS, Per1 and Per2 in the SCN. This effect
was more robust at ZT16 than ZT22 (Varcoe et al., 2003).
The issue of species differences has been addressed by
direct comparison of mouse and hamster responses to a
variety of 5HT1A, 1B, 2 or 7 agonists. The results are complex
with differing phase and spontaneous Fos-induction
responses in each species, plus differing effects on lightinduced phase shifts and Fos-induction. In addition, response
has also been shown to be related to whether the drug was
injected centrally (SCN) or administered peripherally (Antle et
al., 2003b). One aspect of the complexity of serotonergic
activity relative to the effects of light is the demonstration
that phase response to agonists can vary enormously depend-
39
ing upon prior light exposure (Knoch et al., 2004). Hamsters
exposed to LL for as little as 1 day can have very large phase
shifts in response to intracranial NPY given at CT6, a sleep
deprivation procedure at ZT18 or 8-OH-DPAT at ZT18 or ZT21
(Aschoff type 2 test). LL exposure also eliminates the early
night time peak in peri-SCN serotonin and reduces the overall
level to approximately the typical late afternoon levels seen
under LD conditions (Knoch et al., 2004).
The 5HT1B receptor has received substantial attention with
respect to its role in rhythm regulation. The receptor is
identifiable in the SCN, specifically on terminals of the RHT
(Belenky and Pickard, 2001; Pickard et al., 1999) and there is an
approximately 35% reduction in 5HT1B binding sites in the
SCN after bilateral enucleation (Pickard et al., 1996; Manrique
et al., 1999). 5HT1B receptor agonists markedly attenuate lightinduced phase shifts and the induction of FOS in the SCN,
although cells in a central region that appears to include the
CALB neurons continue to express FOS (Pickard et al., 1996,
1999; Pickard and Rea, 1997; Shimazoe et al., 2004). Lightinduced suppression of nocturnal melatonin is also alleviated
by treatment with 5HT1B agonists (Rea and Pickard, 2000b).
Infusion of receptor agonist directly onto the SCN eliminated
phase response to light (Pickard et al., 1996).
Use of the 5HT1B receptor knockout mouse has yielded
substantial additional data. Phase shifts elicited by light in this
strain are not modified by 5HT1B receptor agonists. Electrical
stimulation of the optic nerve or glutamate application to the
bath elicit excitatory postsynaptic currents (EPSCs) in SCN
neurons recorded in a slice preparation. Agonists for the
5HT1B receptor greatly reduce EPSCs in the wild-type mouse,
but have no effect in the 5HT1B receptor KO mouse (Smith et
al., 2001; Pickard et al., 1999). Although the circadian period of
wild-type mice does not differ from that of the 5HT1B KO mice,
the period is longer in LL consistent with the view that this
strain lacks inhibitory control of retinal input to the circadian
visual system (Sollars et al., 2002). Most recently, bicucullinesensitive miniature inhibitory currents were recorded on SCN
(which are predominantly GABAergic; Moore and Speh, 1993;
Morin and Blanchard, 2001). Frequency of the inhibitory currents is reduced by a 5HT1B receptor agonist, but the agonist
had no effect on the currents in SCN neurons of 5HT1B KO mice
(Bramley et al., 2005). The authors suggest that abundant
5HT1B receptors are present presynaptically on GABA terminals, and thereby modulate GABA release within the SCN.
9.3.
Regulation and consequences of serotonin release
Extracellular serotonin increases abruptly from daytime levels
to higher night time levels in both the peri-SCN and peri-IGL
region (Dudley et al., 1998; Grossman et al., 2004). Similarly,
extracellular serotonin increases in both the SCN and IGL in
response to certain nonphotic stimuli (Dudley et al., 1998;
Grossman et al., 2000, 2004). Serotonin availability in these
nuclei is also augmented by electrical stimulation of the dorsal
raphe nucleus (Grossman et al., 2004; Dudley et al., 1999; Glass
et al., 2000b, 2003b).
Despite these effects on serotonin release, it is not yet
known to what extent the stimulation-induced transmitter
increase in either the SCN or IGL contributes to circadian
rhythm phase control. Studies involving electrical stimulation
40
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
of the dorsal or median raphe in which serotonin neurons have
been destroyed have not been performed. Arguing against a
normal effect of serotonin on rhythm phase control is the
observation that treatment of hamsters with a 5HT1A receptor
antagonist elevates serotonin release in the peri-SCN region to
levels comparable to those seen during novel wheel access or
raphe electrical stimulation without augmenting phase shifts
in response to novel wheel activity (Antle et al., 2000). Nor do
systemic or SCN injections of 5HT1/2, 5HT1A and 5HT2/7
receptor antagonists alter novel wheel-induced phase shifts
(Antle et al., 1998). Consistent with those results is the
observation that the serotonergic median raphe projections
appear to be essential for triazolam-induced phase shifts, but
not for those induced by novel wheel activity (Meyer-Bernstein
and Morin, 1998).
It is possible that the electrical stimulation activates
nonserotonergic raphe cells projecting to the SCN or IGL
(Meyer-Bernstein and Morin, 1996). The most directly related
data on the topic come from the mouse in which a serotoninspecific neurotoxin was injected intracranially into the region
of the SCN (Marchant et al., 1997; Edgar et al., 1997) and, in
combination with IGL lesions, block activity-related phase
control in the mouse. The suggestion is that the serotonergic
projections to the SCN destroyed by the neurotoxin are
necessary for such phase control. However, in the experiments,
serotonin depletion occurred widely within the diencephalon
and localization to the SCN cannot be assured. In the hamster,
small injections of neurotoxin that destroyed nearly all
serotonergic innervation to the SCN without damaging most
of the surrounding hypothalamus had very little effect on
circadian rhythmicity compared to the consequences of loss of
all median raphe projections (Meyer-Bernstein and Morin, 1996;
Meyer-Bernstein et al., 1997; Bobrzynska et al., 1996). There are
two implications of these studies. One is that median raphe
projections to non-SCN targets (also destroyed by intraraphe
neurotoxin injection) may contribute to the regulation of
hamster circadian rhythmicity. The other is that the dense
serotonergic projection to the SCN (destroyed by the intra-SCN
neurotoxin) may be minimally involved in such regulation.
One perplexing issue has been the fact that electrical
stimulation of the dorsal or the median raphe nuclei yields
similar release of serotonin in the SCN (Dudley et al., 1999) and
stimulation of either nucleus during the subjective day elicits
equivalent magnitude phase advances (Meyer-Bernstein and
Morin, 1999). A model has been developed (Glass et al., 2003b)
that provides a reasonable reconciliation of behavioral,
pharmacological and anatomical data (Fig. 5; see Glass et al.,
2003b for an extensive explanation and presentation of the
data implicating GABA). The similar effectiveness of stimulation occurring in either dorsal or median raphe may be
possible because of the serotonergic–nonserotonergic interconnection between the two nuclei (Tischler and Morin, 2003).
9.4.
Serotonergic potentiation of light-induced phase shifts
It is generally the case that a manipulation during the
subjective night that modifies phase advances will also
modify phase delays. Such is not true for the drug BMY 7378.
Administered at CT19, this drug potentiates the phase shifting
effect of light given shortly thereafter. However, no such effect
Fig. 5 – Diagrammatic conceptualization of the functional
serotonergic circuitry governing circadian rhythm regulation.
Based on an unpublished update provided courtesy of Dr. J.D.
Glass, modified from the previous version (Glass et al.,
2003b). The presumptive intra-IGL GABA connection is based
upon unpublished work from the Glass laboratory. There are
data for (Glass et al., 2003b) and against (Meyer-Bernstein and
Morin, 1998) the likelihood that behavioral input induces
phase shifts by acting through the serotonin neurons of the
dorsal raphe nucleus.
occurs when the drug/light combination occurs at CT14. The
reason for this is not clear, although it may result from
cancellation of light-induced phase delays by drug-induced
phase advances when each is administered at CT14. At CT19,
BMY 7378 alone induces small (1 h) phase advances, but the
CT19 combination of light and drug yields shifts more than
300% larger than those induced by light alone (light-induced
shifts are about 1.5 h; potentiated shifts are about 5 h; Gannon,
2003). This phenomenon is similar to what has been obtained
with other drugs with different effects on phase response in
the absence of light (Gannon, 2003; Rea et al., 1995; Moriya et
al., 1998). For example, NAN-190 administered alone at CT14 or
CT19-20 has essentially no effect on rhythm phase (Rea et al.,
1995; Moriya et al., 1996), but potentiates both phase advances
and delays when administered at those times in combination
with light (Rea et al., 1995). One interpretation (Gannon, 2003)
is that the drugs, which have mixed agonist/antagonist
properties, facilitate light-induced phase shifts by acting as
agonists on presynaptic 5HT1A receptors of median raphe
serotonin cell dendrites to inhibit release of serotonin in the
SCN. Elsewhere in the brain, the drugs act on postsynaptic
5HT1A receptors which are, presumably, antagonized by the
drugs. WAY 100635, a 5HT1A antagonist, has no effect on
light-induced phase advances (Smart and Biello, 2001), and it
is thought that the particular combination of raphe cell
autoreceptor activation plus the antagonism of SCN postsynaptic receptors is necessary for the large light-induced phase
advances potentiated by the mixed agonist/antagonists
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
41
(Gannon, 2003). If this hypothesis is correct, then it may be
possible to mimic the effect of the mixed agonist/antagonist
drugs by concomitant injection of a 5HT1A antagonist and a
5HT1A agonist known to act on raphe autoreceptors.
The complex control of serotonergic involvement in
rhythm regulation is further suggested by the observation
that depletion of brain serotonin (88% lower in the peri-SCN
region) by systemic pCPA treatment greatly augments phase
shifts in response to SCN application of serotonin or the
5HT1A/7 receptor agonist, 8-OH-DPAT (shifts are about 100 min
with pCPA treatment vs. 30 min without) (Ehlen et al., 2001).
The results suggest that a necessary condition for normal
phase response to agonists is lack of activity at the postsynaptic receptors. Short-term changes in postsynaptic receptor
number or sensitivity to endogenous or exogenous agonists
may be critical to their effect on rhythm phase (see Ehlen et al.,
2001 for a discussion).
In animals pretreated with pCPA, phase advances resulting
from 8-OH-DPAT infusion onto the SCN at CT6 were reduced by
about 68% in animals treated with 5HT7 receptor antagonists
(Ehlen et al., 2001). On the one hand, the result supports the
view that 5HT7 receptors mediate the effect of serotonin
agonists applied to the SCN. On the other, there is a significant
residual phase shift effect of 8-OH-DPAT that may be
attributable to its affinity for the 5HT1A receptor. It is also
interesting that light stimulation simultaneous with 8-OHDPAT (i.e., CT6) in pCPA treated animals completely blocks the
phase shifting effect of the 5HT1A/7 agonist (Ehlen et al., 2001).
The in vivo result is consistent with observations that
glutamate application to the SCN slice preparation, which
presumably mimics the excitatory effect of light exposure,
blocks the phase shifting effect of NPY or serotonin (Biello et
al., 1997a; Prosser, 1998). The presumption is that the action of
serotonin takes place in the SCN, although there is evidence for
IGL involvement in the response to 8-OH-DPAT as well (Challet
et al., 1998b).
10.
Failure to re-entrain
It is axiomatic that an entrained animal will re-entrain to a
shifted light–dark cycle. In this era of transgenic animals
specifically designed to show abnormal behavioral traits that
will allow novel probing of the way things usually work, it is
ironic that one of the more provocative developments during
the last decade has been the discovery that a standard
laboratory rodent, the Siberian hamster, is a model for
failure to re-entrain. The initial observation occurred in both
young and old animals entrained initially to an LD 16:8
photoperiod, then subjected to a 5 h delay of the photoperiod. About 90% of animals had free-running locomotor and
body temperature rhythms for many weeks after the shift or
became arrhythmic (Ruby et al., 1996, 1997) (Fig. 6).
Melatonin treatment, known to modify the phase angle of
entrainment in Siberian hamsters (Margraf and Lynch, 1993),
reduced the percentage of animals failing to re-entrain to a
shifted photoperiod (Ruby et al., 1997). Analysis of melatonin
levels has shown that animals subjected to a 5 h phase delay
have very low circulating melatonin at a time that it is
expected to be high (Ruby et al., 2000), suggesting that the
Fig. 6 – Failure to re-entrain to a shifted photoperiod by
Siberian hamsters. The initial photoperiod was LD16:8 (white
and black bars at the top of each record; lights off = ZT12)
which, on the day of the arrowhead, was changed. On that
day at ZT17, the lights were turned on for 2 h. The next day,
the LD16:8 photoperiod was delayed by 3 h. Shading areas
indicate the times of lights off under the modified
photoperiod. This combination of light manipulations
prevents re-entrainment in most animals. The expected
re-entrainment (A) sometimes follows the phase shift, but is
less likely to occur than the lack of re-entrainment (B, C). One
case (C) demonstrates neither re-entrainment nor a clear
free-running circadian rhythm, and a significant percentage
of animals become arrhythmic. Unpublished data courtesy of
Dr. N.F. Ruby.
42
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
absence of melatonin may contribute to the inability to
achieve a stable phase angle of entrainment after the shift.
Siberian hamsters re-entrain properly to 1 or 3 h delays of
photoperiod phase, and to 1 h advances, but much less well to 3
h advances. Generally, these animals do not re-entrain properly following 5 h advance or delay shifts in the photoperiod,
although the effect is modifiable by the exact manner in which
the photoperiod is shifted (Ruby et al., 1998, 2004). If Siberian
hamsters are exposed to a 5 h phase delay in the photoperiod,
then immediately placed in DD for 14 days, all animals will reentrain when the shifted photoperiod is resumed. If DD
exposure is delayed for several days, re-entrainment probability is reduced to less than 50% (Ruby et al., 2002b).
The Siberian hamster is noted for the existence of
individuals in the general population that fail to undergo gonadal regression when exposed to a short day photoperiod and
breeding has selected this trait in a strain of animals identified
as ‘nonresponders.’ Photoperiodic time measurement for reproduction control relies on the use of a circadian clock (Elliott
et al., 1972; Rusak and Morin, 1976; Stetson and WatsonWhitmyre, 1976). Comparison of the responder and nonresponder strains with the phase shift paradigm has shown that
they also have the traits of re-entrainment and nonreentrainment, respectively (Ruby et al., 2004).
Most recently, Siberian hamsters that fail to re-entrain to a 5
h photoperiod shift have been exposed to light to induce SCN
gene activity. In animals that re-entrain, Fos and Per1 gene
activity is induced, as expected, in SCN neurons. In contrast,
gene induction by light apparently does not occur in animals
that fail to re-entrain (Barakat et al., 2004). This is conceivably
related to the absence of light-induced gene expression in
golden hamsters showing split rhythms in LL (De la Iglesia et al.,
2000). The collective results of these studies indicate that the
lighting conditions dictate subsequent response of the Siberian
hamster circadian system to light. The most extreme conclusion is that a consequence of the 5 h phase shift renders the
circadian rhythm system unresponsive to the effects of light.
11.
Masking
Mrosovsky (1999) is synonymous with masking and has
provided a substantial review of the topic. Masking is
important to the study of circadian rhythms for several
reasons. One is purely technical to the extent that, under a
variety of circumstances in which there are questions
concerning whether the evident rhythm is imposed by the
environment or is from endogenous sources, tests must be
conducted to obtain an explicit answer. For example, mice
lacking the genes coding for cryptochrome1 (Cry1) and
cryptochrome2 (Cry2) show nocturnal locomotor activity
under LD conditions, but are arrhythmic under DD (Albus
et al., 2002; Van der Horst et al., 1999). The transfer into DD is
the test of persistent rhythmicity with the same initial phase
as seen under LD conditions. In the case of the Cry1/2 KO
mice, the result of the transfer demonstrated that the
absence of wheel running during the light portion of the
previous daily photoperiod occurred because the running
was strongly suppressed by the negative masking effects of
light.
A second major reason for studying masking concerns the
anatomy of the visual system. Light-induced masking is a
visual function, but the visual system anatomy contributing to
it has not been satisfactorily elucidated.
Normal mice generate a characteristic irradiance response
curve when tested for light-induced masking of running wheel
revolutions. Through a middle range of intensities, the
stronger the stimulus, the greater the extent of negative
masking. With very dim stimuli, activity level actually
increases slightly (positive masking; see Mrosovsky, 1999;
Redlin and Mrosovsky, 1999a). At the level of the retina, both
classical and melanopsin photoreceptors have been shown to
contribute to masking (Hattar et al., 2003; Panda et al., 2003).
Melanopsin cells influence masking responses to the higher
light intensities (Mrosovsky and Hattar, 2003). Loss of rod
function in mice eliminates the positive masking seen at dim
illumination levels, but for some strains, there may also be a
significant reduction in the masking response to higher
intensities (Mrosovsky et al., 1999, 2000).
The analysis of visual projections contributing to negative
masking remains incomplete. Mrosovsky and colleagues have
evaluated contributions of the visual cortex, superior colliculus, lateral geniculate and SCN without obtaining data
demonstrating that one of these areas is necessary for negative
masking. In fact, the most common effect of lesions is actually
increased masking. This is true for lesions of the visual cortex,
superior colliculus, dorsal lateral geniculate and IGL, although
it has not yet been possible to distinguish the effects of
dorsolateral geniculate lesions from lesions aimed at the IGL
(Edelstein et al., 2001; Redlin et al., 1999, 2003). Lesions of the
SCN render animals arrhythmic and masking experiments
must, of necessity, be conducted differently from those in
which animals have normal circadian locomotor rhythms.
When SCN lesioned animals are given access to running
wheels in a 3.5 h light:3.5 h dark schedule, about 90% of their
wheel revolutions occur during the dark intervals and the
same animals show a strong preference for darkened vs. a
lighted chamber (Redlin and Mrosovsky, 1999b). These results
indicate that negative masking is not abolished by SCN lesions.
By inference, it would also appear that retinal projections
passing through the SCN to the subparaventricular zone and
dorsomedial hypothalamus are not necessary for masking
because they would also be destroyed by the lesions. However,
these conclusions must be tempered by a recent report
indicating loss of negative masking in SCN-lesioned hamsters
(Li et al., 2005). This study did not address possible masking
functions of retinal projections destroyed while passing
through and around the SCN to other hypothalamic sites.
Despite the absence of a demonstration that a particular
structure is necessary for masking, the Mrosovsky work
collectively supports the view that two visual processes
govern the behavior. One, involving thalamus, midbrain and
cortical visual structures, provides a normal inhibitory effect
on the magnitude of the masking response to light. The other,
presumably in one of the two dozen or so other retinorecipient
brain regions (Morin and Blanchard, 1999; Horowitz et al.,
2004) not yet tested, or in the SCN (Li et al., 2005), actually
controls the negative masking response to light.
As indicated above, the rhythmicity of Cry1 and Cry2 double
KO mice under LD conditions has been explained as a
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
phenomenon of negative masking because these mice are arrhythmic under DD conditions (Albus et al., 2002; Van der Horst
et al., 1999). This issue has also been tested in the standard
fashion (1 h light pulse at various levels of illumination). The
double KO animals show masking which is no different from
the wild-type controls, i.e., they are not more sensitive to the
activity suppressing effects of light than the wild-type animals
(Mrosovsky, 2001). These mice show bouts of “predark”
locomotion the magnitude of which may be related to the
prevailing light intensity. These animals may have an altered
phase angle of entrainment that requires specific plotting
methods to discern (see Fig. 3 in Mrosovsky, 2001).
12.
The future
Circadian rhythmicity is a pervasive feature of mammalian
physiology and behavior. Thus, it should not be surprising that
discoveries concerning regulation of the circadian visual
system have importance with respect to understanding the
workings of the brain and body. In this context, the discovery
of melanopsin as a photoreceptive molecule in a subset of
ganglion cells is one of the great strides forward in the study of
the circadian visual system during the last decade. However,
as is already being demonstrated (Dacey et al., 2005), it is
unlikely that the importance of these ganglion cells lies
exclusively with photic regulation of circadian rhythmicity.
The melanopsin cells of the retina are likely to project widely
in the visual system and influence most, if not all, retinorecipient regions.
It is clear that the visual regulation of circadian rhythms
is influenced by the classical visual photoreceptors as well
as the intrinsically photoreceptive ganglion cells. There is an
opportunity for the circadian rhythm field to contribute to
understanding of retinal function by establishing the anatomical and functional relationships between rods, cones
and melanopsin cells with respect to rhythm response to
light.
The most obvious influence of circadian rhythm research
on the understanding of normal bodily function may be
derived from the large amount of research (most of which
has not been considered by this review) concerned with the
molecular regulation of clock-like activity in the SCN. These
studies have discovered a new window into the exploration of
mammalian physiology through the demonstration of oscillating clock genes in peripheral tissues. While not necessarily
circadian oscillations themselves (but see Yoo et al., 2004 to
the contrary), under normal conditions, the genes do oscillate
with many of the same characteristics as those in the SCN
(Yagita et al., 2001). The widespread presence of oscillating
presumptive clock genes undoubtedly has enormous ramifications for the normal functioning of organs as well as in
disease states.
Within the SCN, systematic exploration of the relationship
between cell phenotype and function has been initiated. Part of
this effort has focused on cell level analysis of gene expression.
Another has sought to describe the rhythmic activity of
particular cell types. Both have been directed, at least partially,
toward the intrinsic organization of the whole SCN. The overall
research effort has barely addressed a small fraction of the cell
43
types and already there is typical scientific confusion with one
method showing an unexpected absence of rhythmicity in
certain cells (Hamada et al., 2001), while another method
demonstrates an unexpected form of rhythmicity in the same
type of cell (Hamada et al., 2003). This topic is rich and likely to
contribute substantially to such issues as the bimodal rhythmicity observed in horizontal slices through the hamster SCN,
function-specific rhythm regulation and the general problem
of specifying which cells are pacemakers.
Research focused on the IGL component of the circadian
visual system has refined the view that it contributes to both
photic and nonphotic regulation of rhythmicity. However, the
research on IGL connectivity has pointed toward a function for
this nucleus that may extend well beyond its role in circadian
rhythm regulation. In particular, the IGL appears to have
widespread, bilateral and often reciprocal connections with
brain regions involved in the regulation of sleep, visuomotor
and vestibular system activities. The nature of the functional
relationships among these systems remains to be seen, as is
the circadian system role of the IGL with respect to their
regulation.
The serotonergic system influence on rhythm regulation has
been substantially elucidated, but large parts of that influence
remain a mystery. One characteristic, in particular, is compelling and not yet understood. Certain manipulations of the
serotonergic system contribute to very large, rapid phase shifts
to photic stimuli. How the circadian clock can overcome the
inertia of a stable, entrained system is not understood, but the
presence of such large shifts gives hope that there may be
forthcoming manipulations that can reliably overcome the
detrimental effects of jet lag or modify the severity of timing
disorders such as advanced sleep phase syndrome.
The last decade of research on the circadian visual system
has been extremely exciting and productive. The current
decade will see this trend continue unabated with a much
enhanced recognition of the contributions made by investigators in the field of circadian visual system research to the
understanding of general brain function and the normal
physiology of bodily systems.
Acknowledgments
Supported by NIH grants NS22168 and MH64471 and National
Space Biomedical Research Institute grant HPF0027 via
Cooperative Agreement NCC9-58 with the National Aeronautics and Space Administration (to LPM) and NIH grants
NS36607 and NS40782 (to CNA).
REFERENCES
Abe, H., Rusak, B., 1992. Stimulation of the hamster ventral lateral
geniculate nucleus induces Fos-like immunoreactivity in
suprachiasmatic nucleus cells. Neurosci. Lett. 148, 185–189.
Abe, H., Rusak, B., Robertson, H.A., 1992. NMDA and non-NMDA
receptor antagonists inhibit photic induction of Fos protein in
the hamster suprachiasmatic nucleus. Brain Res. Bull. 28,
831–835.
Abe, H., Honma, S., Shinohara, K., Honma, K., 1996. Substance P
receptor regulates the photic induction of Fos-like protein in
44
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
the suprachiasmatic nucleus of Syrian hamsters. Brain Res.
708, 135–142.
Abe, M., Herzog, E.D., Block, G.D., 2000. Lithium lengthens the
circadian period of individual suprachiasmatic nucleus
neurons. NeuroReport 11, 3261–3264.
Abrahamson, E.E., Moore, R.Y., 2001. Suprachiasmatic nucleus in
the mouse: retinal innervation, intrinsic organization and
efferent projections. Brain Res. 916, 172–191.
Abrahamson, E.E., Leak, R.K., Moore, R.Y., 2001. The
suprachiasmatic nucleus projects to posterior hypothalamic
arousal systems. NeuroReport 12, 435–440.
Aggelopoulos, N.C., Meissl, H., 2000. Responses of neurones of
the rat suprachiasmatic nucleus to retinal illumination
under photopic and scotopic conditions. J. Physiol. 523,
211–222.
Agostino, P.V., Ferreyra, G.A., Murad, A.D., Watanabe, Y.,
Golombek, D.A., 2004. Diurnal, circadian and photic regulation
of calcium/calmodulin-dependent kinase II and neuronal nitric
oxide synthase in the hamster suprachiasmatic nuclei.
Neurochem. Int. 44, 617–625.
Aguilar-Roblero, R., Morin, L.P., Moore, R.Y., 1994. Morphological
correlates of circadian rhythm restoration induced by
transplantation of the suprachiasmatic nucleus in hamsters.
Exp. Neurol. 130, 250–260.
Aida, R., Moriya, T., Araki, M., Akiyama, M., Wada, K., Wada, E.,
Shibata, S., 2002. Gastrin-releasing peptide mediates photic
entrainable signals to dorsal subsets of suprachiasmatic
nucleus via induction of Period gene in mice. Mol. Pharmacol.
61, 26–34.
Aioun, J., Chambille, I., Peytevin, J., Martinet, L., 1998. Neurons
containing gastrin-releasing peptide and vasoactive intestinal
polypeptide are involved in the reception of the photic signal in
the suprachiasmatic nucleus of the Syrian hamster: an
immunocytochemical ultrastructural study. Cell Tissue Res.
291, 239–253.
Akasu, T., Shoji, S., Hasuo, H., 1993. Inward rectifier and
low-threshold calcium currents contribute to the spontaneous
firing mechanism in neurons of the rat suprachiasmatic
nucleus. Pflugers Arch. 425, 109–116.
Albers, H.E., Ferris, C.F., 1984. Neuropeptide Y: role in light–dark
entrainment of hamster circadian rhythms. Neurosci. Lett. 50,
163–168.
Albers, H.E., Ferris, C.F., Leeman, S.E., Goldman, B.D., 1984. Avian
pancreatic polypeptide phase shifts hamster circadian
rhythms when microinjected into the suprachiasmatic region.
Science 223, 833–835.
Albers, H.E., Liou, S.-Y., Stopa, E.G., Zoeller, R.T., 1991. Interaction
of colocalized neuropeptides: functional significance in the
circadian timing system. J. Neurosci. 11, 846–851.
Albus, H., Bonnefont, X., Chaves, I., Yasui, A., Doczy, J., Van der
Horst, G.T.J., Meijer, J.H., 2002. Cryptochrome-deficient mice
lack circadian electrical activity in the suprachiasmatic nuclei.
Curr. Biol. 12, 1130–1133.
Alleva, J.J., Waleski, M.V., Alleva, F.R., 1971. A biological clock
controlling the estrous cycles of the hamster. Endocrinology
88, 1368–1379.
Altman, J., Bayer, S.A., 1979a. Development of the diencephalon in
the rat: V. Thymidine-radiographic observations on
internuclear and intranuclear gradients in the thalamus.
J. Comp. Neurol. 188, 473–500.
Altman, J., Bayer, S.A., 1979b. Development of the diencephalon in
the rat: IV. Quantitative study of the time of origin of neurons
and the internuclear chronological gradients in the thalamus.
J. Comp. Neurol. 188, 455–472.
Amir, S., 1992. Blocking NMDA receptors or nitric oxide production
disrupts light transmission to the suprachiasmatic nucleus.
Brain Res. 586, 336–339.
Amir, S., Edelstein, K., 1997. A blocker of nitric oxide synthase,
NG-nitro-L-arginine methyl ester, attenuates light-induced Fos
protein expression in rat suprachiasmatic nucleus. Neurosci.
Lett. 224, 29–32.
Antle, M.C., Mistlberger, R.E., 2000. Circadian clock resetting
by sleep deprivation without exercise in the Syrian
hamster. J. Neurosci. 20, 9326–9332.
Antle, M.C., Marchant, E.G., Niel, L., Mistlberger, R.E., 1998.
Serotonin antagonists do not attenuate activity-induced phase
shifts of circadian rhythms in the Syrian hamster. Brain Res.
813, 139–149.
Antle, M.C., Glass, J.D., Mistlberger, R.E., 2000. 5-HT1A autoreceptor
antagonist-induced 5-HT release in the hamster
suprachiasmatic nuclei: effects on circadian clock resetting.
Neurosci. Lett. 282, 97–100.
Antle, M.C., Foley, D.K., Foley, N.C., Silver, R., 2003a. Gates and
oscillators: a network model of the brain clock. J. Biol. Rhythms
18, 339–350.
Antle, M.C., Ogilvie, M.D., Pickard, G.E., Mistlberger, R.E., 2003b.
Response of the mouse circadian system to serotonin 1A/2/7
agonists in vivo: surprisingly little. J. Biol. Rhythms 18, 145–158.
Antle, M.C., Kriegsfeld, L.J., Silver, R., 2005. Signaling within the
master clock of the brain: localized activation of
mitogen-activated protein kinase by gastrin-releasing peptide.
J. Neurosci. 25, 2447–2454.
Artinian, L.R., Ding, J.M., Gillette, M.U., 2001. Carbon monoxide and
nitric oxide: interacting messengers in muscarinic signaling to
the brain's circadian clock. Exp. Neurol. 171, 293–300.
Arvanitogiannis, A., Amir, S., 1999. Resetting the rat circadian
clock by ultra-short light flashes. Neurosci. Lett. 261, 159–162.
Arvanitogiannis, A., Robinson, B., Beaulé, C., Amir, S., 2000.
Calbindin-D28k immunoreactivity in the suprachiasmatic
nucleus and the circadian response to constant light in the rat.
Neuroscience 99, 397–401.
Aston-Jones, G., Chen, S., Zhu, Y., Oshinsky, M.L., 2001. A neural
circuit for circadian regulation of arousal. Nat. Neurosci. 4,
732–738.
Aton, S.J., Colwell, C.S., Harmar, A.J., Waschek, J., Herzog, E.D.,
2005. Vasoactive intestinal polypeptide mediates circadian
rhythmicity and synchrony in mammalian clock neurons. Nat.
Neurosci. 8, 476–483.
Aujard, F., Herzog, E.D., Block, G.D., 2001. Circadian rhythms in
firing rate of individual suprachiasmatic nucleus neurons
from adult and middle-aged mice. Neuroscience 106,
255–261.
Azmitia, E.C., Segal, M., 1978. An autoradiographic analysis of the
differential ascending projections of the dorsal and median
raphe nuclei in the rat. J. Comp. Neurol. 179, 641–668.
Bamshad, M., Aoki, V.T., Adkison, M.G., Warren, W.S., Bartness,
T.J., 1998. Central nervous system origins of the sympathetic
outflow to white adipose tissue. Am. J. Physiol. 275, R291–R299.
Bamshad, M., Song, C.K., Bartness, T.J., 1999. CNS origins of the
sympathetic nervous system outflow to brown adipose tissue.
Am. J. Physiol. 276, R1569–R1578.
Barakat, M.T., O'Hara, B.F., Cao, V.H., Larkin, J.E., Heller, H.C., Ruby,
N.F., 2004. Light pulses do not induce c-fos or per1 in the SCN of
hamsters that fail to reentrain to the photocycle. J. Biol.
Rhythms 19, 287–297.
Bartness, T.J., Song, C.K., Demas, G.E., 2001. SCN efferents to
peripheral tissues: implications for biological rhythms. J. Biol.
Rhythms 16, 196–204.
Baslow, M.H., 2000. Functions of N-acetyl-L-aspartate and
N-acetyl-L-aspartylglutamate in the vertebrate brain: role in
glial cell-specific signaling. J. Neurochem. 75, 453–459.
Battaglia, G., Sharkey, J., Kuhar, M.J., de Souza, E.B., 1991.
Neuroanatomic specificity and time course of alterations in rat
brain serotonergic pathways induced by MDMA
(3,4-methylenedioxymethamphetamine): assessment using
quantitative autoradiography. Synapse 8, 249–260.
Beaulé, C., Amir, S., 2002. Effect of 192 IgG-saporin on circadian
activity rhythms, expression of p75 neurotrophin receptors,
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
calbindin-D28K, and light-induced fos in the suprachiasmatic
nucleus in rats. Exp. Neurol. 176, 377–389.
Beaulé, C., Arvanitogiannis, A., Amir, S., 2001a. Entrainment to a
discrete light pulse at dawn is associated with acute
suppression of FOS in the shell region of the suprachiasmatic
nucleus. Soc. Neurosci. (abstract 182.6).
Beaulé, C., Arvanitogiannis, A., Amir, S., 2001b. Light suppresses
fos expression in the shell region of the suprachiasmatic
nucleus at dusk and dawn: implications for photic entrainment
of circadian rhythms. Neuroscience 106, 249–254.
Beaule, C., Houle, L.M., Amir, S., 2003a. Expression profiles of
PER2 immunoreactivity within the shell and core regions of
the rat suprachiasmatic nucleus: lack of effect of photic
entrainment and disruption by constant light. J. Mol.
Neurosci. 21, 133–147.
Beaulé, C., Robinson, B., Lamont, E.W., Amir, S., 2003b. Melanopsin
in the circadian timing system. J. Mol. Neurosci. 21, 73–89.
Belenky, M.A., Pickard, G.E., 2001. Subcellular distribution of 5-HT
(1B) and 5-HT(7) receptors in the mouse suprachiasmatic
nucleus. J. Comp. Neurol. 432, 371–388.
Belenky, M.A., Smeraski, C.A., Provencio, I., Sollars, P.J.,
Pickard, G.E., 2003. Melanopsin retinal ganglion cells receive
bipolar and amacrine cell synapses. J. Comp. Neurol. 460,
380–393.
Bergstrom, A.L., Hannibal, J., Hindersson, P., Fahrenkrug, J., 2003.
Light-induced phase shift in the Syrian hamster (Mesocricetus
auratus) is attenuated by the PACAP receptor antagonist
PACAP6-38 or PACAP immunoneutralization. Eur. J. Neurosci.
18, 2552–2562.
Berson, D.M., Dunn, F.A., Takao, M., 2002. Phototransduction by
retinal ganglion cells that set the circadian clock. Science 295,
1070–1073.
Best, J.D., Maywood, E.S., Smith, K.L., Hastings, M.H., 1999. Rapid
resetting of the mammalian circadian clock. J. Neurosci. 19,
828–835.
Biello, S.M., 1995. Enhanced photic phase shifting after treatment
with antiserum to neuropeptide Y. Brain Res. 673, 25–29.
Biello, S.M., Dafters, R.I., 2001. MDMA and fenfluramine alter the
response of the circadian clock to a serotonin agonist in vitro.
Brain Res. 920, 202–209.
Biello, S.M., Mrosovsky, N., 1993. Circadian phase-shifts induced
by chlordiazepoxide without increased locomotor activity.
Brain Res. 622, 58–62.
Biello, S.M., Mrosovsky, N., 1995. Blocking the phase-shifting effect
of neuropeptide Y with light. Proc. R. Soc. Lond., B Biol. Sci. 259,
179–187.
Biello, S.M., Janik, D., Mrosovsky, N., 1994. Neuropeptide Y and
behaviorally induced phase shifts. Neuroscience 62, 273–279.
Biello, S.M., Golombek, D.A., Harrington, M.E., 1997a. Neuropeptide
Y and glutamate block each other's phase shifts in the
suprachiasmatic nucleus in vitro. Neuroscience 77, 1049–1057.
Biello, S.M., Golombek, D.A., Schak, K.M., Harrington, M.E., 1997b.
Circadian phase shifts to neuropeptide Y in vitro: cellular
communication and signal transduction. J. Neurosci. 17,
8468–8475.
Bina, K.G., Rusak, B., 1996. Muscarinic receptors mediate
carbachol-induced phase shifts of circadian activity rhythms
in Syrian hamsters. Brain Res. 743, 202–211.
Bina, K.G., Rusak, B., Semba, K., 1993. Localization of cholinergic
neurons in the forebrain and brainstem that project to the
suprachiasmatic nucleus of the hypothalamus in rat. J. Comp.
Neurol. 335, 295–307.
Bina, K.G., Rusak, B., Semba, K., 1997. Sources of p75-nerve growth
factor receptor-like immunoreactivity in the rat
suprachiasmatic nucleus. Neuroscience 77, 461–472.
Bittman, E.L., Goldman, B.D., 1979. Serum levels of gonadotrophins
in hamsters exposed to short photoperiods: effects of
adrenalectomy and ovariectomy. J. Endocrinol. 83, 113–118.
Blasiak, T., Lewandowski, M.H., 2003. Dorsal raphe nucleus
45
modulates neuronal activity in rat intergeniculate leaflet.
Behav. Brain Res. 138, 179–185.
Blasiak, T., Lewandowski, M.H., 2004a. Blockade of GABAA
receptors disrupts isoperiodic neuronal oscillations in the
intergeniculate leaflet of the rat. Brain Res. 1009, 82–87.
Blasiak, A., Lewandowski, M.H., 2004b. In vitro extracellular
recording of spontaneous activity of the intergeniculate leaflet
neurons. Brain Res. 1015, 82–86.
Bobrzynska, K.J., Vrang, N., Mrosovsky, N., 1996. Persistence of
nonphotic phase shifts in hamsters after serotonin depletion
in the suprachiasmatic nucleus. Brain Res. 741, 205–214.
Boer, G.J., van Esseveldt, K.E., van der Geest, B.A., Duindam, H.,
Rietveld, W.J., 1999. Vasopressin-deficient suprachiasmatic
nucleus grafts re-instate circadian rhythmicity in
suprachiasmatic nucleus-lesioned arrhythmic rats.
Neuroscience 89, 375–385.
Bos, N.P.A., Mirmiran, M., 1993. Effects of excitatory and inhibitory
amino acids on neuronal discharges in the cultured
suprachiasmatic nucleus. Brain Res. Bull. 31, 67–72.
Botchkina, G.I., Morin, L.P., 1993. Development of the hamster
serotoninergic system: cell groups and diencephalic
projections. J. Comp. Neurol. 338, 405–431.
Botchkina, G.I., Morin, L.P., 1995. Specialized neuronal and glial
contributions to development of the hamster lateral geniculate
complex and circadian visual system. J. Neurosci. 15, 190–201.
Boulos, Z., Morin, L.P., 1985. Entrainment of split circadian activity
rhythms in hamsters. J. Biol. Rhythms 1, 1–15.
Boulos, Z., Rusak, B., 1982. Circadian phase response curves for
dark pulses in the hamster. J. Comp. Physiol. 146, 411–417.
Bouskila, Y., Dudek, F.E., 1995. A rapidly activating type of outward
rectifier K+ current and A-current in rat suprachiasmatic
nucleus neurones. J. Physiol. 488 (Pt. 2), 339–350.
Bradbury, M.J., Dement, W.C., Edgar, D.M., 1997.
Serotonin-containing fibers in the suprachiasmatic
hypothalamus attenuate light-induced phase delays in mice.
Brain Res. 768, 125–134.
Bramley, J.R., Sollars, P.J., Pickard, G.E., Dudek, F.E., 2005. 5-HT1B
receptor-mediated presynaptic inhibition of GABA release in
the suprachiasmatic nucleus. J. Neurophysiol. 93, 3157–3164.
Bryant, D.N., LeSauter, J., Silver, R., Romero, M.T., 2000. Retinal
innervation of calbindin-D28K cells in the hamster
suprachiasmatic nucleus: ultrastructural characterization.
J. Biol. Rhythms 15, 103–111.
Buijs, R.M., Chun, S.J., Niijima, A., Romijn, H.J., Nagai, K., 2001.
Parasympathetic and sympathetic control of the pancreas: a
role for the suprachiasmatic nucleus and other hypothalamic
centers that are involved in the regulation of food intake.
J. Comp. Neurol. 431, 405–423.
Burgoon, P.W., Lindberg, P.T., Gillette, M.U., 2004. Different
patterns of circadian oscillation in the suprachiasmatic
nucleus of hamster, mouse, and rat. J. Comp. Physiol., A 190,
167–171.
Byku, M., Gannon, R.L., 2000a. SNC 80, a delta-opioid agonist,
elicits phase advances in hamster circadian activity rhythms.
NeuroReport 11, 1449–1452.
Byku, M., Gannon, R.L., 2000b. Opioid induced non-photic phase
shifts of hamster circadian activity rhythms. Brain Res. 873,
189–196.
Byku, M., Legutko, R., Gannon, R.L., 2000. Distribution of δ opioid
receptor immunoreactivity in the hamster suprachiasmatic
nucleus and intergeniculate leaflet. Brain Res. 857, 1–7.
Cagampang, F.R., Piggins, H.D., Sheward, W.J., Harmar, A.J., Coen,
C.W., 1998a. Circadian changes in PACAP type 1 (PAC1) receptor
mRNA in the rat suprachiasmatic and supraoptic nuclei. Brain
Res. 813, 218–222.
Cagampang, F.R., Sheward, W.J., Harmar, A.J., Piggins, H.D., Coen,
C.W., 1998b. Circadian changes in the expression of vasoactive
intestinal peptide 2 receptor mRNA in the rat suprachiasmatic
nuclei. Brain Res. Mol. Brain Res. 54, 108–112.
46
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
Caillol, M., Devinoy, E., Lacroix, M.C., Schirar, A., 2000. Endothelial
and neuronal nitric oxide synthases are present in the
suprachiasmatic nuclei of Syrian hamsters and rats. Eur. J.
Neurosci. 12, 649–661.
Canbeyli, R.S., Lehman, M.N., Silver, R., 1991. Tracing SCN graft
efferents with Dil. Brain Res. 554, 15–21.
Card, J.P., Moore, R.Y., 1982. Ventral lateral geniculate nucleus
efferents to the rat suprachiasmatic nucleus exhibit avian
pancreatic polypeptide-like immunoreactivity. J. Comp.Neurol.
206, 390–396.
Card, J.P., Moore, R.Y., 1984. The suprachiasmatic nucleus of the
golden hamster: immunohistochemical analysis of cell and
fiber distribution. Neuroscience 13, 390–396.
Card, J.P., Moore, R.Y., 1989. Organization of lateral
geniculate–hypothalamic connections in the rat. J. Comp.
Neurol. 284, 135–147.
Card, J.P., Brecha, N., Moore, R.Y., 1983. Immunohistochemical
localization of avian pancreatic polypeptide-like
immunoreactivity in the rat hypothalamus. J. Comp. Neurol.
217, 123–136.
Cassone, V.M., Speh, J.C., Card, J.P., 1988. Comparative anatomy of
the mammalian hypothalamic suprachiasmatic nucleus. J. Biol.
Rhythms 3, 71–91.
Castel, M., Morris, J.F., 2000. Morphological heterogeneity of the
GABAergic network in the suprachiasmatic nucleus, the brain's
circadian pacemaker. J. Anat. 196, 1–13.
Castel, M., Belenky, M., Cohen, S., Ottersen, O.P., Storm-Mathisen,
J., 1993. Glutamate-like immunoreactivity in retinal terminals
of the mouse suprachiasmatic nucleus. Eur. J. Neurosci. 5,
368–381.
Challet, E., Naylor, E., Metzger, J.M., MacIntyre, D.E., Turek, F.W.,
1998a. An NK1 receptor antagonist affects the circadian
regulation of locomotor activity in golden hamsters. Brain Res.
800, 32–39.
Challet, E., Scarbrough, K., Penev, P.D., Turek, F., 1998b. Roles of
suprachiasmatic nuclei and intergeniculate leaflets in
mediating the phase-shifting effects of a serotonergic agonist
and their photic modulation during subjective day. J. Biol.
Rhythms 12, 410–421.
Challet, E., Dugovic, C., Turek, F.W., Olivier, V.R., 2001. The
selective neurokinin 1 receptor antagonist R116301 modulates
photic responses of the hamster circadian system.
Neuropharmacology 40, 408–415.
Chambille, I., Doyle, S., Servière, J., 1993. Photic induction and
circadian expression of Fos-like protein.
Immunohistochemical study in the retina and
suprachiasmatic nuclei of hamster. Brain Res. 612, 138–150.
Chen, D., Hurst, W.J., Ding, J.M., Faiman, L.E., Mayer, B., Gillette,
M.U., 1997. Localization and characterization of nitric oxide
synthase in the rat suprachiasmatic nucleus: evidence for a
nitrergic plexus in the biological clock. J. Neurochem. 68,
855–861.
Chen, D., Buchanan, G.F., Ding, J.M., Hannibal, J., Gillette, M.U.,
1999. Pituitary adenylyl cyclase-activating peptide: a pivotal
modulator of glutamatergic regulation of the
suprachiasmatic circadian clock. Proc. Natl. Acad. Sci.
U. S. A. 96, 13468–13473.
Chevassus, A.L., Cooper, H.M., 1998. Is there a
geniculohypothalamic tract in primates? A comparative
immunohistochemical study in the circadian system of
strepsirhine and haplorhine species. Brain Res. 805, 213–219.
Christian, C.A., Harrington, M.E., 2002. Three days of novel
wheel access diminishes light-induced phase delays in vivo
with no effect on per1 induction by light. Chronobiol. Int.
19, 671–682.
Cloues, R.K., Sather, W.A., 2003. Afterhyperpolarization regulates
firing rate in neurons of the suprachiasmatic nucleus.
J. Neurosci. 23, 1593–1604.
Colbron, S., Jones, M., Biello, S.M., 2002. MDMA alters the response
of the circadian clock to a photic and non-photic stimulus.
Brain Res. 956, 45–52.
Colwell, C.S., 2000. Circadian modulation of calcium levels in cells
in the suprachiasmatic nucleus. Eur. J. Neurosci. 12, 571–576.
Colwell, C.S., Menaker, M., 1992. NMDA as well as non-NMDA
receptor antagonists can prevent the phase-shifting effects of
light on the circadian system of the golden hamster. J. Biol.
Rhythms 7, 125–136.
Colwell, C.S., Ralph, M.R., Menaker, M., 1990. Do NMDA receptors
mediate the effects of light on circadian behavior. Brain Res.
523, 117–120.
Colwell, C.S., Foster, R.G., Menaker, M., 1991. NMDA receptor
antagonists block the effects of light on circadian behavior in
the mouse. Brain Res. 554, 105–110.
Colwell, C.S., Kaufman, C.M., Menaker, M., 1993a. Photic induction
of Fos in the hamster suprachiasmatic nucleus is inhibited by
baclofen but not by diazepam or bicucullin. Neurosci. Lett. 163,
177–181.
Colwell, C.S., Kaufman, C.M., Menaker, M., Ralph, M.R., 1993b.
Light-induced phase shifts and Fos expression in the hamster
circadian system: the effects of anesthetics. J. Biol. Rhythms 8,
179–188.
Colwell, C.S., Kaufman, C.M., Menaker, M., 1993c. Phase-shifting
mechanisms in the mammalian circadian system: new light on
the carbachol paradox. J. Neurosci. 13, 1454–1459.
Colwell, C.S., Michel, S., Itri, J., Rodriguez, W., Tam, J., Lelievre, V.,
Hu, Z., Liu, X., Waschek, J.A., 2003. Disrupted circadian rhythms
in VIP- and PHI-deficient mice. Am. J. Physiol.: Regul., Integr.
Comp. Physiol. 285, R939–R949.
Colwell, C.S., Michel, S., Itri, J., Rodriguez, W., Tam, J., Lelievre, V.,
Hu, Z., Waschek, J.A., 2004. Selective deficits in the circadian
light response in mice lacking PACAP. Am. J. Physiol.: Regul.,
Integr. Comp. Physiol. 287, R1194–R1201.
Coogan, A.N., Rawlings, N., Luckman, S.M., Piggins, H.D., 2001.
Effects of neurotensin on discharge rates of rat suprachiasmatic
nucleus neurons in vitro. Neuroscience 103, 663–672.
Cooper, H.M., Parvopassu, F., Herbin, M., Magnin, M., 1994.
Neuroanatomical pathways linking vision and olfaction in
mammals. Psychoneuroendocrinology 19, 623–639.
Coyle, J.T., 1997. The nagging question of the function of
N-acetylaspartylglutamate. Neurobiol. Dis. 4, 231–238.
Crill, W.E., 1996. Persistent sodium current in mammalian central
neurons. Annu. Rev. Physiol. 58, 349–362.
Cutler, D.J., Munekawa, K., Mason, R., 1999. Electrophysiological
effects of opioid receptor activation on Syrian hamster
suprachiasmatic nucleus neurones in vitro. Brain Res. Bull. 50,
119–125.
Cutler, D.J., Hardin, H., Rees, H., Maywood, E., Hastings, M.H.,
Shena, H.M., Martens, J.M., Harrer, C., Piggins, H.D., 2002. The
mouse VPAC2 receptor confers SCN cellular rhythmicity and
responsiveness to vasoactive intestinal polypeptide (VIP) in
vitro. Soc. Res. Biol. Rhythms 8, 37.
Cutler, D.J., Haraura, M., Reed, H.E., Shen, S., Sheward, W.J.,
Morrison, C.F., Marston, H.M., Harmar, A.J., Piggins, H.D., 2003.
The mouse VPAC2 receptor confers suprachiasmatic nuclei
cellular rhythmicity and responsiveness to vasoactive
intestinal polypeptide in vitro. Eur. J. Neurosci. 17, 197–204.
Dacey, D.M., Liao, H.W., Peterson, B.B., Robinson, F.R., Smith, V.C.,
Pokorny, J., Yau, K.W., Gamlin, P.D., 2005.
Melanopsin-expressing ganglion cells in primate retina signal
colour and irradiance and project to the LGN. Nature 433,
749–754.
Daikoku, S., Hisano, S., Kagotani, Y., 1992. Neuronal associations in
the rat suprachiasmatic nucleus demonstrated by
immunoelectron microscopy. J. Comp. Neurol. 325, 559–571.
Dardente, H., Poirel, V.J., Klosen, P., Pevet, P., Masson-Pevet, M.,
2002. Per and neuropeptide expression in the rat
suprachiasmatic nuclei: compartmentalization and
differential cellular induction by light. Brain Res. 958, 261–271.
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
Decker, K., Reuss, S., 1994. Nitric oxide-synthesizing neurons in
the hamster suprachiasmatic nucleus: a combined NOS- and
N. Brain Res. 666, 284–288.
De Jeu, M., Pennartz, C.M.A., 1997. Functional characterization of
the H-current in SCN neurons in subjective day and night: a
whole-cell patch-clamp study in acutely prepared brain slices.
Brain Res. 767, 72–80.
De Jeu, M., Pennartz, C.M.A., 2002. Circadian modulation of GABA
function in the rat suprachiasmatic nucleus: excitatory effects
during the night phase. J. Neurophysiol. 87, 834–844.
De Jeu, M., Hermes, N., Pennartz, C.M.A., 1998. Circadian
modulation of membrane properties in slices of rat
suprachiasmatic nucleus. NeuroReport 9, 3725–3729.
De Jeu, M., Geurtsen, A., Pennartz, C.M.A., 2002. A Ba(2+)-sensitive
K(+) current contributes to the resting membrane potential of
neurons in rat suprachiasmatic nucleus. J. Neurophysiol. 88,
349–878.
De la Iglesia, H.O., Schwartz, W.J., 2002. A subpopulation of
efferent neurons in the mouse suprachiasmatic nucleus is also
light responsive. NeuroReport 13, 857–860.
De la Iglesia, H.O., Blaustein, J.D., Bittman, E.L., 1995. The
suprachiasmatic area in the female hamster projects to
neurons containing estrogen receptors and GnRH.
NeuroReport 6, 1715–1722.
De la Iglesia, H.O., Meyer, J., Carpino Jr., A., Schwartz, W.J., 2000.
Antiphase oscillation of the left and right suprachiasmatic
nuclei. Science 290, 799–801.
De la Iglesia, H.O., Meyer, J., Schwartz, W.J., 2003. Lateralization of
circadian pacemaker output: activation of left- and right-sided
luteinizing hormone-releasing hormone neurons involves a
neural rather than a humoral pathway. J. Neurosci. 23,
7412–7414.
De Vries, M.J., Meijer, J.H., 1991. Aspartate injections into the
suprachiasmatic region of the Syrian hamster do not mimic
the effects of light on the circadian activity rhythm. Neurosci.
Lett. 127, 215–218.
De Vries, M.J., Nunes Cardozo, B., Van der Want, J., De Wolf, A.,
Meijer, J.H., 1993. Glutamate immunoreactivity in terminals of
the retinohypothalamic tract of the brown Norwegian rat.
Brain Res. 612, 231–237.
Devries, M.J., Lakke, E.J.F., 1995. Retrograde labeling of retinal
ganglion-cells and brain neuronal subsets by [H-3]D-aspartate
injection in the Syrian-hamster hypothalamus. Brain Res. Bull.
38, 349–354.
Ding, J.M., Chen, D., Weber, E.T., Faiman, L.E., Rea, M.A., Gillette,
M.U., 1994. Resetting the biological clock: mediation of nocturnal
circadian shifts by glutamate and NO. Science 266, 1713–1717.
Ding, J.M., Faiman, L.E., Hurst, W.J., Kuriashkina, L.R., Gillette, M.U.,
1997. Resetting the biological clock: mediation of nocturnal
CREB phosphorylation via light, glutamate, and nitric oxide.
J. Neurosci. 17, 667–675.
Dkhissi-Benyahya, O., Sicard, B., Cooper, H.M., 2000. Effects of
irradiance and stimulus duration on early gene expression
(Fos) in the suprachiasmatic nucleus: temporal summation
and reciprocity. J. Neurosci. 20, 7790–7797.
Doan, A., Urbanski, H.F., 1994. Diurnal expression of Fos in
luteinizing hormone-releasing hormone neurons of Syrian
hamsters. Biol. Reprod. 50, 301–308.
Donaldson, J.A., Stephan, F.K., 1982. Entrainment of circadian
rhythms: retinofugal pathways and unilateral suprachiasmatic
nucleus lesions. Physiol. Behav. 29, 1161–1169.
Dudley, T.E., DiNardo, L.A., Glass, J.D., 1998. Endogenous
regulation of serotonin release in the hamster suprachiasmatic
nucleus. J. Neurosci. 18, 5045–5052.
Dudley, T.E., DiNardo, L.A., Glass, J.D., 1999. In vivo assessment of
the midbrain raphe nuclear regulation of serotonin release in
the hamster suprachiasmatic nucleus. J. Neurophysiol. 81,
1468–1477.
Duncan, M.J., Short, J., Wheeler, D.L., 1999. Comparison of the
47
effects of aging on 5-HT7 and 5-HT1A receptors in discrete
regions of the circadian timing system in hamsters. Brain Res.
829, 39–45.
Duncan, M.J., Jennes, L., Jefferson, J.B., Brownfield, M.S., 2000.
Localization of serotonin5A receptors in discrete regions of the
circadian timing system in the Syrian hamster. Brain Res. 869,
178–185.
Duncan, M.J., Grear, K.E., Hoskins, M.A., 2004. Aging and
SB-269970-A, a selective 5-HT7 receptor antagonist, attenuate
circadian phase advances induced by microinjections of
serotonergic drugs in the hamster dorsal raphe nucleus. Brain
Res. 1008, 40–48.
Dwyer, S.M., Rosenwasser, A.M., 2000. Effects of light intensity and
restraint on dark-pulse-induced circadian phase shifting during
subjective night in Syrian hamsters. J. Biol. Rhythms 15, 491–500.
Ebling, F.J.P., 1996. The role of glutamate in the photic regulation of
the suprachiasmatic nucleus. Prog. Neurobiol. 50, 109–123.
Edelstein, K., Amir, S., 1998. Glutamatergic antagonists do not
attenuate light-induced Fos protein in rat intergeniculate
leaflet. Brain Res. 810, 264–268.
Edelstein, K., Amir, S., 1999a. The intergeniculate leaflet does not
mediate the disruptive effects of constant light on circadian
rhythms in the rat. Neuroscience 90, 1093–1101.
Edelstein, K., Amir, S., 1999b. The role of the intergeniculate leaflet
in entrainment of circadian rhythms to a skeleton
photoperiod. J. Neurosci. 19, 372–380.
Edelstein, K., Mrosovsky, N., 2001. Behavioral responses to light in
mice with dorsal lateral geniculate lesions. Brain Res. 918,
107–112.
Edelstein, K., De la Iglesia, H.O., Schwartz, W.J., Mrosovsky, N.,
2003a. Behavioral arousal blocks light-induced phase advances
in locomotor rhythmicity but not light-induced Per1 and fos
expression in the hamster suprachiasmatic nucleus.
Neuroscience 118, 253–261.
Edelstein, K., De la Iglesia, H.O., Mrosovsky, N., 2003b. Period
gene expression in the suprachiasmatic nucleus of
behaviorally decoupled hamsters. Brain Res. Mol. Brain Res.
114, 40–45.
Edgar, D.M., Reid, M.S., Dement, W.C., 1997. Serotonergic afferents
mediate activity-dependent entrainment of the mouse
circadian clock. Am. J. Physiol. 273, R265–R269.
Ehlen, J.C., Grossman, G.H., Glass, J.D., 2001. In vivo resetting of the
hamster circadian clock by 5-HT7 receptors in the
suprachiasmatic nucleus. J. Neurosci. 21, 5351–5357.
Elliott, J.A., Stetson, M.H., Menaker, M., 1972. Regulation of testis
function in golden hamsters: a circadian clock measures
photoperiodic time. Science 178, 771–773.
Enaker, M., Roberts, R., Elliott, J., Underwood, H., 1970. Extraretinal
light perception in the sparrow: 3. The eyes do not participate
in photoperiodic photoreception. Proc. Natl. Acad. Sci. U. S. A.
67, 320–325.
Erhardt, C., Galani, R., Jeltsch, H., Cassel, J.C., Klosen, P., Menet, J.S.,
Pevet, P., Challet, E., 2004. Modulation of photic resetting in rats
by lesions of projections to the suprachiasmatic nuclei
expressing p75 neurotrophin receptor. Eur. J. Neurosci. 19,
1773–1788.
Erlander, M.G., Tillakaratne, N.J., Feldblum, S., Patel, N., Tobin, A.J.,
1991. Two genes encode distinct glutamate decarboxylases.
Neuron 7, 91–100.
Fedorkova, L., Rutishauser, U., Prosser, R., Shen, H., Glass, J.D.,
2002. Removal of polysialic acid from the SCN potentiates
nonphotic circadian phase resetting. Physiol. Behav. 77,
361–369.
Ferguson, S.A., Kennaway, D.J., 1999. Emergence of altered
circadian timing in a cholinergically supersensitive rat line.
Am. J. Physiol. 277, R1171–R1178.
Ferguson, S.A., Kennaway, D.J., Moyer, R.W., 1999. Nicotine phase
shifts the 6-sulphatoxymelatonin rhythm and induces c-Fos in
the SCN of rats. Brain Res. Bull. 48, 527–538.
48
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
Ferreyra, G.A., Cammarota, M.P., Golombek, D.A., 1998. Photic
control of nitric oxide synthase activity in the hamster
suprachiasmatic nuclei. Brain Res. 797, 190–196.
Field, M.D., Maywood, E.S., O'Brien, J.A., Weaver, D.R., Reppert, S.M.,
Hastings, M.H., 2000. Analysis of clock proteins in mouse SCN
demonstrates phylogenetic divergence of the circadian
clockwork and resetting mechanisms. Neuron 25, 437–447.
Fite, K.V., Janusonis, S., 2001. Retinal projection to the dorsal raphe
nucleus in the Chilean degus (Octodon degus). Brain Res. 895,
139–145.
Fite, K.V., Janusonis, S., Foote, W., Bengston, L., 1999. Retinal
afferents to the dorsal raphe nucleus in rats and Mongolian
gerbils. J. Comp. Neurol. 414, 469–484.
Fite, K.V., Birkett, M.A., Smith, A., Janusonis, S.F., McLaughlin, S.,
2003. Retinal ganglion cells projecting to the dorsal raphe and
lateral geniculate complex in Mongolian gerbils. Brain Res. 973,
146–150.
Foote, W.E., Taber-Pierce, E., Edwards, L., 1978. Evidence for a
retinal projection to the midbrain raphe of the cat. Brain Res.
156, 135–140.
Foster, R.G., Provencio, I., Hudson, D., Fiske, S., De Grip, W.,
Menaker, M., 1991. Circadian photoreception in the retinally
degenerate mouse (rd/rd). J. Comp. Physiol., A 169, 39–50.
Foster, R.G., Provencio, I., Bovee-Geurts, P.H., DeGrip, W.J., 2003.
The photoreceptive capacity of the developing pineal gland
and eye of the golden hamster (Mesocricetus auratus).
J. Neuroendocrinol. 15, 355–363.
Freedman, M.S., Lucas, R.J., Soni, B., Von Schantz, M., Munoz, M.,
David-Gray, Z., Foster, R.G., 1999. Regulation of mammalian
circadian behavior by non-rod, non-cone, ocular
photoreceptors. Science 284, 502–504.
Freeman, M.E., Kanyicska, B., Lerant, A., Nagy, G., 2000. Prolactin:
structure, function, and regulation of secretion. Physiol. Rev.
80, 1523–1631.
Fujiyama, F., Hioki, H., Tomioka, R., Taki, K., Tamamaki, N.,
Nomura, S., Okamoto, K., Kaneko, T., 2003. Changes of
immunocytochemical localization of vesicular glutamate
transporters in the rat visual system after the retinofugal
denervation. J. Comp. Neurol. 465, 234–249.
Fukuhara, C., Brewer, J.M., Dirden, J.C., Bittman, E.L., Tosini, G.,
Harrington, M.E., 2001. Neuropeptide Y rapidly reduces Period 1
and Period 2 mRNA levels in the hamster suprachiasmatic
nucleus. Neurosci. Lett. 314, 119–122.
Gamble, K.L., Novak, C.M., Paul, K.N., Albers, H.E., 2003.
Tetrodotoxin blocks the circadian effects of NMDA during the
day but not at night. NeuroReport 14, 641–644.
Gamble, K.L., Novak, C.M., Albers, H.E., 2004. Neuropeptide Y and
N-methyl-D-aspartic acid interact within the suprachiasmatic
nuclei to alter circadian phase. Neuroscience 126, 559–565.
Gannon, R.L., 2001. 5HT7 receptors in the rodent suprachiasmatic
nucleus. J. Biol. Rhythms 16, 19–24.
Gannon, R.L., 2003. Serotonergic serotonin1a mixed agonists/
antagonists elicit large-magnitude phase shifts in hamster
circadian wheel-running rhythms. Neuroscience 119, 567–576
(Ref Type: Generic).
Gaston, S., Menaker, M., 1968. Pineal function: the biological clock
in the sparrow? Science 160, 1125–1127.
Gerfen, C.R., Sawchenko, P.E., 1984. An anterograde
neuroanatomical tracing method that shows the detailed
morphology of neurons, their axons and terminals:
immunohistochemical localization of an axonally
transported plant lectin, Phaseolus vulgaris leucoagglutinin
(PHA-L). Brain Res. 290, 219–238.
Geusz, M.E., Fletcher, C., Block, G.D., Straume, M., Copeland, N.G.,
Jenkins, N.A., Kay, S.A., Day, R.N., 1997. Long-term monitoring
of circadian rhythms in c-fos gene expression from
suprachiasmatic nucleus cultures. Curr. Biol. 7, 758–766.
Gillespie, C.F., Huhman, K.L., Babagbemi, T.O., Albers, H.E., 1996.
Bicuculline increases and muscimol reduces the
phase-delaying effects of light and VIP/PHI/GRP in the
suprachiasmatic region. J. Biol. Rhythms 11, 137–144.
Gillespie, C.F., Mintz, E.M., Marvel, C.L., Huhman, K.L., Albers, H.E.,
1997. GABA(A) and GABA(B) agonists and antagonists alter the
phase-shifting effects of light when microinjected into the
suprachiasmatic region. Brain Res. 759, 181–189.
Gillespie, C.F., Van der Beek, E.M., Mintz, E.M., Mickley, N.C.,
Jasnow, A.M., Huhman, K.L., Albers, H.E., 1999. GABAergic
regulation of light-induced c-Fos immunoreactivity within
the suprachiasmatic nucleus. J. Comp. Neurol. 411,
683–692.
Gillette, M.U., 1991. SCN electrophysiology in vitro: rhythmic
activity and endogenous clock properties. In: Klein, D.C.,
Moore, R.Y., Reppert, S.M. (Eds.), Suprachiasmatic Nucleus: The
Mind's Clock. Oxford Univ. Press, New York, pp. 125–143.
Gillette, M.U., DeMarco, S.J., Ding, J.M., Gallman, E.A., Faiman, L.E.,
Liu, C., McArthur, A.J., Medanic, M., Richard, D., Tcheng, T.K.,
Weber, E.T., 1993. The organization of the suprachiasmatic
circadian pacemaker of the rat and its regulation by
neurotransmitters and modulators. J. Biol. Rhythms 8, S53–S58.
Gillette, M.U., Buchanan, G.F., Artinian, L., Hamilton, S.E.,
Nathanson, N.M., Liu, C., 2001. Role of the M1 receptor in
regulating circadian rhythms. Life Sci. 68, 2467–2472.
Ginty, D.D., Kornhauser, J.M., Thompson, M.A., Bading, H., Mayo,
K.E., Takahashi, J.S., Greenberg, M.E., 1993. Regulation of CREB
phosphorylation in the suprachiasmatic nucleus by light and
a circadian clock. Science 260, 238–241.
Glass, J.D., Chen, L., 1999. Serotonergic modulation of astrocytic
activity in the hamster suprachiasmatic nucleus. Neuroscience
94, 1253–1259.
Glass, J.D., Hauser, U.E., Blank, J.L., Selim, M., Rea, M.A., 1993.
Differential timing of amino acid and 5-HIAA rhythms in
suprachiasmatic hypothalamus. Am. J. Physiol. 265,
R504–R511.
Glass, J.D., Lee, W., Shen, H., Watanabe, M., 1994. Expression of
immunoreactive polysialylated neural cell adhesion molecule
in the suprachiasmatic nucleus. Neuroendocrinology 60,
87–95.
Glass, J.D., Shen, H., Fedorkova, L., Chen, L., Tomasiewicz, H.,
Watanabe, M., 2000a. Polysialylated neural cell adhesion
molecule modulates photic signaling in the mouse
suprachiasmatic nucleus. Neurosci. Lett. 280, 207–210.
Glass, J.D., DiNardo, L.A., Ehlen, J.C., 2000b. Dorsal raphe nuclear
stimulation of SCN serotonin release and circadian
phase-resetting. Brain Res. 859, 224–232.
Glass, J.D., Watanabe, M., Fedorkova, L., Shen, H., Ungers, G.,
Rutishauser, U., 2003a. Dynamic regulation of polysialylated
neural cell adhesion molecule in the suprachiasmatic nucleus.
Neuroscience 117, 203–211.
Glass, J.D., Grossman, G.H., Farnbauch, L., DiNardo, L., 2003b.
Midbrain raphe modulation of nonphotic circadian clock
resetting and 5-HT release in the mammalian suprachiasmatic
nucleus. J. Neurosci. 23, 7451–7460.
Goel, N., Lee, T.M., Smale, L., 1999. Suprachiasmatic nucleus and
intergeniculate leaflet in the diurnal rodent Octodon degus:
retinal projections and immunocytochemical characterization.
Neuroscience 92, 1491–1509.
Gooley, J.J., Saper, C.B., 2003. A broad role for melanopsin in
non-visual photoreception based on neuroanatomical
evidence in rats. J. Neurosci. 23, 7093–7106.
Gooley, J.J., Lu, J., Chou, T.C., Scammell, T.E., Saper, C.B., 2001.
Melanopsin in cells of origin of the retinohypothalamic tract.
Nat. Neurosci. 4, 1165.
Gorman, M.R., Kendall, M., Elliott, J.A., 2005. Scotopic illumination
enhances entrainment of circadian rhythms to lengthening
light:dark cycles. J. Biol. Rhythms 20, 38–48.
Graham, E.S., Turnbull, Y., Fotheringham, P., Nilaweera, K.,
Mercer, J.G., Morgan, P.J., Barrett, P., 2003. Neuromedin U and
Neuromedin U receptor-2 expression in the mouse and rat
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
hypothalamus: effects of nutritional status. J. Neurochem. 87,
1165–1173.
Graham, E.S., Littlewood, P., Turnbull, Y., Mercer, J.G., Morgan, P.J.,
Barrett, P., 2005. Neuromedin-U is regulated by the circadian
clock in the SCN of the mouse. Eur. J. Neurosci. 21, 814–819.
Gribkoff, V.K., Pieschl, R.L., Wisialowski, T.A., Park, W.K., Strecker,
G.J., De Jeu, M.T.G., Pennartz, C.M.A., Dudek, F.E., 1999. A
reexamination of the role of GABA in the mammalian
suprachiasmatic nucleus. J. Biol. Rhythms 14, 126–130.
Grossman, G.H., Mistlberger, R.E., Antle, M.C., Ehlen, J.C., Glass, J.D.,
2000. Sleep deprivation stimulates serotonin release in the
suprachiasmatic nucleus. NeuroReport 11, 1929–1932.
Grossman, G.H., Farnbauch, L., Glass, J.D., 2004. Regulation of
serotonin release in the Syrian hamster intergeniculate leaflet
region. NeuroReport 15, 103–106.
Guido, M.E., Goguen, D., De Guido, L., Robertson, H.A., Rusak, B.,
1999. Circadian and photic regulation of immediate-early gene
expression in the hamster suprachiasmatic nucleus.
Neuroscience 90, 555–571.
Haak, L.L., Heller, H.C., van den Pol, A.N., 1997. Metabrotropic
glutamate receptor activation modulates kainate and serotonin
calcium response in astrocytes. J. Neurosci. 17, 1825–1837.
Hamada, T., Yamanouchi, S., Watanabe, A., Shibata, S., Watanabe,
S., 1999. Involvement of glutamate release in substance
P-induced phase delays of suprachiasmatic neuron activity
rhythm in vitro. Brain Res. 836, 190–193.
Hamada, T., LeSauter, J., Venuti, J.M., Silver, R., 2001. Expression of
Period genes: rhythmic and non-rhythmic compartments of
the suprachiasmatic nucleus pacemaker. J. Neurosci. 21,
7742–7750.
Hamada, T., LeSauter, J., Lokshin, M., Romero, M.T., Yan, L.L.,
Venuti, J.M., Silver, R., 2003. Calbindin influences response to
photic input in suprachiasmatic nucleus. J. Neurosci. 23,
8820–8826.
Hamada, T., Antle, M.C., Silver, R., 2004a. Temporal and spatial
expression patterns of canonical clock genes and
clock-controlled genes in the suprachiasmatic nucleus. Eur. J.
Neurosci. 19, 1741–1748.
Hamada, T., Antle, M.C., Silver, R., 2004b. The role of Period1 in
non-photic resetting of the hamster circadian pacemaker in
the suprachiasmatic nucleus. Neurosci. Lett. 362, 87–90.
Hanada, R., Teranishi, H., Pearson, J.T., Kurokawa, M., Hosoda, H.,
Fukushima, N., Fukue, Y., Serino, R., Fujihara, H., Ueta, Y.,
Ikawa, M., Okabe, M., Murakami, N., Shirai, M., Yoshimatsu, H.,
Kangawa, K., Kojima, M., 2004. Neuromedin U has a novel
anorexigenic effect independent of the leptin signaling
pathway. Nat. Med. 10, 1067–1073.
Hannibal, J., 2002a. Neurotransmitters of the retino-hypothalamic
tract. Cell Tissue Res. 309, 73–88.
Hannibal, J., 2002b. Pituitary adenylate cyclase-activating
peptide in the rat central nervous system: an
immunohistochemical and in situ hybridization study.
J. Comp. Neurol. 453, 389–417.
Hannibal, J., Fahrenkrug, J., 2002. Immunoreactive substance P is
not part of the retinohypothalamic tract in the rat. Cell Tissue
Res. 309, 293–299.
Hannibal, J., Fahrenkrug, J., 2004. Target areas innervated by
PACAP-immunoreactive retinal ganglion cells. Cell Tissue Res.
316, 99–113.
Hannibal, J., Mikkelsen, J.D., Clausen, H., Holst, J.J., Wulff, B.S.,
Fahrenkrug, J., 1995. Gene expression of pituitary adenylate
cyclase activating polypeptide (PACAP) in the rat
hypothalamus. Regul. Pept. 55, 133–148.
Hannibal, J., Ding, J.M., Chen, D., Fahrenkrug, J., Larsen, P.J.,
Gillette, M.U., Mikkelsen, J.D., 1997. Pituitary adenylate cyclaseactivating peptide (PACAP) in the retinohypothalamic tract: a
potential daytime regulator of the biological clock. J. Neurosci.
17, 2637–2644.
Hannibal, J., Moller, M., Ottersen, O.P., Fahrenkrug, J., 2000. PACAP
49
and glutamate are co-stored in the retinohypothalamic tract. J.
Comp. Neurol. 418, 147–155.
Hannibal, J., Vrang, N., Card, J.P., Fahrenkrug, J., 2001a. Lightdependent induction of cFos during subjective day and night in
PACAP-containing ganglion cells of the retinohypothalamic
tract. J. Biol. Rhythms 16, 457–470.
Hannibal, J., Jamen, F., Nielsen, H.S., Journot, L., Brabet, P.,
Fahrenkrug, J., 2001b. Dissociation between light-induced
phase shift of the circadian rhythm and clock gene expression
in mice lacking the pituitary adenylate cyclase activating
polypeptide type 1 receptor. J. Neurosci. 21, 4883–4890.
Hannibal, J., Hindersson, P., Knudsen, S.M., Georg, B., Fahrenkrug,
J., 2002. The photopigment melanopsin is exclusively present
in pituitary adenylated cyclase-activating
polypeptide-containing retinal ganglion cells of the
retinohypothalamic tract. J. Neurosci. 22, RC191 (1-7).
Harmar, A.J., Marston, H.M., Shen, S., Spratt, C., West, K.M.,
Sheward, W.J., Morrison, C.F., Dorin, J.R., Piggins, H.D., Reubi,
J.-C., Kelly, J.S., Maywood, E., Hastings, M.H., 2002. The VPAC2
receptor is essential for circadian function in the mouse
suprachiasmatic nuclei. Cell 109, 497–508.
Harrington, M.E., 1997. The ventral lateral geniculate nucleus and
the intergeniculate leaflet: interrelated structures in the visual
and circadian systems. Neurosci. Biobehav. Rev. 21, 705–727.
Harrington, M.E., Rusak, B., 1986. Lesions of the thalamic
intergeniculate leaflet alter hamster circadian rhythms.
J. Biol. Rhythms 1, 309–325.
Harrington, M.E., Schak, K.M., 2000. Neuropeptide Y phase
advances the in vitro hamster circadian clock during the
subjective day with no effect on phase during the subjective
night. Can. J. Physiol. Pharm. 78, 87–92.
Harrington, M.E., Hoque, S., Hall, A., Golombek, D.A., Biello, S.,
1999. Pituitary adenylate cyclase activating peptide phase
shifts circadian rhythms in a manner similar to light.
J. Neurosci. 19, 6637–6642.
Hartwich, M., Kalsbeek, A., Pevet, P., Nurnberger, F., 1994. Effects of
illumination and enucleation on substance-P-immunoreactive
structures in subcortical visual centers of golden-hamster and
Wistar rat. Cell Tissue Res. 277, 351–361.
Hastings, M.H., Mead, S.M., Vindlacheruvu, R.R., Ebling, F.J.P.,
Maywood, E.S., Grosse, J., 1992. Non-photic phase shifting of
the circadian activity rhythm of Syrian hamsters: the relative
potency of arousal and melatonin. Brain Res. 591, 20–26.
Hastings, M.H., Field, M.D., Maywood, E.S., Weaver, D.R., Reppert,
S.M., 1999. Differential regulation of mPER1 and mTIM proteins
in the mouse suprachiasmatic nuclei: new insights into a core
clock mechanism. J. Neurosci. 19, RC11.
Hattar, S., Liao, H.W., Takao, M., Berson, D.M., Yau, K.W., 2002.
Melanopsin-containing retinal ganglion cells: architecture,
projections, and intrinsic photosensitivity. Science 295,
1065–1070.
Hattar, S., Lucas, R.J., Mrosovsky, N., Thompson, S., Douglas, R.H.,
Hankins, M.W., Lem, J., Biel, M., Hofmann, F., Foster, R.G., Yau,
K.W., 2003. Melanopsin and rod-cone photoreceptive systems
account for all major accessory visual functions in mice.
Nature 424, 76–81.
Hay-Schmidt, A., Vrang, N., Larsen, P.J., Mikkelsen, J.D., 2003.
Projections from the raphe nuclei to the suprachiasmatic
nucleus of the rat. J. Chem. Neuroanat. 25, 293–310.
Hendrickson, A.E., Wagoner, N., Cowan, W.M., 1972. An
autoradiographic and electron microscopic study of
retinohypothalamic connections. Z. Zellforsch. 135, 1–26.
Hertz, L., 2004. Intercellular metabolic compartmentation in the
brain: past, present and future. Neurochem. Int. 45, 285–296.
Herzog, E.D., Geusz, M.E., Khalsa, S.B., Straume, M., Block, G.D.,
1997. Circadian rhythms in mouse suprachiasmatic nucleus
explants on multimicroelectrode plates. Brain Res. 757,
285–290.
Herzog, E.D., Takahashi, J.S., Block, G.D., 1998. Clock controls
50
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
circadian period in isolated suprachiasmatic nucleus neurons.
Nat. Neurosci. 1, 708–713.
Honma, S., Shirakawa, T., Katsuno, Y., Namihira, M., Honma, K.,
1998a. Circadian periods of single suprachiasmatic neurons in
rats. Neurosci. Lett. 250, 157–160.
Honma, S., Ikeda, M., Abe, H., Tanahashi, Y., Namihira, M., Honma,
K., Nomura, M., 1998b. Circadian oscillation of BMAL1, a partner
of a mammalian clock gene Clock, in rat suprachiasmatic
nucleus. Biochem. Biophys. Res. Commun. 250, 83–87.
Honrado, G.I., Johnson, R.S., Golombek, D.A., Spiegelman, B.M.,
Papaioannou, V.E., Ralph, M.R., 1996. The circadian system of
c-fos deficient mice. J. Comp. Physiol., A 178, 563–570.
Horikawa, K., Yokota, S., Fuji, K., Akiyama, M., Moriya, T.,
Okamura, H., Shibata, S., 2000. Nonphotic entrainment by
5-HT1A/7 receptor agonists accompanied by reduced Per1 and
Per2 mRNA levels in the suprachiasmatic nuclei. J. Neurosci.
20, 5867–5873.
Horowitz, S.S., Blanchard, J.H., Morin, L.P., 2004. Intergeniculate
leaflet and ventral lateral geniculate nucleus afferent
connections: an anatomical substrate for functional input from
the vestibulo-visuomotor system. J. Comp. Neurol. 474, 227–245.
Horowitz, S.S., Blanchard, J.H., Morin, L.P., 2005. Medial vestibular
connections with the hypocretin (orexin) system. J. Comp.
Neurol. 487, 127–146.
Horvath, T.L., 1998. An alternate pathway for visual signal
integration into the hypothalamo-pituitary axis:
retinorecipient intergeniculate neurons project to various
regions of the hypothalamus and innervate neuroendocrine
cells including those producing dopamine. J. Neurosci. 18,
1546–1558.
Hsiao, K., Sachs, G.M., Schneider, G.E., 1984. A minute fraction of
Syrian golden hamster retinal ganglion cells project bilaterally.
J. Neurosci. 4, 359–367.
Huang, R.C., 1993. Sodium and calcium currents in acutely
dissociated neurons from rat suprachiasmatic nucleus.
J. Neurophysiol. 70, 1692–1703.
Hut, R.A., van der Zee, E.A., Jansen, K., Gerkema, M.P., Daan, S.,
2002. Gradual reappearance of post-hibernation circadian
rhythmicity correlates with numbers of
vasopressin-containing neurons in the suprachiasmatic
nuclei of European ground squirrels. J. Comp. Physiol., B 172,
59–70.
Ichikawa, T., Hirata, Y., 1986. Organization of choline
acetyltransferase-containing structure in the forebrain of the
rat. J. Neurosci. 6, 281–292 (Ref Type: Abstract).
Ikeda, M., Sugiyama, T., Wallace, C.S., Gompf, H.S., Yoshioka, T.,
Miyawaki, A., Allen, C.N., 2003. Circadian dynamics of cytosolic
and nuclear Ca2+ in single suprachiasmatic nucleus neurons.
Neuron 38, 253–263.
Ingram, C.D., Ciobanu, R., Coculescu, I.L., Tanasescu, R., Coculescu,
M., Mihai, R., 1998. Vasopressin neurotransmission and the
control of circadian rhythms in the suprachiasmatic nucleus.
Prog. Brain Res. 119, 351–364.
Inouye, S.T., Kawamura, H., 1979. Persistence of circadian
rhythmicity in a mammalian hypothalamic “island”
containing the suprachiasmatic nucleus. Proc. Natl. Acad. Sci.
U. S. A. 76, 5962–5966.
Isoldi, M.C., Rollag, M.D., de Lauro Castrucci, A.M., Provencio, I.,
2005. Rhabdomeric phototransduction initiated by the
vertebrate photopigment melanopsin. Proc. Natl. Acad. Sci.
U. S. A. 102, 1217–1221.
Itri, J., Colwell, C.S., 2003. Regulation of inhibitory synaptic
transmission by vasoactive intestinal peptide (VIP) in the
mouse suprachiasmatic nucleus. J. Neurophysiol. 90,
1589–1597.
Jackson, A.C., Yao, G.L., Bean, B.P., 2004. Mechanism of spontaneous
firing in dorsomedial suprachiasmatic nucleus neurons.
J. Neurosci. 24, 7985–7998.
Jagota, A., De la Iglesia, H.O., Schwartz, W.J., 2000. Morning and
evening circadian oscillations in the suprachiasmatic nucleus
in vitro. Nat. Neurosci. 3, 372–376.
Janik, D., Mrosovsky, N., 1994. Intergeniculate leaflet lesions and
behaviorally-induced shifts of circadian rhythms. Brain Res.
651, 174–182.
Jhaveri, S., Edwards, M.A., Schneider, G.E., 1991. Initial stages of
retinofugal axon development in the hamster: evidence for two
distinct modes of growth. Exp. Brain Res. 87, 371–382.
Jiang, Z.-G., Allen, C.N., North, R.A., 1995. Presynaptic inhibition by
baclofen of retinohypothalamic excitatory synaptic
transmission in rat suprachiasmatic nucleus. Neuroscience 64,
813–819.
Jiang, Z.-G., Yang, Y.-Q., Liu, Z.P., Allen, C.N., 1997. Membrane
properties and synaptic inputs of suprachiasmatic nucleus
neurons in rat brain slices. J. Physiol. 499, 141–159.
Jobst, E.E., Allen, C.N., 2002. Calbindin neurons in the hamster
suprachiasmatic nucleus do not exhibit a circadian variation in
spontaneous firing rate. Eur. J. Neurosci. 16, 2469–2474.
Jobst, E.E., Robinson, D.W., Allen, C.N., 2004. Potential pathways
for intercellular communication within the calbindin
subnucleus of the hamster suprachiasmatic nucleus.
Neuroscience 123, 87–99.
Johnson, R.F., Morin, L.P., Moore, R.Y., 1988a. Retinohypothalamic
projections in the hamster and rat demonstrated using cholera
toxin. Brain Res. 462, 301–312.
Johnson, R.F., Smale, L., Moore, R.Y., Morin, L.P., 1988b. Lateral
geniculate lesions block circadian phase shift responses to a
benzodiazepine. Proc. Natl. Acad. Sci. U. S. A. 85, 5301–5304.
Kallo, I., Kalamatianos, T., Wiltshire, N., Shen, S., Sheward, W.J.,
Harmar, A.J., Coen, C.W., 2004. Transgenic approach reveals
expression of the VPAC2 receptor in phenotypically defined
neurons in the mouse suprachiasmatic nucleus and in its
efferent target sites. Eur. J. Neurosci. 19, 2201–2211.
Kalsbeek, A., Teclemariam-Mesbah, R., Pevet, P., 1993. Efferent
projections of the suprachiasmatic nucleus in the golden
hamster (Mesocricetus auratus). J. Comp. Neurol. 332, 293–314.
Kalsbeek, A., Fliers, E., Franke, A.N., Wortel, J., Buijs, R.M., 2000.
Functional connections between the suprachiasmatic nucleus
and the thyroid gland as revealed by lesioning and viral tracing
techniques in the rat. Endocrinology 141, 3832–3841.
Karatsoreos, I.N., Yan, L., LeSauter, J., Silver, R., 2004. Phenotype
matters: identification of light-responsive cells in the mouse
suprachiasmatic nucleus. J. Neurosci. 24, 68–75.
Kawaguchi, C., Tanaka, K., Isojima, Y., Shintani, N., Hashimoto, H.,
Baba, A., Nagai, K., 2003. Changes in light-induced phase shift
of circadian rhythm in mice lacking PACAP. Biochem. Biophys.
Res. Commun. 310, 169–175.
Kawamoto, K., Nagano, M., Kanda, F., Chihara, K., Shigeyoshi, Y.,
Okamura, H., 2003. Two types of VIP neuronal components in
rat suprachiasmatic nucleus. J. Neurosci. Res. 74, 852–857.
Kennaway, D.J., Moyer, R.W., 1998. Serotonin 5-HT2c agonists
mimic the effect of light pulses on circadian rhythms. Brain
Res. 806, 257–270.
Kennaway, D.J., Moyer, R.W., Voultsios, A., Varcoe, T.J., 2001.
Serotonin, excitatory amino acids and the photic control of
melatonin rhythms and SCN c-FOS in the rat. Brain Res. 897,
36–43.
Kennaway, D.J., Voultsios, A., Varcoe, T.J., Moyer, R.W., 2003.
Melatonin and activity rhythm responses to light pulses in
mice with the Clock mutation. Am. J. Physiol.: Regul., Integr.
Comp. Physiol. 284, R1231–R1240.
Kim, Y.I., Dudek, F.E., 1992. Intracellular electrophysiological
study of suprachiasmatic nucleus neurons in rodents:
inhibitory synaptic mechanisms. J. Physiol. (London) 458,
247–260.
Kim, Y.I., Dudek, F.E., 1993. Membrane properties of rat
suprachiasmatic nucleus neurons receiving optic nerve input.
J. Physiol. (London) 464, 229–243.
Kim, Y.I., Kim, S.H., Kim, D.Y., Lee, H.W., Shin, H.C., Chung, J.M.,
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
Han, H.C., Na, H.S., Hong, S.K., 1999. Electrophysiological
evidence for the role of substance P in retinohypothalamic
transmission in the rat. Neurosci. Lett. 274, 99–102.
Kim, D.Y., Kang, H.C., Shin, H.C., Lee, K.J., Yoon, Y.W., Han, H.C.,
Na, H.S., Hong, S.K., Kim, Y.I., 2001. Substance P plays a critical
role in photic resetting of the circadian pacemaker in the rat
hypothalamus. J. Neurosci. 21, 4026–4031.
King, V.M., Chahad-Ehlers, S., Shen, S., Harmar, A.J., Maywood, E.S.,
Hastings, M.H., 2003. A hVIPR transgene as a novel tool for the
analysis of circadian function in the mouse suprachiasmatic
nucleus. Eur. J. Neurosci. 17, 822–832.
Kiss, J., Halasz, B., 1996. Synaptic contacts between cholinergic
afferents and suprachiasmatic neurones of the rat.
NeuroReport 7, 1961–1964.
Knoch, M.E., Gobes, S.M., Pavlovska, I., Su, C., Mistlberger, R.E.,
Glass, J.D., 2004. Short-term exposure to constant light
promotes strong circadian phase-resetting responses to
nonphotic stimuli in Syrian hamsters. Eur. J. Neurosci. 19,
2779–2790.
Koibuchi, N., Sakai, M., Watanabe, K., Yamaoka, S., 1992. Change in
Fos-like immunoreactivity in the suprachiasmatic nucleus in
the adult male rat. NeuroReport 3, 501–504.
Kononenko, N.I., Dudek, F.E., 2004. Mechanism of irregular firing of
suprachiasmatic nucleus neurons in rat hypothalamic slices.
J. Neurophysiol. 91, 267–273.
Kononenko, N.I., Shao, L.R., Dudek, F.E., 2004. Riluzole-sensitive
slowly inactivating sodium current in rat suprachiasmatic
nucleus neurons. J. Neurophysiol. 91, 710–718.
Kopp, M.D., Schomerus, C., Dehghani, F., Korf, H.W., Meissl, H.,
1999. Pituitary adenylate cyclase-activating polypeptide and
melatonin in the suprachiasmatic nucleus: effects on the
calcium signal transduction cascade. J. Neurosci. 19, 206–219.
Kornhauser, J.M., Nelson, D.E., Mayo, K.E., Takahashi, J.S., 1990.
Photic and circadian regulation of c-fos gene expression in the
hamster suprachiasmatic nucleus. Neuron 5, 127–134.
Kriegsfeld, L.J., Demas, G.E., Lee Jr., S.E., Dawson, T.M., Dawson, V.
L., Nelson, R.J., 1999. Circadian locomotor analysis of male mice
lacking the gene for neuronal nitric oxide synthase (nNOS−/−).
J. Biol. Rhythms 14, 20–27.
Kriegsfeld, L.J., Drazen, D.L., Nelson, R.J., 2001. Circadian organization in male mice lacking the gene for endothelial nitric
oxide synthase (eNOS−/−). J. Biol. Rhythms 16, 142–148.
Kriegsfeld, L.J., LeSauter, J., Silver, R., 2004. Targeted microlesions
reveal novel organization of the hamster suprachiasmatic
nucleus. J. Neurosci. 24, 2449–2457.
Kromer, L.F., Moore, R.Y., 1980. A study of the organization of the
locus coeruleus projections to the lateral geniculate nucleus in
the albino rat. Neuroscience 5, 255–271.
Krout, K.E., Kawano, J., Mettenleiter, T.C., Loewy, A.D., 2002. CNS
inputs to the suprachiasmatic nucleus of the rat. Neuroscience
110, 73–92.
Kuhlman, S.J., McMahon, D.G., 2004. Rhythmic regulation of
membrane potential and potassium current persists in SCN
neurons in the absence of environmental input. Eur. J.
Neurosci. 20, 1113–1117.
Kuhlman, S.J., Quintero, J.E., McMahon, D.G., 2000. GFP
fluorescence reports Period 1 circadian gene regulation in the
mammalian biological clock. NeuroReport 11, 1479–1482.
Kuhlman, S.J., Silver, R., LeSauter, J., Bult-Ito, A., McMahon, D.G.,
2003. Phase resetting light pulses induce Per1 and persistent
spike activity in a subpopulation of biological clock neurons.
J. Neurosci. 23, 1441–1450.
Ladenheim, E.E., Jensen, R.T., Mantey, S.A., Moran, T.H., 1992.
Distinct distributions of two bombesin receptor subtypes in the
rat central nervous system. Brain Res. 593, 168–178.
Ladenheim, E.E., Jensen, R.T., Mantey, S.A., Taylor, J.E., Coy, D.H.,
Moran, T.H., 1993. Bombesin receptor antagonists differentiate
receptor subtypes in rat brain. Eur. J. Pharmacol. 235, 121–125.
Laemle, L.K., Ottenweller, J.E., 2001. The relationship between
51
circadian rhythmicity and vasoactive intestinal polypeptide in
the suprachiasmatic nucleus of congenitally anophthalmic
mice. Brain Res. 917, 105–111.
Lall, G.S., Biello, S.M., 2002. Attenuation of phase shifts to light by
activity or neuropeptide Y: a time course study. Brain Res. 957,
109–116.
Lall, G.S., Biello, S.M., 2003a. Attenuation of circadian light induced
phase advances and delays by neuropeptide Y and a
neuropeptide Y Y1/Y5 receptor agonist. Neuroscience 119,
611–618.
Lall, G.S., Biello, S.M., 2003b. Neuropeptide Y, GABA and circadian
phase shifts to photic stimuli. Neuroscience 120, 915–921.
Land, P.W., Kyonka, E., Shamalla-Hannah, L., 2004. Vesicular
glutamate transporters in the lateral geniculate nucleus:
expression of VGLUT2 by retinal terminals. Brain Res. 996,
251–254.
Larsen, P.J., 1999. Tracing autonomic innervation of the rat pineal
gland using viral transneuronal tracing. Microsc. Res. Tech. 46,
296–304.
Larsen, P.J., Enquist, L.W., Card, J.P., 1998. Characterization of
the multisynaptic neuronal control of the rat pineal gland
using viral transneuronal tracing. Eur. J. Neurosci. 10,
128–145.
Leak, R.K., Moore, R.Y., 2001. Topographic organization of
suprachiasmatic nucleus projection neurons. J. Comp. Neurol.
433, 312–334.
Leak, R.K., Card, J.P., Moore, R.Y., 1999. Suprachiasmatic
pacemaker organization analyzed by viral transynaptic
transport. Brain Res. 819, 23–32.
Leanza, G., Nilsson, O.G., Wiley, R.G., Bjorklund, A., 1995.
Selective lesioning of the basal forebrain cholinergic system
by intraventricular 192 IgG-saporin: behavioural, biochemical
and stereological studies in the rat. Eur. J. Neurosci. 7,
329–343.
Leathers, V.L., Linse, S., Forsen, S., Norman, A.W., 1990.
Calbindin-D28K, a 1 alpha,25-dihydroxyvitamin D3-induced
calcium-binding protein, binds five or six Ca2+ ions with high
affinity. J. Biol. Chem. 265, 9838–9841.
Lee, H.S., Nelms, J.L., Nguyen, M., Silver, R., Lehman, M.N., 2003.
The eye is necessary for a circadian rhythm in the
suprachiasmatic nucleus. Nat. Neurosci. 6, 111–112.
Lehman, M.N., Silver, R., Gladstone, W.R., Kahn, R.M., Gibson, M.,
Bittman, E.L., 1987. Circadian rhythmicity restored by neural
transplant. Immunocytochemical characterization of the graft
and its integration with the host brain. J. Neurosci. 7,
1626–1638.
Lehman, M.N., LeSauter, J., Silver, R., 1998. Fiber outgrowth from
anterior hypothalamic and cortical xenografts in the third
ventricle. J. Comp. Neurol. 391, 133–145.
LeSauter, J., Silver, R., 1998. Output signals of the SCN. Chronobiol.
Int. 15, 535–550.
LeSauter, J., Silver, R., 1999. Localization of a suprachiasmatic
nucleus subregion regulating locomotor rhythmicity.
J. Neurosci. 19, 5574–5585.
LeSauter, J., Lehman, M.N., Silver, R., 1996. Restoration of circadian
rhythmicity by transplants of SCN “micropunches”. J. Biol.
Rhythms 11, 163–171.
LeSauter, J., Romero, P., Cascio, M., Silver, R., 1997. Attachment site
of grafted SCN influences precision of restored circadian
rhythm. J. Biol. Rhythms 12, 327–338.
LeSauter, J., Stevens, P., Jansen, H., Lehman, M.N., Silver, R., 1999.
Calbindin expression in the hamster SCN is influenced by
circadian genotype and by photic conditions. NeuroReport 10,
3159–3163.
LeSauter, J., Kriegsfeld, L.J., Hon, J., Silver, R., 2002. Calbindin-D28K
cells selectively contact intra-SCN neurons. Neuroscience 111,
575–585.
LeSauter, J., Yan, L., Vishnubhotla, B., Quintero, J.E., Kuhlman, S.J.,
McMahon, D.G., Silver, R., 2003. A short half-life GFP mouse
52
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
model for analysis of suprachiasmatic nucleus organization.
Brain Res. 964, 279–287.
Lewandowski, M.H., Usarek, A., 2002. Effects of intergeniculate
leaflet lesions on circadian rhythms in the mouse. Behav. Brain
Res. 128, 13–17.
Lewandowski, M.H., Blasiak, T., Domoslawski, J., Wolkowska, A.,
2000. Ultradian rhythmic neuronal oscillation in the
intergeniculate leaflet. NeuroReport 11, 317–321.
Lewandowski, M.H., Blasiak, T., Blasiak, A., 2002. Are ultra-slow
isoperiodic oscillations in rat intergeniculate leaflet neurons
dependent on reciprocal connection with its contralaterally
located counterpart? Neurosci. Lett. 330, 243–246.
Li, X., Gilbert, J., Davis, F.C., 2005. Disruption of masking by
hypothalamic lesions in Syrian hamsters. J. Comp. Physiol., A
Neuroethol. Sens. Neural Behav. Physiol. 191, 23–30.
Ling, C., Schneider, G.E., Jhaveri, S., 1998. Target-specific
morphology of retinal axon arbors in the adult hamster. Vis.
Neurosci. 15, 559–579.
Liou, S.Y., Albers, H.E., 1991. Single unit response of neurons
within the hamster suprachiasmatic nucleus to neuropeptide
Y. Brain Res. Bull. 27, 825–828.
Liou, S.-Y., Shibata, S., Iwasaki, K., Ueki, S., 1986. Optic-nerve
stimulation-induced increase of release of H-3 glutamate and
H-3 aspartate but not H-3 GABA from the suprachiasmatic
nucleus in slices of rat hypothalamus. Brain Res. Bull. 16,
527–531.
Liou, S.-Y., Shibata, S., Albers, H.E., Ueki, S., 1990. Effects of GABA
and anxiolytics on the single unit discharge of
suprachiasmatic neurons in rat hypothalamic slices. Brain
Res. Bull. 25, 103–107.
Liu, C., Gillette, M.U., 1996. Cholinergic regulation of the
suprachiasmatic nucleus circadian rhythm via a muscarinic
mechanism at night. J. Neurosci. 16, 744–751.
Liu, C., Reppert, S.M., 2000. GABA synchronizes clock cells within
the suprachiasmatic circadian clock. Neuron 25, 123–128.
Liu, C., Weaver, D.R., Strogatz, S.H., Reppert, S.M., 1997a. Cellular
construction of a circadian clock: period determination in the
suprachiasmatic nuclei. Cell 91, 855–860.
Liu, C., Ding, J.M., Faiman, L.E., Gillette, M.U., 1997b. Coupling of
muscarinic cholinergic receptors and cGMP in nocturnal
regulation of the suprachiasmatic circadian clock. J. Neurosci.
17, 659–666.
Liu, Q.S., Xu, Q., Arcuino, G., Kang, J., Nedergaard, M., 2004.
Astrocyte-mediated activation of neuronal kainate receptors.
Proc. Natl. Acad. Sci. U. S. A. 101, 3172–3177.
Lovenberg, T.W., Baron, B.M., De Lecea, L., Miller, J.D., Prosser, R.A.,
Rea, M.A., Foye, P.E., Racke, M., Slone, A.L., Siegel, B.W.,
Danielson, P.E., Sutcliffe, J.G., Erlander, M.G., 1993. A novel
adenylyl cyclase-activating serotonin receptor (5-HT7)
implicated in the regulation of mammalian circadian rhythms.
Neuron 11, 449–458.
Lucas, R.J., Foster, R.G., 1999. Neither functional rod
photoreceptors nor rod or cone outer segments are required
for the photic inhibition of pineal melatonin. Endocrinology
140, 1520–1524.
Lucas, R.J., Freedman, M.S., Munoz, M., Garcia-Fernández, J.-M.,
Foster, R.G., 1999. Regulation of the mammalian pineal by
non-rod, non-cone, ocular photoreceptors. Science 284,
505–507.
Lucas, R.J., Douglas, R.H., Foster, R.G., 2001. Characterization of an
ocular photopigment capable of driving pupillary constriction
in mice. Nat. Neurosci. 4, 621–626.
Lucas, R.J., Hattar, S., Takao, M., Berson, D.M., Foster, R.G., Yau, K.
W., 2003. Diminished pupillary light reflex at high irradiances
in melanopsin-knockout mice. Science 299, 245–247.
Madeira, M.D., Pereira, P.A., Silva, S.M., Cadete-Leite, A.,
Paula-Barbosa, M.M., 2004. Basal forebrain neurons modulate
the synthesis and expression of neuropeptides in the rat
suprachiasmatic nucleus. Neuroscience 125, 889–901.
Mahoney, M.M., Nunez, A.A., Smale, L., 2000. Calbindin and Fos
within the suprachiasmatic nucleus and the adjacent
hypothalamus of Arvicanthis niloticus and Rattus norvegicus.
Neuroscience 99, 565–575.
Mai, L.M., Pan, J.T., 1993. Central administration of bombesin
blocks the estrogen-induced afternoon prolactin surge.
Neuroendocrinology 57, 40–44.
Mai, L.M., Pan, J.T., 1995. Bombesin acts in the suprachiasmatic
nucleus to affect circadian changes in tuberoinfundibular
dopaminergic neuron activity and prolactin secretion.
Endocrinology 136, 4163–4167.
Major, D.E., Rodman, H.R., Libedinsky, C., Karten, H.J., 2003. Pattern
of retinal projections in the California ground squirrel
(Spermophilus beecheyi): anterograde tracing study using cholera
toxin. J. Comp. Neurol. 463, 317–340.
Manrique, C., Héry, F., Faudon, M., François-Bellan, A.M., 1999.
Indirect evidence for an association of 5-HT1B binding sites
with retinal and geniculate axon terminals in the rat
suprachiasmatic nucleus. Synapse 33, 314–323.
Marchant, E.G., Morin, L.P., 1999. The hamster circadian rhythm
system includes nuclei of the subcortical visual shell.
J. Neurosci. 19, 10482–10493.
Marchant, E.G., Watson, N.V., Mistlberger, R.E., 1997. Both
neuropeptide Y and serotonin are necessary for entrainment of
circadian rhythms in mice by daily treadmill running
schedules. J. Neurosci. 17, 7974–7987.
Margraf, R.R., Lynch, G.R., 1993. Melatonin injections affect
circadian behavior and SCN neurophysiology in Djungarian
hamsters. Am. J. Physiol. 264, R615–R621.
Martinet, L., Bonnefond, C., Peytevin, J., Monnerie, R., Marcilloux, J.C.,
1995. Vasoactive intestinal polypeptide in the suprachiasmatic
nucleus of the mink (Mustela vison) could play a key role in photic
induction. J. Neuroendocrinol. 7, 69–79.
Maywood, E.S., Mrosovsky, N., 2001. A molecular explanation of
interactions between photic and non-photic circadian
clock-resetting stimuli. Gene Expr. Patterns 1, 27–31.
Maywood, E.S., Smith, E., Hall, S.J., Hastings, M.H., 1997. A thalamic
contribution to arousal-induced, non-photic entrainment of
the circadian clock of the Syrian hamster. Eur. J. Neurosci. 9,
1739–1747.
Maywood, E.S., Mrosovsky, N., Field, M.D., Hastings, M.H., 1999.
Rapid down-regulation of mammalian Period genes during
behavioral resetting of the circadian clock. Proc. Natl. Acad.
Sci. U. S. A. 96, 15211–15216.
Maywood, E.S., Okamura, H., Hastings, M.H., 2002. Opposing
actions of neuropeptide Y and light on the expression of
circadian clock genes in the mouse suprachiasmatic nuclei.
Eur. J. Neurosci. 15, 216–220.
McArthur, A.J., Coogan, A.N., Ajpru, S., Sugden, D., Biello, S.M.,
Piggins, H.D., 2000. Gastrin-releasing peptide phase-shifts
suprachiasmatic nuclei neuronal rhythms in vitro. J. Neurosci.
20, 5496–5502.
Mead, S., Ebling, F.J.P., Maywood, E.S., Humby, T., Herbert, J.,
Hastings, M.H., 1992. A nonphotic stimulus causes
instantaneous phase advances of the light-entrainable
circadian oscillator of the Syrian hamster but does not induce
the expression of c-fos in the suprachiasmatic nuclei.
J. Neurosci. 12, 2516–2522.
Meijer, J.H., van der Zee, E.A., Dietz, M., 1988. Glutamate phase
shifts circadian activity rhythms in hamsters. Neurosci. Lett.
86, 177–183.
Meijer, J.H., Schaap, J., Watanabe, K., Albus, H., 1997. Multiunit
activity recordings in the suprachiasmatic nuclei: in vivo vs. in
vitro models. Brain Res. 753, 322–327.
Meijer, J.H., Ruijs, A.C.J., Albus, H., Van de Geest, B., Duindam, H.,
Zwinderman, A.H., Dahan, A., 2000. Fentanyl, a μ-opioid
receptor agonist, phase shifts the hamster circadian
pacemaker. Brain Res. 868, 135–140.
Melyan, Z., Tarttelin, E.E., Bellingham, J., Lucas, R.J., Hankins, M.W.,
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
2005. Addition of human melanopsin renders mammalian cells
photoresponsive. Nature 433, 741–745.
Menaker, M., 1968. Extraretinal light perception in the sparrow: I.
Entrainment of the biological clock. Proc. Natl. Acad. Sci.
U. S. A. 59, 414–421.
Menaker, M., Keatts, H., 1968. Extraretinal light perception in the
sparrow: II. Photoperiodic stimulation of testis growth. Proc.
Natl. Acad. Sci. U. S. A. 60, 146–151.
Menaker, M., Underwood, H., 1976. Extraretinal photoreception in
birds. Photophysiology 23, 299–306.
Menet, J.S., Vuillez, P., Pevet, P., 2003. Calbindin expression in the
hamster suprachiasmatic nucleus depends on day-length.
Neuroscience 122, 591–598.
Meyer-Bernstein, E.L., Morin, L.P., 1996. Differential serotonergic
innervation of the suprachiasmatic nucleus and the
intergeniculate leaflet and its role in circadian rhythm
modulation. J. Neurosci. 16, 2097–2111.
Meyer-Bernstein, E.L., Morin, L.P., 1998. Destruction of
serotonergic neurons in the median raphe nucleus blocks
circadian rhythm phase shifts to triazolam, but not to novel
wheel access. J. Biol. Rhythms 13, 494–505.
Meyer-Bernstein, E.L., Morin, L.P., 1999. Electrical stimulation of
the median or dorsal raphe nuclei reduces light-induced FOS
protein in the suprachiasmatic nucleus and causes circadian
activity rhythm phase shifts. Neuroscience 92, 267–279.
Meyer-Bernstein, E.L., Blanchard, J.H., Morin, L.P., 1997. The
serotonergic projection from the median raphe nucleus to the
suprachiasmatic nucleus modulates activity phase onset, but
not other circadian rhythm parameters. Brain Res. 755,
112–120.
Meyer-Bernstein, E.L., Jetton, A.E., Matsumoto, S.I., Markuns, J.F.,
Lehman, M.N., Bittman, E.L., 1999. Effects of suprachiasmatic
transplants on circadian rhythms of neuroendocrine function
in golden hamsters. Endocrinology 140, 207–218.
Meyer-Spasche, A., Reed, H.E., Piggins, H.D., 2002. Neurotensin
phase-shifts the firing rate rhythm of neurons in the rat
suprachiasmatic nuclei in vitro. Eur. J. Neurosci. 16, 339–344.
Mick, G., Maeno, H., Kiyama, H., Tohyama, M., 1994. Marginal
topography of neurons expressing the substance P receptor
in the rat suprachiasmatic nucleus. Mol. Brain Res. 21,
157–161.
Mick, G., Shigemoto, R., Kitahama, K., 1995. Localization of
substance P receptors in central neural structures controlling
daily rhythms in nocturnal rodents. C. R. Acad. Sci. III 318,
209–217.
Mikkelsen, J.D., 1990. Projections from the lateral geniculate
nucleus to the hypothalamus of the Mongolian gerbil (Meriones
unguiculatus): an anterograde and retrograde tracing study.
J. Comp. Neurol. 299, 493–508.
Mikkelsen, J.D., Larsen, P.J., 1993. Substance P in the
suprachiasmatic nucleus of the rat: an immunohistochemical
and in situ hybridization study. Histochemistry 100, 3–16.
Mikkelsen, J.D., Vrang, N., 1994. A direct pretectosuprachiasmatic
projection in the rat. Neuroscience 62, 497–505.
Miller, J.D., Fuller, C.A., 1992. Isoperiodic neuronal activity in
suprachiasmatic nucleus of the rat. Am. J. Physiol. 263,
R51–R58.
Miller, J.D., Morin, L.P., Schwartz, W.J., Moore, R.Y., 1996. New
insights into the mammalian circadian clock [Review] [230 refs]
Sleep 19, 641–667.
Mintz, E.M., Albers, H.E., 1997. Microinjection of NMDA into the
SCN region mimics the phase shifting effect of light in
hamsters. Brain Res. 758, 245–249.
Mintz, E.M., Marvel, C.L., Gillespie, C.F., Price, K.M., Albers, H.E.,
1999. Activation of NMDA receptors in the suprachiasmatic
nucleus produces light-like phase shifts of the circadian clock
in vivo. J. Neurosci. 19, 5124–5130.
Mintz, E.M., van den Pol, A.N., Casano, A.A., Albers, H.E., 2001.
Distribution of hypocretin-(orexin) immunoreactivity in the
53
central nervous system of Syrian hamsters (Mesocricetus
auratus). J. Chem. Neuroanat. 21, 225–238.
Mintz, E.M., Jasnow, A.M., Gillespie, C.F., Huhman, K.L., Albers, H.
E., 2002. GABA interacts with photic signaling in the
suprachiasmatic nucleus to regulate circadian phase shifts.
Neuroscience 109, 773–778.
Mistlberger, R.E., Antle, M.C., 1998. Behavioral inhibition of
light-induced circadian phase resettting is phase and
serotonin dependent. Brain Res. 786, 31–38.
Mistlberger, R.E., Landry, G.J., Marchant, E.G., 1997. Sleep
deprivation can attenuate light-induced phase shifts of
circadian rhythms in hamsters. Neurosci. Lett. 238, 5–8.
Mistlberger, R.E., Antle, M.C., Glass, J.D., Miller, J.D., 2000.
Behavioral and serotonergic regulation of circadian rhythms.
Biol. Rhythm Res. 31, 240–283.
Mistlberger, R.E., Belcourt, J., Antle, M.C., 2002. Circadian clock
resetting by sleep deprivation without exercise in Syrian
hamsters: dark pulses revisited. J. Biol. Rhythms 17,
227–237.
Mistlberger, R.E., Antle, M.C., Webb, I.C., Jones, M., Weinberg, J.,
Pollock, M.S., 2003. Circadian clock resetting by arousal in
Syrian hamsters: the role of stress and activity. Am. J. Physiol.:
Regul., Integr. Comp. Physiol. 285, R917–R925.
Moffett, J.R., 2003. Reductions in N-acetylaspartylglutamate and
the 67 kDa form of gluamic acid decarboxylase
immunoreactivities in the visual system of albino and
pigmented rats after optic nerve transections. J. Comp. Neurol.
458, 221–239.
Moffett, J.R., Williamson, L., Palkovits, M., Namboodiri, M.A.A.,
1990. N-acetylaspartylglutamate: a transmitter candidate for
the retinohypothalamic tract. Proc. Natl. Acad. Sci. U. S. A. 87,
8065–8069.
Moffett, J.R., Williamson, L.C., Neale, J.H., Palkovits, M.,
Namboodin, M.A.A., 1991. Effect of optic nerve transection on
N-acetylaspartylglutamate immunoreactivity in the primary
and accessory optic projection systems in the rat. Brain Res.
538, 86–94.
Moga, M.M., 1998. 192 IgG-saporin abolishes p75 neurotrophin
receptor immunoreactivity in rat SCN. NeuroReport 9,
3197–3200.
Moga, M.M., Moore, R.Y., 1996. Putative excitatory amino acid
projections to the suprachiasmatic nucleus in the rat. Brain
Res. 743, 171–177.
Moga, M.M., Moore, R.Y., 1997. Organization of neural inputs to the
suprachiasmatic nucleus in the rat. J. Comp. Neurol. 389,
508–534.
Moore, R.Y., 1983. Organization and function of a central nervous
system oscillator: the suprachiasmatic hypothalamic nucleus.
Fed. Proc. 42, 2783–2789.
Moore, R.Y., 1989. The geniculohypothalamic tract in monkey and
man. Brain Res. 486, 190–194.
Moore, R.Y., 1993. Organization of the primate circadian system.
J. Biol. Rhythms 8, S3–S9.
Moore, R.Y., 1996. Entrainment pathways and the functional
organization of the circadian timing system. In: Buijs, R.,
Kalsbeek, A., Romijn, H.J., Pennartz, C.M.A., Mirmiran, M. (Eds.),
Hypothalamic Integration of Circadian Rhythms. Elsevier,
Amsterdam, pp. 101–117.
Moore, R.Y., Card, J.P., 1994. Intergeniculate leaflet: an
anatomically and functionally distinct subdivision of the
lateral geniculate complex. J. Comp. Neurol. 344, 403–430.
Moore, R.Y., Lenn, N.J., 1972. A retinohypothalamic projection in
the rat. J. Comp. Neurol. 146, 1–14.
Moore, R.Y., Speh, J.C., 1993. GABA is the principal
neurotransmitter of the circadian system. Neurosci. Lett. 150,
112–116.
Moore, R.Y., Gustafson, E.L., Card, J.P., 1984. Identical
immunoreactivity of afferents to the rat suprachiasmatic
nucleus with antisera against avian pancreatic polypeptide,
54
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
molluscan cardiaoexcitatory peptide and neuropeptide Y.
Cell Tissue Res. 236, 41–46.
Moore, R.Y., Speh, J.C., Card, J.P., 1995. The retinohypothalamic
tract originates from a distinct subset of retinal ganglion cells.
J. Comp. Neurol. 352, 351–366.
Moore, R.Y., Weis, R., Moga, M.M., 2000. Efferent projections of the
intergeniculate leaflet and the ventral lateral geniculate
nucleus in the rat. J. Comp. Neurol. 420, 398–418.
Moore, R.Y., Speh, J.C., Leak, R.K., 2002. Suprachiasmatic nucleus
organization. Cell Tissue Res. 309, 89–98.
Mori, K., Miyazato, M., Ida, T., Murakami, N., Serino, R., Ueta, Y.,
Kojima, M., Kangawa, K., 2005. Identification of neuromedin S
and its possible role in the mammalian circadian oscillator
system. EMBO J. 24, 325–335.
Morin, L.P., 1994. The circadian visual system. Brain Res. Rev. 67,
102–127.
Morin, L.P., 1999. Serotonin and the regulation of mammalian
circadian rhythmicity. Ann. Med. 31, 12–33.
Morin, L.P., Blanchard, J.H., 1991. Depletion of brain serotonin by
5,7-DHT modifies hamster circadian rhythm response to light.
Brain Res. 566, 173–185.
Morin, L.P., Blanchard, J.H., 1995. Organization of the hamster
intergeniculate leaflet: NPY and ENK projections to the
suprachiasmatic nucleus, intergeniculate leaflet and posterior
limitans nucleus. Vis. Neurosci. 12, 57–67.
Morin, L.P., Blanchard, J.H., 1998. Interconnections among nuclei
of the subcortical visual shell: the intergeniculate leaflet is a
major constituent of the hamster subcortical visual system.
J. Comp. Neurol. 396, 288–309.
Morin, L.P., Blanchard, J.H., 1999. Forebrain connections of the
hamster intergeniculate leaflet: comparison with those of
ventral lateral geniculate nucleus and retina. Vis. Neurosci. 16,
1037–1054.
Morin, L.P., Blanchard, J.H., 2001. Neuromodulator content of
hamster intergeniculate leaflet neurons and their projection to
the suprachiasmatic nucleus or visual midbrain. J. Comp.
Neurol. 437, 79–90.
Morin, L.P., Blanchard, J.H., 2005. Descending projections of the
hamster intergeniculate leaflet: relationship to the sleep/
arousal and visuomotor systems. J. Comp. Neurol. 487, 204–216.
Morin, L.P., Pace, L., 2002. The intergeniculate leaflet, but not the
visual midbrain, mediates hamster circadian rhythm response
to constant light. J. Biol. Rhythms 17, 217–226.
Morin, L.P., Blanchard, J.H., Moore, R.Y., 1992. Intergeniculate
leaflet and suprachiasmatic nucleus organization and
connections in the hamster. Vis. Neurosci. 8, 219–230.
Morin, L.P., Goodless-Sanchez, N., Smale, L., Moore, R.Y., 1994.
Projections of the suprachiasmatic nuclei, subparaventricular
zone and retrochiasmatic area in the golden hamster.
Neuroscience 61, 391–410.
Morin, L.P., Blanchard, J.H., Provencio, I., 2003. Retinal ganglion
cell projections to the hamster suprachiasmatic nucleus,
intergeniculate leaflet and visual midbrain: bifurcation and
melanopsin immunoreactivity. J. Comp. Neurol. 465,
401–416.
Moriya, T., Yamanouchi, S., Fukushima, T., Shimazoe, T., Shibata,
S., Watanabe, S., 1996. Involvement of 5-HT1A receptor
mechanisms in the inhibitory effects of methamphetamine on
photic responses in the rodent suprachiasmatic nucleus. Brain
Res. 740, 261–267.
Moriya, T., Yoshinobu, Y., Ikeda, M., Yokota, S., Akiyama, M.,
Shibata, S., 1998. Potentiating action of MKC-242, a selective
5-HT1A receptor agonist, on the photic entrainment of the
circadian activity rhythm in hamsters. Br. J. Pharmacol. 125,
1281–1287.
Moyer, R.W., Kennaway, D.J., 1999. Immunohistochemical
localization of serotonin receptors in the rat suprachiasmatic
nucleus. Neurosci. Lett. 271, 147–150.
Moyer, R.W., Kennaway, D.J., 2000. Serotonin depletion decreases
light induced c-fos in the rat suprachiasmatic nucleus.
NeuroReport 11, 1021–1024.
Mrosovsky, N., 1996. Locomotor activity and non-photic
influences on circadian clocks. Biol. Rev. Camb. Philos. Soc. 71,
343–372.
Mrosovsky, N., 1999. Masking: history, definitions, and
measurement. Chronobiol. Int. 16, 415–429.
Mrosovsky, N., 2001. Further characterization of the phenotype of
mCry1/mCry2-deficient mice. Chronobiol. Int. 18, 613–625.
Mrosovsky, N., 2003. Contribution of classic photoreceptors to
entrainment. J. Comp. Physiol., A 189, 69–73.
Mrosovsky, N., Hattar, S., 2003. Impaired masking responses to
light in melanopsin-knockout mice. Chronobiol. Int. 20,
989–999.
Mrosovsky, N., Foster, R.G., Salmon, P.A., 1999. Thresholds for
masking responses to light in three strains of retinally
degenerate mice. J. Comp. Physiol., A 184, 423–428.
Mrosovsky, N., Salmon, P.A., Foster, R.G., McCall, M.A., 2000.
Responses to light after retinal degeneration. Vision Res. 40,
575–578.
Mrugala, M., Zlomanczuk, P., Jagota, A., Schwartz, W.J., 2000.
Rhythmic multiunit neural activity in slices of hamster
suprachiasmatic nucleus reflect prior photoperiod. Am. J.
Physiol.: Regul., Integr. Comp. Physiol. 278, R987–R994.
Munch, I.C., Moller, M., Larsen, P.J., Vrang, N., 2002. Light-induced
c-fos expression in suprachiasmatic nuclei neurons targeting
the paraventricular nucleus of the hamster hypothalamus:
phase dependence and immunochemical identification.
J. Comp. Neurol. 442, 48–62.
Muscat, L., Morin, L.P., in press. Bionocular contributions to the
responsiveness and integrative capacity of the circadian
rhythm system to light. J. Biol. Rhythms.
Muscat, L., Huberman, A.D., Jordan, C.L., Morin, L.P., 2003. Crossed
and uncrossed retinal projections to the hamster circadian
system. J. Comp. Neurol. 466, 513–524.
Muscat, L., Tischler, R.C., Morin, L.P., 2005. Functional analysis of
the role of the median raphe as a regulator of hamster
circadian system sensitivity to light. Brain Res. 1044 (1), 59–66.
Nagano, M., Adachi, A., Nakahama, K., Nakamura, T., Tamada, M.,
Meyer-Bernstein, E., Sehgal, A., Shigeyoshi, Y., 2003. An abrupt
shift in the day/night cycle causes desynchrony in the
mammalian circadian center. J. Neurosci. 23, 6141–6151.
Nakahara, K., Hanada, R., Murakami, N., Teranishi, H., Ohgusu, H.,
Fukushima, N., Moriyama, M., Ida, T., Kangawa, K., Kojima, M.,
2004. The gut-brain peptide neuromedin U is involved in the
mammalian circadian oscillator system. Biochem. Biophys.
Res. Commun. 318, 156–161.
Nakamura, H., Itoh, K., 2004. Cytoarchitectonic and connectional
organization of the ventral lateral geniculate nucleus in the cat.
J. Comp. Neurol. 473, 439–462.
Nakamura, W., Honma, S., Shirakawa, T., Honma, K., 2002. Clock
mutation lengthens the circadian period without damping
rhythms in individual SCN neurons. Nat. Neurosci. 5, 399–400.
Neale, J.H., Bzdega, T., Wroblewska, B., 2000.
N-Acetylaspartylglutamate: the most abundant peptide
neurotransmitter in the mammalian central nervous system.
J. Neurochem. 75, 443–452.
Negroni, J., Nevo, E., Cooper, H.M., 1997. Neuropeptidergic
organization of the suprachiasmatic nucleus in the blind mole
rat (Spalax ehrenbergi). Brain Res. Bull. 44, 633–639.
Nelson, D.E., Takahashi, J.S., 1991. Sensitivity and integration in a
visual pathway for circadian entrainment in the hamster
(Mesocricetus auratus). J. Physiol. (London) 439, 115–145.
Nelson, D.E., Takahashi, J.S., 1999. Integration and saturation
within the circadian photic entrainment pathway of hamsters.
Am. J. Physiol.: Regul., Integr. Comp. Physiol. 277, R1351–R1361.
Newman, L.A., Walker, M.T., Brown, R.L., Cronin, T.W., Robinson,
P.R., 2003. Melanopsin forms a functional short-wavelength
photopigment. Biochemistry 42, 12734–12738.
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
Novak, C.M., Albers, H.E., 2004a. Novel phase-shifting effects of
GABAA receptor activation in the suprachiasmatic nucleus of a
diurnal rodent. Am. J. Physiol.: Regul., Integr. Comp. Physiol.
286, R820–R825.
Novak, C.M., Albers, H.E., 2004b. Circadian phase alteration by
GABA and light differs in diurnal and nocturnal rodents during
the day. Behav. Neurosci. 118, 498–504.
Novak, C.M., Ehlen, J.C., Huhman, K.L., Albers, H.E., 2004. GABA(B)
receptor activation in the suprachiasmatic nucleus of diurnal
and nocturnal rodents. Brain Res. Bull. 63, 531–535.
Nunez, A.A., Bult, A., McElhinny, T.L., Smale, L., 1999. Daily
rhythms of Fos expression in hypothalamic targets of the
suprachiasmatic nucleus in diurnal and nocturnal rodents.
J. Biol. Rhythms 14, 300–306.
O'Hara, B.F., Edgar, D.M., Cao, V.H., Wiler, S.W., Heller, H.C., Kilduff,
T.S., Miller, J.D., 1998. Nicotine and nicotinic receptors in the
circadian system. Psychoneuroendocrinology 23, 161–173.
Oliver, K.R., Kinsey, A.M., Wainwright, A., Sirinathsinghji, D.J.,
2000. Localization of 5-ht(5A) receptor-like immunoreactivity
in the rat brain. Brain Res. 867, 131–142.
Otori, Y., Tominaga, K., Fukuhara, C., Yang, J., Yamazaki, S.,
Cagampang, F.R.A., Okamura, H., Inouye, S.-I.T., 1993.
Substance P-like immunoreactivity in the suprachiasmatic
nucleus of the rat. Brain Res. 619, 271–277.
Panda, S., Sato, T.K., Castrucci, A.M., Rollag, M.D., DeGrip, W.J.,
Hogenesch, J.B., Provencio, I., Kay, S.A., 2002. Melanopsin
(Opn4) requirement for normal light-induced circadian phase
shifting. Science 298, 2213–2216.
Panda, S., Provencio, I., Tu, D.C., Pires, S.S., Rollag, M.D., Castrucci,
A.M., Pletcher, M.T., Sato, T.K., Wiltshire, T., Andahazy, M., Kay,
S.A., Van Gelder, R.N., Hogenesch, J.B., 2003. Melanopsin is
required for non-image-forming photic responses in blind
mice. Science 301, 525–527.
Panda, S., Nayak, S.K., Campo, B., Walker, J.R., Hogenesch, J.B.,
Jegla, T., 2005. Illumination of the melanopsin signaling
pathway. Science 307, 600–604.
Panula, P., Yang, H.Y., Costa, E., 1984. Comparative distribution of
bombesin/GRP- and substance-P-like immunoreactivities in
rat hypothalamus. J. Comp. Neurol. 224, 606–617.
Paul, K.N., Fukuhara, C., Tosini, G., Albers, H.E., 2003.
Transduction of light in the suprachiasmatic nucleus:
evidence for two different neurochemical cascades
regulating the levels of Per1 mRNA and pineal melatonin.
Neuroscience 119, 137–144.
Paul, K.N., Gamble, K.L., Fukuhara, C., Novak, C.M., Tosini, G.,
Albers, H.E., 2004. Tetrodotoxin administration in the
suprachiasmatic nucleus prevents NMDA-induced reductions
in pineal melatonin without influencing Per1 and Per2 mRNA
levels. Eur. J. Neurosci. 19, 2808–2814.
Penev, P.D., Zee, P.C., Wallen, E.P., Turek, F.W., 1995. Aging alters
the phase-resetting properties of a serotonin agonist on
hamster circadian rhythmicity. Am. J. Physiol. 268, R293–R298.
Pennartz, C.M., Bierlaagh, M.A., Geurtsen, A.M., 1997. Cellular
mechanisms underlying spontaneous firing in rat
suprachiasmatic nucleus: involvement of a slowly inactivating
component of sodium current. J. Neurophysiol. 78, 1811–1825.
Pennartz, C.M., Bos, N.P., Jeu, M.T., Geurtsen, A.M., Mirmiran, M.,
Sluiter, A.A., Buijs, R.M., 1998. Membrane properties and
morphology of vasopressin neurons in slices of rat
suprachiasmatic nucleus. J. Neurophysiol. 80, 2710–2717.
Pennartz, C.M., de Jeu, M.T., Bos, N.P., Schaap, J., Geurtsen, A.M.,
2002. Diurnal modulation of pacemaker potentials and calcium
current in the mammalian circadian clock. Nature 416,
286–290.
Peterson, B.B., Liao, H.W., Dacey, D.M., Yau, K.-W., Gamlin, P.D.R.,
Robinson, F.R., Marshak, D.W., 2003. Functional architecture of
the photoreceptive ganglion cell in primate retina:
morphology, mosaic organization and central targets of
melanopsin immunostained cells. ARVO Abst. 5182.
55
Pickard, G.E., 1982. The afferent connections of the
suprachiasmatic nucleus of the golden hamster with emphasis
on the retinohypothalamic projection. J. Comp. Neurol. 211,
65–83.
Pickard, G.E., 1985. Bifurcating axons of retinal ganglion cells
terminate in the hypothalamic suprachiasmatic nucleus and
the intergeniculate leaflet of the thalamus. Neurosci. Lett. 55,
211–217.
Pickard, G.E., 1994. Intergeniculate leaflet ablation alters circadian
rhythms in the mouse. NeuroReport 5, 2186–2188.
Pickard, G.E., Rea, M.A., 1997. TFMPP, a 5HT1B receptor agonist,
inhibits light-induced phase shifts of the circadian activity
rhythm and c-Fos expression in the mouse suprachiasmatic
nucleus. Neurosci. Lett. 231, 95–98.
Pickard, G.E., Silverman, A.J., 1981. Direct retinal projections to
hypothalamus, piriform cortex and accessory optic nuclei in
the golden hamster as demonstrated by a sensitive
anterograde horseradish peroxidase technique. J. Comp.
Neurol. 196, 155–172.
Pickard, G.E., Turek, F.W., 1983. The suprachiasmatic nuclei: two
circadian clocks? Brain Res. 268, 201–210.
Pickard, G.E., Ralph, M.R., Menaker, M., 1987. The intergeniculate
leaflet partially mediates effects of light on circadian rhythms.
J. Biol. Rhythms 2, 35–56.
Pickard, G.E., Weber, E.T., Scott, P.A., Riberdy, A.F., Rea, M.A., 1996.
5HT1B receptor agonists inhibit light-induced phase shifts of
behavioral circadian rhythms and expression of the
immediate-early gene c-fos in the suprachiasmatic nucleus.
J. Neurosci. 16, 8208–8220.
Pickard, G.E., Smith, B.N., Belenky, M., Rea, M.A., Dudek, F.E.,
Sollars, P.J., 1999. 5-HT1B receptor-mediated presynaptic
inhibition of retinal input to the suprachiasmatic nucleus.
J. Neurosci. 19, 4034–4045.
Pickard, G.E., Smeraski, C.A., Tomlinson, C.C., Banfield, B.W.,
Kaufman, J., Wilcox, C.L., Enquist, L.W., Sollars, P.J., 2002.
Intravitreal injection of the attenuated pseudorabies virus PRV
Bartha results in infection of the hamster suprachiasmatic
nucleus only by retrograde transsynaptic transport via
autonomic circuits. J. Neurosci. 22, 2701–2710.
Piggins, H.D., Rusak, B., 1993. Electrophysiological effects of
pressure-ejected bombesin-like peptides on hamster
suprachiasmatic nucleus neurons in vitro. J. Neuroendocrinol.
5, 575–581.
Piggins, H.D., Rusak, B., 1997. Effects of microinjections of
substance P into the suprachiasmatic nucleus region on
hamster wheel-running rhythms. Brain Res. Bull. 42,
451–455.
Piggins, H.D., Cutler, D.J., Rusak, B., 1994. Effects of
ionophoretically applied bombesin-like peptides on hamster
suprachiasmatic nucleus neurons in vitro. Eur. J. Pharmacol.
271, 413–419.
Piggins, H.D., Cutler, D.J., Rusak, B., 1995a. Ionophoretically applied
substance P activates hamster suprachiasmatic nucleus
neurons. Brain Res. Bull. 37, 475–479.
Piggins, H.D., Antle, M.C., Rusak, B., 1995b. Neuropeptides phase
shift the mammalian circadian pacemaker. J. Neurosci. 15,
5612–5622.
Piggins, H.D., Stamp, J.A., Burns, J., Rusak, B., Semba, K., 1996.
Distribution of pituitary adenylate cyclase activating
polypeptide (PACAP) immunoreactivity in the hypothalamus
and extended amygdala of the rat. J. Comp. Neurol. 376,
278–294.
Piggins, H.D., Marchant, E.G., Goguen, D., Rusak, B., 2001a.
Phase-shifting effects of pituitary adenylate cyclase activating
polypeptide on hamster wheel-running rhythms. Neurosci.
Lett. 305, 25–28.
Piggins, H.D., Samuels, R.E., Coogan, A.N., Cutler, D.J., 2001b.
Distribution of substance P and neurokinin-1 receptor
immunoreactivity in the suprachiasmatic nuclei and
56
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
intergeniculate leaflet of hamster, mouse, and rat. J. Comp.
Neurol. 438, 50–65.
Porter, J.T., McCarthy, K.D., 1997. Astrocytic neurotransmitter
receptors in situ and in vivo. Prog. Neurobiol. 51, 439–455.
Prosser, R.A., 1998. Neuropeptide Y blocks serotonergic phase
shifts of the suprachiasmatic circadian clock in vitro. Brain Res.
808, 31–41.
Prosser, R.A., Dean, R.R., Edgar, D.M., Heller, H.C., Miller, J.D., 1993.
Serotonin and the mammalian circadian system: I. In vitro
phase shifts by serotonergic agonists and antagonists. J. Biol.
Rhythms 8, 1–16.
Prosser, R.A., Macdonald, E.S., Heller, H.C., 1994. c-fos mRNA in
the suprachiasmatic nuclei in vitro shows a circadian rhythm
and responds to a serotonergic agonist. Mol. Brain Res. 25,
151–156.
Prosser, R.A., Rutishauser, U., Ungers, G., Fedorkova, L., Glass, J.D.,
2003. Intrinsic role of polysialylated neural cell adhesion
molecule in photic phase resetting of the mammalian
circadian clock. J. Neurosci. 23, 652–658.
Provencio, I., Foster, R.G., 1995. Circadian rhythms in mice can be
regulated by photoreceptors with cone-like characteristics.
Brain Res. 694, 183–190.
Provencio, I., Wong, S., Lederman, A.B., Argamaso, S.M., Foster,
R.G., 1994. Visual and circadian responses to light in aged
retinally degenerate mice. Vision Res. 34, 1799–1806.
Provencio, I., Jiang, G., De Grip, W.J., Hayes, W.P., Rollag, M.D.,
1998a. Melanopsin: an opsin in melanophores, brain, and eye.
Proc. Natl. Acad. Sci. U. S. A. 95, 340–345.
Provencio, I., Cooper, H.M., Foster, R.G., 1998b. Retinal
projections in mice with inherited retinal degeneration:
implications for circadian photoentrainment. J. Comp.
Neurol. 395, 417–439.
Provencio, I., Rodriguez, I.R., Jiang, G., Hayes, W.P., Moreira, E.F.,
Rollag, M.D., 2000. A novel human opsin in the inner retina.
J. Neurosci. 20, 600–605.
Provencio, I., Rollag, M.D., Castrucci, A.M., 2002. Photoreceptive net
in the mammalian retina. Nature 415, 493.
Pu, M., 1999. Dendritic morphology of cat retinal ganglion cells
projecting to suprachiasmatic nucleus. J. Comp. Neurol. 414,
267–274.
Pu, M., 2000. Physiological response properties of cat retinal
ganglion cells projecting to suprachiasmatic nucleus. J. Biol.
Rhythms 15, 31–36.
Pu, M.L., Pickard, G.E., 1996. Ventral lateral geniculate nucleus
afferents to the suprachiasmatic nucleus in the cat. Brain Res.
725, 247–251.
Qiu, X., Kumbalasiri, T., Carlson, S.M., Wong, K.Y., Krishna, V.,
Provencio, I., Berson, D.M., 2005. Induction of photosensitivity
by heterologous expression of melanopsin. Nature 433,
745–749.
Quintero, J.E., Kuhlman, S.J., McMahon, D.G., 2000. Mouse
Period1-driven GFP gene expression cycles in SCN slices and in
individual SCN cells. Soc. Res. Biol. Rhythms 7, 87.
Quintero, J.E., Kuhlman, S.J., McMahon, D.G., 2003. The biological
clock nucleus: a multiphasic oscillator network regulated by
light. J. Neurosci. 23, 8070–8076.
Ralph, M.R., Menaker, M., 1987. Bicuculline blocks circadian phase
delays but not advances. Neurochem. Int. 11, 55–62.
Ralph, M.R., Menaker, M., 1989. GABA regulation of circadian
responses to light: I. Involvement of GABAA-benzodiazepine
and GABAB receptors. J. Neurosci. 9, 2858–2865.
Ralph, M.R., Mrosovsky, N., 1992. Behavioral inhibition of circadian
responses to light. J. Biol. Rhythms 7, 353–359.
Ralph, M.R., Foster, R.G., Davis, F.C., Menaker, M., 1990.
Transplanted suprachiasmatic nucleus determines circadian
period. Science 247, 975–978.
Rea, M.A., 1989. Light increases Fos-related protein
immunoreactivity in the rat suprachiasmatic nuclei. Brain Res.
Bull. 23, 577–581.
Rea, M.A., 1992. Different populations of cells in the
suprachiasmatic nuclei express c-fos in association with
light-induced phase delays and advances of the free-running
activity rhythm in hamsters. Brain Res. 579, 107–112.
Rea, M.A., Pickard, G.E., 2000a. Serotonergic modulation of photic
entrainment in the Syrian hamster. Biol. Rhythm Res. 31,
284–314.
Rea, M.A., Pickard, G.E., 2000b. A 5-HT1B receptor agonist inhibits
light-induced suppression of pineal melatonin production.
Brain Res. 858, 424–428.
Rea, M.A., Ferriera, S., Randolph, W., Glass, J.D., 1993a. Daily profile
of the extracellular concentration of glutamate in the
suprachiasmatic region of the Siberian hamster. Proc. Soc. Exp.
Biol. Med. 204, 104–109.
Rea, M.A., Buckley, B., Lutton, L.M., 1993b. Local administration of
EAA antagonists blocks light-induced phase shifts and c-fos
expression in hamster SCN. Am. J. Physiol. 265, R1191–R1198.
Rea, M.A., Barrera, J., Glass, J.D., Gannon, R.L., 1995. Serotonergic
potentiation of photic phase shifts of the circadian activity
rhythm. NeuroReport 6, 1289–1292.
Reddy, A.B., Field, M.D., Maywood, E.S., Hastings, M.H., 2002.
Differential resynchronisation of circadian clock gene
expression within the suprachiasmatic nuclei of mice
subjected to experimental jet lag. J. Neurosci. 22,
7326–7330.
Redlin, U., Mrosovsky, N., 1999a. Masking of locomotor activity in
hamsters. J. Comp. Physiol., A 184, 429–437.
Redlin, U., Mrosovsky, N., 1999b. Masking by light in hamsters with
SCN lesions. J. Comp. Physiol., A 184, 439–448.
Redlin, U., Vrang, N., Mrosovsky, N., 1999. Enhanced masking
response to light in hamsters with IGL lesions. J. Comp.
Physiol., A 184, 449–456.
Redlin, U., Cooper, H.M., Mrosovsky, N., 2003. Increased masking
response to light after ablation of the visual cortex in mice.
Brain Res. 965, 1–8.
Reebs, S.G., Lavery, R.J., Mrosovsky, N., 1989. Running activity
mediates the phase-advancing effects of dark pulses on
hamster circadian rhythms. J. Comp. Physiol., A 165,
811–818.
Reed, H.E., Cutler, D.J., Brown, T.M., Brown, J., Coen, C.W., Piggins,
H.D., 2002. Effects of vasoactive intestinal polypeptide on
neurones of the rat suprachiasmatic nuclei in vitro.
J. Neuroendocrinol. 14, 639–646.
Reuss, S., 1991. Photoperiod effects on bombesin- and
cholecystokinin-like immunoreactivity in the suprachiasmatic
nuclei of the Djungarian hamster (Phodopus sungorus).
Neurosci. Lett. 128, 13–16.
Reuss, S., Bürger, K., 1994. Substance P-like immunoreactivity in
the hypothalamic suprachiasmatic nucleus of Phodopus
sungorus—Relation to daytime, photoperiod, sex and age.
Brain Res. 638, 189–195.
Reuss, S., Decker, K., Rosseler, L., Layes, E., Schollmayer, A.,
Spessert, R., 1995. Nitric oxide synthase in the hypothalamic
suprachiasmatic nucleus of rat: evidence from histochemistry,
immunohistochemistry and Western blot; and colocalization
with VIP. Brain Res. 695, 257–262.
Rhoades, R.W., Hsu, L., Parfett, G., 1979. An electronmicroscopic
analysis of the optic nerve in the golden hamster. J. Comp.
Neurol. 186, 491–504.
Roberts, W.M., 1994. Localization of calcium signals by a mobile
calcium buffer in frog saccular hair cells. J. Neurosci. 14,
3246–3262.
Rosenwasser, A.M., Dwyer, S.M., 2002. Phase shifting the hamster
circadian clock by 15-minute dark pulses. J. Biol. Rhythms 17,
238–247.
Ruby, N.F., Saran, A., Kang, T., Franken, P., Heller, H.C., 1996.
Siberian hamsters free run or become arrhythmic after a phase
delay of the photocycle. Am. J. Physiol. 271, R881–R890.
Ruby, N.F., Kang, T., Heller, H.C., 1997. Melatonin attenuates photic
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
disruption of circadian rhythms in Siberian hamsters. Am. J.
Physiol. 273, R1540–R1549.
Ruby, N.F., Joshi, N., Heller, H.C., 1998. Phase shift magnitude and
direction determine whether Siberian hamsters reentrain to
the photocycle. J. Biol. Rhythms 13, 506–517.
Ruby, N.F., Dubocovich, M.L., Heller, H.C., 2000. Siberian hamsters
that fail to reentrain to the photocycle have suppressed
melatonin levels. Am. J. Physiol.: Regul., Integr. Comp. Physiol.
278, R757–R762.
Ruby, N.F., Brennan, T.J., Xie, X., Cao, V., Franken, P., Heller, H.C.,
2002a. Role of melanopsin in circadian responses to light.
Science 298, 2211–2213.
Ruby, N.F., Joshi, N., Heller, H.C., 2002b. Constant darkness restores
entrainment to phase-delayed Siberian hamsters. Am. J.
Physiol.: Regul., Integr. Comp. Physiol. 283, R1314–R1320.
Ruby, N.F., Barakat, M.T., Heller, H.C., 2004. Phenotypic differences
in reentrainment behavior and sensitivity to nighttime light
pulses in Siberian hamsters. J. Biol. Rhythms 19, 530–541.
Rusak, B., Morin, L.P., 1976. Testicular responses to photoperiod
are blocked by lesions of the suprachiasmatic nuclei in golden
hamsters. Biol. Reprod. 15, 366–374.
Rutishauser, U., Landmesser, L., 1996. Polysialic acid in the
vertebrate nervous system: a promoter of plasticity in cell–cell
interactions. Trends Neurosci. 19, 422–427.
Saeb-Parsy, K., Dyball, R.E., 2003. Defined cell groups in the rat
suprachiasmatic nucleus have different day/night rhythms of
single-unit activity in vivo. J. Biol. Rhythms 18, 26–42.
Saper, C.B., Chou, T.C., Scammell, T.E., 2001. The sleep switch:
hypothalamic control of sleep and wakefulness. Trends
Neurosci. 24, 726–731.
Sato, T., Kawamura, H., 1984. Circadian rhythms in multiple unit
activity inside and outside the suprachiasmatic nucleus in the
diurnal chipmunk (Eutamias sibiricus). Neurosci. Res. 1, 45–52.
Schaap, J., Bos, N.P., de Jeu, M.T., Geurtsen, A.M., Meijer, J.H.,
Pennartz, C.M., 1999. Neurons of the rat suprachiasmatic
nucleus show a circadian rhythm in membrane properties that
is lost during prolonged whole-cell recording. Brain Res. 815,
154–166.
Schaap, J., Albus, H., Eilers, P.H., Detari, L., Meijer, J.H., 2001. Phase
differences in electrical discharge rhythms between neuronal
populations of the left and right suprachiasmatic nuclei.
Neuroscience 108, 359–363.
Schaap, J., Albus, H., VanderLeest, H.T., Eilers, P.H., Detari, L.,
Meijer, J.H., 2003. Heterogeneity of rhythmic suprachiasmatic
nucleus neurons: implications for circadian waveform and
photoperiodic encoding. Proc. Natl. Acad. Sci. U. S. A. 100,
15994–15999.
Schwartz, W.J., Takeuchi, J., Shannon, W., Davis, E.M., Aronin, N.,
1994. Temporal regulation of light-induced Fos and Fos-like
protein expression in the ventrolateral subdivision of the rat
suprachiasmatic nucleus. Neuroscience 58, 573–583.
Schwartz, W.J., Peters, R.V., Aronin, N., Bennett, M.R., 1996.
Unexpected c-fos gene expression in the suprachiasmatic
nucleus of mice entrained to a skeleton photoperiod. J. Biol.
Rhythms 11, 35–44.
Schwartz, W.J., Carpino Jr., A., De la Iglesia, H.O., Baler, R., Klein,
D.C., Nakabeppu, Y., Aronin, N., 2000. Differential regulation
of fos family genes in the ventrolateral and dorsomedial
subdivisions of the rat suprachiasmatic nucleus.
Neuroscience 98, 535–547.
Schwartz, M.D., Nunez, A.A., Smale, L., 2004. Differences in
the suprachiasmatic nucleus and lower subparaventricular
zone of diurnal and nocturnal rodents. Neuroscience 127,
13–23.
Semo, M., Munoz, L.M., Foster, R.G., Jeffery, G., 2005. Melanopsin
(Opn4) positive cells in the cat retina are randomly distributed
across the ganglion cell layer. Vis. Neurosci. 22, 111–116.
Shearman, L.P., Zylka, M.J., Weaver, D.R., Kolakowski Jr., L.F.,
Reppert, S.M., 1997. Two period homologs: circadian
57
expression and photic regulation in the suprachiasmatic
nuclei. Neuron 19, 1261–1269.
Shearman, L.P., Sriram, S., Weaver, D.R., Maywood, E.S., Chaves,
I., Zheng, B., Kume, K., Lee, C.C., van der Horst, G.T., Hastings,
M.H., Reppert, S.M., 2000. Interacting molecular loops in the
mammalian circadian clock. Science 288, 1013–1019.
Shen, H., Semba, K., 1994. A direct retinal projection to the dorsal
raphe nucleus in the rat. Brain Res. 635, 159–168.
Shen, H., Watanabe, M., Tomasiewicz, H., Rutishauser, U.,
Magnuson, T., Glass, J.D., 1997. Role of neural cell adhesion
molecule and polysialic acid in mouse circadian clock function.
J. Neurosci. 17, 5221–5229.
Shen, H., Glass, J.D., Seki, T., Watanabe, M., 1999. Ultrastructural
analysis of polysialylated neural cell adhesion molecule in the
suprachiasmatic nuclei of the adult mouse. Anat. Rec. 256,
448–457.
Shen, S., Spratt, C., Sheward, W.J., Kallo, I., West, K., Morrison, C.F.,
Coen, C.W., Marston, H.M., Harmar, A.J., 2000. Overexpression
of the human VPAC2 receptor in the suprachiasmatic nucleus
alters the circadian phenotype of mice. Proc. Natl. Acad. Sci.
U. S. A. 97, 11575–11580.
Shen, H., Watanabe, M., Tomasiewicz, H., Glass, J.D., 2001. Genetic
deletions of NCAM and PSA impair circadian function in the
mouse. Physiol. Behav. 73, 185–193.
Shi, H., Bartness, T.J., 2001. Neurochemical phenotype of
sympathetic nervous system outflow from brain to white fat.
Brain Res. Bull. 54, 375–385.
Shibata, S., Shiratsuchi, A., Liou, S.Y., Ueki, S., 1984. The role of
calcium ions in circadian rhythm of suprachiasmatic nucleus
neuron activity in rat hypothalamic slices. Neurosci. Lett. 52,
181–184.
Shibata, S., Liou, S.Y., Ueki, S., 1986. Influence of excitatory amino
acid receptor antagonists and of baclofen on synaptic
transmission in the optic nerve to the suprachiasmatic nucleus
in slices of rat hypothalamus. Neuropharmacology 25, 403–409.
Shibata, S., Tsuneyoshi, A., Hamada, T., Tominaga, K., Watanabe,
S., 1992. Effect of substance P on circadian rhythms of firing
activity and the 2-deoxyglucose uptake in the rat
suprachiasmatic nucleus in vitro. Brain Res. 597, 257–263.
Shibata, S., Watanabe, A., Hamada, T., Watanabe, S., 1994.
Protein-synthesis inhibitor blocks
(R,S)-α-amino-3-hydroxy-5-methylisoxazole-4-propionic acid
(AMPA)- or substance P-induced phase shift of the circadian
rhythm of neuronal activity in the rat suprachiasmatic nucleus
in vitro. Neurosci. Lett. 168, 159–162.
Shigeyoshi, Y., Taguchi, K., Yamamoto, S., Takekida, S., Yan, L.,
Tei, H., Moriya, T., Shibata, S., Loros, J.J., Dunlap, J.C., Okamura,
H., 1997. Light-induced resetting of a mammalian circadian
clock is associated with rapid induction of the mPer1
transcript. Cell 91, 1043–1053.
Shimazoe, T., Nakamura, S., Kobayashi, K., Watanabe, S.,
Miyasaka, K., Kono, A., Funakoshi, A., 2004. Role of 5-HT1B
receptors in entrainment disorder of Otsuka Long Evans
Tokushima fatty (OLETF) rats. Neuroscience 123, 201–205.
Shirakawa, T., Moore, R.Y., 1994. Responses of rat suprachiasmatic
nucleus neurons to substance P and glutamate in vitro. Brain
Res. 642, 213–220.
Shirakawa, T., Honma, S., Katsuno, Y., Oguchi, H., Honma, K.,
2000. Synchronization of circadian firing rhythms in
cultured rat suprachiasmatic neurons. Eur. J. Neurosci. 12,
2833–2838.
Shiromani, P.J., Overstreet, D., Levy, D., Goodrich, C.A., Campbell,
S.S., Gillin, J.C., 1988. Increased REM sleep in rats selectively
bred for cholinergic hyperactivity. Neuropsychopharmacology
1, 127–133.
Shivers, K., Muscat, L., 2004. Relative distribution of crossed and
uncrossed retinohypothalamic tract (RHT) projections to the
suprachiasmatic nucleus (SCN) in rat and mouse. Soc.
Neurosci. 765.14.
58
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
Silver, J., Brand, S., 1979. A route for direct retinal input to the
preoptic hypothalamus: dendritic projections into the optic
chiasm. Am. J Anat. 155, 391–401.
Silver, R., Lehman, M.N., Gibson, M., Gladstone, W.R., Bittman, E.L.,
1990. Dispersed cell suspensions of fetal SCN restore circadian
rhythmicity in SCN-lesioned adult hamsters. Brain Res. 525,
45–58.
Silver, R., Romero, M.T., Besmer, H.R., Leak, R.K., Nunez, J.M.,
LeSauter, J., 1996a. Calbindin-D28K cells in the hamster SCN
express light-induced Fos. NeuroReport 7, 1224–1228.
Silver, R., LeSauter, J., Tresco, P.A., Lehman, M.N., 1996b. A
diffusible coupling signal from the transplanted
suprachiasmatic nucleus controlling circadian locomotor
rhythms. Nature 382, 810–813.
Silver, R., Sookhoo, A.I., LeSauter, J., Stevens, P., Jansen, H.T.,
Lehman, M.N., 1999. Multiple regulatory elements result in
regional specificity in circadian rhythms of neuropeptide
expression in mouse SCN. NeuroReport 10, 3165–3174.
Smale, L., Boverhof, J., 1999. The suprachiasmatic nucleus and
intergeniculate leaflet of Arvicanthis niloticus, a diurnal murid
rodent from East Africa. J. Comp. Neurol. 403, 190–208.
Smale, L., Michels, K.M., Moore, R.Y., Morin, L.P., 1990. Destruction
of the hamster serotonergic system by 5,7-DHT: effects on
circadian rhythm phase, entrainment and response to
triazolam. Brain Res. 515, 9–19.
Smale, L., Blanchard, J.H., Moore, R.Y., Morin, L.P., 1991.
Immunocytochemical characterization of the suprachiasmatic
nucleus and the intergeniculate leaflet in the diurnal ground
squirrel, Spermophilus lateralis. Brain Res. 563, 77–86.
Smale, L., Castleberry, C., Nunez, A.A., 2001. Fos rhythms in the
hypothalamus of Rattus and Arvicanthis that exhibit nocturnal
and diurnal patterns of rhythmicity. Brain Res. 899, 101–105.
Smale, L., Lee, T., Nunez, A.A., 2003. Mammalian diurnality: some
facts and gaps. J. Biol. Rhythms 18, 356–366.
Smart, C.M., Biello, S.M., 2001. WAY-100635, a specific 5-HT1A
antagonist, can increase the responsiveness of the
mammalian circadian pacemaker to photic stimuli. Neurosci.
Lett. 305, 33–36.
Smeraski, C.A., Sollars, P.J., Ogilvie, M.D., Enquist, L.W., Pickard,
G.E., 2004. Suprachiasmatic nucleus input to autonomic
circuits identified by retrograde transsynaptic transport of
pseudorabies virus from the eye. J. Comp. Neurol. 471, 298–313.
Smith, R.D., Inouye, S.-I.T., Turek, F.W., 1989. Central
administration of muscimol phase-shifts the mammalian
circadian clock. J. Comp. Physiol. 164, 805–814.
Smith, B.N., Sollars, P.J., Dudek, F.E., Pickard, G.E., 2001.
Serotonergic modulation of retinal input to the mouse
suprachiasmatic nucleus mediated by 5-HT1B and 5-HT7
receptors. J. Biol. Rhythms 16, 25–38.
Sollars, P.J., Pickard, G.E., 1993. Time course of fiber outgrowth
from fetal anterior hypothalamic heterografts. Brain Res. 614,
212–219.
Sollars, P.J., Kimble, D.P., Pickard, G.E., 1995. Restoration of
circadian behavior by anterior hypothalamic heterografts.
J. Neurosci. 15, 2109–2122.
Sollars, P.J., Ogilvie, M.D., Rea, M.A., Pickard, G.E., 2002. 5-HT1B
receptor knockout mice exhibit an enhanced response to
constant light. J. Biol. Rhythms 17, 428–437.
Sollars, P.J., Smeraski, C.A., Kaufman, J.D., Ogilvie, M.D., Provencio,
I., Pickard, G.E., 2003. Melanopsin and non-melanopsin
expressing retinal ganglion cells innervate the hypothalamic
suprachiasmatic nucleus. Vis. Neurosci. 20, 601–610.
Speh, J.C., Moore, R.Y., 1990. Synaptogenesis and
retinohypothalamic tract (RHT) development in the hamster
suprachiasmatic nucleus (SCN). Society for Neuroscience 16,
602 (Ref Type: Abstract).
Speh, J.C., Moore, R.Y., 1993. Retinohypothalamic tract
development in the hamster and rat. Dev. Brain Res. 76,
171–181.
Spoelstra, K., Oklejewicz, M., Daan, S., 2002. Restoration of
self-sustained circadian rhythmicity by the mutant clock allele
in mice in constant illumination. J. Biol. Rhythms 17, 520–525.
Sprouse, J., Reynolds, L., Braselton, J., Schmidt, A., 2004a.
Serotonin-induced phase advances of SCN neuronal firing in
vitro: a possible role for 5-HT5A receptors? Synapse 54,
111–118.
Sprouse, J., Reynolds, L., Li, X., Braselton, J., Schmidt, A., 2004b.
8-OH-DPAT as a 5-HT7 agonist: phase shifts of the circadian
biological clock through increases in cAMP production.
Neuropharmacology 46, 52–62.
Stamp, J.A., Piggins, H.D., Rusak, B., Semba, K., 1997. Distribution of
ionotropic glutamate receptor subunit immunoreactivity in
the suprachiasmatic nucleus and intergeniculate leaflet of the
hamster. Brain Res. 756, 215–224.
Starkey, S.J., 1996. Melatonin and 5-hydroxytryptamine
phase-advance the rat circadian clock by activation of nitric
oxide synthesis. Neurosci. Lett. 211, 199–202.
Stephan, F.K., Donaldson, J.A., Gellert, J., 1982.
Retinohypothalamic tract symmetry and phase shifts of
circadian rhythms in rats and hamsters. Physiol. Behav. 29,
1153–1159.
Stetson, M.H., Watson-Whitmyre, M., 1976. Nucleus
suprachiasmaticus: the biological clock in the hamster?
Science 191, 197–199.
Summer, T.L., Ferraro, J.S., McCormack, C.E., 1984. Phase-response
and Aschoff illuminance curves for locomotor activity rhythm
of the rat. Am. J. Physiol. 246, R299–R304.
Sumova, A., Illnerova, H., 1996. Melatonin instantaneously resets
intrinsic circadian rhythmicity in the rat suprachiasmatic
nucleus. Neurosci. Lett. 218, 181–184.
Sumova, A., Travnickova, Z., Mikkelsen, J.D., Illnerova, H., 1998.
Spontaneous rhythm in c-Fos immunoreactivity in the
dorsomedial part of the rat suprachiasmatic nucleus. Brain
Res. 801, 254–258.
Sumova, A., Travnickova, Z., Illnerova, H., 2000. Spontaneous c-Fos
rhythm in the rat suprachiasmatic nucleus: location and effect
of photoperiod. Am. J. Physiol.: Regul., Integr. Comp. Physiol.
279, R2262–R2269.
Sumova, A., Bendova, Z., Sladek, M., Kovacikova, Z., Illnerova, H.,
2004. Seasonal molecular timekeeping within the rat circadian
clock. Physiol. Res. 53 (Suppl. 1), S167–S176.
Swann, J.M., Turek, F.W., 1985. Multiple circadian oscillators
regulate the timing of behavioral and endocrine rhythms in
female golden hamsters. Science 228, 898–900.
Takahashi, J.S., DeCoursey, P.J., Bauman, L., Menaker, M., 1984.
Spectral sensitivity of a novel photoreceptive system
mediating entrainment of mammalian circadian rhythms.
Nature 308, 186–188.
Takatsuji, K., Miguel-Hidalgo, J.-J., Tohyama, M., 1991. Substance
P-immunoreactive innervation from the retina to the
suprachiasmatic nucleus in the rat. Brain Res. 568, 223–229.
Takatsuji, K., Senba, E., Mantyh, P.W., Tohyama, M., 1995. A
relationship between substance P receptor and retinal fibers in
the rat suprachiasmatic nucleus. Brain Res. 698, 53–61.
Takeuchi, Y., Takahashima, M., Katoh, Y., Nishikawa, T.,
Takahashi, K., 1991. N-Methyl-D-aspartate, quisqualate and
kainate receptors are all involved in transmission of photic
stimulation in the suprachiasmatic nucleus in rats. Brain Res.
563, 127–131.
Tang, I.H., Murakami, D.M., Fuller, C.A., 2002. Unilateral optic
nerve transection alters light response of suprachiasmatic
nucleus and intergeniculate leaflet. Am. J. Physiol. 282,
R569–R577.
Teclemariam-Mesbah, R., ter Horst, G.J., Postema, F., Wortel, J.,
Buijs, R.M., 1999. Anatomical demonstration of the
suprachiasmatic nucleus-pineal pathway. J. Comp. Neurol.
406, 171–182.
Teshima, K., Kim, S.H., Allen, C.N., 2003. Characterization of an
BR A I N R ES E A RC H R EV IE W S 5 1 (2 0 0 6) 1 –60
apamin-sensitive potassium current in suprachiasmatic
nucleus neurons. Neuroscience 120, 65–73.
Thomson, A.M., 1984. Slow, regular discharge in suprachiasmatic
neurons is calcium dependent, in slices of rat brain.
Neuroscience 13, 761–767.
Thomson, A.M., West, D.C., 1990. Factors affecting slow regular
firing in the suprachiasmatic nucleus in vitro. J. Biol. Rhythms
5, 59–75.
Tiao, Y.-C., Blakemore, C., 1976. Regional specialization in the
golden hamster's retina. J. Comp. Neurol. 168, 439–458.
Tieman, S.B., Tieman, D.G., 1996. N-Acetylaspartylglutamate
immunoreactivity in human retina. Vision Res. 36, 941–947.
Tieman, S.B., Moffett, J.R., Irtenkauf, S.M., 1991. Effect of eye
removal on N-acetylaspartylglutamate immunoreactivity in
retinal targets of the cat. Brain Res. 562, 318–322.
Tierno, A., Fiore, P., Gannon, R.L., 2002. Delta opioid inhibition of
light-induced phase advances in hamster circadian activity
rhythms. Brain Res. 937, 66–73.
Tischler, R.C., Morin, L.P., 2003. Reciprocal serotonergic
connections between the hamster median and dorsal raphe
nuclei. Brain Res. 981, 126–132.
Tokunaga, A., Ono, K., Kondo, S., Tanaka, H., Kurose, K., Nagai, H.,
1992. Retinal projections in the house musk shrew, Suncus
murinus, as determined by anterograde transport of WGA-HRP.
Brain Behav. Evol. 40, 321–329.
Tominaga, K., Shibata, S., Ueki, S., Watanabe, S., 1992. Effects of
5-HT1A receptor agonists on the circadian rhythm of
wheel-running activity in hamsters. Eur. J. Pharmacol. 214,
79–84.
Tournier, B.B., Menet, J.S., Dardente, H., Poirel, V.J., Malan, A.,
Masson-Pevet, M., Pevet, P., Vuillez, P., 2003. Photoperiod
differentially regulates clock genes' expression in the
suprachiasmatic nucleus of Syrian hamster. Neuroscience 118,
317–322.
Tousson, E., Meissl, H., 2004. Suprachiasmatic nuclei grafts restore
the circadian rhythm in the paraventricular nucleus of the
hypothalamus. J. Neurosci. 24, 2983–2988.
Trachsel, L., Heller, H.C., Miller, J.D., 1995. Nicotine
phase-advances the circadian neuronal activity rhythm in rat
suprachiasmatic nuclei explants. Neuroscience 65, 797–803.
Treep, J.A., Abe, H., Rusak, B., Goguen, D.M., 1995. Two distinct
retinal projections to the hamster suprachiasmatic nucleus.
J. Biol. Rhythms 10, 299–307.
Ueyama, T., Krout, K.E., Van Nguyen, X., Karpitskiy, V., Kollert, A.,
Mettenleiter, T.C., Loewy, A.D., 1999. Suprachiasmatic nucleus:
a central autonomic clock. Nat. Neurosci. 2, 1051–1053.
Underwood, H., Menaker, M., 1970. Extraretinal light perception:
entrainment of the biological clock controlling lizard
locomotor activity. Science 170, 190–193.
Urbanski, H.F., Doan, A., Pierce, M., 1991. Immunocytochemical
investigation of luteinizing hormone- releasing hormone
neurons in Syrian hamsters maintained under long or short
days. Biol. Reprod. 44, 687–692.
van den Pol, A.N., 1980. The hypothalamic suprachiasmatic
nucleus of rat: intrinsic anatomy. J. Comp. Neurol. 191,
661–702.
van den Pol, A.N., 1991. The suprachiasmatic nucleus:
morphological and cytochemical substrates for cellular
interaction. In: Klein, D.C., Moore, R.Y., Reppert, S.M. (Eds.),
Suprachiasmatic Nucleus: The Mind's Clock. Oxford Univ.
Press, New York, pp. 17–50.
van den Pol, A.N., Gorcs, T., 1986. Synaptic relationships between
neurons containing vasopressin, gastrin-releasing peptide,
vasoactive intestinal polypeptide, and glutamate
decarboxylase immunoreactivity in the suprachiasmatic
nucleus: dual ultrastructural immunocytochemistry with
gold-substituted silver peroxidase. J. Comp. Neurol. 252,
507–521.
van den Pol, A.N., Tsujimoto, K.L., 1985. Neurotransmitters of
59
the hypothalamic suprachiasmatic nucleus:
immunocytochemical analysis of 25 neuronal antigens.
Neuroscience 15, 1049–1086.
van den Pol, A.N., Cao, V., Heller, H.G., 1998. Circadian system of
mice integrates brief light stimuli. Am. J. Physiol.: Regul., Integr.
Comp. Physiol. 275, R654–R657.
Van der Beek, E.M., Horvath, T.L., Wiegant, V.M., Buijs, R.M.,
1997. Evidence for a direct neuronal pathway from the
suprachiasmatic nucleus to the gonadotropin-releasing
hormone system: combined tracing and light and electron
microscopic immunocytochemical studies. J. Comp. Neurol.
384, 569–579.
Van der Gucht, E., Hof, P.R., Arckens, L., 2003. Neurochemical
organization, architectonic subdivision and three-dimensional
reconstruction of cat ventral lateral geniculate nucleus and
monkey pregeniculate nucleus. Soc. Neurosci. 33, 68.7.
Van der Horst, G.T.J., Muijtjens, M., Kobayashi, K., Takano, R.,
Kanno, S., Takao, M., De Wit, J., Verkerk, A., Eker, A.P.M., Van
Leenen, D., Buijs, R., Bootsma, D., Hoeijmakers, J.H.J., Yasui, A.,
1999. Mammalian Cry1 and Cry2 are essential for maintenance
of circadian rhythms. Nature 398, 627–630.
Van Reeth, O., Turek, F.W., 1989. Stimulated activity mediates
phase shifts in the hamster circadian clock induced by dark
pulses or benzodiazepines. Nature 339, 49–51.
Van Reeth, O., Hinch, D., Tecco, J.M., Turek, F.W., 1991. The effects
of short periods of immobilization on the hamster circadian
clock. Brain Res. 545, 208–214.
Vansteensel, M.J., Yamazaki, S., Albus, H., Deboer, T., Block, G.D.,
Meijer, J.H., 2003a. Dissociation between circadian Per1 and
neuronal and behavioral rhythms following a shifted
environmental cycle. Curr. Biol. 13, 1538–1542.
Vansteensel, M.J., Deboer, T., Dahan, A., Meijer, J.H., 2003b. Onset
and offset following GABA-ergic and opioid receptor activation.
J. Biol. Rhythms 18, 297–306.
Varcoe, T.J., Kennaway, D.J., Voultsios, A., 2003. Activation of
5-HT2C receptors acutely induces Per gene expression in the
rat suprachiasmatic nucleus at night. Brain Res. Mol. Brain Res.
119, 192–200.
Vertes, R.P., 1991. A PHA-L analysis of ascending projections of
the dorsal raphe nucleus in the rat. J. Comp. Neurol. 313,
643–668.
Vidal, L., Morin, L.P., 2005. Hypothalamic and zona incerta neurons
expressing hypocretin, but not melanin concentrating
hormone, project to the hamster intergeniculate leaflet.
Neuroscience 134, 1081–1090.
Vincent, S.R., Kimura, H., 1992. Histochemical mapping of nitric
oxide synthase in the rat brain. Neuroscience 46, 755–784.
Vindlacheruvu, R.R., Ebling, F.J.P., Maywood, E.S., Hastings, M.H.,
1992. Blockade of glutamatergic neurotransmission in the
suprachiasmatic nucleus prevents cellular and behavioural
responses of the circadian system to light. Eur. J. Neurosci. 4,
673–679.
Vitaterna, M.H., King, D.P., Chang, A.-M., Kornhauser, J.M., Lowrey,
P.L, McDonald, J.D., Dove, W.F., Pinto, L.H., Turek, F.W.,
Takahashi, J.S., 1994. Mutagenesis and mapping of a mouse
gene, Clock, essential for circadian behavior. Science 264,
719–725.
Vrang, N., Larsen, P.J., Moller, M., Mikkelsen, J.D., 1995a.
Topographical organization of the rat
suprachiasmatic-paraventricular projection. J. Comp. Neurol.
353, 585–603.
Vrang, N., Larsen, P.J., Mikkelsen, J.D., 1995b. Direct projection
from the suprachiasmatic nucleus to hypophysiotrophic
corticotropin-releasing factor immunoreactive cells in the
paraventricular nucleus of the hypothalamus demonstrated by
means of Phaseolus vulgaris-leucoagglutinin tract tracing. Brain
Res. 684, 61–69.
Vrang, N., Mrosovsky, N., Mikkelsen, J.D., 2003. Afferent
projections to the hamster intergeniculate leaflet demonstrated
60
BR A I N R ES E A RC H R EV IE W S 5 1 ( 2 0 0 6) 1 –60
by retrograde and anterograde tracing. Brain Res. Bull. 59,
267–288.
Wagner, S., Castel, M., Gainer, H., Yarom, Y., 1997. GABA in the
mammalian suprachiasmatic nucleus and its role in diurnal
rhythmicity. Nature 387, 598–603.
Wagner, S., Sagiv, N., Yarom, Y., 2001. GABA-induced current and
circadian regulation of chloride in neurones of the rat
suprachiasmatic nucleus. J. Physiol. 537, 853–869.
Wang, H., Morris, J.F., 1996. Presence of neuronal nitric oxide
synthase in the suprachiasmatic nuclei of mouse and rat.
Neuroscience 74, 1059–1068.
Warren, E.J., Allen, C.N., Brown, R.L., Robinson, D.W., 2003.
Intrinsic light responses of retinal ganglion cells projecting to
the circadian system. Eur. J. Neurosci. 17, 1727–1735.
Watanabe, A., Ono, M., Shibata, S., Watanabe, S., 1995. Effect of a
nitric oxide synthase inhibitor, N-nitro-L-arginine methylester,
on light-induced phase delay of circadian rhythm of
wheel-running activity in golden hamsters. Neurosci. Lett. 192,
25–28.
Watanabe, K., Vanecek, J., Yamaoka, S., 2000. In vitro entrainment
of the circadian rhythm of vasopressin-releasing cells in
suprachiasmatic nucleus by vasoactive intestinal polypeptide.
Brain Res. 877, 361–366.
Watts, A.G., Swanson, L.W., 1987. Efferent projections of the
suprachiasmatic nucleus: II. Studies using retrograde transport
of fluorescent dyes and simultaneous peptide
immunohistochemistry in the rat. J. Comp. Neurol. 258,
230–252.
Watts, A.G., Swanson, L.W., Sanchez-Watts, G., 1987. Efferent
projections of the suprachiasmatic nucleus: I. Studies using
anterograde transport of Phaseolus vulgaris leucoagglutinin in
the rat. J. Comp. Neurol. 258, 204–229.
Weber, E.T., Rea, M.A., 1997. Neruopeptide Y blocks light-induced
phase advances but not delays of the circadian activity rhythm
in hamsters. Neurosci. Lett. 231, 159–162.
Weber, E.T., Gannon, R.L., Michel, A.M., Gillette, M.U., Rea, M.A.,
1995. Nitric oxide synthase inhibitor blocks light-induced
phase shifts of the circadian activity rhythm, but not c-fos
expression in the suprachiasmatic nucleus of the Syrian
hamster. Brain Res. 692, 137–142.
Welsh, D.K., Logothetis, D.E., Meister, M., Reppert, S.M., 1995.
Individual neurons dissociated from rat suprachiasmatic
nucleus express independently phased circadian firing
rhythms. Neuron 14, 697–706.
Wheal, H.V., Thomson, A.M., 1984. The electrical properties of
neurones of the rat suprachiasmatic nucleus recorded
intracellularly in vitro. Neuroscience 13, 97–104.
Wickland, C., Turek, F.W., 1994. Lesions of the thalamic
intergeniculate leaflet block activity-induced phase shifts in
the circadian activity rhythm of the golden hamster. Brain Res.
660, 293–300.
Wollnik, F., Brysch, W., Uhlmann, E., Gillardon, F., Bravo, R.,
Zimmermann, M., Schlingensiepen, K.H., Herdegen, T., 1995.
Block of c-Fos and JunB expression by antisense
oligonucleotides inhibits light-induced phase shifts of the
mammalian circadian clock. Eur. J. Neurosci. 7, 388–393.
Yagita, K., Tamanini, F., van der Horst, G.T., Okamura, H., 2001.
Molecular mechanisms of the biological clock in cultured
fibroblasts. Science 292, 278–281.
Yamaguchi, S., Isejima, H., Matsuo, T., Okura, R., Yagita, K.,
Kobayashi, M., Okamura, H., 2003. Synchronization of cellular
clocks in the suprachiasmatic nucleus. Science 302, 1408–1412.
Yamazaki, S., Goto, M., Menaker, M., 1999. No evidence for
extraocular photoreceptors in the circadian system of the
Syrian hamster. J. Biol. Rhythms 14, 197–201.
Yamazaki, S., Numano, R., Abe, M., Hida, A., Takahashi, R., Ueda,
M., Block, G.D., Sakaki, Y., Menaker, M., Tei, H., 2000. Resetting
central and peripheral circadian oscillators in transgenic rats.
Science 288, 682–685.
Yan, L., Okamura, H., 2002. Gradients in the circadian expression
of Per1 and Per2 genes in the rat suprachiasmatic nucleus. Eur.
J. Neurosci. 15, 1153–1162.
Yan, L., Silver, R., 2004. Resetting the brain clock: time course and
localization of mPER1 and mPER2 protein expression in
suprachiasmatic nuclei during phase shifts. Eur. J. Neurosci. 19,
1105–1109.
Yan, L., Takekida, S., Shigeyoshi, Y., Okamura, H., 1999. Per1 and
Per2 gene expression in the rat suprachiasmatic nucleus:
circadian profile and the compartment-specific response to
light. Neuroscience 94, 141–150.
Yannielli, P.C., Harrington, M.E., 2000. Neuropeptide Y applied in
vitro can block the phase shifts induced by light in vivo.
NeuroReport 11, 1587–1591.
Yannielli, P.C., Harrington, M.E., 2001. The neuropeptide YY5
receptor mediates the blockade of “photic-like”
NMDA-induced phase shifts in the golden hamster.
J. Neurosci. 21, 5367–5373.
Yannielli, P., Harrington, M.E., 2004. Let there be “more” light:
enhancement of light actions on the circadian system through
non-photic pathways. Prog. Neurobiol. 74, 59–76.
Yannielli, P.C., McKinley, B.J., Harrington, M.E., 2002. Is novel
wheel inhibition of per1 and per2 expression linked to phase
shift occurrence? Neuroscience 112, 677–685.
Yannielli, P.C., Brewer, J.M., Harrington, M.E., 2004. Blockade of the
NPY Y5 receptor potentiates circadian responses to light:
complementary in vivo and in vitro studies. Eur. J. Neurosci. 19,
891–897.
Yokota, S.I., Horikawa, K., Akiyama, M., Moriya, T., Ebihara, S.,
Komuro, G., Ohta, T., Shibata, S., 2000. Inhibitory action of
brotizolam on circadian and light-induced per1 and per2
expression in the hamster suprachiasmatic nucleus.
Br. J. Pharmacol. 131, 1739–1747.
Yoo, S.H., Yamazaki, S., Lowrey, P.L., Shimomura, K., Ko, C.H.,
Buhr, E.D., Siepka, S.M., Hong, H.K., Oh, W.J., Yoo, O.J., Menaker,
M., Takahashi, J.S., 2004. PERIOD2∷LUCIFERASE real-time
reporting of circadian dynamics reveals persistent circadian
oscillations in mouse peripheral tissues. Proc. Natl. Acad. Sci.
U. S. A. 101, 5339–5346.
Yoshimura, T., Ebihara, S., 1996. Spectral sensitivity of
photoreceptors mediating phase shifts of circadian rhythms in
retinally degenerate CBA/J (rd/rd) and normal CBA/N(+/+) mice.
J. Comp. Physiol., A 178, 797–802.
Yoshimura, T., Nishio, M., Goto, M., Ebihara, S., 1994. Differences in
circadian photosensitivity between retinally degenerate CBA/J
mice (rd/rd) and normal CBA/N mice (+/+). J. Biol. Rhythms 9,
51–60.
Zhang, Y., Zee, P.C., Kirby, J.D., Takahashi, J.S., Turek, F.W., 1993. A
cholinergic antagonist, mecamylamine, blocks light-induced
Fos immunoreactivity in specific regions of the hamster
suprachiasmatic nucleus. Brain Res. 615, 107–112.
Zhang, Y., Kornhauser, J.M., Zee, P.C., Mayo, K.E., Takahashi, J.S.,
Turek, F.W., 1996. Effects of aging on light-induced
phase-shifting of circadian behavioral rhythms, Fos expression
and CREB phosphorylation in the hamster suprachiasmatic
nucleus. Neuroscience 70, 951–961.
Zylka, M.J., Shearman, L.P., Weaver, D.R., Reppert, S.M., 1998.
Three period homologs in mammals: differential light
responses in the suprachiasmatic circadian clock and
oscillating transcripts outside of brain. Neuron 20,
1103–1110.