UNIVERSITY OF CINCINNATI
Date:______
I, ______, hereby submit this work as part of the requirements for the degree of: in:
It is entitled:
This work and its defense approved by:
Chair: ______
Rapid regulation of the hypothalamus-pituitary-adrenal axis by glutamate and glucocorticoids
A dissertation submitted to the
Division of Research and Advanced Studies of the University of Cincinnati
in partial fulfillment of the requirements for the degree of
Doctor of Philosophy (Ph.D.)
in the Graduate Program in Neuroscience of the College of Medicine
2008
by Nathan K. Evanson B.S. Biochemistry, Nutritional Science Brigham Young University, Provo, UT 2002
Committee Chair James P. Herman
Abstract
The hypothalamus-pituitary-adrenal (HPA) axis is the main neuroendocrine arm of the stress response, activation of which leads to the production of glucocorticoid hormones.
Glucocorticoids are steroid hormones that are secreted from the adrenal cortex, and have a variety of effects on the body, including modulation of the immune system, suppression of reproductive hormones, and maintenance of blood glucose levels, and maintenance of blood pressure. Activity of the HPA axis is known to be modulated by signaling through molecules like glutamate and glucocorticoids. Although it is clear that these signals play a role in HPA axis regulation, their roles in regulating the HPA axis at the paraventricular nucleus of the hypothalamus (PVN) are not yet entirely clear. In this work, we demonstrate HPA regulatory roles for the ionotropic glutamate receptor GluR5, the group I metabotropic glutamate receptors, and glucocorticoids at the PVN. GluR5 inhibits the HPA axis response to restraint stress when acting near the cell bodies in the PVN proper, and plays an excitatory role at axon terminals in the median eminence. Group I metabotropic glutamate receptor signaling has an inhibitory effect on the HPA axis response to restraint stress at the PVN. Glucocorticoids also exert an inhibitory influence on HPA axis activity at the level of the PVN. This negative regulation of the
HPA axis by glucocorticoids is likely nongenomic in nature and occurs on a very rapid time scale. This fast feedback is mediated by cannabinoid CB1 receptor signaling at the PVN. Thus, glutamate and glucocorticoids acting at the PVN can inhibit the HPA axis response to restraint stress in rats.
3 4 Acknowledgments
This work has been made possible by the contributions of many people. In particular, I
thank the members of the Herman lab who have contributed to this work: Yve Ulrich-Lai,
Helmer Figuieredo, Nancy Mueller, Michelle Ostrander, Ben Packard, Ryan Jankord, Matia
Solomon, Rong Zhang, Kenny Jones, Amanda Jones, Ingrid Thomas, Mark Dolgas, Miyuki
Tauchi, Amy Furay, Dennis Choi, Jon Flak, and Anne Christianson. An experiment using
animals takes a lot of people, and all of these have given invaluable assistance in helping me do
the experiments described in this work. I especially thank Nancy for teaching me to do
stereotaxic surgery, Yve for teaching me to do adrenalectomy surgeries (or, more accurately, for
doing almost all of them for me), and Yve, Amy, and Anne for their help in designing and
interpreting some of these studies. I thank Ben for making sure that there was as little dullness
as possible during the lab work, and for helping me to keep my (often very late) Thursday
evenings occupied. I thank Daniella Van Hooren for the work she has done to lay the foundation for the GluR5 studies that are described in this work.
Beyond the nitty-gritty details of the lab work, I thank Jim and Susan Herman for their support and help in this work. Without Susan, it is clear that the lab couldn’t function, so I thank her for all her work to keep us in business. Thank you also for helping me get my dissertation defense (and many other things) scheduled. Jim’s mentoring has likewise been invaluable to my growth as a graduate student and as a scientist. Thank you for all your advice, helpful comments
on my writing, and, of course, for providing the funding that has been necessary for this work to
be done. Thank you for helping me feel like I might become a scientist after all.
I owe a debt of gratitude to the programs that I have called home during this graduate
experience. The Physician Scientist Training Program has given me the chance to move beyond
5 my undergraduate ways (which can seem oh so sophomoric now, looking back). I thank Les
Myatt for believing in me enough to bring me into the program in spite of my “evil Canadian”
status, and for both Terri Berning and Laurie Mayleben for making the program run smoothly so
it was easy to stay here. I also thank the neuroscience graduate program for getting me interested
in the brain. My thanks go to Mike Lehman, who was the first program director I knew here, and
whose enthusiasm for neuroscience was so infectious. I thank Bruce Giffin for running Brain
and Behavior I in a way that helped me see that the brain is not only a fascinating organ, but also
tractable to scientific inquiry. I also thank Jim Herman (again) for being the second director of
the neuroscience program, but still finding time to help me with my research projects. Thanks
also go to Deb Cummins, who has showed me unfailing kindness and helped me out whenever I
needed her during graduate school.
I could not be where I am without my family. I thank my parents, Kent and Iris Evanson,
for encouraging me throughout my childhood to realize my potential. They taught me how to
work hard, and they taught me that I could go out and find answers to any questions that I had
(even before the time of the internet). It was by their encouragement and gentle (and occasionally otherwise) guidance and direction that I have been able to persist in my schooling for so many years. I also thank my children, Adam and Joshua for loving and supporting me through this time, and for understanding that sometimes Daddy has to go to GRI to check rats at odd and not always convenient times. I also thank (the presumptive) Caleb for not being born before I wrote this dissertation. I thank my beautiful wife Tricia Harris Evanson for sticking with me for so many years of starving studenthood, when everyone else we used to know has had a job for many years now. Thank you for believing in me and supporting me when I have been gone or distracted, or when I have been up all hours of the night writing (like tonight, for
6 example) or reading, or when I have been gone to conferences. You are the love of my life, and I
couldn’t have come this far without you. Finally, I thank God for providing me with a brain,
which is a very interesting thing to study as well as a very necessary instrument for doing the studying.
7 Table of Contents
Abstract...... 3 Acknowledgments...... 5 List of Figures...... 10 Abbreviations...... 11 1. Introduction...... 12 HPA Axis...... 13 Rapid regulation of the HPA axis...... 14 Glutamate...... 15 Ionotropic Glutamate receptors...... 16 Metabotropic glutamate receptors...... 17 Glutamate receptor signaling ...... 18 Glutamate and the HPA axis ...... 19 Glucocorticoids...... 20 Non-genomic effects of glucocorticoids in non-neural Tissue ...... 21 Signaling Mechanisms: receptors ...... 25 Intracellular signaling systems...... 32 Conclusion ...... 41 Figure Legends...... 42 2. Glutamate—GluR5 ...... 48 Introduction...... 48 Materials and Methods...... 50 Animals:...... 50 Microinjections and restraint stress challenge:...... 51 Radioimmunoassay:...... 52 Immunohistochemistry: ...... 52 In situ hybridization ...... 54 Imaging: ...... 54 Statistics:...... 54 Results...... 55 GluR5 expression...... 55 Injection site confirmation ...... 56 GluR5 Antagonist Infusion: PVN ...... 56 GluR5 Antagonist Infusion: Median Eminence...... 57 GluR5 expression in regions projecting to the PVN...... 58 Discussion...... 58 Figure Legends...... 63 3. Glucocorticoids and Cannabinoids ...... 75 Introduction...... 75 Materials and Methods...... 76 Animals:...... 76 Cannula Surgeries:...... 77 Drugs:...... 77 Restraint Challenge:...... 78 Radioimmunoassay:...... 78
8 Imaging ...... 78 Statistics:...... 79 Results:...... 79 Cannula Placement...... 79 Intra-paraventricular dexamethasone rapidly inhibits the HPA axis...... 79 Fast Feedback with peripheral administration of AM-251 ...... 80 Time dependence of AM-251 effects...... 81 Intra-PVN infusion of AM-251 blocks fast feedback...... 81 CB1 agonism is not sufficient to cause fast feedback ...... 82 Discussion...... 82 Figure Legends...... 89 4. Glutamate—group I metabotropic glutamate receptors...... 99 Introduction...... 99 Materials and Methods...... 101 Animals...... 101 Stereotactic surgery...... 101 Drugs...... 102 Injection and Restraint Challenge...... 102 Nissl stain for cannula placement ...... 103 Radioimmunoassay...... 103 Statistical analysis...... 104 Results...... 104 Metabotropic glutamate receptor activation inhibits the HPA axis response to restraint ... 104 Blocking mGluR signaling potentiates HPA axis response to restraint...... 105 Dexamethasone treatment reverses the antagonist-induced HPA potentiation...... 105 Discussion...... 106 Figure Legends...... 110 5. General Discussion ...... 115 Timing of effects: genomic vs. nongenomic...... 115 Fast feedback ...... 116 Metabotropic glutamate receptors...... 120 GluR5...... 121 Receptor signaling in HPA axis regulation ...... 122 Glutamate:...... 122 Glucocorticoids...... 125 Endocannabinoid receptor signaling in HPA axis regulation...... 130 Multiple CB receptors involved in HPA axis regulation in PVN?...... 130 General Considerations...... 132 Methods...... 132 Necessity vs. sufficiency of corticosteroids in fast feedback...... 136 Dissociation between ACTH and corticosterone ...... 137 Future Directions ...... 139 Conclusion ...... 140 Figure Legends...... 143 References...... 145
9 List of Figures Figure 1-1. HPA axis overview...... 44 Figure 1-2 - Glutamate Receptors...... 45 Figure 1-3 - Cannabinoid Structures...... 46 Figure 1-4 - Glucocorticoid/endocannabinoid synaptic signaling...... 47 Figure 2-1 GluR5 mRNA in the PVN...... 66 Figure 2-2 Antigen Retrieval ...... 67 Figure 2-3 GluR5/CRH Immunohistochemistry...... 68 Figure 2-4 Cannula Placement...... 69 Figure 2-5 LY382884 in the PVN...... 70 Figure 2-6 LY382884 in the Median Eminence...... 71 Figure 2-7 GluR5 Protein Expression in Areas Projecting to the PVN ...... 72 Figure 2-8 Model for GluR5 Signaling at the PVN...... 73 Figure 3-1 Cannula Placement...... 92 Figure 3-2 Fast Feedback in the PVN...... 93 Figure 3-3 Peripheral AM-251 Does Not Block Fast Feedback...... 94 Figure 3-4 Potentiation of HPA Axis Response After CB1 Blockade ...... 95 Figure 3-5 AM-251 Blocks Fast Feedback ...... 96 Figure 3-6 Win 55,212-2 in Fast Feedback...... 97 Figure 3-7 Model for Fast Feedback Signaling ...... 98 Figure 4-1 Group I Metabotropic Glutamate Receptor Agonist ...... 112 Figure 4-2 Group I Metabotropic Glutamate Receptor Antagonist ...... 113 Figure 4-3 Model for Group I Metabotropic Glutamate Receptor Signaling at the PVN ...... 114 Figure 5-1 Model for Homer-2 Mediation of Membrane GR Signaling ...... 144
10 Abbreviations
2-AG, 2-arachidonoyl glycerol ACTH, adrenocorticotropic hormone AEA, anandamide AMPA, alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid ANOVA, analysis of variance AUC, area under the curve AVP, arginine vasopressin CB1, cannabinoid receptor 1 CB2, cannabinoid receptor 2 COX, cyclooxygenase CRH, corticotropin releasing hormone DAB, 3,3' diaminobenzidine Dex:BSA, dexamethasone covalently bound to bovine serum albumin DHPG, (S)-3,5-dihydroxyphenylglycine DMSO, dimethyl sulfoxide ERK, extracellular signal-regulated kinase FAAH, fatty acid amide hydrolase GABA, gamma-aminobutyric acid GC, glucocorticoid GFAP, glial fibrillary acidic protein GPCR, G-protein coupled receptor GR, glucocorticoid receptor hexylHIBO, (S)-hexylhomoibotenic acid HPA, hypothalmus-pituitary-adrenal KPB, potassium phosphate buffer KPBS, potassium phosphate buffered saline MPEP, 2-methyl-6-(phenylethynyl) pyridine MR, mineralocorticoid receptor NMDA, N-methyl-D-aspartate NO, nitric oxide PKC, protein kinase C PLA2, phospholipase A2 PLC, phospholipase C PVN, paraventricular nucleus of the hypothalamus SEM, standard error of the mean THC, Δ9-tetrahydrocannabinol VGLUT, vesicular glutamate transporter
11 1. Introduction
The diversity of ecosystems that contain living organisms on our planet is staggering.
Life is found in the arctic and Antarctic ice, in the Sahara desert, at hot water vents at the bottom
of the ocean, and even in the earth's crust at least 4 km deep (Gold, 1992). Many of the
organisms that live in such extreme environments are single-celled organisms, including
eukaryotes, bacteria, and archaea. These organisms have evolved specialized characteristics that allow them to survive in their chosen extreme environments, and such adaptation is one of the strategies that life on this planet has used to expand into all of these ecosystems.
Another strategy, which is used by multicellular organisms in many of these same extreme environments, is to change the internal conditions of the organism to suit the needs of their cells, rather than adapting the characteristics of the cells to the demands of the external environment. This ability of a complex organism to adapt its internal environment to the needs of its individual cells was carefully investigated by Walter Cannon, who called the maintenance of a relatively constant internal environment “homeostasis” (Cannon, 1939). Since the time of
Cannon, biologists have used the word homeostasis to describe the result of a multitude of adaptations that complex organisms use to adapt to their environment and survive the challenges that are presented by changes in the external environment.
The mammalian body has evolved a number of systems that are specialized to maintain homeostasis in the face of various challenges. Such systems include the hemostasis system for responding to hemorrhage, the insulin/glucagon system for responding to alterations in blood glucose levels, brown adipose tissue/shivering for responding to decreased body temperature, etc. In addition to these specialized homeostatic mechanisms, however, there are also generalized systems for maintaining homeostasis.
12 The existence of a generalized response to varied homeostatic challenges was first
described by the Canadian endocrinologist, Hans Selye. Selye described a constellation of responses common to a variety of challenges, such as heat, cold, toxins, and surgery, and called the response to these stimuli the “general adaptation syndrome” (Selye, 1936). Animals that were exposed to noxious stimuli at first adapted to the challenge, but eventually experienced physiological responses including weight loss, immune suppression, and gastrointestinal ulcers.
The responses that Selye noted were common to a wide variety of seemingly unrelated noxious
stimuli. The responses involved in mediating the general adaptation syndrome eventually
became known as “stress” (Selye, 1956), and the noxious stimuli that cause a stress response are
often termed “stressors”. Stress responses are mediated by physiological systems, such as the
adrenal medullary system, the sympathetic nervous system, and the hypothalamus-pituitary-
adrenal (HPA) axis.
HPA Axis
The HPA axis is a neuroendocrine system, the core of which is made up of the
paraventricular nucleus of the hypothalamus (PVN), the anterior lobe of the pituitary gland, and
the adrenal cortex. In response to stress, the PVN integrates signals from multiple levels of the
brain and brainstem into a peptide response made up primarily of corticotropin releasing
hormone (CRH) and vasopressin (AVP), which are released by neurons of the PVN into the
hypophyseal portal vasculature. From there, these hormones are carried to the anterior pituitary,
where they stimulate the synthesis and release of adrenocorticotropin (ACTH). ACTH is
released into the general circulation, and acts on the adrenal cortex to induce the synthesis of
glucocorticoid hormones (mainly cortisol in humans, corticosterone in rodents). Figure 1
13 outlines these core elements of the HPA axis.
Glucocorticoid hormones that are released in response to activation of the HPA axis are
involved in a number of processes that help an organism to maintain homeostasis. For example,
glucocorticoids act to elevate blood glucose levels, to inhibit reproductive activities of an
organism, to regulate the immune system, and to maintain or regulate blood pressure (such as in
preventing shock). These activities are thought to help the animal to survive acute challenges to
homeostasis, and to recover following the survival of such an acute challenge (Sapolsky et al.,
2000). Glucocorticoids also serve to regulate the HPA axis response to subsequent stressors,
through negative feedback (Keller-Wood and Dallman, 1984).
Rapid regulation of the HPA axis
Since the HPA axis is involved in the response of an organism to stress, it is clear that
activity of the HPA axis needs to be regulated in a rapid manner. Although the HPA axis is
typically activated more slowly than other parts of the stress response, such as the sympathetic
nervous system and the adrenal medullary catecholamine system (Sapolsky et al., 2000), the
HPA axis is still activated relatively quickly in response to stress. ACTH secretion is detectably
increased within 5 minutes after the onset of restraint stress (Vahl et al., 2005), while
corticosterone secretion is detectably increased within less than 15 minutes after the onset of
stress (Tauchi et al., 2008). In contrast to this fast regulation, the HPA axis is also regulated by
slower signals. For example, intermediate negative feedback inhibition regulates HPA axis activity on the scale of about 2-6 hours (Keller-Wood and Dallman, 1984). In addition, there is an abundant literature about organizing influences on HPA axis activity, such as maternal separation, which regulate long-term activity of the axis (Hennessy, 1997; Anisman et al., 1998).
14 This slow HPA axis regulation takes place over a long period of time, up to the entire lifespan of
the organism, and has been linked to adult diseases including the metabolic syndrome and
depression (Kajantie, 2006).
Thus, both fast and slow signaling mechanisms are responsible for regulating HPA axis
activity. This dissertation will focus on some of the rapid regulatory mechanisms. There are a
number of signaling modalities that have been explored as being involved in such rapid
regulation, but this work mainly explores signals involving glutamate (through ionotropic and
metabotropic receptors) and glucocorticoid hormones (in a signaling pathway that involves
endocannabinoid signaling).
Glutamate
Glutamate is the major excitatory neurotransmitter in the central nervous system (Brann,
1995; Ozawa et al., 1998). It signals pre-synaptically and post-synaptically, through a family of receptors that includes ionotropic and metabotropic members. The ionotropic glutamate receptors act as glutamate-gated ion channels, and are divided into three general families, based on homology and pharmacological properties of the receptor proteins. These families are the N- methyl-D-aspartate (NMDA) receptors, the α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors, and the kainate receptors. Metabotropic receptors signal through G- proteins, and are also divided into three groups, known as Groups I, II, and III. Figure 2 outlines the members of these glutamate receptor families.
15 Ionotropic Glutamate receptors
Ionotropic glutamate receptors include the NMDA, AMPA, and kainate subgroups of
receptors. They function as ligand-gated ion channels, which are permeable to Na+ and K+ ions when bound to ligand (Mayer and Armstrong, 2004). Some of the ionotropic glutamate receptors are permeable to or blocked by Ca++ and Mg++ as well. Typically, the ionotropic receptors are
expressed post-synaptically (Petralia and Wenthold, 1992; Petralia et al., 1994a; Petralia et al.,
1994b), although some of them have been reported presynaptically as well (Chittajallu et al.,
1999; Brasier and Feldman, 2008). The ionotropic glutamate receptors mediate most of the fast
excitatory signaling in the mammalian central nervous system (Ozawa et al., 1998), and appear
to have some functions in peripheral, non-neural tissue as well (Dingledine et al., 1999).
NMDA receptors are expressed throughout the brain (Watanabe et al., 1993) and are
involved in signaling processes such as synaptic plasticity (Mori and Mishina, 1995). The
NMDA receptor family is divided into three types of subunit, known as NR1, NR2, and NR3,
each of which subtype is made up of a number of individual receptor proteins (See figure 2)
(Paoletti and Neyton, 2007). The NMDA receptor complex is most likely made up of a tetramer
consisting of either two NR1 and two NR2, or a combination of the three subtypes. Binding of
glutamate to the NMDA receptor complex allows the ion channel part of the receptor to open,
allowing the passage of K+ and Na+ ions (Mayer and Armstrong, 2004). Once the channel is
opened, it is also susceptible to being blocked by compounds including Mg+ and ketamine
(Paoletti and Neyton, 2007). Flux of K+ and Na+ ions through the receptor ion channel leads to
depolarization of a cell, and thus to propagation of an action potential after glutamate signaling at
a synapse.
Like NMDA receptors, AMPA receptors are glutamate-gated ion channel proteins (Ozawa
16 et al., 1998). The AMPA receptor family is made up of GluR1, GluR2, GluR3, and GluR4,
which are assembled in hetero- and homo-tetrameric ion channel pores (Rosenmund et al., 1998).
Each of these receptor proteins exists in two forms, termed “flip” and “flop,” which are products of alternative splicing (Sommer et al., 1990). Most of the fast excitatory neurotransmission in the central nervous system is thought to be transmitted by signaling through AMPA receptors
(Lees, 2000).
Kainate receptors are relatively less understood than AMPA or NMDA receptors. Like the other ionotropic glutamate receptors, kainate receptors function as post-synaptic receptors and convey excitatory signals (Huettner, 2003). In addition, presynaptic kainate receptors have been described (Lerma, 1997; Chittajallu et al., 1999). The kainate receptor family is made up of
GluR5, GluR6, GluR7, KA1, and KA2, which assemble as homomeric (GluR5-7 only) or heteromeric tetramers (Dingledine et al., 1999). A role for kainate receptors has been postulated in synaptic plasticity as well as in conveying excitatory synaptic signaling (Huettner, 2001).
Metabotropic glutamate receptors
In addition to ionotropic glutamate receptors, there are also a group of glutamate
receptors that signal metabotropically, rather than acting as ligand gated ion channels. The
metabotropic glutamate receptor family is made up of 8 members, termed mGluR1-8. These
receptors are further subdivided into group I (mGluR1 and mGluR5), group II (mGluR2 and
mGluR3), and group III (mGluR4,6,7,8) (Pin and Duvoisin, 1995). The metabotropic glutamate receptors are coupled to G-proteins, through which they initiate cellular signaling (Pin and
Duvoisin, 1995). Group I metabotropic receptors are coupled to PLC signaling and activation of
L-type Ca++ channels, while group II receptors inhibit adenylate cyclase. Group II and III
17 receptors also signal through inhibition of voltage gated Ca++ channels and activation of K+ channels (Benarroch, 2008). Group I mGluR signaling is typically excitatory, while group II and
III receptors usually act as presynaptic autoreceptors that inhibit glutamate release from presynaptic terminals (Benarroch, 2008).
Glutamate receptor signaling
Glutamate is released from a presynaptic axon terminal by a signal-dependent release of
glutamate vesicles into the synapse. This release is mediated by a Ca++ signal, usually in response to depolarization of the axon terminal (Nicholls and Attwell, 1990). Glutamate released into the synapse binds to post-synaptic glutamate receptors, thus conveying an excitatory signal onto the post-synaptic neuron. Binding of glutamate to ionotropic glutamate receptors leads to depolarization of the target cell, by Na+ and K+ ion flux through the receptor complex, and thus
to propagation of an action potential. Metabotropic glutamate receptors, on the other hand, are
known to be found outside the synaptic zone on post-synaptic cells (Van den Pol, 1994; Smith et
al., 2000), and mediate excitatory signaling through second messenger cascades. Kainate
receptors and group II and III metabotropic glutamate receptors can act as presynaptic
autoreceptors, decreasing the release of glutamate from these terminals (Jaskolski et al., 2005;
Benarroch, 2008).
Glutamate that is released during synaptic signaling is inactivated by reuptake into
neurons and astrocytes. Most of this reuptake is performed by astrocytes that are associated with the synapse (Hertz, 1979), and is mediated by excitatory amino acid transporters 1-5 (Shigeri et al., 2004). Glutamate that is taken up by glutamatergic neurons is repackaged into vesicles by vesicular glutamate transporters (VGLUT 1-3) (Shigeri et al., 2004), while glutamate taken up by
18 astrocytes is returned to the glutamatergic neurons by means of glutamate-glutamine cycles
(Hertz et al., 1999).
Glutamate and the HPA axis
Since glutamate is the major excitatory central nervous system neurotransmitter, it is not surprising that glutamate signaling is an important regulatory influence in the HPA axis. Cells in many areas involved in activation of the HPA axis express glutamate receptors, suggesting that glutamatergic terminals innervate these areas. These include the PVN, hippocampus, prefrontal cortex, and the anterior pituitary gland (Petralia and Wenthold, 1992; Petralia et al., 1994a;
Petralia et al., 1994b; Mahesh et al., 1999), suggesting an anatomic basis for glutamate signaling in activation of the HPA axis. Furthermore, neurons in the PVN proper and in areas that send afferent projections to the PVN express glutamate transporter proteins such as VGLUT1,
VGLUT2, and VGLUT3 (Fremeau et al., 2002; Kaneko et al., 2002; Ziegler et al., 2002; Danik et al., 2005; Poulin et al., 2008). These expression studies also suggest a role for glutamate in regulation of the HPA axis, which has been functionally confirmed on both acute and chronic time scales.
Acutely, glutamate infusion into the third ventricle (Makara and Stark, 1975) or PVN
(Feldman and Weidenfeld, 1997) leads to increased corticosterone secretion. Direct PVN infusion causes increased ACTH secretion (Darlington et al., 1989) and depletion of CRH from the median eminence (Feldman and Weidenfeld, 1997). Acute excitatory effects of ionotropic glutamate receptor signaling are known to be mediated by NMDA receptors (Pechnick et al.,
1989; Jezova et al., 1995) and non-NMDA receptors (Tokarev and Jezova, 1997). In addition, metabotropic glutamate receptors mediate both excitatory (Johnson et al., 2001) and inhibitory
19 (Bradbury et al., 2003; Scaccianoce et al., 2003) effects on HPA axis activity, on an acute time scale.
Chronic treatment with an NMDA antagonist increases the corticosterone response in mice (Pistovcakova et al., 2005), while chronic concurrent inhibition of NMDA and AMPA receptors leads to decreased HPA axis response to immobilization stress (Zelena et al., 1999).
Chronic stress exposure leads to alteration of glutamate receptor expression, such as increased metabotropic glutamate receptor expression in the CA1 region of the hippocampus (Wieronska et al., 2001), decreased NMDA receptor expression in the PVN (Ziegler et al., 2005), and altered expression of the AMPA receptors GluR1 (decreased) and GluR2 (increased) in the hippocampus
(Rosa et al., 2002). Consistent with alterations in glutamate receptor expression, chronic stress also alters glutamatergic signaling in the hippocampus (Joels et al., 2004). Taken together, these expression and activity data suggest that glutamate is involved in a number of different ways in regulation of the HPA axis, both acutely and chronically.
Glucocorticoids
The second focus of this work is rapid signaling by glucocorticoid hormones. Classically, glucocorticoid signaling has been thought of as a genomically-mediated signaling pathway, with effects of such signaling becoming evident as early as 1-2 hours after treatment, and occurring over hours to days. This genomic signaling is generally considered to be mediated by the cytosolic steroid hormone receptors called the type I or mineralocorticoid receptor (MR) and the type II or glucocorticoid receptor (GR). However, classic effects of glucocorticoids and other steroid hormones on gene expression cannot account for all of the effects that are caused by these hormones. Some of the rapid effects of glucocorticoids, which have been noted to occur within
20 time frames as short as seconds, must be mediated by extra-genomic mechanisms. These
nongenomic effects have been described in in vitro systems, as well as on physiological
outcomes and behavior in animals.
Non-genomic effects of glucocorticoids in non-neural Tissue
Historically, the most important pharmacological use of glucocorticoids has been in
immunosuppression. Many aspects of the immunosuppressive action of glucocorticoids are
explained by genomic actions, such as inhibition of cytokine gene expression (Liberman et al.,
2007); however, there are other effects that are most likely mediated by nongenomic actions, including rapid effects on second messenger systems and cell membranes in immune cells
(Buttgereit and Scheffold, 2002; Lowenberg et al., 2005). These changes are thought to lead to observed rapid anti-inflammatory effects of high-dose steroid treatment (Buttgereit et al., 1998).
In addition to these effects in the immune system, nongenomic glucocorticoid effects have been
documented in cells from other peripheral tissues, including endometrium (Koukouritaki et al.,
1996), lung (Urbach et al., 2002; Verriere et al., 2005), liver (Daufeldt et al., 2003) and intestine and kidney (Wang et al., 2007).
Non-genomic effects of glucocorticoids in the Nervous System: behavior and physiology
Behavioral. Behaviors elicited in a nongenomic fashion by glucocorticoids are evidence of
neural effects of glucocorticoid hormones. Some of the most well characterized behavioral
effects of glucocorticoids are found in the amphibian Taricha granulosa, in which
21 glucocorticoids induce sexual clasping behavior when acutely administered (Orchinik et al.,
1991). Nongenomic effects of glucocorticoids have also been described in avian species. For
example, in the white crowned sparrow, corticosterone increases locomotor activity within 15
minutes after administration (Breuner et al., 1998). Other work done with the migratory Red
Knot is consistent with nongenomic effects of corticosterone on locomotor activity, although
more work is necessary to confirm that these effects are nongenomic (Landys et al., 2004).
In addition to non-mammalian species, there have been reports of possible or probable
non-genomic actions of glucocorticoids in rats. Novelty-related locomotor activity in rats is
increased in a nongenomic fashion by corticosterone treatment (Sandi et al., 1996b). This increase in activity may relate to risk-assessment behavior, which is rapidly increased after
treatment with corticosterone, without change in anxiety-like behavior or general locomotion
(Mikics et al., 2005). In contrast to locomotor activity, corticosterone rapidly inhibits the
acoustic startle response in rats (Sandi et al., 1996a). Also, intrathecal injection of RU486 into
rats with spinal cord injury decreases the hyperalgesic effects of the lesion, in a manner that is
consistent with nongenomic signaling (Wang et al., 2005). The experimental design in this case
does not rule out effects by genomic signaling; however, it is on a time course (within 30 minutes
of treatment) that is consistent with nongenomic signaling. In addition, the effects of RU486
treatment mirror the effects of NMDA receptor antagonism, suggesting that the mechanism of
action for the two may be similar. Glucocorticoid treatment rapidly inhibits retrieval of long-
term memory, in a manner that is insensitive to protein synthesis inhibition (Sajadi et al., 2006).
There is even evidence for a rapid, presumably nongenomic role for glucocorticoids in regulating
feeding, at the level of the hypothalamus (Tasker, 2006). Thus, there appear to be nongenomic
effects of glucocorticoids on behavior in a wide array of vertebrates.
22
HPA Axis.
One of the important systems in which glucocorticoids are known to exert nongenomic
effects is the HPA axis, where glucocorticoids rapidly induce negative feedback inhibition (see
figure 1). The first evidence for fast feedback inhibition of the HPA axis by glucocorticoids was
published in the late 1940s (Sayers and Sayers, 1947). Treatment of rats with adrenal extract
immediately before a variety of stressors blocked the decrease in adrenal ascorbic acid content
(an indirect index of corticosterone production), indicative of decreased corticosterone synthesis.
When ACTH was administered, there was no difference between adrenal extract of treated and
control animals, suggesting that the negative feedback signaling was occurring proximal to the
adrenal gland. Since then, evidence has accumulated that nongenomic negative feedback
regulation of the HPA axis by glucocorticoids occurs at the anterior pituitary gland. In vitro
work has shown that corticosterone inhibits CRH-induced ACTH release from fragments of
pituitary explants in culture in less than 20 minutes (Widmaier and Dallman, 1984), suggesting a
nongenomic mechanism of action. In cultured pituicytes, this inhibition persists in the presence
of cycloheximide, thus strengthening the assertion that this inhibition represents a nongenomic
action of glucocorticoids (Abou-Samra et al., 1986).
In addition to in vitro studies, there is in vivo evidence for nongenomic glucocorticoid
actions at the pituitary. Rats with mediobasal hypothalamus lesions, which destroy the afferent connections from the brain to the pituitary, display rapid inhibition of ACTH release in response
to corticosterone treatment. (Jones et al., 1977) Furthermore, glucocorticoids rapidly inhibit
CRH-induced ACTH secretion in anesthetized rats (Hinz and Hirschelmann, 2000). This effect persists despite prior actinomycin D treatment, thereby implicating a nongenomic signaling
23 pathway. These data strongly suggest that negative feedback inhibition of the HPA axis can
occur at the pituitary gland.
Similarly, there is evidence that the hypothalamus is a site for nongenomic negative feedback. Glucocorticoids rapidly inhibit CRH release from hypothalamic slices (Jones and
Hillhouse, 1976; Jones et al., 1977) and synaptosomes (Edwardson and Bennett, 1974).
Treatment with dexamethasone or corticosterone, but not cholesterol or isopregnanolone rapidly decreases the frequency of excitatory post synaptic currents in CRH-containing hypothalamic cells (Di et al., 2003). The glutamatergic effect suggests a rapid inhibitory action of glucocorticoids on CRH release, although CRH was not measured in this study. Furthermore, infusion of dexamethasone directly into the PVN of rats 5 minutes before restraint stress blunts the ACTH and corticosterone responses (see Chapter 3). This suppression of HPA activation is also caused by dexamethasone covalently bound to bovine serum albumin (dex:BSA), thus implicating a membrane glucocorticoid receptor in the nongenomic feedback signaling pathway.
Taken together, these studies suggest that fast feedback inhibition of the HPA axis occurs via signaling at the PVN as well as at the pituitary.
Other parts of the brain have not yet been implicated in fast, nongenomic feedback inhibition of the HPA axis. However, because other brain sites including the hippocampus
(Feldman and Weidenfeld, 1999; Mizoguchi et al., 2003), prefrontal cortex (Akana et al., 2001) and paraventricular thalamus (Jaferi and Bhatnagar, 2006) are known to be involved in negative feedback inhibition of the HPA axis, it is reasonable to expect that at least some of these areas may be involved in negative feedback through nongenomic signaling. Further studies are needed to explore this possibility.
24 Electrophysiology/neuronal activity.
There is a wealth of information available about rapid effects of glucocorticoids on
neuron electrophysiology. Glucocorticoids rapidly exert both excitatory and inhibitory
influences on neurons in the hypothalamus (Dafny et al., 1973; Mandelbrod et al., 1974) (Ruf
and Steiner, 1967), hippocampus (Pfaff et al., 1971; Dafny et al., 1973; Karst et al., 2005;
Wiegert et al., 2006), and brainstem (Ruf and Steiner, 1967; Dafny et al., 1973) in rodents.
These effects appear between milliseconds and about 30 minutes after application of
glucocorticoids, suggesting nongenomic actions. In addition, because the hippocampus and
hypothalamus are involved in regulation of the HPA axis, these effects on neuronal excitability may be involved in nongenomic negative feedback. These effects could be caused by action of glucocorticoids on ion channels or membranes (see below) or via modulation of neurotransmitter
systems. Corticosterone rapidly increases both aspartate and glutamate levels in the
hippocampus (Venero and Borrell, 1999). Glucocorticoid treatment also rapidly increases
glutamate uptake into hippocampal synaptosomes (Zhu et al., 1998) and tryptophan uptake into
whole brain synaptosomes (Neckers and Sze, 1975), suggesting that some of the nongenomic
effects of glucocorticoids on neuronal signaling may occur through rapidly altering
neurotransmitter metabolism.
Signaling Mechanisms: receptors
There is a great deal of information available about receptors and intracellular signaling
pathways that are involved in the nongenomic effects of glucocorticoids. One of the major
questions that remains open, with respect to the receptors that are operative in nongenomic effects of these steroids, is whether the effects are mediated by the classical intracellular
25 receptors GR and MR (whether in the normal intracellular form, or as a modified, membrane- bound version) or by novel, as yet unidentified receptors. There are data supporting both of these positions, and this evidence suggests that nongenomic effects of glucocorticoids can be classified three ways, based on receptors mediating these effects: 1) unspecific effects, or those not mediated by a receptor, 2) effects mediated by the cytosolic GR, and 3) effects mediated by a membrane-bound receptor.
Unspecific/non-receptor mediated.
Various steroid hormones, including glucocorticoids, have specific nongenomic actions that are mediated by direct effects of these molecules on cell membrane properties. Sex steroids, for example, can alter membrane fluidity and permeability to hydrophilic molecules, as well as inducing membrane vesicle fusion (Shivaji and Jagannadham, 1992; Whiting et al., 2000). There is also evidence that steroids alter membrane fluidity in a manner that is specific for individual steroids (Massa et al., 1975). Specifically in this study, cortisol increased membrane fluidity, while progesterone decreased it. In fact, membrane effects of individual steroids have even shown specificity for membrane preparations from different organs, such as brain vs. muscle
(Whiting et al., 2000).
Alterations in the physicochemical properties of the plasma membrane can change the activity of membrane-bound proteins (Stallkamp et al., 1999) or result in altered permeability of the membrane to ions (Buttgereit et al., 1998). Direct effects of glucocorticoids on cell membranes are hypothesized to mediate some of the rapid therapeutic effects of glucocorticoids, such as those achieved by pulse glucocorticoid treatment of severe inflammatory disorders
(Buttgereit et al., 1998; Lipworth, 2000). Indeed, direct effects on permeability of the
26 mitochondrial membrane may underlie the induction of apoptosis by glucocorticoids in immune cells (Gavrilova-Jordan and Price, 2007). These results suggest that physicochemical interactions between steroids and cell membranes may be important mediators of the pharmacological effects of steroids. The relatively high concentration of steroids needed, however, suggests that these effects may not be important in physiological steroid signaling.
Cytosolic GR.
Nongenomic actions of glucocorticoids mediated through the classical GR are suggested by the association of kinases, such as the Src kinase, with the GR complex in the cytoplasm of the cell (Buttgereit and Scheffold, 2002). Binding of glucocorticoids to GR leads to dissociation of other proteins in the complex from GR (Buckingham, 2006). In the case of Src and other putative signaling molecules operating nongenomically through the cytosolic GR, dissociation of the kinase from the GR complex would free it to phosphorylate its targets, and thus to mediate nongenomic signaling. In support of this putative signaling pathway, estrogen is known to signal through Src (Wong et al., 2002). This SRC-mediated signaling involves the scaffold protein
MNAR/PELP, which is extensively co-expressed with GR in the nervous system (Khan et al.,
2006), suggesting that GR may signal through a similar pathway. Glucocorticoids are also known to signal nongenomically by causing dissociation of the GR complex in T-cells, where
GR forms part of the T-cell receptor complex (Lowenberg et al., 2007). Binding of glucocorticoids to GR in these cells causes disruption of the T-cell receptor complex in a nongenomic fashion, thus leading to impaired T-cell receptor signaling.
27 Membrane-bound Receptors.
Although cytosolic GR may be involved in nongenomic glucocorticoid signaling, in most cases the receptor mediating nongenomic effects is apparently membrane-bound. The earliest evidence for this conclusion was supplied by studies revealing corticosteroid binding to brain cell membranes in vitro (Towle and Sze, 1983). Assays comparing the binding of estrogen, progesterone, testosterone, and corticosterone suggested that there are specific binding sites for each of these steroids. None of these hormones was able to compete with the others at their respective binding sites, indicating that there are specific receptors for each steroid.
Plasma membrane binding sites for glucocorticoids in synaptic membranes and pituitary cells often have characteristics that are compatible with these sites being G protein-coupled receptors (GPCRs) (Orchinik et al., 1992; Guo et al., 1995; Di et al., 2003; Maier et al., 2005).
However, there is also evidence of other membrane receptors, such as the GABAA receptor, that
mediate some of the nongenomic effects of glucocorticoids. Thus, we subdivide the
classification of membrane-bound receptor-mediated effects into three classes: effects mediated
by a membrane-bound form of GR, effects mediated by a membrane bound, non-GR
glucocorticoid receptor, and effects mediated by interactions with proteins that are not primarily
glucocorticoid receptors.
Membrane-bound glucocorticoid receptors: GR or non-GR? Although the presence of
receptors for glucocorticoids in the membranes of cells is relatively undisputed, the identity of
the membrane receptor is still controversial. There is a great deal of evidence supporting a form
of GR as the receptor through which nongenomic effects are mediated, especially in the immune
system. A presumably modified version of the classical GR has been identified by
28 immunocytochemistry as being bound to the membrane of immune cells (Gametchu, 1987;
Gametchu et al., 1999). Membrane-bound GR is associated with a specific alternative transcript of GR (Chen et al., 1999), and appears to mediate at least some of the rapid effects of glucocorticoids in these cells (Lowenberg et al., 2005; Bartis et al., 2006). GR has been
identified in caveolae in the cell membrane (Jain et al., 2005), where it can elicit transcriptional
signaling. It is not clear whether this caveolae-associated GR is also able to mediate nongenomic
signaling, but this seems likely given that caveolae are membrane specializations associated with
signal transduction (Anderson, 1998). Furthermore, GR has been shown by yeast two-hybrid
screening to directly interact with G protein beta (Kino et al., 2005), and by co-
immunoprecipitation to interact with the guanine exchange factor Brx (Kino et al., 2006).
Dexamethasone causes rapid phosphorylation of ZAP kinase (Bartis et al., 2006), and GR interacts with this kinase (Bartis et al., 2007). In aggregate, it appears that nongenomic glucocorticoid signaling in the immune system is usually mediated by a membrane-associated form of GR. This is in contrast to liver cells, in which the membrane part of the signaling cascade is apparently a steroid transport protein, which ties into GR signaling intracellularly
(Daufeldt et al., 2003; Daufeldt et al., 2006).
In the nervous system, current evidence suggests that most nongenomic and putative nongenomic effects of glucocorticoids may be mediated by receptors other than GR. For example, the relative potencies for synthetic glucocorticoid-induced nongenomic effects are significantly different from their relative potencies at causing GR-mediated genomic effects
(Buttgereit et al., 1999). In addition, hippocampal cells cultured so as to eliminate GR expression in the cells retain their ability to rapidly activate signaling through intracellular kinase cascades in response to acute glucocorticoid treatment (Xiao et al., 2005). Finally, many of the described
29 nongenomic effects of glucocorticoids in the central nervous system are insensitive to inhibition
by the GR antagonist RU486 (Han et al., 2002; Di et al., 2003; Maier et al., 2005). Together,
these results appear to support the existence of a novel receptor that mediates the nongenomic
actions of glucocorticoid.
As a caveat to this conclusion, there is some evidence that higher concentrations of
RU486 are able to at least partially antagonize effects that were not sensitive to inhibition at
lower doses used in other studies (Liu and Chen, 1995). This could be caused by an alteration in
the conformation of GR due to membrane binding per se, or because of structural alterations
necessary to allow membrane localization of the receptor. There is evidence, for example, that
changing the hydrophobicity of an enzyme’s environment can significantly alter the activity or
other chemical properties of the protein (Stallkamp et al., 1999) (Wu and Gorenstein, 1993). In
fact, altering the lipid content of brain cell membranes through treatment with phospholipase C
or A2 can abolish the binding of corticosterone to the membranes (Towle and Sze, 1983). These
data suggest that placing GR into the membrane could lead to alterations in the ligand binding domain that are significant enough to alter the affinity of the receptor for ligand and inhibitors.
That said, however, a non-GR, membrane-bound, GPCR-associated receptor for corticosterone has been partially characterized in the salamander T. granulosa (Evans et al.,
2000). This receptor, based on physical characteristics of the partially purified protein, is most
likely an acidic glycoprotein, with a size of about 63 kilodaltons. The physical and chemical
characteristics of this protein are incompatible with the classical GR, although the possibility that the protein is a modified form of GR has not been unequivocally ruled out. Binding of corticosterone to this receptor is insensitive to RU486 (Orchinik et al., 1991) and appears to mediate the behavioral effects of corticosterone on sexual clasping behavior in this animal (Rose
30 et al., 1993; Moore and Orchinik, 1994). Interestingly, the rapid behavioral effects mediated through the Taricha membrane receptor appear to be regulated by endocannabinoid signaling
(Coddington et al., 2007), similar to the rapid inhibitory effects of glucocorticoids on CRH- containing neurons in the hypothalamus (Di et al., 2003).
Another possibility is that some of the nongenomic actions of glucocorticoids are mediated through a GPCR-associated receptor for a neurosteroid or similar molecule. A recent paper reported a GPCR-associated receptor for the neurosteroid dehydroepiandrosterone
(DHEA) (Charalampopoulos et al., 2006), which also has affinity for corticosterone.
Corticosterone binding to this receptor antagonized the effects of DHEA, suggesting the possibility that some of the nongenomic actions of glucocorticoids in the nervous system occur by antagonizing actions of neurosteroids.
On the other hand, there is evidence that GR is expressed in the membranes of hippocampal and hypothalamic neurons (Liposits and Bohn, 1993). In the lateral amygdala, the presence of GR at post-synaptic membranes and in astrocytic processes has been demonstrated by electron microscopy (Johnson et al., 2005). In addition, several synaptic proteins, including
Vesl-2, LIM, and SH-3, have been identified as interacting with GR (Hedman et al., 2006).
Binding of GR to these proteins suggests a mechanism by which GR could be localized to the synapse. Vesl-2 (also known as Homer-2) also binds to metabotropic glutamate receptors
(Brakeman et al., 1997), indicating a route by which GR could affect presynaptic signaling by modulating presynaptic glutamate receptors.
Non-glucocorticoid-receptor membrane proteins. Alternatively, glucocorticoids may have rapid effects in the mammalian brain by acting directly on ion channels. For example, some
31 steroids, especially neurosteroids, are known to directly bind to and modulate the activity of the
GABAA receptor (Orchinik et al., 1994; Stromberg et al., 2005). Although glucocorticoids most
likely do not have a strong influence on the GABAA receptor (Orchinik et al., 1994), many of the
neurosteroids that do are products of glucocorticoid metabolism in the brain. Therefore,
glucocorticoids could have nongenomic effects on GABA transmission indirectly, through metabolite actions. It is also possible that effects on ion transmission that have been attributed to unspecific glucocorticoid effects on cell membranes could be explained by this type of direct effect on ion channels.
Intracellular signaling systems
Another question that remains unanswered is, once glucocorticoids have acted on
whatever receptor mediates their effects, what downstream signaling systems are activated to
convey this signal? One answer that has been suggested by in vitro work is that
endocannabinoid signaling mediates the nongenomic signal initiated by glucocorticoid actions at
the PVN (Di et al., 2003).
Cannabinoids
Cannabinoids and endocannabinoid signaling have recently gained a great deal of
attention, both in the popular press (medical marijuana, for example), and in the scientific
literature. Since endocannabinoid signaling is one of the proposed pathways for rapid actions of
glucocorticoids in fast negative feedback regulation of the HPA axis, this signaling system will
be examined more closely.
32 Cannabinoids. Cannabinoids are substituted monoterpenoids derived from the plant Cannabis sativa (Mechoulam, 1970), also known as marijuana or Indian Hemp. This plant has a long history of use by humans, as a source for fibers, as a medicine, and as a recreational drug (Todd,
1946; Mikuriya, 1969; Mechoulam and Feigenbaum, 1987; Zuardi, 2006). Recorded mention of
Cannabis cultivation was made at least 6000 years ago in China, where it was originally grown as a source of fibers (Li and Lin, 1974). Fiber from the Cannabis plant has been used to make rope, fabric, and even paper.
Cannabis has also been used both medicinally and in religious rites. Cannabis was used medicinally by about 2000 B.C. in China (Li and Lin, 1974). Written accounts of its use in
Assyria, Egypt, Greece and Rome, and India have also been discovered (Rouyer, 1810;
Mechoulam and Feigenbaum, 1987). Traditional medicinal uses for Cannabis cover a wide variety of afflictions, including fever, arthritis, intestinal disorders, “female disorders,” malaria, convulsions, emesis, headache, and earache (Mechoulam and Feigenbaum, 1987; Zuardi, 2006).
Although Cannabis appears to have been widely used for medicinal purposes in Eastern civilizations, Western medicine was generally silent on the use of Cannabis as a medicine up to the 19th century. Around this time, the Irish physician William B. O'Shaughnessy used Cannabis, and advocated for its use by other physicians (O'Shaugnessy, 1843; Zuardi, 2006).
O'Shaughnessy discovered Cannabis while serving in India with the British. He then began to test it in animals, and treat patients with it. He successfully used the plant to treat rheumatism, convulsions, and apparently even tetanus and rabies (Mikuriya, 1969). After the time of
O’Shaughnessy, Cannabis drugs enjoyed more widespread use. However, mainly because of difficulties with standardization, and inconsistency in effects, the medicinal use of this plant gradually fell into disfavor. It was removed from the British pharmacopoeia in 1932 (Todd,
33 1946), and from the American pharmacopoeia in 1941 (Zuardi, 2006).
Cannabinoid chemistry. The unique chemical compounds found in the Cannabis plant are
called cannabinoids, and these chemicals are thought to underlie the pharmacological actions of
Cannabis. Beginning at the end of the 19th century, work on the chemistry of Cannabis began to reveal the identities of cannabinoids. Cannabinol was the first to be isolated (Wood et al., 1896), although its structure was not elucidated until the work of Todd (Todd, 1946) and Adams
(Adams, 1941). After the 1940s little cannabinoid chemistry was done until Δ9- tetrahydrocannabinol (THC) was isolated and its structure determined (Gaoni and Mechoulam,
1964). Since then much more work on the chemistry and pharmacology of cannabinoids has been done, and many of the traditional uses of Cannabis have been validated and the active cannabinoid(s) responsible for these actions discovered (Mechoulam and Feigenbaum, 1987).
The two major known pharmacological agents in Cannabis are cannabidiol and THC
(Mechoulam and Feigenbaum, 1987), of which THC is the main psychotropic cannabinoid.
Cannabidiol is also known to have pharmacological effects, including anticonvulsant activity.
Cannabinoids are known to have antiemetic, anticonvulsant/antiepileptic, bronchodilatory, antipyretic, and analgesic activity, and to reduce intraoptic pressure (Mikuriya, 1969;
Mechoulam and Feigenbaum, 1987; Zuardi, 2006). Figure 3 illustrates chemical structures of some of the cannabinoids.
The brain endocannabinoid system: Cannabinoid receptors. Early in the investigation of cannabinoid chemistry, it was widely believed that the effects of cannabinoids were mediated by nonspecific effects of the chemicals on cell membranes, because of the lipophilic nature of the
34 cannabinoids. However, it became clear that many of the observed cellular effects of
cannabinoids were consistent with actions of a G-protein coupled receptor system (Martin,
1986). In 1988, an endogenous pharmacological entity that was sensitive to the synthetic
cannabinoid CP 55,940 was described in rats (Devane et al., 1988), and the cloning of the first
cannabinoid receptor, CB1, was reported in 1990 (Matsuda et al., 1990). The second known
cannabinoid receptor, CB2, was cloned in 1993 (Munro et al., 1993). In addition to these two cannabinoid receptors, cannabinoids are also known to interact with other proteins, including
Ca++ and K+ channels, and vanilloid receptors (Di Marzo and De Petrocellis, 2006).
Furthermore, there is evidence for the existence of an as-yet undiscovered receptor related to
CB1 and CB2 (Sagan et al., 1999; Di Marzo et al., 2000; Breivogel et al., 2001).
CB1 is generally considered to be the brain cannabinoid receptor while CB2 is thought of
as a peripheral cannabinoid receptor. CB1 is highly expressed in all parts of the brain
(Herkenham et al., 1991; Moldrich and Wenger, 2000), and at lower levels in peripheral tissues
including the pituitary gland and immune tissues (Galiegue et al., 1995). CB2 expression is
mainly confined to the periphery, being expressed most highly in immune tissues (Galiegue et al., 1995). However, CB2 is also found in glial cells (Nunez et al., 2004; Sheng et al., 2005) and neurons (Onaivi et al., 2006) in the brain, and this neural expression appears to have functional implications in some animal studies.
The brain endocannabinoid system: Endocannabinoids. The presence in the body of receptors that mediate the effects of cannabinoids, combined with the strict structure-activity relationship between cannabinoids and activation of the cannabinoid receptor strongly suggested that natural endogenous ligands for the receptor must be made in the body (Mechoulam et al.,
35 1988). The search for endogenous molecules with activity profiles similar to cannabinoids
yielded the discovery of arachidonoyl ethanolamide (anandamide or AEA) in 1992 (Devane et
al., 1992). AEA is a derivative of arachidonic acid, and takes its name from ananda, the Sanskrit
word for bliss. The other major endocannabinoid, 2-arachidonyl glycerol (2-AG), was discovered several years later (Mechoulam et al., 1995; Sugiura et al., 1995). In addition to these two compounds, two other endogenous compounds with endocannabinoid activity are known: 2- arachidonoyl glyceryl ether or noladin ether (Hanus et al., 2001), and virodhamine or arachidonoyl ethanolamine (Porter et al., 2002). AEA and 2-AG both activate both CB1 and
CB2, although both CB1 and CB2 most likely have greater affinity for 2-AG than for AEA
(Sugiura et al., 2002). Noladin ether is selective for CB1, and virodhamine is an agonist at CB2
and a partial agonist/antagonist at CB1. In addition to these endocannabinoids, there are related
molecules, such as N-arachidonoyl-dopamine, palmitoylethanolamide, and oleamide that have
effects related to those of the endocannabinoids (Hanus, 2005).
Endocannabinoid synthesis, release, reuptake, metabolism. AEA and 2-AG are synthesized
from arachidonic acid through non-oxidative biosynthetic pathways (Sugiura et al., 2006;
Malcher-Lopes et al., 2008). Although the synthesis of endocannabinoids is regulated, there are
multiple synthetic pathways leading to their production. AEA can be synthesized from
arachidonic acid and ethanolamine by a direct N-acylation reaction. This reaction is thought to
be catalyzed by a fatty acid amide hydrolase (FAAH), which also mediates the degradation of
AEA. It is unclear whether this pathway for AEA synthesis is of physiological relevance, as high
concentrations of substrate are needed to drive this reaction forward (Di Marzo et al., 1998).
Alternatively, AEA can be synthesized from the precursor N-arachidonoyl
36 phosphatidylethanolamine, through the action of phospholipase D (Di Marzo et al., 1994). The
precursor itself is made by a Ca++-dependent transacylase which transfers arachidonate onto
phosphatidylethanolamine. 2-AG, on the other hand is produced upon cleavage of
diacylglycerol, which can occur through phospholipase C-dependent or -independent pathways.
The precise synthetic pathway for 2-AG production appears to depend on the tissue or cell in
which it is being produced (Sugiura and Waku, 2002).
Endocannabinoids are synthesized “on demand” rather than being stored and released like
many neurotransmitters. They are released upon synthesis, probably via transporter proteins
(Adermark and Lovinger, 2007), and are removed from the synapse by neurons and astrocytes.
This uptake is mediated by specific uptake proteins, and is followed by enzymatic degradation
(Bisogno et al., 2001; Deutsch et al., 2001). AEA is metabolized by FAAH, while 2-AG is
metabolized by monoacylglycerol lipase, and possibly by FAAH (Sugiura and Waku, 2002).
There is also evidence that 2-AG may be metabolized by cyclooxygenase enzymes (Jonsson et
al., 2006). Reuptake and metabolism of endocannabinoids by these pathways terminates their
synaptic signaling activity.
Synaptic actions of endocannabinoids. Presynaptic signaling by endocannabinoids in the
central nervous system is initiated by activation of post-synaptic group I metabotropic glutamate
receptors (Maejima et al., 2001; Varma et al., 2001; Mackie, 2005). These receptors directly
activate PLC and elevate intracellular Ca++ concentrations (Houamed et al., 1991; Abe et al.,
1992), both of which are known to initiate endocannabinoid synthesis. Cannabinoids are released from the post-synaptic cell, diffuse in a retrograde direction across the synapse, and act
on cannabinoid receptors on the presynaptic terminal. Most published reports indicate that
37 ++ activation of presynaptic CB1 closes voltage-gated calcium channels, leading to decreased Ca flux into the terminal (Hoffman and Lupica, 2006). Decreased Ca++ flux causes inhibition of neurotransmitter release from the presynaptic terminal. This signaling sequence is known to account for the decreased release of glutamate and GABA in response to cannabinoid receptor
++ signaling. In addition to this Ca -mediated pathway, CB1 activation is also known to interact
with various K+ channels, also leading to decreased neurotransmitter release (Hoffman and
Lupica, 2006). Figure 4 outlines a general model of endocannabinoid synaptic signaling.
Endocannabinoids in the HPA axis. Since cannabinoid receptors are expressed nearly
ubiquitously in the brain, and cannabinoid signaling affects widely disparate neurotransmitter
systems in the central nervous system, it is reasonable to think that endocannabinoid signaling
might regulate the HPA axis. This is borne out by available evidence. Acute and repeated
restraint both result in increased levels of the endocannabinoid anandamide in the amygdala and
forebrain (Patel et al., 2005), and both of these areas have been implicated in regulating the HPA
axis (Herman et al., 2003). In vivo, signaling through CB1 decreases the magnitude of the HPA
response to stress (Patel et al., 2004), while antagonizing CB1 leads to an enhanced HPA
response (Ginsberg et al., 2006). In addition, CB1 knockout mice mount enhanced ACTH and
corticosterone responses to novelty stress (Barna et al., 2004) and increased ACTH response
after exposure to elevated plus maze (Haller et al., 2004). CB1 expression has been noted in
areas important for HPA axis activity, including the hypothalamus (Tsou et al., 1998; Moldrich
and Wenger, 2000) and anterior pituitary (Galiegue et al., 1995). Chronic treatment of rats with
the synthetic cannabinoid CP-55,940 leads to increased expression of CRH in the PVN and
increased expression of POMC in the anterior pituitary (Corchero et al., 1999).
38 These results support a role for cannabinoids in regulating the magnitude of an HPA axis
response, which is consistent with fast negative feedback limiting of the HPA response to stress
(Keller-Wood and Dallman, 1984). In fact, electrophysiological evidence suggests that signaling
through CB1 may mediate the nongenomic actions of glucocorticoids in fast feedback regulation
of the HPA axis (Di et al., 2003). In vivo experiments in the amphibian T. granulosa suggest that
nongenomic actions of corticosterone on mating behavior may be mediated by endocannabinoid
signaling (Coddington et al., 2007). However, the evidence supporting a role for
endocannabinoid signaling in mediating nongenomic negative feedback on the HPA axis is as yet
weak. A role for CB1 receptor signaling underlying nongenomic negative feedback in the HPA axis has never yet been demonstrated in vivo, and in vitro experiments showing nongenomic
effects of glucocorticoids being mediated by CB1 signaling have not shown functional outputs of
cells, in the form of CRH secretion (Di et al., 2003). Therefore, it is important to establish
whether cannabinoid signaling is involved in the described phenomenon of nongenomic negative
feedback regulation of the HPA axis by glucocorticoid hormones.
Other signaling pathways
Other signaling pathways that may mediate nongenomic glucocorticoid actions have been
exhaustively reviewed elsewhere (Borski, 2000; Makara and Haller, 2001; Haller et al., 2008).
Briefly, some of the systems that have been implicated in these actions include endocannabinoids
(Alger, 2002; Di et al., 2003; Freund et al., 2003; Barna et al., 2004), protein kinase A (PKA)
(Verriere et al., 2005), protein kinase C (PKC) (Han et al., 2002; Di et al., 2003), mitogen activated protein kinases (MAPK) (Lowenberg et al., 2005; Hedman et al., 2006), cAMP
(Koukouritaki et al., 1996; Borski et al., 2002), intracellular Ca++ signaling (Borski et al., 2002;
39 Urbach et al., 2002), phospholipase A2 (PLA2) (Towle and Sze, 1983; Hibbeln et al., 1989),
phospholipase C (PLC) (Towle and Sze, 1983; Borski et al., 2002), nitric oxide (NO) (Sandi et
al., 1996a), and scaffolding proteins such as MNAR/PELP(Khan et al., 2006). Many of these
signaling pathways are generic intracellular signaling pathways, which could have effects in
nearly any cell type that they might be present in. However, some of these pathways are of
particular interest in the neuroendocrine system. For example, cannabinoids in the hypothalamus
(as discussed above) and the annexin 1 system in the anterior pituitary have both been implicated in nongenomic glucocorticoid regulation of HPA axis activity.
Annexin 1 (formerly known as lipocortin 1 or lipomodulin) is a Ca++ and phospholipid
binding protein in the annexin family. It was first described in mouse macrophages as a
glucocorticoid-induced protein, which mediates many of the anti-inflammatory effects of
glucocorticoids in these cells (Buckingham et al., 2006). Of interest in the neuroendocrine
system, annexin 1 is expressed in the folliculostellate cells in the anterior pituitary (Solito et al.,
2003), where it appears to play a role in nongenomic negative feedback regulation of the HPA
axis. There is clear evidence that glucocorticoid exposure causes transcription independent
translocation of annexin 1 to the plasma membrane (Taylor et al., 1993). From there it appears to
act in a juxtacrine manner, binding to putative receptors on pituicytes (Christian et al., 1997;
Chapman et al., 2002; John et al., 2002). Signaling through annexin 1 appears to mediate the
rapid inhibitory effects of glucocorticoids on ACTH secretion in vitro (Taylor et al., 1993). This signaling system can be reconstituted in a co-culture of folliculostellate cells and pituicytes, but not in single culture of either one (Tierney et al., 2003).
These data are interesting in light of the fact that annexin 1 is expressed in the hypothalamus (Philip et al., 1997), mainly in glial cells (Philip et al., 1995), as well as
40 ependymal cells and tanycytes (Strijbos et al., 1991). These expression data suggest the
possibility of an annexin-1 mediated signaling pathway in the hypothalamus, similar to that found in the pituitary gland. This possibility is supported by a study showing that annexin 1 inhibits cytokine-induced release of CRH from the hypothalamus in vitro and corticosterone secretion in vitro (Loxley et al., 1993). Nongenomic glucocorticoid signaling involving annexin
1 has also been described in A549 lung fibroblasts (Croxtall et al., 2000), suggesting that annexin
1 may be a player in nongenomic glucocorticoid signaling in peripheral tissues. Although there is a possibility that annexin I may be involved in regulating the HPA axis at the level of the hypothalamus, however, there is not yet any evidence that glucocorticoid signaling is involved in this signaling. Further, it is not clear how annexin I could be involved with the endocannabinoid-mediated fast feedback system described below. Thus, the actions of annexin I are beyond the scope of this work.
Conclusion
It is clear from the discussion above that rapid regulation of the HPA axis is an important
modality for controlling the HPA axis response to stress. As discussed above, the HPA axis
response to restraint is turned on quickly, and can also be terminated rapidly. Rapid signaling in
the brain is known to occur due to the actions of glutamate, glucocorticoids, and cannabinoids, as
discussed above. Nongenomic signaling by glucocorticoids occurs in multiple areas of the brain,
including some that are involved in regulation of the HPA axis, and there is in vitro evidence that
nongenomic actions of glucocorticoids at the PVN regulate HPA axis activity. In addition, there
is abundant evidence for a role of glutamate signaling (both ionotropic and metabotropic) in
regulation of the HPA axis. The proceeding studies investigate the roles of glutamate (through
41 GluR5 and group I metabotropic glutamate receptors), glucocorticoids, and cannabinoids in rapid
regulation of the HPA axis response to restraint stress.
Figure Legends
Figure 1. Overview of the HPA axis. The main elements of the HPA axis are the
paraventricular nucleus of the hypothalamus (PVN), the anterior pituitary gland, and the adrenal
cortex. The PVN secretes CRH into the hypophyseal portal vasculature, which induces the
synthesis and release of ACTH from the anterior lobe of the pituitary gland. ACTH causes the
synthesis of glucocorticoids by the adrenal cortex. The PVN is regulated in part by inputs from
other parts of the brain, including the hypothalamus, prefrontal cortex, and paraventricular
thalamus.
Figure 2. Glutamate receptor families. Glutamate receptors are divided roughly into ionotropic (A) and metabotropic (B) glutamate receptors. Each of these is subdivided into three groups. The members of each group are outlined.
Figure 3. Chemical structures of some cannabinoids. A) Endocannabinoids, or naturally occurring endogenous molecules in the body that have activity at cannabinoid receptors. B)
Naturally occurring cannabinoids found in the Cannabis sativa plant. C) Synthetic cannabinoids.
Figure 4. Model of endocannabinoid synaptic signaling. Glucocorticoids (GCs) act on a putative G-protein coupled receptor, by which they induce production of endocannabinoids
(CBs) in the postsynaptic neuron. CBs act on CB1 receptors in the presynaptic terminal, which
42 mediate a decreased release of glutamate onto the postsynaptic cell. Decreased glutamatergic
inputs onto the CRH-containing postsynaptic cell leads to decreased CRH release and therefore to decreased HPA axis activity.
43 Figure 1-1. HPA axis overview
44 Figure 1-2 - Glutamate Receptors
45 Figure 1-3 - Cannabinoid Structures
46 Figure 1-4 - Glucocorticoid/endocannabinoid synaptic signaling
47 2. Glutamate—GluR5
Introduction
The HPA axis is the main neuroendocrine arm of the stress response, and is activated in
response to stimuli that threaten the homeostasis of an animal. Central drive of the HPA axis is
mediated by the paraventricular nucleus of the hypothalamus (PVN). When activated, medial
parvocellular PVN neurons release ACTH secretagogues, including CRH and vasopressin (AVP),
into the hypophyseal portal vasculature in the external zone of the median eminence. These
hormones travel through the portal system to the anterior pituitary gland and stimulate
corticotrophs to release ACTH (Vale and River, 1977; Vale et al., 1981). ACTH is then released
into the circulation, and induces the adrenal cortex to synthesize and release glucocorticoid
hormones (corticosterone is the main glucocorticoid in rats).
A significant portion of the excitatory control of HPA axis activity is thought to be mediated by glutamate signaling (Herman et al., 2004; Jezova, 2005). Glutamate signals through
presynaptic and postsynaptic ionotropic and metabotropic glutamate receptors, thereby
regulating neuron excitation and excitability (Brann, 1995). With respect to the HPA axis, glutamate infusion into the third ventricle increases ACTH release, an effect that is blocked by deafferentation of the medial basal hypothalamus (Makara and Stark, 1975). This suggests that glutamate may regulate the HPA axis at the PVN or above. Because injection of glutamate
directly into the PVN also induces ACTH release (Darlington et al., 1989), depletion of median
eminence CRH content (Feldman and Weidenfeld, 1997), and release of CRH from
hypothalamic slices (Joanny et al., 1997), it is likely that at least some of the effects of glutamate
on the HPA axis are mediated through the PVN itself. This idea is supported by evidence that
48 blockade of ionotropic glutamate receptors by injection of the glutamate receptor antagonist
kynurenate into the PVN decreases corticosterone responses to restraint stress (Ziegler and
Herman, 2000).
These HPA axis actions of glutamate are most likely mediated solely by actions on the
central nervous system. Although some glutamate receptor subtypes are expressed in both the
adrenal cortex (Kristensen, 1993) and the pituitary gland (Petralia et al., 1994b; Mahesh et al.,
1999), there is little evidence that glutamate acting on these tissues increases secretion of ACTH or glucocorticoids (Hinoi et al., 2004). Some studies have shown activation of the HPA axis in response to peripherally-administered glutamate agonists (Gay and Plant, 1987; Carlson et al.,
1989), but these effects are likely to have been caused by actions on the central nervous system, rather than on peripheral tissues (Zelena et al., 2005).
It is clear that glutamate acts in regulating the HPA axis at the PVN; however, it is not known which glutamate receptors are responsible for these regulatory actions. Anatomical evidence suggests that receptors from the n-methyl-d-aspartate (NMDA), alpha-amino-3- hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA), and kainate receptor families, as well as metabotropic receptors, are localized in CRH-containing regions of the PVN (Herman et al.,
2000; Scaccianoce et al., 2003; Eyigor et al., 2005; Ziegler et al., 2005). Antagonist studies have demonstrated functional roles for ionotropic (Ziegler and Herman, 2000) and metabotropic
(Johnson et al., 2001; Scaccianoce et al., 2003) glutamate receptors in regulation of the HPA axis. It is not clear, however, which individual receptors from these classes are involved in regulating the HPA axis.
One possibility is suggested by the preferential expression of mRNA from the kainate family of receptors in hypophysiotrophic neurons in the PVN (Herman et al., 2000). In addition,
49 kainate has an excitatory influence on the HPA axis when administered directly into the PVN
(Zelena et al., 2005), suggesting a role for kainate receptors in HPA axis regulation. A possible role of kainate receptors in HPA axis regulation is further supported by increased expression of
GluR5 and KA2 kainate receptors in response to hypoglycemia stress (Koenig and Cho, 2005).
Because GluR5 mRNA and protein are potentially expressed and regulated in the PVN (Herman et al., 2000; Eyigor et al., 2005), the current study investigates the role of GluR5 signaling in regulation of the HPA axis. These experiments were designed to test the hypothesis that GluR5 mediates stress-induced excitation of the PVN and activation of the HPA axis.
Materials and Methods
Animals:
Adult male Sprague-Dawley rats weighing 250-300 g (Harlan, Indianapolis, IN) were
used in these experiments. All experimental protocols were approved by the University of
Cincinnati institutional animal care and use committee, and conform to the guidelines published in the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Care was
taken to minimize the number and suffering of animals used in these experiments. Animals were
kept singly housed, on a 12h:12h light:dark cycle (light phase from 6:00 AM to 6:00 PM) and
given ad libitum access to water and chow. Animals were allowed to adapt to the new
environment for at least one week prior to undergoing surgery. Rats were anesthetized using a mix of ketamine and xylazine (87-90 mg/kg ketamine and 10-13 mg/kg xylazine) and given butorphanol as an analgesic. They were implanted with 26-gauge unilateral chronic guide cannulas (inner diameter = 0.24 mm, outer diameter = 0.46 mm, Plastics One, Roanoke, VA) into
50 the median eminence, or with bilateral guide cannulas into the right and left PVN. Stereotactic
coordinates used for the guide cannulas were (from bregma) PVN: 1.9 mm posterior, 7.3 mm
ventral (from the dura mater at the site of implant), and centered over the sagittal sinus; and
median eminence: 2.2 mm posterior, 0.3 mm lateral, and 9 mm ventral (from the skull at
bregma). For the PVN injections, internal cannulas projected 0.5 mm past the guide cannula.
Internal cannulas for the median eminence injections were 1 mm longer than the guide cannula.
Cannula placement was verified by Nissl stain.
Microinjections and restraint stress challenge:
Starting two days after surgery, the animals were handled daily to accustom them to
having their cannulas handled, removed and re-inserted. On the day of the experiment, five to
seven days after surgery, the obturators were removed from the animal and an internal guide
cannula (33 gauge, inner diameter = 0.10 mm, outer diameter = 0.20 mm) was inserted in the
guide cannula. The animals were given a 100 nl injection of either drug or vehicle (sterile 0.9%
saline with 1% Chicago sky blue dye), using a PHD 2000 motorized syringe pump (Harvard
Apparatus, Holliston, MA). Animals were given either 0.35 μg/μl (35 ng/injection) for the PVN
or median eminence injections or 0.035 µg/µl (low treatment group in the median eminence, 3.5 ng/injection) of LY382884, a GluR5-selective glutamate antagonist. Fifteen minutes post- injection, the animals were placed in plastic restraining tubes, and were restrained for 45 minutes
(30 minutes for the lateral ventricle group). Blood samples were taken by tail clips at 15, 30, 60,
and 120 minutes post-injection for measurement of plasma hormones. Chicago Sky Blue dye
was purchased from Sigma (St. Louis, MO). LY382884 was generously provided by Eli Lilly.
Blood samples were kept on ice until plasma was isolated. Plasma was isolated from blood by
centrifugation, and samples were stored at –20°C until being assayed.
51 Radioimmunoassay:
Plasma corticosterone levels were measured using a commercial 125I radioimmunoassay
kit (ICN, Costa Mesa CA). ACTH radioimmunoassay was performed as previously described
(Engeland et al., 1989), using 125I-labeled ACTH as tracer. The ACTH antiserum was a generous
gift of W. Engeland (University of Minnesota).
Immunohistochemistry:
After the final blood sample was taken from the rats (120 minutes post-injection), they
were killed by barbiturate overdose. Animals were perfused intracardially with 0.9% saline
solution, then with 4% paraformaldehyde in 50 mM potassium phosphate buffer. Brains were
removed and placed in the paraformaldehyde solution and post-fixed overnight at 4°C. After
post-fixation, the paraformaldehyde solution was removed and the brains were kept in 30%
sucrose until sectioning. Perfused brains were frozen on dry ice, then sectioned at 25 µm using a
sliding microtome (Leica, Bannockburn, IL), then stored in cryoprotectant (30 % sucrose, w/v;
1% polyvinylpyrrolidone (PVP-40), w/v; 30% ethylene glycol, v/v; in 50mM sodium phosphate
buffer, pH 7.4) at -20°C until staining.
For sections to be stained using GluR5 antibody, antigen retrieval was performed as
described (Eyigor et al., 2005). Briefly, after being washed in 50 mM potassium phosphate
buffered saline (KPBS), sections were placed in 100 ml plastic beakers containing 50 ml of 50
mM citrate buffer (pH 6.0), then microwaved at high power until the temperature of the buffer
reached 80-90°C (about two minutes for 4 beakers). Sections were allowed to sit in the heated
buffer for 15 minutes, and immunohistochemistry was continued as outlined below. For sections
not stained for GluR5, antigen retrieval was not used.
52 Sections were washed in KPBS, then native peroxide activity was blocked with 1.5%
H2O2 in KPBS for 10 minutes. After washing again, tissue sections were incubated in a blocking
solution (KPBS with 0.1% bovine serum albumin and 0.2% triton X-100). Primary antibodies
were added and sections were incubated overnight at 4°C (48 hours at room temperature for
GluR5). The sections were washed, then incubated in biotinylated secondary antibody (1:500
dilution) for one hour at room temperature, then washed again. For sections visualized using
3,3'-diaminobenzidine (DAB) or amplified using biotinylated tyramide, the tissue was then
treated with ABC (Vector Laboratories, Burlingame, CA) at 1:1000 in KPBS for one hour,
according to the manufacturer’s instructions. Sections immunostained for CRH were further amplified using biotinylated tyramide at 1:250 in KPBS, according to the manufacturer’s
instructions. The sections were then developed in DAB (0.4 mg/ml in KPBS) with hydrogen
peroxide (0.167 μl of 30%/ml KPBS) for 10 minutes, washed, then mounted on glass slides.
Mounted sections were dehydrated in graded ethanol, cleared in xylene, and coverslipped using
DPX mountant (Sigma, St. Louis, MO).
For sections visualized using immunofluorescence, after the incubation with biotinylated secondary antibody, sections were washed, then incubated in streptavidin-conjugated Cy3
(1:500), washed, then mounted on glass slides. For double-stained sections, the second secondary antibody was conjugated to Alexa-488. These slides were coverslipped using gelvatol. Primary antibodies used were rabbit anti-GFAP (DAKO, Carpinteria, CA) at 1:20,000 dilution, rabbit anti-fos (Calbiochem, La Jolla, CA) at 1:20,000, goat anti-GluR5 (Santa Cruz
Biotechnology, Santa Cruz, CA) at 1:1000 (DAB) or 1:100 (immunofluorescence), and rabbit anti-CRH (courtesy Dr. Wylie Vale) at 1:100000. The CRH immunostain was amplified using biotinylated tyramide.
53 In situ hybridization
In situ hybridization was performed as described previously (Ziegler et al., 2002, 2005).
Briefly, 35S-labeled antisense RNA probes were made against a cDNA for GluR5. Tissue
sections mounted on glass slides were hybridized using standard hybridization protocols as
described. Following hybridization and washing, slides were coated with Kodak photographic
emulsion NTB2, developed, and counterstained with cresyl violet Nissl stain. These slides were
then coverslipped using DPX mountant.
Imaging:
DAB-stained slides were photographed using an Axioplan 2 microscope (Zeiss,
Thornwood, NY). c-Fos activation was measured using immunostaining visualized with DAB.
The number of c-Fos positive cells in the PVN was determined by counting particles, using the
ImageJ software (Rasband, 1997-2007). Two sections were counted for each PVN. Confocal
pictures of immunofluorescence-stained slides were taken using an LSM 510 microscopy system
(Zeiss, Thornwood, NY), with an image size of 1024 x 1024 pixels, and 0.48 μm optical slice
depth. Digital images were cropped, and brightness and contrast were adjusted to improve
clarity. No other alterations were made.
Statistics:
Plasma hormone concentrations were analyzed using two-way or three-way analysis of
variance (ANOVA) with repeated measures. c-Fos data were analyzed using one-way ANOVA
or an unpaired t test, as appropriate. Because there were no differences in c-Fos expression in
the right PVN vs. the left PVN, both sides were pooled in each animal for these analyses (except
where otherwise indicated). Outliers were identified using Grubb’s test (Barnett and Lewis,
54 1994). Significant main effects shown upon ANOVA were further analyzed using the Fisher least significant difference post-hoc test. Statistical analyses were performed using GB Stat
(Dynamic Microsystems, Silver Spring, MD) and the Prism software program (GraphPad, San
Diego, CA). Significance was set at p < 0.05.
Results
GluR5 expression
In situ hybridization for GluR5 mRNA in the PVN agreed with previous studies that document expression of GluR5 mRNA in the parvocellular PVN (Figure 1). We performed immunohistochemistry to confirm that GluR5 protein is also expressed in the PVN. Similar to the experience of Eyigor et al. (Eyigor et al., 2005), we found that GluR5 immunoreactivity could not be detected without using an antigen retrieval technique (Figure 2A). We also did not see any immunoreactivity after antigen retrieval in the absence of the anti-GluR5 primary antibody (Figure 2B). However, when the tissues were subjected to antigen retrieval prior to immunostaining, immunoreactivity was revealed. We observed GluR5-like immunoreactivity in both the PVN (Figure 2C) and median eminence, especially in the external lamina (Figure 2D).
Overall, the pattern of expression for GluR5 immunoreactivity was similar to that seen for
GluR5 mRNA expression using in situ hybridization. For example, large numbers of GluR5 immunoreactive cells were detected in regions known to express high levels of GluR5 mRNA, including the nucleus reuniens of the thalamus and hippocampus (not shown).
The presence of GluR5-like immunoreactivity in both the PVN and the external lamina of the median eminence suggested the possibility of co-localization with the ACTH secretagogue
55 CRH. Therefore, we performed dual-label confocal microscopy to determine whether GluR5
immunoreactivity was co-localized with CRH in median eminence terminals. Nearly all of the
nerve terminals in the external lamina of the median eminence that contained CRH also contained GluR5-like immunoreactivity, although not all GluR5-containing terminals contained
CRH (Figure 3A-C).
Injection site confirmation
We confirmed the sites of injection in the PVN and median eminence cannulation
experiments by two methods. First, we injected Chicago Sky Blue dye with the drug or vehicle,
which allowed us to visualize the location of the injection grossly while cutting the brain.
Although the dye was not visible when the tissue was immunostained and mounted on slides, the dye allowed us to confirm the presence and approximate location of an injection. We also used
Nissl stain (Figure 4) to visualize the cannula tracks and injection site. Injections were counted
as a hit if the center of the injection site was within 0.1 mm of the targeted structure (PVN or median eminence). Missed injections were omitted from the analyses of plasma hormones and c-Fos expression.
GluR5 Antagonist Infusion: PVN
Previous data from our group indicates that the corticosterone response to restraint is
blunted by intracerebroventricular injection of LY382884 (van Hooren and Herman, unpublished
observations), suggesting that there is a role for GluR5 in excitatory regulation of the HPA axis.
Since GluR5 mRNA and protein are expressed in the PVN, we hypothesized that paraventricular
GluR5 is responsible for a major part of this excitatory input. To test this hypothesis, we injected
vehicle (n=10) or LY382884 (35 ng per side, n=8) into the PVN via cannula 15 minutes before
56 subjecting the rats to restraint. We then measured plasma ACTH and corticosterone, and c-Fos
expression in the PVN. In contrast to the results of the third ventricle infusion, intra-PVN infusion of LY382884 led to a significant increase in plasma ACTH (F1,71 = 5.38, p < 0.05).
Post-hoc analysis revealed that this increase was significant at the 60 min time point. There was
also a significant main effect of LY382884 treatment on corticosterone (F1,71 = 6.00, p <
0.05)(see figure 5A,B), consistent with an overall increase in the integrated corticosterone response. However, post-hoc tests did not reveal a significant difference at any given point. Fos
protein expression in the PVN was significantly elevated (p < 0.05, see figure 5C) by LY382884
treatment.
GluR5 Antagonist Infusion: Median Eminence
The intense staining of the external lamina of the hypothalamic median eminence with
the GluR5 antibody suggested that there is also GluR5 protein present on nerve terminals there.
Thus, we hypothesized that GluR5 may also influence release of CRH into the portal circulation,
and thus act to modulate activation of the HPA axis at the median eminence. To test this
hypothesis, we implanted rats with cannulas aimed at the median eminence. Rats injected with
either 35 ng (high, n=6) or 3.5 ng (low, n=7) of LY382884, or vehicle (n=7) were subjected to
restraint stress challenge after the injection. LY382884 injection significantly reduced the ACTH
responses to restraint stress (treatment x time interaction F6,51 = 2.356, p < 0.05) at 60 minutes
after the injection (the end of the restraint challenge, Figure 6A). There was also a trend toward
a decreased corticosterone response to the restraint in LY382884 treated animals, but this failed
to reach statistical significance (Figure 6B). There was no effect of the LY382884 treatment on
c-Fos protein expression in the PVN, either between the drug and vehicle treated animals (Figure
57 6C), or between the right and left PVN (not shown). There also did not appear to be a dose- response effect of LY382884 in the median eminence, on either ACTH or corticosterone responses to restraint, at the doses tested.
GluR5 expression in regions projecting to the PVN
The pattern of HPA axis inhibition caused by paraventricular GluR5 blockade suggested that GluR5 may be signaling presynaptically in this nucleus. Therefore, we evaluated the expression of GluR5-like immunoreactivity in areas known to project to the PVN (Silverman et al., 1981; Sawchenko and Swanson, 1983). Table 1 summarizes the areas examined, while
Figure 7 shows photomicrographs of some of these areas, immunostained for GluR5.
Discussion
Previous studies suggested a role for paraventricular glutamate signaling in regulation of the HPA axis (Darlington et al., 1989; Feldman and Weidenfeld, 1997; Ziegler and Herman,
2000; Zelena et al., 2005). The current studies tested the hypothesis that GluR5 mediates effects of glutamate on the HPA axis. Our anatomical data confirm that GluR5 protein is present in both the PVN and median eminence, and co-localizes with CRH immunoreactivity in median eminence terminals, suggesting the capacity for GluR5 signaling in PVN CRH neurons.
Functional tests revealed that injection of a GluR5 antagonist into the PVN increases ACTH and
PVN Fos induction after restraint stress, consistent with an inhibitory role for GluR5 in modulating central activation of the HPA axis. In contrast, injection of LY382884 directly into the median eminence reduces HPA axis stress responses, consistent with an excitatory role for
58 glutamate in ACTH secretagogue release. Together, the results suggest that GluR5 receptors play
a role in both activation and inhibition of HPA axis stress responses.
Glutamate is an excitatory amino acid neurotransmitter, so it is interesting that blocking
its actions in the PVN led to excitation of the HPA axis response to restraint. Reduced HPA axis
activation may be mediated by presynaptic inhibition via GluR5-containing receptors, as has
been seen in other brain regions. In the hippocampus, for example, GluR5 is found
presynaptically (Pinheiro et al., 2005), and presynaptic GluR5-mediated signaling leads to
decreased excitatory activity (Clarke et al., 1997). One possible explanation for this
phenomenon is that GluR5 is an autoreceptor, leading to decreased glutamate release from the
synapse. Presynaptic regulation of neurotransmitter release has been reported in the amygdala
and neocortex, where high concentrations of GluR5 agonists suppress GABA release from
interneurons (Braga et al., 2003; Campbell et al., 2007). If this is the case in the PVN,
presynaptic GluR5 may decrease excitatory neurotransmission by glutamate at PVN synapses.
The result of inhibiting this GluR5-mediated signaling may be to release presynaptic inhibition, leading to enhanced HPA axis activity and enhanced c-Fos activation in the PVN, consistent with our observed data.
An alternative and perhaps more likely explanation is that activation of presynaptic
GluR5-containing receptors increases GABA release onto parvocellular neurons in the PVN.
GluR5-mediated signaling is known to facilitate the release of neurotransmitters from presynaptic terminals (Xu et al., 2006; Aroniadou-Anderjaska et al., 2007; Campbell et al., 2007;
Wu et al., 2007). Therefore, inhibition of presynaptic GluR5 would lead to enhanced HPA axis activation and PVN c-Fos expression by removal of inhibition, consistent with the results that we observed.
59 A role for GluR5 in presynaptic excitation is also consistent with the results of our
LY382884 infusion into the median eminence. Because inhibiting GluR5 in the median
eminence decreased the HPA axis response to restraint, glutamate signaling through GluR5
appears to be excitatory. However, the absence of cell bodies in the median eminence requires
that GluR5 in the median eminence is presynaptic. An excitatory action of GluR5 in the median
eminence suggests that GluR5 signaling here enhances CRH release, similar to its role in
presynaptic facilitation of neurotransmitters release. Thus, the opposing outcomes of GluR5
inhibition in the PVN and median eminence may result from similar mechanisms of action for
GluR5, in regulating GABA and CRH release, respectively. A possible model for actions of
GluR5 in the HPA axis is outlined in Figure 8.
A possible presynaptic role for GluR5 in regulating the HPA axis at the PVN raises the
question of where GluR5-containing inputs into the PVN originate. To address this problem, we
examined the expression of GluR5-like immunoreactivity in areas known to project to the PVN.
Areas examined included those shown in Table 1. Of the regions noted, the bed nucleus of the stria terminalis and dorsomedial hypothalamus are known to regulate HPA axis activity (Evans et al., 2004; Choi et al., 2007) and may be reasonable candidates for regulation of PVN excitation through presynaptic GluR5. Although GluR5 expression in areas known to project to the PVN hints at a possible source of presynaptic GluR5 in the PVN, this evidence is suggestive rather than confirmatory. Further work is needed to determine which of these areas supply GluR5 that is responsible for the inhibitory effects on HPA axis activity that we have observed.
The GluR5-like immunoreactivity in the median eminence suggests a mechanism by which glutamate can excite the HPA axis at the level of the neurosecretory terminal. CRH is the primary hypothalamic secretagogue for ACTH release from the anterior lobe of the pituitary
60 (Vale et al., 1981; Rivier and Vale, 1983), and is secreted into the hypophyseal portal vasculature
in the external lamina of the median eminence. The co-localization of GluR5 with CRH in the
external lamina suggests that presynaptic GluR5 may regulate CRH release, such that glutamate
signaling through GluR5 in the median eminence enhances activity of the HPA axis. The fact
that GluR5 antagonist injection into the median eminence reduced HPA axis stress responses
without affecting PVN c-Fos induction further supports a local action of GluR5 signaling at the
nerve terminal.
The lack of a dose-response to LY382884 in the animals injected into the median
eminence suggests that the dose of LY382884 given in these experiments may be higher than the
maximally efficacious concentration. The incomplete inhibition of HPA axis activation by local
LY382884 treatment suggests that glutamate effects mediated by GluR5 modulate stress
excitation of the HPA axis, but are not required for the initiation of the response.
Our studies could not determine whether drugs injected into the median eminence were able to diffuse out of the median eminence and reach the pituitary gland. This could potentially present a problem, as there are glutamate receptors expressed in the anterior pituitary (Hinoi et al., 2004), and glutamate binds to the anterior lobe of the pituitary, albeit weakly (Meeker et al.,
1994). However, kainate does not compete with glutamate for binding to pituitary membranes in binding assays (Yoneda and Ogita, 1989), suggesting that functional kainate receptors, including
GluR5, are not found in the anterior pituitary. Additionally, peripherally administered glutamatergic drugs that affect the HPA axis are thought to exert these effects via brain areas deficient in blood-brain barrier, rather than through the pituitary gland (Jezova, 2005).
Therefore, even if some of the LY382884 did diffuse to the pituitary gland, it does not seem likely that it affected HPA axis signaling in the pituitary gland. We can not, however, completely
61 rule out effects on the pituitary, nor on structures near the median eminence, such as the arcuate
nucleus.
The external lamina of the median eminence contains axon terminals from neurons
located almost exclusively in the PVN and other parts of the hypothalamus (Wiegand and Price,
1980), although there may be afferent fibers from the medial septum and diagonal band of Broca
(Silverman et al., 1987). There is evidence that CRH-expressing neurons in the PVN also
express VGLUT2 protein in the median eminence (Hrabovszky et al., 2005), suggesting that glutamate acting on GluR5 in the median eminence could be released by CRH nerve terminals.
Indeed, neuropeptidergic neurons almost uniformly co-release classical neurotransmitters, including glutamate (Hokfelt et al., 2000). This suggests that presynaptic glutamate signaling
through GluR5 in the median eminence could be acting as a feed-forward signal in activation of
the HPA axis response to stress.
In conclusion, the current report indicates novel, GluR5-mediated presynaptic
mechanisms for glutamatergic regulation of the HPA axis. Our data support a role for kainate-
preferring receptors in gating of CRH release at median eminence terminals, as well as in local
inhibition of PVN activation. Presynaptic modulation of HPA activation provides a mechanism
whereby release of CRH can be adjusted at the level of the nerve terminal via glutamatergic
projections to the median eminence. Importantly, the localized blood-brain barrier deficiency in
the median eminence provides a potential route for specific pharmacological modulation of HPA
stress reactivity in disorders characterized by glucocorticoid hypersecretion.
62 Figure Legends
Figure 1. GluR5 expression in PVN by in situ hybridization. A) Bright field image of the
PVN on an emulsion-dipped slide after in-situ hybridization. The PVN is outlined. Scale bar
represents 100 μm. B) Higher magnification of the boxed area in A) above. Cells associated
with silver grains representing GluR5 mRNA expression are indicated with arrowheads. The
third ventricle is indicated as 3V. Scale bar represents 50 μm.
Figure 2. Antigen retrieval. Antigen retrieval was necessary in order for GluR5-like
immunoreactivity to be detected. A) Section of the hypothalamus at the level of the PVN stained
by DAB immunohistochemistry for GluR5, without antigen retrieval. B) Immunostained
section at the same level without primary antibody, for comparison. C) Section immunostained
for GluR5, after antigen retrieval. D) Section immunostained for GluR5, at the level of the
median eminence. 3V- third ventricle. Scale bar indicates 100 μm.
Figure 3. GluR5/CRH co-localization. Tissue was double-labeled for GluR5 and CRH, and
imaged using confocal microscopy. Pictures shown are photomicrographs of the median
eminence, taken at 400 X magnification and are 0.48 μm optical slices. GluR5 is colored green
and CRH is red. A) GluR5-like immunoreactivity in the median eminence. B) CRH
immunoreactivity. C) Overlay. Areas co-expressing GluR5 and CRH are colored yellow.
Terminals co-expressing CRH and GluR5 are indicated with arrowheads. Scale bar represents 20
μm.
63 Figure 4. PVN cannula placement. Cannula placement was verified by microscopic
examination of Nissl stained slices. A) Photomicrograph of Nissl stained section, showing the
injection site for a PVN injection. The track made by the guide cannula is indicated with an
asterisk, and the injection site is outlined. B) Photomicrograph of Nissl stained section, showing
the injection site for a median eminence injection. The injection site is outlined. 3V- third
ventricle. The scale bar represents 100 μm.
Figure 5. PVN injection of LY382884. Animals were injected bilaterally into the PVN with
LY382884, and subjected to restraint stress. A) Plasma ACTH levels. There was a significant main effect of LY382884 treatment on plasma ACTH. B) Plasma corticosterone levels. There was a significant main effect of LY382884 treatment on corticosterone, although post-hoc tests did not reveal significant differences at any individual time points. C) c-Fos immunoreactive cells in the PVN. Results are expressed as mean ± SEM. * denotes p < 0.05 vs. vehicle.
Figure 6. Median eminence injection of LY382884. Animals were injected with LY382884
into the median eminence, and subjected to restraint stress. A) Plasma ACTH levels. LY382884
treatment significantly attenuated the ACTH response to restraint stress at 60 minutes after the
injection, vs. vehicle, in both low and high treatment groups. B) Plasma corticosterone levels.
There was no significant main effect of LY382884 treatment on the corticosterone response. C)
Particle counts from the PVN of c-Fos-immunostained slices. There were no significant
differences between treatment groups. Low treatment group rats received 3.5 ng of LY382884, while high treatment group rats received 35 ng. There was no significant difference between these two doses. Results are expressed as mean ± SEM. * denotes p < 0.05.
64
Figure 7. Expression of GluR5 in brain areas with glutamatergic projections to the PVN.
Immunohistochemistry for GluR5 was performed, and areas with known glutamatergic afferents
to the PVN were examined. A) Ventral bed nucleus of the stria terminalis. B) Medial Preoptic
Area. C) Subfornical organ. D) Dorsomedial hypothalamic nucleus. E) Ventromedial hypothalamic nucleus. F) Lateral Hypothalamus. ac, anterior commissure; 3V third ventricle; ot
optic tract; zi, zona incerta; fx, fornix. Scale bars indicate 100 um.
Figure 8. Model of GluR5-mediated effects on the HPA axis. GluR5 activation in the PVN
and at the median eminence have opposite actions on activation of the HPA axis. We propose
that this is due to presynaptic GluR5 activation leading to enhanced release of neurotransmitters;
GABA at the PVN proper and CRH at the median eminence. A) Activation of GluR5 in presynaptic terminals in the PVN leads to increased GABA release onto parvocellular neurons in
the PVN, leading to inhibition of the neuron. The net effect of GluR5 signaling in this area is a decreased HPA axis response to stress. B) Activation of GluR5 in presynaptic terminals in the median eminence leads to enhanced release of CRH into the portal capillaries, leading to enhanced activation of the HPA axis.
65 Figure 2-1 GluR5 mRNA in the PVN
66 Figure 2-2 Antigen Retrieval
67 Figure 2-3 GluR5/CRH Immunohistochemistry
68 Figure 2-4 Cannula Placement
69 Figure 2-5 LY382884 in the PVN
70 Figure 2-6 LY382884 in the Median Eminence
71 Figure 2-7 GluR5 Protein Expression in Areas Projecting to the PVN
72 Figure 2-8 Model for GluR5 Signaling at the PVN
73 Table 2-1 GluR5 Expression in Areas Projecting to the PVN Brain region GluR5 expression Notes Arcuate - BST (anterior) ++ BST (ventral) ++ Central Amygdala ++ Dorsal Medial Hypothalamus ++ Lateral Hypothalamus -/+ only in some scattered cells Lateral Septum ++ Medial Septum +++ Medial Amygdala +++ Median Preoptic Area +++ Subfornical Organ +++ Supraoptic Nucleus + not in cell bodies Ventromedial Hypothalamus -/+ only in some scattered cells Suprachiasmatic Nucleus -
Table 1 Expression of GluR5 in sources of afferent glutamatergic innervation to the PVN.
Areas that send afferent fibers to the PVN and express either VGLUT1 or VGLUT2 were assessed for their content of GluR5-like immunoreactivity. The amount of GluR5 immunoreactivity was scored qualitatively, from – (no reactivity) to +++ (heavy reactivity).
74 3. Glucocorticoids and Cannabinoids
Introduction
Like many other endocrine axes, the hypothalamus-pituitary-adrenal (HPA) axis is
regulated in part by negative feedback. Negative feedback regulation of the HPA axis occurs on
fast and slow time frames (Keller-Wood and Dallman, 1984). Fast feedback occurs with
glucocorticoid exposures of less than about 10 minutes, and has thus been assumed to be mediated by non-classical, nongenomic signaling pathways. In addition, there is evidence that rapid inhibition of the HPA axis by glucocorticoids on this fast time scale occurs independently of protein and mRNA synthesis (Dayanithi and Antoni, 1989; Hinz and Hirschelmann, 2000).
This contrasts with classical glucocorticoid signaling, which involves binding of glucocorticoids to cytosolic receptors which then influence gene expression by interaction either with glucocorticoid response elements in gene promoters or with other transcription factors (Heitzer et al., 2007).
Fast feedback in the HPA axis was first demonstrated in the 1940s (Sayers and Sayers,
1947), and since then several groups have investigated this phenomenon. It is established that repeat stressing of an animal as early as 2 minutes after a previous stressor leads to profoundly inhibited adrenocorticotropic hormone (ACTH) secretion to the second stressor (Sakakura et al.,
1976), and that treatment of animals with glucocorticoids before stressing them inhibits the corticosterone response to the stressor (Yates et al., 1961). However, the site(s) of action of these effects and the signaling pathways that operate in causing this effect are not entirely clear. In vitro evidence suggests that fast feedback occurs at the pituitary gland (Widmaier and Dallman,
1984; Abou-Samra et al., 1986) and hypothalamus (Edwardson and Bennett, 1974; Jones and
75 Hillhouse, 1976), independent from new gene expression in both cases. In vivo, treatment of animals with glucocorticoids inhibits corticotropin releasing hormone (CRH)-induced ACTH release (Hinz and Hirschelmann, 2000), suggesting that fast feedback does occur at the pituitary gland. In the case of the hypothalamus, there is electrophysiological evidence that fast feedback may occur through inhibition of glutamate release onto CRH containing cells in the paraventricular nucleus of the hypothalamus (PVN) in a manner that is dependent on signaling through cannabinoid CB1 receptors (Di et al., 2003). However, it is not yet clear whether this inhibition of glutamate release is associated with inhibition of CRH release or with reduced activity of the HPA axis.
We undertook the current studies to test the hypothesis that glucocorticoid-mediated negative feedback occurs at the PVN. We further attempted to determine whether any rapid alterations of the HPA axis by glucocorticoids are mediated through CB1 receptor signaling at the
PVN.
Materials and Methods
Animals:
Adult male Sprague-Dawley rats (Harlan, Indianapolis, IN) weighing between 275 and
350 g were used for all the described studies. Animals were singly housed and kept in temperature- and humidity-controlled rooms with a 12h:12h light:dark cycle (lights on at 6:00
AM). Animals were given ad libitum access to water and standard rat chow. After arrival in the lab, rats were allowed to adapt to the new surroundings for at least 1 week before surgery or other experiments were performed. All procedures were done in accordance with the National
76 Institutes of Health Guide for the Care and Use of Laboratory Animals, and were approved by
the University of Cincinnati Institutional Animal Care and Use Committee.
Cannula Surgeries:
Rats were anesthetized using ketamine (90 mg/kg) and xylazine (10 mg/kg), and given
butorphanol as a preemptive analgesic. Under anesthesia, they were implanted with 26 gauge
bilateral guide cannulas (1.0 mm separation between sides, center to center, Plastics One,
Roanoke, VA). Stereotactic coordinates used were 1.9mm posterior from bregma, centered over
the sagittal sinus, and 6.3 mm ventral from the dura mater (final depth for the injector is 7.3
mm). After surgery, the animals were allowed to recover for at least 5-7 days before restraint
challenge. Beginning 2 days after surgery, the cannulas were handled daily to maintain patency
of the guide, and to accustom the rats to handling. After the stress challenges, cannula placement
was verified using a cresyl violet Nissl stain.
Drugs:
Dexamethasone 21-phosphate was purchased from Sigma (St. Louis, MO), and dissolved
in 0.9% saline, then diluted into the appropriate vehicle for the experiment. AM251 was
purchased from Tocris (Ellisville, MO), and dissolved in dimethyl sulfoxide (DMSO) to make a
stock solution, then diluted into vehicle (10% DMSO in saline for peripheral injection, and 0.4%
DMSO in saline for intra-PVN injections). SR141716 was obtained from the NIDA drug supply program (Research Triangle Park, NC) and dissolved in DMSO, then diluted to final concentration in saline (1:10 dilution for 10% DMSO final). Neither SR141716 nor AM251
77 were soluble in 10% DMSO at 4 mg/kg, so for the intraperitoneal injections, the drugs were
diluted immediately before administration, and the resulting slurry was injected.
Restraint Challenge:
Restraint challenges were performed in the morning, between 8:00 and 12:00 AM (the
nadir of circadian HPA axis activity). Each animal was brought from the animal housing room
into the procedure room immediately before beginning the restraint challenge. Rats were given intraperitoneal or intra-PVN injections, then subjected to restraint for 30 minutes by being placed in transparent Plexiglas tubes. Blood samples (~300 μl) were taken by tail clip at the indicated times. At the end of the restraint challenge, rats were given lethal injections of Fatal Plus, then perfused with 3.7% formaldehyde in potassium phosphate buffer (KPB, 50 mM potassium phosphate, pH 7.4), after which brains were collected for immunohistochemistry.
Radioimmunoassay:
Plasma corticosterone was measured using a commercial radioimmunoassay kit (MP
Biomedicals, Solon, OH). ACTH was measured by 125I radioimmunoassay, as described
(Engeland et al., 1989). The ACTH antibody used was a generous gift of W. Engeland.
Imaging
Nissl-stained sections were photographed using an Axioplan 2 microscope (Zeiss,
Thornwood, NY). Micrographs were cropped, and brightness and contrast were adjusted when
needed, but no other manipulations were performed on digital images.
78 Statistics:
Plasma hormone data were analyzed by two- or three-way ANOVA, with repeated
measures for the time-course data. Area under the curve (AUC) calculations were made using the
trapezoidal method and analyzed by ANOVA. Hormone levels were square root or log10 transformed if necessary to obtain homogeneity of variance, and significant main effects and interactions were further examined using Fisher's LSD post-hoc tests. Because our intention was only to compare the treatments against each other, we decided a priori to only compare treatments within each time point. Fos data were analyzed by two-way ANOVA, with main effects further analyzed with Fisher's post-hoc test. Significance was set at p < 0.05. Nonlinear regression analysis for the radioimmunoassays was performed using AssayZap (Biosoft,
Cambridge, UK) and Prism (GraphPad, San Diego, CA) software packages. Statistical analysis was done using GBStat and Prism.
Results:
Cannula Placement
After each of the experiments described, cannula placement was confirmed by Nissl stain
(Figure 1). Only animals with correctly placed cannulas were included in the analyses of data in
these studies.
Intra-paraventricular dexamethasone rapidly inhibits the HPA axis
Published data suggest that glutamatergic signaling into the CRH-containing
79 parvocellular PVN is rapidly reduced by glucocorticoids in the PVN (Di et al., 2003). However,
it has been unclear whether this alteration in glutamate signaling has any functional significance
on activation of the HPA axis. In order to test the hypothesis that glucocorticoid treatment
delivered locally to the PVN leads to fast negative feedback inhibition of the HPA axis response
to stress, we injected bilaterally cannulated rats with dexamethasone (10 ng per side) or dex:BSA
(1.25 μg per side) before subjecting the animals to restraint stress challenge. Animals treated
with dexamethasone or dex:BSA had significantly decreased ACTH (main effect of treatment,
F2,107 = 4.117, p < 0.05) and corticosterone (main effect of treatment, F2,107 = 12.869, p < 0.05)
time course responses to restraint. There was a trend toward reduced ACTH AUC (F2,24 = 2.951,
p = 0.07) and a significant reduction in corticosterone AUC (F2,25 = 4.785, p < 0.05). Figure 2
illustrates the results of this experiment.
Fast Feedback with peripheral administration of AM-251
Decreased glutamatergic signal caused by dexamethasone treatment in hypothalamic slices is mediated by endocannabinoid signaling through CB1 receptors (Di et al., 2003).
Therefore, we attempted to block dexamethasone-induced fast feedback in the PVN by pre-
treating rats with the CB1 antagonist AM-251, 15 minutes before administration of
dexamethasone into the PVN. Consistent with our earlier results, dexamethasone inhibited the
time course ACTH response to restraint (main effect of dexamethasone F1,131 = 13.498, p < 0.05),
as well as the AUC (main effect of dexamethasone F1,29 = 23.059, p < 0.05). The effect on corticosterone did not achieve statistical significance in the time course or AUC. AM-251 alone did not affect HPA axis responses to restraint, and did not block the dexamethasone-mediated
HPA axis inhibition. Figure 3 summarizes these results.
80 Time dependence of AM-251 effects
In contrast with our results, other studies have suggested that peripheral infusion of CB1 antagonists leads to potentiation of the HPA axis response to restraint (Ginsberg et al., 2006).
Comparison of our data with others suggested that there is a time dependence of this effect.
Since administration of AM-251 failed to potentiate the HPA axis responses to restraint, we gave intra-peritoneal injections of the CB1 receptor antagonists AM-251 or SR141716A (4mg/kg) then
subjected rats to restraint for 15 minutes. At the end of the restraint, animals had elevated ACTH
responses to the restraint (main effect of CB1 antagonist treatment F2,53 = 17.436, p < 0.05).
There was also a significant elevation of the corticosterone response in the antagonist-treated
animals (antagonist x time interaction F2,53 = 3.826, p < 0.05). Figure 4 illustrates the results of
this experiment.
Intra-PVN infusion of AM-251 blocks fast feedback
Although the results of the previous experiment suggested that there is a strong time
dependence of the effects of CB1 receptor antagonism, the potentiation of HPA axis response to
restraint presents a strong confound to any effect of AM-251 treatment on dexamethasone-
induced fast feedback. In order to mitigate this confound, we examined the effect on fast
feedback of AM-251 treatment given locally at the PVN. We mixed dexamethasone and AM-
251 (20 pmol per side) and injected the cocktail immediately before the onset of restraint stress.
AM-251 treatment alone had no effect on either ACTH or corticosterone responses, suggesting
that local administration of this drug into the PVN avoids the confounding HPA axis excitation
seen with a peripheral administration of this drug. Dexamethasone alone caused a significant
inhibition of the ACTH response to restraint (main effect of dexamethasone treatment F1,123 =
81 10.156, p < 0.05), as well as the AUC of the ACTH response (main effect of dexamethasone
treatment F1,30 = 5.929, p < 0.05). Likewise, there was a main effect of dexamethasone treatment on the time course of corticosterone secretion (F1,127 = 7.328, p < 0.05) and a
dexamethasone x AM-251 interaction on the area under the curve of corticosterone secretion
(F1,31 = 5.326, p < 0.05). AM-251 completely blocked the dexamethasone-induced inhibition of
the HPA axis response to restraint. Figure 5 shows these results.
CB1 agonism is not sufficient to cause fast feedback
Since inhibition of CB1-mediated signaling blocked dexamethasone-induced negative
feedback signaling, we attempted to mimic the dexamethasone-mediated effect using the
cannabinoid receptor agonist Win 55,212-2 (20 pmol/side). Contrary to our predictions, agonist
treatment elevated the ACTH time-course response to restraint (Main effect of Win, F1,127 =
5.813, P < 0.05). There was also a main effect of Win treatment on the AUC of the ACTH response (F1,28 = 6.287, p < 0.05). There was not a main effect of Win treatment on either the
corticosterone time course or the AUC; however, in both cases, the corticosterone secretion in
the Win-treated group was significantly higher than the response in dexamethasone-treated
animals (main effect of dexamethasone treatment F1,127 = 5.081, p < 0.05). In addition,
dexamethasone treatment reduced the AUC in both ACTH and corticosterone vs. Win 55,212-2,
when combined with Win 55,212-2 treatment. Figure 6 summarizes the results of these studies.
Discussion
The current studies demonstrate that fast negative feedback regulation of the HPA axis
82 response to restraint is mediated by glucocorticoid signaling in the hypothalamus. CB1-mediated endocannabinoid signaling is necessary for this fast negative feedback to occur. Fast feedback apparently is initiated by glucocorticoid actions at a membrane glucocorticoid receptor, and occurs on a rapid time scale.
Treatment of animals with dexamethasone directly in the PVN led to a rapid inhibition of the HPA axis response to restraint. This effect was completely reversed by co-treatment of the animals with the CB1 selective inhibitor AM-251, suggesting that CB1 receptor signaling is necessary for this fast feedback pathway to be activated. This is consistent with the model of fast feedback signaling suggested by Di et al (Di et al., 2003). In this model, nongenomic glucocorticoid actions at a CRH containing neuron in the PVN lead to activation of endocannabinoid synthetic enzymes. Endocannabinoids produced in response to the glucocorticoid signal act on presynaptic endocannabinoid receptors to reduce the release of glutamate from the presynaptic terminal. Reduced glutamatergic tone onto the CRH neuron leads to decreased excitation-release coupled CRH release, and thus to decreased HPA axis activity in response to a stressor. This model is consistent with known actions of endocannabinoids in reducing neurotransmitter release from axon terminals (Hoffman and
Lupica, 2006).
Interestingly, blocking of fast feedback by AM-251 was not observed with peripheral treatment of animals with AM-251 20 minutes before the onset of restraint. It is not clear whether the difference between our two experiments with AM-251 was caused by peripheral vs. intra-PVN infusion, or by the different time of application. However, there is evidence that there are time-dependent differences in the effects of CB1 receptor antagonism on the HPA axis response to restraint stress (Ginsberg et al., 2006). Further, the differences we saw between the
83 HPA axis response to restraint between animals given intraperitoneal AM-251 20 minutes vs. 2
minutes before the onset of restraint appears to confirm this time-dependent effect of CB1
antagonism. The mechanism behind this time-dependent effect of CB1 antagonism is not clear.
One possibility is that the neuronal system rapidly adapts to the loss of cannabinoid tone, thus
rapidly reversing the effects of CB1 antagonism. More work is necessary to elucidate the
mechanism of this time-dependent effect.
In these experiments, the CB1 antagonist AM-251 blocked the fast feedback effect of
dexamethasone, while the CB1 agonist Win 55,212-2 failed to mimic the effects of
dexamethasone. Thus, CB1 receptor signaling is necessary for fast feedback inhibition of the
HPA axis at the PVN, but is not sufficient for this effect, at least in this experimental system.
One explanation for this phenomenon is that the CB1 receptor could be involved in both
excitatory and inhibitory inputs into CRH-containing cells in the PVN. Presynaptic
endocannabinoid signaling typically leads to decreased neurotransmitter release from the affected synapse (Hoffman and Lupica, 2006). Dexamethasone treatment leads, through endocannabinoid signaling, to decreased glutamatergic inputs into parvocellular neurons (Di et
al., 2003). However, GABAergic synapses also contain CB1 receptors in the PVN, and GABA
release is decreased by CB1 signaling in the PVN (Castelli et al., 2007; Oliet et al., 2007). Our
results with Win 55,212-2 support the existence of a tonic “braking” system in the hypothalamus,
which is negatively regulated by endocannabinoid signaling, but not by rapid glucocorticoid
actions. This inhibitory system lacks disinhibitory endocannabinoid tone during restraint, as
evidenced by the lack of effect of AM-251 alone on the HPA axis responses. However, when a
cannabinoid agonist is introduced, the activation of CB1 leads to inhibition of GABA release and
thus disinhibition. The magnitude of the disinhibitory effect on GABA synapses appears to be
84 greater than the inhibitory effect on glutamatergic synapses, thus leading to an enhanced HPA
axis response. Figure 7 illustrates this proposed model for actions of dexamethasone and Win
55,212-2 in HPA axis regulation at the PVN.
A second possible explanation for this disconnect between the actions of CB1 agonist and
antagonist in the PVN is that the effects of Win 55,212-2 might be mediated through CB2, since
this compound is not specific for CB1 (Felder et al., 1995). CB1 is generally considered to be the
major cannabinoid receptor in the central nervous system, although CB2 is also known to be
expressed in some parts of the brain (Nunez et al., 2004; Onaivi et al., 2006). Although CB2 has
been reported to be absent from the PVN (Gong et al., 2006), it is probably expressed at low
levels elsewhere in the hypothalamus (Onaivi et al., 2006). Thus, it is possible, if unlikely, that
CB2 in the PVN or peri-PVN region is responsible for the excitatory effects of Win 55,212-2 on the HPA axis.
CB1 is located almost exclusively in the presynaptic terminals of central nervous system neurons (Freund et al., 2003). Thus, it is very likely that effects on the HPA axis mediated by
CB1 are through a presynaptic mechanism. However, presynaptic regulation of the HPA axis by
CB1 raises the question of where the CB1 containing projections to the PVN originate. There are
a number of areas that are known to project to the PVN, including hypothalamic areas such as the dorsomedial hypothalamus, ventromedial hypothalamus, bed nucleus of the stria terminalis, and lateral hypothalamic area (Silverman et al., 1981). If fast feedback is controlled at glutamatergic synapses, it is likely that the neurons supplying the CB1 innervation also express
vesicular glutamate transporters. Thus, some areas that may be supplying these fibers to the
PVN could include the lateral hypothalamus, bed nucleus of the stria terminalis, ventromedial
hypothalamus, dorsomedial hypothalamus, and lateral septum (Kaneko et al., 2002; Ziegler et al.,
85 2002). All of these are known or suspected to play a role in regulating or modulating the HPA
axis response to stress (Suemaru et al., 1995; Thrivikraman et al., 2000; Evans et al., 2004; Choi
et al., 2007). The lateral septum, lateral hypothalamus, bed nucleus of the stria terminalis, and paraventricular nucleus also express CB1 (Tsou et al., 1998), and so are candidates for further
study.
Because dex:BSA treatment was sufficient to cause rapid inhibition of the HPA axis
response to restraint, it is possible that this response in the PVN is mediated by a membrane-
bound receptor for glucocorticoids. BSA-conjugated steroids have been used as non-membrane-
permeable compounds in a variety of studies, both with glucocorticoids (Di et al., 2003) and
other steroids such as estrogen (Pappas et al., 1995). These steroids are extensively purified by
the supplier, and are thought to be stable in vitro, although this does not appear to have been
directly tested. The approximate half life for hydrolysis of hydrocortisone 21-hemisuccinate at
approximately physiological temperature and pH is >24 hours (Garrett, 1962). Further, large
molecules attached to a steroid by a hemisuccinate linker appear to sterically hinder enzymatic
hydrolysis of the hemisuccinate linker (Mehvar et al., 2000). Thus, even if dex:BSA is not
entirely stable in vivo, the amount of dexamethasone released within 15 minutes of injection of
dex:BSA is likely to be very small. However, we cannot claim with absolute certainty that the
fast feedback effects of dexamethasone in the PVN are mediated by a membrane bound receptor,
rather than by a receptor that could be acted on by free dexamethasone liberated from the
dex:BSA compound. Further studies are required to confirm the role of a membrane receptor for
glucocorticoids.
We have postulated that the fast feedback effects of dexamethasone occur at least in part
at the PVN. The PVN is the most important integration point for relaying central stress-response
86 signaling to the pituitary-adrenal axis (Herman et al., 2002). However, the size of our injection would lead to diffusion of the drugs beyond the PVN proper. Thus, it is possible that these fast feedback responses have a location of action other than the PVN. Based on the response of missed dexamethasone injections, however, it appears that the site of action must be close to the
PVN, such as in the peri-PVN (data not shown). Further, based on the spread of dex:BSA from the injection site, as measured by anti-BSA immunohistochemistry, we can rule out the pituitary as a site for this fast feedback activity (data not shown).
We have noted throughout these studies that the magnitude of the change in ACTH secretion in response to dexamethasone treatment is greater than that of the corticosterone responses. It is not clear why the magnitude of the corticosterone response is not greater than that of the ACTH response, as would be expected, but there are several possible explanations for this. One possibility is that the corticosterone response has reached a ceiling; that is, the greater
ACTH secretion seen in vehicle treated animals is unable to cause greater corticosterone secretion because the animal is not physiologically capable of secreting more corticosterone.
However, past studies from our laboratory have measured much higher corticosterone responses than what we have observed in the current studies (we have observed corticosterone levels of around 800-900 ng/ml, compared with the peaks in these experiments of 500-600 ng/ml.
(Ostrander et al., 2006).
Alternatively, our sampling times could have missed the peak corticosterone secretion. If this is the case, then we might have observed greater effects with a different sampling schedule.
While we can not rule out this explanation, it seems unlikely that a large difference in corticosterone secretion could be hidden inside a time of ~30 minutes, which is the longest window that is available for this in our time course.
87 A third explanation for this phenomenon is that there are compensatory changes in the
HPA axis that mitigate the changed ACTH secretion. This seems to be a likely explanation, since
our experimental design results in a localized increase in glucocorticoid concentrations at the
hypothalamus, but not in other areas. Since many locations are known to be involved in regulating the HPA axis, including the pituitary (Dayanithi and Antoni, 1989; Hinz and
Hirschelmann, 2000), hypothalamus (Herman et al., 2003), hippocampus (Feldman and
Weidenfeld, 1999), paraventricular thalamus (Jaferi and Bhatnagar, 2006), and prefrontal cortex
(Akana et al., 2001), it seems reasonable to suspect that some of these areas may be involved in compensating for a decreased ACTH response to stress. This could occur through changes in adrenal sensitivity to ACTH, such as that caused by sympathetic signals to the adrenal through the splanchnic nerve (Jasper and Engeland, 1997).
In conclusion, we present evidence that glucocorticoid signaling within the PVN leads to fast feedback inhibition of the HPA axis response to restraint stress in a time frame that is consistent with nongenomic glucocorticoid signaling. CB1-mediated signaling is necessary for
this fast feedback to occur, and this pathway may be initiated by glucocorticoid actions on a
membrane bound receptor for glucocorticoids.
88 Figure Legends
Figure 1. Bilateral cannula placement in the PVN. Cannula placement was verified using
Nissl-staining. The injection locations are outlined by dashed lines, and the end of the guide cannulas are indicated with asterisks. The scale bar indicates 100 μm.
Figure 2. Intra-PVN administration of dexamethasone causes rapid inhibition of the HPA axis response to restraint. Animals were given bilateral intra-PVN injections of dexamethasone
(10 ng per side) or dex:BSA (1.25 μg per side), and plasma hormone responses to 25 minutes of restraint stress were measured. A) ACTH response to restraint. B) AUC for ACTH response. C)
Corticosterone response to restraint. D) AUC of corticosterone response. * p < 0.05 vs. vehicle.
Figure 3. Peripheral administration of CB1 antagonist does not block fast feedback.
Animals were pretreated with the CB1 receptor antagonist AM-251 (5 mg/kg) by intraperitoneal injection 20 minutes before the onset of restraint stress. A, B) Dexamethasone significantly decreased the ACTH response to restraint, both in the time course and area under the curve.
Peripheral AM-251 did not block fast negative feedback by dexamethasone. C, D) There was not a statistically significant effect of any treatment on the corticosterone response to restraint, either in the time course or the integrated response. * p < 0.05 vs. vehicle. AUC, area under the curve.
Figure 4. Effects of CB1 antagonism on HPA axis are time dependent. In contrast to treatment with CB1 antagonists 20 minutes prior to restraint, treatment with SR141716 or AM-
251, 2 minutes before restraint led to robust potentiation of the HPA axis response to restraint within 15 minutes. A) ACTH response to restraint. B) Corticosterone response. * p < 0.05 vs.
89 vehicle treated group.
Figure 5. CB1 receptor signaling is necessary for fast feedback. Animals were treated with dexamethasone, the CB1 receptor antagonist AM-251, or both, then subjected to 30 minutes restraint. Plasma ACTH and corticosterone responses were measured for 60 minutes after the injection. A) Plasma ACTH response to restraint. B) Total magnitude of ACTH response, expressed as area under the curve. Dexamethasone treatment decreased ACTH secretion vs. dexamethasone/AM-251 treatment. C) Plasma corticosterone response to restraint. D) Area under the curve of corticosterone secretion. Dexamethasone treatment decreased the corticosterone response relative to all other groups. * p < 0.05 vs. vehicle; † p < 0.05 vs. dexamethasone; ‡ p <
0.05 vs. all other treatments.
Figure 6. CB receptor agonism potentiates the HPA axis response to restraint. Animals were treated with the CB receptor agonist Win 55,212-2 and/or dexamethasone, then subjected to restraint stress. Win 55,212-2 treatment significantly elevated the ACTH response to stress. Win
55,212-2 treatment also resulted in a significantly higher corticosterone AUC response than dexamethasone, while the combination of both of these led to a similar AUC response to dexamethasone treatment alone. A) ACTH time course response to restraint. B) AUC of the
ACTH response. C) Corticosterone time course response. D) AUC of the corticosterone response. * p < 0.05 vs. vehicle within time points. Significant differences in the AUC responses are indicated with lines.
Figure 7. Model for cannabinoid-mediated actions of the PVN on the HPA axis. CB1
90 agonists and antagonists applied directly to the PVN do not have opposite actions; both appear to
have “excitatory” actions. One possible explanation for this is that dexamethasone-mediated fast
feedback acts through excitatory glutamatergic synapses, while CB1 receptor-mediated signaling
regulates both glutamatergic and GABAergic inputs into the parvocellular PVN. A) Under glucocorticoid treatment, CB1 receptor signaling leads to decreased glutamate release onto the
CRH containing neuron. This leads to decreased CRH release from the cell. Blocking CB1 receptors leads to normalization of the glutamate release, and thus, normalization of the CRH output of the cells. B) CB1 agonism, like dexamethasone treatment, leads to a decrease in
glutamate release from excitatory synapses. In addition, GABA release from inhibitory synapses is also reduced. These GABA synapses contain CB1, but do not experience tonic
endocannabinoid tone under the conditions in our experiments. The net effect of this signaling is
disinhibition of the neuron, leading to increased CRH output.
91 Figure 3-1 Cannula Placement
92 Figure 3-2 Fast Feedback in the PVN
93 Figure 3-3 Peripheral AM-251 Does Not Block Fast Feedback
94 Figure 3-4 Potentiation of HPA Axis Response After CB1 Blockade
95 Figure 3-5 AM-251 Blocks Fast Feedback
96 Figure 3-6 Win 55,212-2 in Fast Feedback
97 Figure 3-7 Model for Fast Feedback Signaling
98 4. Glutamate—group I metabotropic glutamate receptors
Introduction
Glutamate is the major excitatory neurotransmitter in the central nervous system, and plays a role in regulating neuroendocrine systems such as the hypothalamus-pituitary-adrenal
(HPA) axis (Brann, 1995). Glutamate signaling in the paraventricular nucleus of the
hypothalamus (PVN) is known to play a role in regulation of HPA axis responses to stress. For
example, direct infusion of glutamate into the PVN leads to increased ACTH secretion
(Darlington et al., 1989), suggesting an excitatory role for glutamate in regulating the HPA axis
at the level of the PVN. However, there are a great number of receptors in the glutamate receptor
family, and signaling through different glutamate receptors can have diverse effects. The
ionotropic glutamate receptor GluR5, for example, appears to inhibit the HPA axis response to
restraint when signaling at the PVN (see chapter 2), while other ionotropic glutamate receptors
typically play an excitatory role in HPA axis regulation (Jezova, 2005). The role of metabotropic receptors in the HPA axis is less certain.
Metabotropic glutamate receptors are generally categorized into 3 different families, based on homology and pharmacology (Benarroch, 2008). Group I consists of mGluR1 and mGluR5, which are almost exclusively post-synaptic in location in the central nervous system
(Van den Pol, 1994; van den Pol et al., 1995). Activation of Group I metabotropic glutamate receptors leads to excitation of the neuron. Groups II and III, on the other hand are typically expressed presynaptically and negatively regulate synaptic release of neurotransmitters
(Benarroch, 2008). In addition to exciting post-synaptic cells, however, group I metabotropic glutamate receptors are also known to reduce presynaptic neurotransmitter release by
99 downstream signaling through endocannabinoid actions on CB1 receptors (Doherty and
Dingledine, 2003). Thus, signaling through metabotropic glutamate receptors could be either excitatory or inhibitory.
These dual roles are supported by the recorded effects of metabotropic glutamatergic drugs on regulation of the HPA axis. Intracerebroventricular treatment of animals with the group
I metabotropic agonist (S)-3,5-dihydroxyphenylglycine (DHPG) leads to increased plasma corticosterone within 1 hour after administration (Johnson et al., 2001), consistent with an excitatory role for these receptors. Peripheral administration of the mGluR5 antagonist 2-methyl-
6-(phenylethynyl) pyridine (MPEP) also leads to increased plasma corticosterone levels within 1 hour after intraperitoneal injection (Bradbury et al., 2003), however, suggesting an inhibitory
HPA axis action for group I mGluRs. It may be that there are different sites of action for these opposite results, but there is currently insufficient evidence to confirm or refute this.
Clearly, more work is needed to understand the roles of metabotropic glutamate signaling in regulating the HPA axis. This is especially true in determining the sites of action for metabotropic glutamate receptors in regulating the HPA axis. Intracerebroventricular administration of a drug is likely to lead to actions relatively close to the ventricular surface
(Francis et al., 2006). Therefore, it is possible that one site of action is at the PVN. Group I metabotropic glutamate receptors are expressed in the PVN (Van den Pol, 1994; van den Pol et al., 1995), but it is currently unknown whether these receptors are located in CRH-containing neurons, or whether they play a role in regulating the HPA axis response to stress.
The purpose of the current studies is to determine the effect of group I metabotropic glutamate receptor signaling in the PVN on HPA axis responsivity to restraint stress. We hypothesized that group I metabotropic receptors would act in an inhibitory fashion in the PVN,
100 due to coupling of metabotropic receptors with endocannabinoid signaling in the PVN. To test
this hypothesis, we gave rats intra-PVN injections of group I metabotropic agonist or antagonist
before restraint stress and measured the plasma hormonal response to restraint.
Materials and Methods
Animals
Male Sprague-Dawley weighing 300-350g were used in these studies. Rats were kept in
dedicated animal holding rooms with controlled temperature and humidity. The animals were kept on a 12:12 hour light:dark cycle, with lights on at 6:00 AM. After arrival at the facility, rats
were allowed to accommodate to the new surroundings for at least 1 week before undergoing
surgery. All animal protocols were approved by the University of Cincinnati Institutional Animal
Care and Use Committee, and were consistent with the guidelines set forth in the National
Institutes of Health Guide for the Care and Use of Laboratory Animals.
Stereotactic surgery
Rats were anesthetized with a mixture of ketamine and xylazine (90 mg/kg and 10 mg/kg,
respectively), with additional dosing as necessary to maintain a surgical plane of anesthesia and
pre-emptive analgesia with butorphanol. They were implanted with 26-gauge bilateral guide
cannulas (1.0 mm center-center distance, Plastics One, Roanoke VA) aimed at the PVN (1.9mm
posterior to bregma, centered over the sagittal sinus, and 6.3 mm ventral from the dura mater at
the site of implantation). Cannulas were anchored to the skull using stainless steel screws and cranioplastic cement (Plastics One, Roanoke, VA). Dummy cannulas (cut to project 0.1 mm
101 beyond then end of the guide cannula) were placed in the guide cannula to maintain patency, and dust caps were placed over the dummy cannulas to hold them in place. Animals were given prophylactic gentamicin post-operatively, and observed during recovery before being returned to their home cages. All animals were observed post-surgery for signs of pain or illness, and
additional analgesic was given as necessary for pain. After several days of recovery, the dummy
cannulas were removed and replaced daily until the day of the experiment, to maintain an open
guide cannula.
Drugs
(S)-3,5-dihydroxyphenylglycine (DHPG) and hexyl-homoibotenic acid (hexyl-HIBO)
were purchased from Tocris (Ellisville, MO). Stock solutions were made by dissolving the drugs
in 0.9% saline alkalinized with NaOH. The pH of the resulting solutions was checked using pH
paper, and neutralized with HCl as necessary. The drugs were then diluted to working
concentrations (10 nmol/μl for DHPG and 1 nmol/μl for hexyl-HIBO) in 0.9% saline. All drug
solutions were made on the morning they were used.
Injection and Restraint Challenge
After cannula implantation, animals were allowed to recover for 6-8 days before restraint
challenge. On the day of the experiment, the dummy cannulas were removed from the guide
cannulas, then internal cannulas (cut to project 1.0 mm beyond the end of the guide cannula)
were placed. The animals were given injections (500 nl per side) through the cannula, using a
PHD 2000 syringe pump (Harvard Apparatus, Holliston, MA). Injections were given over 1
102 minute (500 nl/min injection rate). Immediately following the injection, animals were restrained
in plastic restraining tubes, and blood samples (~250 μl) were collected into microcentrifuge tubes containing EDTA, immediately upon restraint, then 15 and 30 minutes later. After the 30 min blood sample, animals were released into their home cages, and killed by rapid decapitation at 60 minutes post-injection. Trunk blood was collected, and brains removed after decapitation.
Nissl stain for cannula placement
Brains were immersion fixed in 3.7% formaldehyde in 50 mM potassium phosphate
buffer for 5 days. Fixed brains were impregnated with 30% sucrose solution, then sectioned at
25 μm using a sliding microtome (Leica, Bannockburn, IL). Sections were mounted on glass
slides then Nissl stained with cresyl violet. Cannula placement was verified by observing the
cannula tracks in brain sections. Only animals with correctly placed cannulas were included in
the analyses of plasma hormones.
Radioimmunoassay
Plasma was separated from the blood samples by centrifugation. Plasma levels of
corticosterone were assayed using a commercial 125I-labeled corticosterone radioimmunoassay
kit (ICN, Costa Mesa, CA). ACTH levels were assayed with a 125I-labeled ACTH
radioimmunoassay as previously described (Engeland et al., 1989), using tracer from DiaSorin
(Stillwater, MN). ACTH antiserum was a generous gift of W. Engeland (University of
Minnesota).
103 Statistical analysis
Plasma hormone levels were analyzed using 2-way or 3-way ANOVA with repeated
measures, as appropriate. Area under the curve (AUC) was estimated using the trapezoidal
method. Data were square root transformed if necessary to obtained homogeneity of variance,
and outliers were removed using Grubb's test (Barnett and Lewis, 1994). Significant main
effects or interactions were analyzed further using the Fisher's LSD post-hoc test. Statistical
significance was considered to be p < 0.05. Statistical analysis was performed using the GB-Stat
program (Dynamic Microsystems, Silver Spring, MD).
Results
Metabotropic glutamate receptor activation inhibits the HPA axis response
to restraint
CB1 receptor signaling is known to inhibit neurotransmitter release in a presynaptic manner in many brain systems (Hoffman and Lupica, 2006). In at least most of these systems, endocannabinoid synthesis is initiated by signaling through group I metabotropic glutamate receptors. To see if this was the case in fast-feedback regulation of the HPA axis, we bilaterally infused the group I metabotropic glutamate receptor agonist DHPG (5 ng per side) into the PVN, then subjected the animals to restraint challenge. Consistent with a role for metabotropic
glutamate receptor signaling in the fast feedback pathway, DHPG treatment attenuated the HPA
axis response to restraint stress. There was a significant reduction in the ACTH time course response (Time x DHPG interaction F3,67 = 2.942, p < 0.05), and in the corticosterone AUC
response (2 tailed t = 2.277, p < 0.05). These results are summarized in Figure 1.
104 Blocking mGluR signaling potentiates HPA axis response to restraint
We then attempted to block group I metabotropic glutamate receptors, using the group I
selective antagonist hexyl-HIBO (Madsen et al., 2001). Blockade of group I metabotropic
receptors resulted in a significant elevation of the ACTH response to restraint, both in the time
course (time x HIBO x dexamethasone interaction, F3,159 = 23.255, p < 0.05) and the AUC (main
effect of hexyl-HIBO treatment, F1,30 = 6.161, p < 0.05). There was also a significant effect on
corticosterone secretion (hexyl-HIBO treatment x time interaction F3,135 = 3.613, p < 0.05),
although there was a statistically significant elevation in corticosterone seen only at the 0 minute time point. There was not a significant change in the corticosterone AUC measurement as a result of hexyl-HIBO treatment. These results are summarized in Figure 2.
Dexamethasone treatment reverses the antagonist-induced HPA potentiation
Dexamethasone infusion into the PVN causes a rapid inhibition of the HPA axis response
to restraint stress (See chapter 3). Consistent with these results, dexamethasone treatment led to
a rapid inhibition of the ACTH response to restraint (time x hexyl-HIBO x dexamethasone
interaction F3,159 = 23.255, p < 0.05). Dexamethasone treatment also blocked the hexyl-HIBO-
mediated potentiation of the HPA axis response to restraint at 30 minutes after onset of restraint,
while hexyl-HIBO blocked the dexamethasone-mediated attenuation of ACTH secretion at 15
minutes. Figure 2 illustrates these results.
105 Discussion
In the current studies we demonstrate that group I metabotropic glutamate receptor signaling is involved in inhibitory regulation of the HPA axis response to restraint in the PVN.
Group I metabotropic glutamate receptor agonism leads to inhibition of the HPA axis response to restraint, while antagonist treatment of the PVN leads to excitation of the HPA axis response. In addition, antagonist treatment blocks the early (15 minute) attenuation of ACTH secretion caused by dexamethasone treatment at the PVN. These studies add significantly to our understanding of the roles for glutamate in regulation of the HPA axis in the PVN.
It has been understood for a number of years that glutamate regulates HPA axis activity at the PVN. Infusion of glutamate into the PVN leads to excitation of ACTH secretion (Darlington et al., 1989), suggesting that the predominant action of glutamate in the PVN is excitatory.
However, the current results, in conjunction with studies that show an inhibitory role for GluR5 on HPA axis activity (Chapter 2), suggest that the glutamatergic regulation of the HPA axis at the
PVN is complex, with inhibitory as well as excitatory signals involved in control of parvocellular neurons.
Inhibitory signaling through metabotropic glutamate receptors has precedent, with presynaptic group II/III metabotropic glutamate receptors (Scaccianoce et al., 2003). These receptors are located on presynaptic terminals, and have an autoreceptor function, where they decrease glutamate release in response to excess glutamate presence at the synapse (Jaskolski et al., 2005; Benarroch, 2008). However, a presynaptic autoreceptor function is not compatible with group I metabotropic receptors, which are exclusively post-synaptic in location (Van den
Pol, 1994; van den Pol et al., 1995). Further, group I metabotropic receptor signaling is typically excitatory to the neuron (Benarroch, 2008). Thus, possible mechanistic explanations of the
106 actions of group I receptors in the PVN are more limited; however, there remain several possible explanations.
One possible mechanism of action for metabotropic glutamate receptors in this
experimental system is through inhibiting glutamate release onto CRH-containing cells. Group I
metabotropic glutamate receptors are known to be functionally coupled to endocannabinoid
signaling (Doherty and Dingledine, 2003), which leads to decreased neurotransmitter release
from synapses associated with CB1 receptors (Hoffman and Lupica, 2006). If this is the case, signaling through group I metabotropic glutamate receptors would lead to synthesis of endocannabinoids in the post-synaptic cell, which would then signal through presynaptic CB1
receptors, leading to decreased release of glutamate onto the CRHergic cell and thus to decreased secretion of the HPA hormones ACTH and corticosterone (Figure 3B).
An inhibitory role for metabotropic glutamate receptor signaling, mediated by
endocannabinoids, is reminiscent of fast feedback inhibition of the HPA axis by glucocorticoids.
Fast feedback occurs at the PVN through an endocannabinoid-mediated mechanism (Di et al.,
2003) (see Chapter 3). In brief, glucocorticoid signaling is thought to rapidly induce release of
endocannabinoids from the post-synaptic CRH-containing neuron in the PVN. These endocannabinoids cause decreased glutamate release from presynaptic terminals by signaling through the CB1 receptor. There are several points of similarity between fast feedback and the
inhibition caused by metabotropic signaling in the PVN. First, the time frame is similar between
the two. Fast feedback effects of glucocorticoids on ACTH secretion are significant within 15 minutes of infusion of dexamethasone, the same as with DHPG. Second, induction of endocannabinoid production is often initiated by signaling through group I metabotropic glutamate receptors (Doherty and Dingledine, 2003). Further, the dexamethasone-mediated
107 attenuation of the HPA axis response to restraint is completely reversed at 15 minutes after the onset of restraint, a time point where hexyl-HIBO alone has no effect on the ACTH response.
However, there are some shortcomings of this comparison. First, the antagonist hexyl-
HIBO alone causes potentiation of the HPA axis response to restraint. The endocannabinoid receptor antagonist AM-251 alone has no effect on the restraint-induced ACTH or corticosterone responses when infused into the PVN. If endocannabinoid signaling were the mechanism by which metabotropic glutamate receptor activation inhibits the HPA axis response to restraint, it would be expected that AM-251 and hexyl-HIBO treatment would have similar results. In spite of these limitations, though, the fact that hexyl-HIBO alone has no effect at the 15 minute time point but blocks the dexamethasone-induced ACTH attenuation at this time point is consistent with a role for metabotropic glutamate receptor signaling in fast feedback inhibition of the HPA axis response to restraint at the PVN. Further studies are needed to investigate this signaling pathway further.
One question raised by these possible mechanisms of action for the effects of group I metabotropic glutamate receptor modulation involves the source of the glutamate mediating this signaling. For example, if metabotropic glutamate receptor signaling inhibits the HPA axis in the
PVN by activating endocannabinoid signaling, it is likely that the glutamate that initiates this response is released from a local glutamatergic axon terminal. Thus, glutamate release from an axon terminal sufficient to elevate glutamate concentrations at the metabotropic glutamate receptors would lead to activation of endocannabinoid synthesis. Through endocannabinoid- mediated inhibition of glutamate release, the glutamatergic input through that synapse would be dampened.
On the other hand, if the metabotropic glutamate receptors are acting by exciting a
108 presynaptic GABAergic neuron, it is likely that the glutamate initiating this signal is provided
through direct synaptic inputs to the GABAergic cell. In addition, since the glutamatergic drugs in this study were applied locally at the PVN, such a GABAergic neuron would need to be in close proximity to the PVN. Indeed, it is known that GABAergic neurons in the peri-PVN region act to regulate the PVN through GABAergic input onto PVN neurons (Herman et al.,
2002). In this case, the possible candidates for supplying the glutamatergic input to the nucleus for this action are likely to be neurons innervating the peri-PVN rather than the PVN proper.
Further studies are needed to determine the source of glutamatergic synapses responsible for activating the group I metabotropic glutamate receptor system at the PVN.
In conclusion, we have shown that group I mGluRs mediate inhibition of the HPA axis at the level of the PVN. Inhibition of group I metabotropic glutamate receptor signaling at the
PVN is sufficient to reverse dexamethasone-mediated fast inhibition of the HPA axis response to restraint at early time points. However, since metabotropic glutamate receptor inhibition alone causes excitation of the HPA axis response to restraint at later time points, more work is necessary to determine whether metabotropic glutamate receptor signaling is involved in the pathway mediating fast feedback inhibition of the HPA axis by glucocorticoids.
109 Figure Legends
Figure 1. Group I metabotropic glutamate receptor agonist inhibits the HPA axis response
to restraint. Rats were injected with the group I metabotropic glutamate receptor agonist
DHPG. Plasma ACTH and corticosterone levels were decreased in response to restraint in animals treated with DHPG. A) Time course of ACTH secretion in response to restraint. B)
AUC of ACTH secretion in response to restraint. C) Time course of corticosterone secretion in response to restraint. D) AUC of corticosterone secretion in response to restraint. * p < 0.05 vs.
vehicle.
Figure 2. Group I metabotropic glutamate receptor antagonist elevates the HPA axis response to restraint in a manner that is reversed by dexamethasone treatment. Rats were injected with the group I metabotropic glutamate receptor antagonist hexyl-HIBO and/or dexamethasone, then subjected to 30 minutes of restraint. Hexyl-HIBO treatment elevated the
ACTH response to restraint at 30 minutes after onset of restraint, while dexamethasone reversed this effect. Dexamethasone treatment attenuated the ACTH response to restraint at 15 minutes after restraint onset, and this effect was blocked by hexyl-HIBO treatment. A) Time course of
ACTH secretion in response to restraint. B) AUC of ACTH secretion shown in A. C) Time course of corticosterone secretion in response to restraint. D) AUC of corticosterone secretion in
C. * p < 0.05 vs. vehicle.
Figure 3. Model for actions of metabotropic glutamate receptor signaling in the PVN, on the
HPA axis. We present two possible models for group I metabotropic glutamate receptor- induced inhibition of the HPA axis at the PVN. A) Increased inhibition. Metabotropic glutamate
110 receptors are located on a GABAergic neuron presynaptic to the CRH-containing parvocellular
PVN neuron. Glutamate signaling through the receptors excites the inhibitory neuron, leading to
increased GABA release onto the CRH-containing cell. Increased GABAergic signal onto the
CRH neuron leads to decreased CRH release, and thus to decreased ACTH and corticosterone
secretion in response to restraint stress. B) Presynaptic inhibition through endocannabinoid
signaling. Metabotropic glutamate receptors are located on the parvocellular neuron.
Glutamatergic signal through the metabotropic glutamate receptor leads to activation of
endocannabinoid synthesis. The endocannabinoids create a retrograde signal that causes
presynaptic glutamatergic terminals to decrease glutamate release onto the parvocellular neuron.
Decreased glutamatergic signal leads to decreased CRH release from the parvocellular neuron,
which in turn leads to decreased secretion of ACTH and corticosterone in response to restraint stress.
111 Figure 4-1 Group I Metabotropic Glutamate Receptor Agonist
112 Figure 4-2 Group I Metabotropic Glutamate Receptor Antagonist
113 Figure 4-3 Model for Group I Metabotropic Glutamate Receptor Signaling at the PVN
114 5. General Discussion
The preceding chapters describe the results of a series of experiments directed at
understanding the roles of glutamate and glucocorticoids in rapid regulation of the HPA axis.
Significantly, glutamate was found to have both excitatory and inhibitory functions in regulation
of the HPA axis response to restraint stress. This finding represents an important contribution to
understanding how the HPA axis is regulated at the level of the hypothalamus, since glutamate has traditionally been thought of as being an excitatory neurotransmitter. Glucocorticoid
hormones were found to rapidly inhibit the HPA axis response to restraint stress, consistent with
a fast negative feedback function for these hormones. The rapidity of onset of the inhibitory
actions is significant, since glucocorticoids have traditionally been thought of as initiating
genomic signaling pathways that take hours to days cause their effects.
All of the effects documented in this work were found to occur rapidly. The slowest of
the actions described are the effects of signaling through ionotropic GluR5 receptors in the PVN
and median eminence on the HPA axis response to restraint. The experiments discussed in this
work reveal significant changes in ACTH secretion, caused by pharmacological manipulation of
GluR5, group I metabotropic glutamate receptors, and cannabinoid receptors, and by
glucocorticoid treatment. Effects of all of these treatments were evident within 1 hour, with
group I metabotropic glutamate receptor, cannabinoid receptor, and glucocorticoid effects all
being detectable within 15 minutes.
Timing of effects: genomic vs. nongenomic
One of the unifying themes of the work presented here is the time scale on which these
115 systems work. Fast feedback inhibition of ACTH release occurs within 15 minutes of treatment
with glucocorticoids, and the effects of metabotropic receptor manipulation were also obvious
within 15 minutes. In the fast feedback experiments, corticosterone effects were generally seen
by 60 minutes post-treatment, which is also the time when corticosterone effects were seen with
the GluR5 studies. The timing of fast feedback most likely places it in a nongenomically
mediated mechanism, as does that of group I metabotropic glutamate receptor signaling. The
timing of the GluR5-mediated events is also consistent with nongenomic, possibly second messenger-mediated, signaling, but does not rule out a possible genomic mechanism.
Fast feedback
Fast feedback inhibition of the HPA axis by glucocorticoids at the PVN is likely mediated by nongenomic signaling, because of the rapidity with which these effects are detectable. In order for glucocorticoids acting at the PVN to reduce ACTH output, several steps must be affected. Our evidence suggests that endocannabinoid signaling is involved in the signaling pathway, so endocannabinoid synthesis must be initiated, signaling through CB1 carried out, glutamate release decreased, leading to reduced excitation-coupled release of CRH from the median eminence. This reduced CRH release must then be translated through the portal vasculature to the anterior pituitary, leading to decreased cAMP production and finally reduced
ACTH release. The process, from the initiation of a stress response to increased ACTH release takes more than 3 minutes (Vahl et al., 2005). Thus, a significant reduction in ACTH release within 15 minutes of glucocorticoid treatment would leave only approximately 10 minutes for any kind of genomic response to glucocorticoids to be initiated, which is far too little time for a meaningful change in the level of functional protein.
116 Further, the kinetics of the ACTH response seen above suggests a nongenomic action of dexamethasone in fast feedback. By 15 minutes, the ACTH response to restraint is significantly lower in animals treated with dexamethasone than in vehicle treated animals. The current studies were not designed to identify the earliest time point that differences are evident, but since the plasma half-life of ACTH in rats is about 6-7 minutes (Lopez and Negro-Vilar, 1988), glucocorticoid treatment would have to have reduced the initial ACTH response within less than
10 minutes after the initiation of restraint to observe a difference in the ACTH response of the magnitude typically seen. Further, this would only be true if the decrease in ACTH secretion had approached a near maximal magnitude within 10 minutes. This would leave a very short time for a genomic response to develop.
Although glucocorticoid-induced transcriptional changes have been detected within as little as 7.5 minutes (Groner et al., 1983), this was a change in transcription only, and was done in a viral system. Further, 7.5 minutes was the earliest time that a change was detectable, while the maximal response was not evident until more than an hour after treatment. The fastest time frame in which genomic effects have been observed in non-viral systems using glucocorticoids is about 15-30 minutes, depending on cell type (Makara and Haller, 2001). Furthermore, this time frame is from onset of glucocorticoid treatment until the effect was detectable, rather than until the effect was maximal. In the fast feedback experiments, the effects of dexamethasone on
ACTH secretion are maximal within 15 minutes. Since times earlier than 15 minutes were not evaluated, it is not clear how early the effects are detectable. All in all, it appears that the temporal characteristics of fast feedback in the PVN rule out a genomic action of dexamethasone. Thus, fast feedback is most likely mediated through a nongenomic signaling pathway.
117 On the other hand, the presence of a membrane bound receptor in this model of fast feedback regulation of the HPA axis was not clearly demonstrated. Since one criterion that has been suggested for classifying fast feedback as a nongenomically mediated effect of glucocorticoids is that a membrane-bound receptor is involved in mediating this effect (Makara and Haller, 2001), it is important to further examine the evidence supporting a nongenomic mechanism for fast feedback.
The current studies were not able to conclusively prove that dex:BSA is both membrane- impermeable and stable to hydrolysis. However, it has been argued that other evidence supports the hypothesis that the receptor for fast feedback is a membrane-bound receptor. For example, inhibiting GPCR function with GDP-β-S blocks the rapid inhibition of excitatory post synaptic currents onto CRH containing neurons in the PVN (Di et al., 2003). The cytosolic GR inhibitor
RU38486 has failed to inhibit fast feedback-like actions in vitro, such as decreasing glutamatergic currents onto PVN cells (Di et al., 2003) and release of ACTH from pituicytes
(Hinz and Hirschelmann, 2000), suggesting that these effects are mediated through a receptor distinct from GR. Glucocorticoids also rapidly inhibit CRH release from hypothalamic synaptosomes, in the presumed absence of genetic machinery (Edwardson and Bennett, 1974).
Another piece of evidence that argues for a nongenomic mechanism of action in fast feedback is that the rapid effects are preserved after treatment of the system with inhibitors of transcription and translation (Abou-Samra et al., 1986; Hinz and Hirschelmann, 2000), although this is not necessarily true in the pituitary (Dayanithi and Antoni, 1989).
Although this evidence does suggest a nongenomic, membrane receptor-mediated mechanism of action in fast feedback, there are other possible explanations. The blockade of the
electrophysiological effect by GTP-γ-S does indicate that a GPCR most likely is involved in this
118 signaling pathway. However, there is now a demonstrated role for endocannabinoid signaling
through CB1 in mediating fast feedback both in vitro and in vivo. Since CB1 is a GPCR itself, and is downstream from the putative glucocorticoid receptor, blocking GPCR signaling would inhibit the fast feedback response regardless of whether the glucocorticoid receptor were a
GPCR. Evidence for a membrane bound GPCR acting as a glucocorticoid receptor has been published for pituicytes (Maier et al., 2005); however no such evidence is yet in the literature for the PVN. Thus, one cannot state beyond reasonable doubt that the receptor for glucocorticoids in fast feedback is a membrane receptor or a GPCR.
Since the criterion of membrane-receptor-mediation may not be reliable, we must look at other criteria for classifying this as a nongenomic action. In vitro evidence suggesting that
inhibitors of gene transcription and translation fail to block fast feedback effects is supportive of a nongenomic classification for fast feedback (Abou-Samra et al., 1986; Hinz and Hirschelmann,
2000). However, it must be remembered that these inhibitors block the expression of genes but do not affect processes that inhibit gene expression. Transcription inhibitors do not rule out the synthesis of new protein from existing mRNA, for example. Further, gene expression inhibitors do not unequivocally rule out genomic signaling as a mechanism for expression of an effect, since glucocorticoids also inhibit gene expression. This occurs when glucocorticoids bind to negative glucocorticoid response elements in protein promoters, or interact with other transcription factors to decrease transcription (Heitzer et al., 2007). Glucocorticoids are also known to destabilize certain mRNAs (Newton et al., 1998; Dhawan et al., 2007). Thus, in the absence of new gene expression, glucocorticoids could still be acting in a way that could be termed genomic, by blocking expression of genes. There is as yet no satisfactory way of blocking these inhibitory signals of glucocorticoids. That said, however, blocking the synthesis
119 of new proteins will not have an effect on a cell until the existing stocks of that protein are
depleted, a process that is typically slower than expression of new proteins.
In aggregate, then, although not all of the criteria for nongenomic actions of
glucocorticoids, as suggested by Makara (Makara and Haller, 2001) have been met, it appears
likely that the fast feedback effects described above are mediated by nongenomic signaling.
Further work is necessary to determine whether the glucocorticoid receptor mediating fast
feedback is a form of GR and/or membrane-bound.
Metabotropic glutamate receptors
Like fast feedback, metabotropic glutamate receptor signaling led to changes in ACTH
secretion within 15 minutes of treatment. The same arguments outlined above will serve as
evidence that these effects are mediated by nongenomic mechanisms. Further, since DHPG and hexyl-HIBO caused their effects on HPA axis activity within the same time frame as dexamethasone, these results may be consistent with a role for metabotropic glutamate receptor signaling in mediating the fast feedback response itself.
One difference between the timing of fast feedback and group I metabotropic receptor actions is that in the agonist experiment, there was a possible difference in the corticosterone response within 15 minutes. Because there wasn't a significant main effect of DHPG treatment in this experiment, however, it is not possible to determine whether there truly is a difference at this time point. Since there was a significant effect of DHPG treatment on the AUC of corticosterone release, and the only times where there appears to be a difference are the 15 and
30 minute time points, there may have been a change in corticosterone release in this experiment
earlier than we have observed with the fast feedback experiments. However, since there was not
120 a statistical difference, more studies will be needed to rule this possibility in or out.
GluR5
In contrast to fast feedback and metabotropic receptors, the effects of GluR5 treatment at either the median eminence or the PVN occur slow enough to be genomically mediated. There was never a statistically significant change in ACTH before the 60 minute time point in these
experiments. Thus, effects on ACTH occurred somewhere between 30 minutes and 60 minutes.
As discussed above, this is slow enough for changes in gene expression to occur. In contrast to
these results, activation of GluR5 in other systems leads to effects within seconds (Binns et al.,
2003; Braga et al., 2003; Campbell et al., 2007). The absence of any rapid effect of GluR5
antagonism in the PVN on activity of the HPA axis suggests that GluR5 is acting through a
different signaling pathway. This could support a genomic mechanism of action.
In the case of the PVN, a genomically mediated action of GluR5 signaling may be
supported by the significant changes in restraint-induced c-Fos expression in the PVN, in
response to LY382884 treatment. Of course, this alone is not strong evidence for a genomic
effect of GluR5 signaling. If, as postulated, GluR5 is signaling presynaptically, it is likely that
there would be differences in the response of the downstream cell to nongenomic actions of
presynaptic GluR5. Thus, a change in gene expression in PVN cells is not strong evidence for a genomic action of GluR5. The slow response could indicate, for example, that the effects of
GluR5 signaling are mediated by second-messenger or other intracellular signaling pathways.
This hypothesis is supported by evidence that GluR5-containing receptor complexes can signal in a metabotropic manner (Chittajallu et al., 1999).
Like the intra-PVN injection experiments, effects on HPA axis activation in response to
121 intra-median eminence injections of LY382884 were not apparent until the 60 minute time point.
In the median eminence, genomic signaling does not seem to be likely. The external lamina of the median eminence, while containing some cells, is mostly made up of axon terminals at hypophyseal portal capillaries. There is likely to be no gene transcription occurring at these terminals, since the genomic DNA is not present. It is possible that there is synthesis of new protein at axon terminals (Giuditta et al., 2002), which could be considered as a type of genomic signaling, but there is likely not a change in gene expression beyond this. Further work will be required to conclusively show the nature of GluR5-mediated signaling in the HPA axis.
Receptor signaling in HPA axis regulation
Glutamate:
Glutamate is the main excitatory neurotransmitter in the central nervous system, including the HPA axis (Brann, 1995). However, it is apparent from a review of the relevant literature that glutamate does not act as a simple excitatory signal in the PVN (see introduction).
The results of the studies reported in this work further support this assertion. Glutamate working through two different receptor types (GluR5 and group I mGluR) was excitatory and inhibitory to the HPA axis. It is interesting to note that glutamate signaling through GluR5 in the PVN was inhibitory to the HPA axis response to restraint, and that signaling through group I metabotropic glutamate receptors in the PVN was also inhibitory to the HPA axis response to restraint.
Although this inhibitory action is an exception to the rule for glutamate acting as an excitatory signal, this inhibitory signaling is not without precedence. Presynaptic kainate glutamate receptors are known to occasionally act as autoreceptors, leading to decreased release of
122 neurotransmitters from the presynaptic terminals on which they are found (Huettner, 2003).
Group II and III metabotropic glutamate receptors are likewise known to act as inhibitory
autoreceptors at presynaptic terminals (Benarroch, 2008).
One purpose of presynaptic inhibitory signaling through glutamate receptors is negative
feedback regulation of glutamate release from a synapse. At low levels of glutamate release, the
glutamate is retained in the synaptic cleft, where it acts exclusively on post-synaptic glutamate
receptors. However, if glutamate release is significant enough for released glutamate to leave the
synapse, this “escaped” glutamate can then act on peri-synaptic glutamate receptors, such as the
presynaptic autoreceptors. Thus when sufficiently high levels of glutamate are released, the excess glutamate serves to decrease the release of glutamate from the presynaptic terminal.
Since excess levels of glutamate can lead to cellular toxicity (Greenwood and Connolly, 2007), these autoreceptors play an important role in preserving the integrity of the central nervous system under conditions of high glutamate release.
Of course, signaling through a glutamate receptor that leads to inhibition of a physiological effect is not necessarily evidence that glutamate per se is acting in an inhibitory fashion. For example, excitatory glutamatergic signaling onto a GABAergic inhibitory interneuron will lead to increased GABA release onto the target cells of the interneuron; thus, glutamate signaling will lead to decreased activity of the cell that is a target of the GABAergic neuron. This is one possible explanation of why group I metabotropic glutamate receptors mediate an inhibitory signal on the HPA axis, even though these receptors are known to have a post-synaptic location and to generate a primarily excitatory signal. Metabotropic glutamate receptor signaling in one of the many GABAergic neurons in the peri-PVN region, which are known to play a regulatory role in the HPA axis (Herman et al., 2002) could be the mechanism
123 behind this inhibitory signaling. Another possible explanation for these actions is in the mGluR- endocannabinoid signaling pathway, which is discussed below.
In contrast to group I metabotropic glutamate receptors, GluR5 is expressed both presynaptically and post-synaptically (Pinheiro et al., 2005). GluR5 mainly signals in an
excitatory fashion. Although it is conceivable that presynaptic GluR5 is acting as an
autoreceptor, the majority of the evidence for an inhibitory role of GluR5 signaling suggests that
the inhibition is mediated by effects on inhibitory interneurons (Campbell et al., 2007) (Braga et
al., 2003). The literature further suggests that presynaptic GluR5 acts to facilitate
neurotransmitter release from presynaptic terminals (Xu et al., 2006; Aroniadou-Anderjaska et
al., 2007; Campbell et al., 2007; Wu et al., 2007). The results of the current studies on GluR5
actions in the HPA axis support this role. Neurotransmitter release facilitation by presynaptic
GluR5 can explain the effects of GluR5 signaling in both the median eminence and PVN, as
discussed in Chapter 2.
Overall, the results of the studies that I have performed are consistent with known
signaling modalities for these receptors in other parts of the brain. Further, these results support
a complex regulatory role for glutamate signaling in the PVN. Glutamate signaling through
different receptors at different synapses in the PVN apparently causes a wide variety of
regulatory effects on HPA axis activity. Since there are a large number of glutamate receptors
that are known to be expressed in the PVN (Van den Pol, 1994; van den Pol et al., 1995; Herman
et al., 2000), glutamatergic regulation of the HPA axis presents a field that remains rich for
exploration. Much more work is necessary to completely understand the roles that glutamate
plays in the PVN, in regulating the HPA axis.
124 Glucocorticoids
One question not adequately addressed in the studies described is, “What is the receptor
responsible for transducing a glucocorticoid signal into a fast feedback outcome?” The current
results are consistent with a membrane bound receptor mediating fast feedback. However, there
are still several alternate explanations that have not been ruled out, including GR-mediated signaling, non-GR receptor mediated signaling, and receptor-independent signaling.
The first of these possibilities is that glucocorticoids signal through a membrane-bound form of GR that is coupled to a G-protein signaling pathway. This is the type of signaling that is commonly thought to underlie fast feedback inhibition of the HPA axis by glucocorticoids. In this model, binding of glucocorticoid to the membrane receptor initiates the signaling cascade that leads to fast feedback. Note that this signaling modality would be valid whether the actual membrane receptor is a novel, non-GR receptor as has been previously postulated (Di et al.,
2003), or an altered, membrane-bound form of GR as has also been suggested (Gametchu, 1987;
Gametchu et al., 1999; Watson and Gametchu, 2001). In either case, binding of glucocorticoid to the membrane receptor initiates downstream synthesis of endocannabinoids, which cause inhibition of glutamate release onto the CRH-containing cell in the PVN, as outlined in Figure 7 of Chapter 3. This model seems to be held in the most in favor currently, but there are problems with its explanations.
First of all, it is not yet clear how glucocorticoid binding to a membrane receptor would lead to increased endocannabinoid synthesis and release. However, a useful comparison may be found in how group I metabotropic glutamate receptors mediate endocannabinoid release.
Briefly, binding of glutamate to group I metabotropic glutamate receptor proteins causes activation of G-proteins, which cause endocannabinoid synthesis by activating synthetic
125 enzymes phospholipase C (PLC) and diacylglycerol lipase (Jung et al., 2005). This signaling is apparently initiated by the Gq subunit directly interacting with these synthetic proteins. A similar signaling cascade could in theory be initiated by glucocorticoid binding to a G-protein coupled glucocorticoid receptor.
This postulated role for G-protein coupled glucocorticoid signaling is easy to envision for an as-yet undiscovered glucocorticoid GPCR; however, there is some evidence that the receptor initiating fast feedback is a modified form of GR (Liu and Chen, 1995). If this is the case, it is not clear how GR would be modified to cause this effect or how GR could be coupled to a
GPCR. For the first of these problems, there is some evidence in immune cells that the membrane bound form of GR is associated with expression of a specific alternate transcript of the GR gene (Gametchu et al., 1999). However, the correlation between expression of this alternative transcript and membrane GR is not complete, and the alternate mRNA produced is only different in the 3' untranslated region. Thus, it is not clear how this alternative transcript could cause a change in the structure of GR significant enough to change the location and signaling mechanism of the protein.
A mechanism that is able to alter the subcellular location of GR might, however, be sufficient to explain the pharmacological differences between classical glucocorticoid signaling and the putative membrane glucocorticoid receptor. Altering the lipid milieu of a membrane protein is sufficient to alter its chemical properties (Wu and Gorenstein, 1993) and activity
(Stallkamp et al., 1999). Altering membrane characteristics is sufficient to alter the signal transduction efficacy of the thyrotropin releasing hormone receptor (a GPCR), for example
(Ostasov et al., 2007). Thus, moving the classical GR to the membrane could conceivably lead to the observed pharmacological differences between the putative membrane receptor-mediated
126 effects and classical glucocorticoid signaling. Although a change in location could explain the
altered pharmacology of the postulated glucocorticoid receptor mediating fast feedback, it is still
unclear how this receptor could be coupled to a G-protein coupled signaling pathway.
One intriguing possibility for a connection between GR and GPCR signaling is the
reported association between GR and Vesl-2, which is also known as Homer-2 (Hedman et al.,
2006). Homer proteins are synaptic scaffold proteins that are known to associate with metabotropic glutamate receptors (Xiao et al., 1998), and may be necessary for their localization to synapses (Brakeman et al., 1997; Ehrengruber et al., 2004). Homer proteins mediate the association of group I metabotropic glutamate receptors with other proteins that are involved in their signaling cascade, such as the IP3 receptor (Ehrengruber et al., 2004), and functionally link metabotropic glutamate receptors to the mitogen activated protein kinase members ERK 1 and 2
(Mao et al., 2005). Homer proteins bind to and interact with members of the NMDA receptor
scaffold complex (Ehrengruber et al., 2004), and are necessary for the activity of metabotropic
glutamate receptor proteins in processes such as long-term depression (Ronesi and Huber, 2008).
Because of this close linkage, both physical and functional, between Homer proteins and
group I metabotropic glutamate receptors, it is possible that Homer-2 could mediate an
interaction between GR and group I receptors. The resultant localization of GR to the membrane
could lead to the observed changes in pharmacology of the fast feedback effects of
glucocorticoids vs. classical genomic-mediated effects, as discussed above. Further, an indirect,
Homer-mediated association between GR and metabotropic glutamate receptors suggests a way
that glucocorticoid binding to GR could lead to the initiation of endocannabinoid synthesis, and
is consistent with the apparent interaction between metabotropic glutamate receptor signaling
and fast glucocorticoid signaling shown in Chapter 4. Figure 1 outlines how this proposed
127 signaling pathway could work.
A physical interaction between GR and group I metabotropic glutamate receptors
mediated by Homer proteins could also explain the difference between the effects of AM-251
treatment and hexyl-HIBO treatment on acute stress-mediated HPA axis activity. It is reasonable
to believe that if a proposed GR-Homer-metabotropic glutamate receptor complex exists, it does
not include all of the functional metabotropic glutamate receptors present in a given cell. The
elevation of ACTH induced by treatment with the antagonist hexyl-HIBO suggests that there is
inhibitory tone on the HPA axis that is mediated through the group I metabotropic glutamate
receptors. However, a lack of change in the stress-induced HPA axis responses induced by
treatment with AM-251 suggests that this inhibitory tone is not mediated by signaling through
the endocannabinoid pathway. Thus, it may be that metabotropic glutamate receptors are
signaling through multiple pathways (e.g. inhibition through endocannabinoid signaling and
excitation of inhibitory interneurons; see Figure 3), but fast feedback inhibition of the HPA axis
is only signaling through the endocannabinoid pathway.
A second candidate for the receptor mediating fast feedback is the cytosolic GR. For example, glucocorticoid binding to the GR complex could cause dissociation of the GR-heat shock protein complex and subsequent signaling via another member of this complex. This type of signaling pathway has been observed in some immune cells, where GR forms part of the T- cell receptor complex. Binding of glucocorticoid to this complex causes dissociation of the T- cell receptor complex followed by impaired signaling through the T-cell receptor complex
(Lowenberg et al., 2007). Other proteins could also signal through a similar mechanism, thus making the cytosolic GR-HSP complex the receptor mediating fast feedback effects.
Fast feedback effects could also be mediated by a GR-independent signaling pathway.
128 One possibility involves putative GPCR proteins that act as glucocorticoid receptors, but are
unrelated to GR. Such a GPCR protein has been described in the amphibian Taricha granulosa
(Evans et al., 2000), but this receptor has not yet been cloned, and a mammalian cognate receptor
has yet to be discovered. Also, a GPCR for progesterone has been discovered and cloned, which
is unrelated to the cytosolic progesterone steroid receptor (Zhu et al., 2003). Together, these
findings suggest that a GR-unrelated GPCR for glucocorticoids may exist, but such a receptor
has not yet been discovered.
Glucocorticoids could also be acting by binding to and modulating the activity of other
receptors. Glucocorticoid-derived neurosteroids are known to bind to and alter the signaling of
GABAA receptors (Orchinik et al., 1994), and thus change ion flux across the affected
membranes. Although in this case the affected GABAA receptor could be called a glucocorticoid
receptor, it would seem more correct to call glucocorticoids modulators of the GABAA receptor.
A final possibility is that the fast feedback effects of glucocorticoids occur independent of
a receptor, through direct interactions with cell membranes. Cell membranes are made up largely
of phospholipids and embedded proteins. The fluidity of the phospholipid bilayer is partly determined by the characteristics of the molecules comprising the membrane. For example, cholesterol tends to make a membrane more rigid, while unsaturated fatty acids moderate this rigidity (Subczynski and Wisniewska, 2000). Unsaturated fatty acids can also increase the fluidity of a membrane, especially if the membrane is near its phase transition temperature
(Stubbs and Smith, 1984). The fluidity of a membrane affects the properties of the membrane, and can also affect the activity of proteins, such as ion channels, embedded in the membrane
(Tillman and Cascio, 2003). Steroids are thought to act on neuronal cell membranes in a way
that alters membrane fluidity (due perhaps to the similarity in structure between steroid
129 hormones and cholesterol), and thus affects the flux of ions across the membranes (Buttgereit et
al., 1998). These effects can be highly specific for individual steroid hormones (Massa et al.,
1975). Further work is necessary to determine which of these possibilities is responsible for
mediating glucocorticoid signaling in fast feedback inhibition of the HPA axis.
Endocannabinoid receptor signaling in HPA axis regulation
The data described suggest an active role for endocannabinoid signaling in regulating the
HPA axis response to restraint stress. The most direct evidence for this comes from the
demonstration that pharmacological blockade of CB1 receptors antagonizes dexamethasone-
induced fast negative feedback. This result points to an endocannabinoid-mediated step
downstream from the initial action of glucocorticoids on the HPA axis at the PVN, similar to the
model proposed by the Tasker lab (Di et al., 2003). This interpretation is complicated somewhat
by the fact that cannabinoid receptor activation by the cannabinoid Win 55,212-2 fails to mimic the actions of dexamethasone, suggesting that cannabinoid signaling per se is not sufficient to cause fast feedback inhibition of the HPA axis. Some possible explanations for necessity without sufficiency could include multiple cannabinoid receptors being involved in fast feedback, or multiple synapse types (e.g. glutamate and GABA) being under cannabinoid control.
Multiple CB receptors involved in HPA axis regulation in PVN?
Our data on cannabinoid drugs in the HPA axis raise the possibility that multiple receptor
types may be at work in the PVN. The most direct evidence for this is the difference between the
effects of dexamethasone and Win55,212-2 treatment. Dexamethasone rapidly causes
130 attenuation of the ACTH response to restraint by a mechanism that is dependent on signaling
through CB1. Win 55,212-2 on the other hand, leads to a potentiation of the ACTH response to
restraint, even though it is purportedly signaling through the same pathway (CB1) as
dexamethasone. We have advanced an explanation for this in Chapter 4, where the differences in
effects are due to multiple CB1 receptor-containing synapses at CRH cells. According to this
explanation, some of these synapses are excitatory, and are inhibited by cannabinoid signaling
initiated by dexamethasone. Others are inhibitory, and their activity is also decreased by
cannabinoid signaling; however, dexamethasone does not initiate cannabinoid signaling at these
inhibitory synapses, consistent with a literature suggesting that glucocorticoid-mediated effects
in the hypothalamus are mediated by excitatory, rather than inhibitory synapses (Jones et al.,
1977; Di et al., 2003).
An alternate explanation for these data involves multiple types of cannabinoid receptor.
AM-251 is selective for CB1 over CB2, while Win 55,212-2 has activity at both CB1 and CB2.
CB2 inhibition by Win 55,212-2 has typically been ignored by investigators examining
endocannabinoid effects on neuronal transmission, most likely because CB2 expression has been
reported to be absent from the brain (Galiegue et al., 1995). More recent evidence suggests that
CB2 is in fact expressed in brain microglial cells (Nunez et al., 2004), as well as in astrocytes
(Sheng et al., 2005) and some populations of neurons (Gong et al., 2006). CB1 is expressed in the PVN, where it is found presynaptically (Marsicano and Lutz, 1999; Castelli et al., 2007).
CB2, on the other hand has been found grossly in the hypothalamus, where it is mostly a post-
synaptic receptor (Onaivi et al., 2006), but it is currently thought to be absent from the PVN
(Gong et al., 2006). Thus, it does not seem likely that there is a role for CB2 in regulating the
HPA axis at the level of the PVN. Future studies will be needed to rule this out conclusively.
131 General Considerations
Methods.
The described studies were undertaken mainly with an approach that uses awake, cannulated animals receiving injections, followed by restraint challenge and measurement of plasma hormones to deduce information regarding the activity of the HPA axis. These methods bear some commenting, both for their strengths and also their weaknesses.
Injecting awake animals by cannula has several distinct advantages. These include relative anatomic specificity of injections, absence of the interfering influence of anesthetics, and the ability to subject the rats to a restraint challenge immediately after infusion of a drug.
Direct injection of drug into the PVN is a more anatomically specific approach than
others that are available. Many published HPA axis studies have used intraperitoneal,
intravenous, or intracerebroventricular infusions. Peripheral injections are much easier to
administer than central injections, and have the advantage of near uniformity in drug exposure
between subjects. That is, the amount and relative distribution of the drug, and the affected areas
will be very similar from animal to animal. However, it is not possible to narrow down the site
of effect of a drug given in this manner, beyond it being somewhere in the pharmacokinetic
distribution of the drug. Further, in the case of drugs that do not cross the blood-brain barrier, it
is not really possible to determine central effects using a peripheral injection, since very little of
the drug will actually be in the central nervous system. Intracerebroventricular injections suffer
from similar problems. Although the extent of spread of an intracerebroventricular injection is
certainly not uniform over the entire brain, there is still a considerable swath of periventricular
tissue that will likely be affected (Francis et al., 2006). Thus, intracerebral injection is the most
anatomically specific method that we have available for use in vivo at this time.
132 Injecting awake rats by cannula also obviates confounds inherent in injecting an animal
under anesthesia. Although injection of a brain region through a syringe mounted on stereotactic
apparatus is at least as anatomically specific as cannula injections, analysis of these experiments is compounded by the fact that anesthetics can potently induce HPA axis activity (Vahl et al.,
2005). Also, it is not clear whether the same brain pathways are active under anesthesia as in an
awake animal, especially since many anesthetic drugs modulate brain GABA activity. Thus, from experiments performed on anesthetized animals, it is not possible to determine whether an observed effect is due to a direct effect of the drug, an interaction with the anesthetic, or an interaction with the HPA axis itself. Our studies have the advantage that they are given in awake, behaving animals; thus, observed effects are more likely to be due to actual modulatory actions
of the drugs on the HPA axis than to an artifact of anesthesia. Further, since the rats are awake
and able to behave normally, we are able to measure the modulation of responses to restraint or other stress modalities in these experiments.
One of the problems with the approach used in these studies is that, with the exception of the dex:BSA compounds, there is not a good way to determine the exact areas that receive injections. With Nissl staining, as I have used, it is simple to determine the placement of the guide cannulas. It is also relatively easy to find the position of the internal cannula if enough sections are stained. However, it is obvious that the injected drugs will diffuse from the initial injection site, in some cases affecting other nearby areas of the brain. There is also a possibility that injections given close to the third ventricle (in the PVN or median eminence, for example) could enter the ventricular system and thus have effects distant from the injection site. Finally, because of the limitations of the equipment used, it is not always clear whether the drug has
actually been injected (although an effect of treatment suggests that the drug was injected).
133 The degree of diffusion of an injection is dependent on such characteristics as the size of
the injection, the lipophilicity of the drug injected, the length of time from injection to observed
effect, and the size of the molecule. In many of the above experiments, I have injected Chicago
Sky Blue® dye as a surrogate for observing the presence of an injection and the approximate degree of spread of the injection. I have compared the spread of dye to the spread of dex:BSA as
measured by anti-BSA immunostaining and found the two to be closely correlated. I have also injected dex:BSA and dexamethasone into adrenalectomized rats, and the spread of dye once again was similar to the spread of dexamethasone, as observed by GR translocation at the injection site. The spread of GR translocation was also comparable to the spread of BSA immunoreactivity. From these approaches I have been able to approximate the spread of drugs when they are injected into the PVN. In the case of the GluR5 experiments, where the injection volume was 100 nl, the injections through correctly placed cannulas are unlikely to have spread significantly beyond the median eminence or PVN. Thus, I have reason to believe with a fair degree of confidence that the results of the GluR5 experiments are due to actions of the drug in the structure that was examined in a given experiment.
In the case of the feedback and metabotropic experiments, where the injection volume was 500 nl, there is a possibility that observed effects were due to actions of the drugs outside the
PVN. Immunohistochemistry against BSA in the case of dex:BSA injections suggests that the degree of spread into the ventricular system is very small. I was not able to detect BSA staining in the ventricles, except in cases when cannula placement by Nissl stain revealed an injection site in the ventricle itself. This suggests that the brain-cerebrospinal fluid barrier is intact in PVN injected animals. Beyond this, the spread of dye and dex:BSA both suggest that the effects observed in these experiments could possibly be due to actions of the drugs at sites including the
134 peri-PVN and sub-PVN regions, and thalamic nuclei such as the reuniens nucleus that are
immediately dorsal to the PVN. There is precedence for both the peri-PVN (Ziegler and
Herman, 2000) and sub-paraventricular hypothalamus (Roland and Sawchenko, 1993; Boudaba
et al., 1997) being involved in regulation of the HPA axis by signaling through the PVN. Thus,
although it is possible that the drug treatments described in this work are having their effects via
extra-PVN sites, it is very likely that they are occurring by actions very close to the PVN.
Determining whether an injection has occurred is a fairly straightforward problem to
address. Mixing the drug with Chicago Sky Blue® dye allows the direct observation of whether
an injection has taken place. In injected animals, a dye spot is visible upon cutting the brains on
the microtome (in most cases dye spots visible on cutting are no longer visible after the section
has been washed, stained, and mounted on slides). The presence of dye spots was critical for
evaluating injection sites for the GluR5 experiments, because the 100 nl injection size was not
large enough to see the drug moving through the injection tubing. Thus, blue dye was used in all experiments in which this small volume injection was used. In the case of the 500nl injections, on the other hand, it was possible to see the injected fluid entering the clear injection tubing above the top of the injector cannula, then leaving as the injection was given.
One other consideration to keep in mind with these experiments is that injecting an awake
animal as I have done leads to a stress response to the injection itself. This early stress response
has the potential of masking results of the drug treatment. For example, there is recent evidence
that fast feedback has no effect on stress hormone pulses in progress at the time of glucocorticoid
administration (Atkinson et al., 2008). Also, since there is a maximum secretory ability of the
pituitary and adrenals, it is possible to reach a “ceiling” effect, where further stress stimulus is
unable to increase plasma hormones further. The ceiling effect is unlikely to be a consideration
135 in these studies, since the maximum ACTH responses to injection and restraint combined seen in
these studies are usually in the 300 pg/ml range. In other studies in the Herman lab we have
typically seen values around 400 pg/ml in response to restraint, and as high as 600 pg/ml, in
Sprague-Dawley rats (Ziegler and Herman, 2000; Vahl et al., 2005; Tauchi et al., 2008),
suggesting the ceiling ACTH responses are significantly higher than the responses achieved in
these studies..
Necessity vs. sufficiency of corticosteroids in fast feedback
It is clear from both the current work and previous work by others that glucocorticoids
are sufficient to induce rapid negative feedback inhibition of the HPA axis. However, most of
the studies investigating this phenomenon have not attempted to determine whether
glucocorticoids are necessary for this effect. Thus, it is possible that, although exogenous
glucocorticoids rapidly inhibit the HPA axis, fast feedback in vivo is caused by neuronal
signaling or some other glucocorticoid-independent mechanism.
There is evidence suggesting that glucocorticoids are necessary for fast feedback to occur.
For example, immobilization stress causes an almost complete attenuation of the CRH response
to a second immobilization stress occurring 2 minutes after the first (Sakakura et al., 1976).
Although this study did not examine glucocorticoids specifically, it does suggest that fast
feedback is an endogenous process that does occur in the absence of pharmacological treatment
with glucocorticoids. In another experiment, anesthetized rats were given a histamine injection
to activate the HPA axis, while intravenous corticosterone was given at either constant or rising
levels. Only when the plasma corticosterone level was rising at 6 μg/100ml/min or faster did fast feedback occur (Kaneko and Hiroshige, 1978). A final piece of evidence suggesting that
136 glucocorticoids are necessary for fast feedback is that adrenalectomized rats fail to exhibit
attenuated ACTH responses to a second stressor, as compared to intact rats (De Souza and Van
Loon, 1989). Although the complete lack of glucocorticoids in the latter adrenalectomized rats
complicates the analysis of these data, these results still suggest that glucocorticoids are necessary for fast feedback. Further work is still needed to confirm this, however.
Dissociation between ACTH and corticosterone
One of the phenomena observed over much of this work is a dissociation between ACTH
and corticosterone responses to stress, especially when comparing the effects of treatments in the
fast feedback and metabotropic glutamate experiments. Pharmacological manipulation of the
HPA axis using glucocorticoid, cannabinoid, and metabotropic glutamatergic drugs has
consistently led to a more pronounced effect on ACTH than on corticosterone. Since ACTH
induces the corticosterone response, it would seem logical that a difference in ACTH between
two treatments would lead to an approximately equal relative change in the corticosterone
response. This is clearly not the case.
This phenomenon of dissociated ACTH and corticosterone secretion is actually fairly
widespread, but only recently has it begun to be appreciated (see (Bornstein et al., 2008) for a
comprehensive review). Typically what is seen is an increase in corticosterone without a concomitant increase in ACTH, although there is at least one report of ACTH being increased in the absence of a resultant increase in corticosterone secretion (Sutherland et al., 2003). This can be accomplished by altered adrenal sensitivity to ACTH, as well as through other, poorly characterized pathways.
A disconnect between the corticosterone and ACTH responses to a stressor, especially
137 when there is a greater change in the ACTH response than in the corticosterone response, begs the question of whether such a regulatory mechanism is relevant to the organism. Since the main functional output of activation of the HPA axis is glucocorticoid secretion, it is reasonable to
suppose that a change in ACTH secretion in the absence of changes in corticosterone secretion is
functionally irrelevant, even if it does contribute incrementally to an understanding of how the
HPA axis is regulated. If ACTH has effects independent of causing corticosterone secretion,
however, increased ACTH secretion without a resultant increase in corticosterone would be relevant. In fact, there is a small amount of evidence in the literature that ACTH does have actions independent of stimulating glucocorticoid secretion.
Melanocortin 2 receptors, which are the ACTH receptor in the adrenal cortex, are also expressed in adipose tissue of rats (Boston and Cone, 1996; Noon et al., 2006) and mice (Kubo et al., 2004), and pancreatic islet cells (Al-Majed et al., 2004), and ACTH binding sites in the brain have also been identified (Tatro, 1990). Further, several reports of adrenal-independent actions of ACTH must be considered. For example, ACTH is able to cause localized increases in capillary permeability after injection, independent of corticosteroid actions (Menkin, 1957).
ACTH is also able to alter gene expression in cervical ganglia in the absence of adrenal glands
(Serova et al., 2008), to increase insulin release from insulinoma and primary pancreatic islet cells (Al-Majed et al., 2004), and to induce lipolysis in rat adipocytes (Oelofsen and
Ramachandran, 1983). In addition to these, there are also a number of studies detailing actions of ACTH and fragments of ACTH on animal behavior (De Wied and Jolles, 1982). These corticosteroid-independent actions of ACTH suggest that alterations in ACTH secretion are relevant, even in the absence of significant changes in corticosterone.
138 Future Directions
The results of these studies on fast feedback inhibition of the HPA axis reveal a need for
reductionist approaches in teasing out the signaling pathways involved in this process. In order
to do this, however, a more solid connection must be demonstrated between the signaling shown in vitro and the actual in vivo regulation of the HPA axis. For example, the results of these studies, combined with those of the Tasker lab suggest that changes in glutamate signaling
underlie fast feedback in the PVN. However, the currently available evidence is circumstantial.
It is true that glucocorticoid treatment rapidly decreases glutamate signaling onto PVN neurons
and that it also decreases the ACTH response to stress. However there is not yet conclusive
evidence that it is the changes in glutamate signaling that cause the decrease in ACTH release.
In spite of this weakness, it does appear likely that changes in glutamate signaling at the PVN are
responsible for mediating fast feedback inhibition of the HPA axis.
Once this connection between the in vitro models of fast feedback signaling and fast
feedback per se is firmly established, it seems likely that in vitro models will be of great value in
discovering the signaling pathway(s) that mediate(s) nongenomic glucocorticoid actions in
inhibiting the HPA axis response to restraint. The results of the in vivo experiments, while
validating a role for rapid glucocorticoid signaling in the fast feedback response, also
demonstrated a complexity in signaling at the PVN that cannot be overcome with currently
available methods for in vivo study. For example, there is a possibility that multiple synapse
types at the same cell could be mediating opposite effects via similar messengers (like
endocannabinoids). Currently available methods for in vivo study are unable to differentiate
between signaling at a single cell. While it may be that there are subtle differences, such as in
receptor subtypes, between these centers of action, it appears that it will be much easier to
139 discover these differences in vitro, then validate them in vivo, rather than attempting to understand subtle differences by in vivo experimentation.
Conclusion
This work has demonstrated that there are a number of signaling pathways that are involved in rapid regulation of the HPA axis. Glutamate, through both metabotropic and ionotropic glutamate receptor pathways, regulates the HPA axis. Glucocorticoids also rapidly regulate the HPA axis, in an endocannabinoid-mediated signaling pathway. While it is not entirely surprising that glutamate rapidly regulates neural activity, the idea that glucocorticoids can regulate neural activity in a nongenomic fashion is a relatively new field (in spite of scattered reports that have appeared since the 1940s), where a great deal of important work is yet to be done.
Much of the work that has been done examining negative feedback regulation of the HPA axis has focused on signaling mediated by genomic signaling through GR and MR. This type of negative feedback is known to take more than an hour to develop (Keller-Wood and Dallman,
1984). When contemplating the place of negative feedback regulation of the HPA axis in nature, however, it becomes clear that a significant proportion of acute stress experiences are ended long before genomically mediated fast feedback would be able to shut of the response. For example, an antelope who encounters a wolf will mount a fight or flight response, including activation of the HPA axis, in its attempt to escape. In many cases, this will lead to tremendous expenditures of energy as the animals run, but the entire experience is likely to be ended with escape or meal- time within a few minutes, rather than hours after the initiation of the response.
If the main purpose of glucocorticoid secretion is, as has been postulated, to help an
140 animal recover from stress rather than to cope with it directly (Sapolsky et al., 2000), then it would seem apparent that negative feedback of the HPA axis response to stress should operate on the scale of at least hours; in other words, on the intermediate feedback time scale. Thus, our antelope would escape from the wolf on the same time frame as the glucocorticoid response is being initiated in earnest, and would then have several hours of high glucocorticoid levels to aid the animal in recovering energy stores, repairing damage, and preparing for the next such encounter.
The presence of the fast negative feedback signaling pathway suggests that there is another purpose for the HPA axis response, which responds much faster to glucocorticoid hormones than the stress recovery response. The existence of such a rapid system is supported by the wealth of evidence illustrating rapid, nongenomic actions of glucocorticoids in physiological processes. One area in which this response appears to be critical is in the immune response. While much of the immune response is relatively slow when compared with the nervous system, there are some responses, such as anaphylaxis, which develop very rapidly.
Obviously, it would be very detrimental to an antelope's well-being to experience anaphylactic shock while being chased by a wolf; thus, a rapid system for delaying such a response would be of great benefit to an animal. There is evidence that glucocorticoids rapidly inhibit such responses, by blocking histamine release from mast cells, for example (Liu et al., 2007).
Similarly, glucocorticoids induce a number of nongenomically mediated responses in the central nervous system (see introduction). Thus, it seems that, while glucocorticoid secretion is likely to be critical in long-term survival, such as in recovering from stressful experiences, it is also important in rapid responses to stress.
In addition to the physiological role of rapid glucocorticoid actions in HPA axis
141 regulation, the existence of these nongenomic glucocorticoid actions has significance in the way
we treat disease. Nongenomic actions of glucocorticoids indicate that drugs with specificity for
nongenomic or genomic actions could be exploited, especially because there are many conditions
that glucocorticoids appear to play a part in. HPA axis dysfunction is a common finding in major depressive disorders (Holsboer, 2001), and successful treatment of patients with this subtype of depression leads to normalization of the axis. HPA axis dysfunction has also been associated with other neural disorders, including fibromyalgia (Parker et al., 2001), chronic fatigue
syndrome (Parker et al., 2001) and other pain syndromes (Lariviere and Melzack, 2000; Gaab et
al., 2005), and Alzheimer's Disease (Landfield and Eldridge, 1991). Although it is not yet clear
what role glucocorticoids may be playing in these disorders, the apparent involvement of HPA
axis dysfunction suggests that these hormones may be playing a role in these diseases.
Also importantly, it appears that synthetic glucocorticoids already in clinical use vary in
their ability to activate genomic vs. nongenomic effects (Croxtall et al., 2002). These differences
could be directly exploited by tailoring the choice of steroid to the pathways (genomic or
nongenomic) that are primarily at work in the pathology of a given condition. Or it may be that
these differences need to be considered with regard to the side-effect profile of glucocorticoids
used primarily for their genomic activity. Reported differences between already existing drugs
further suggest that better pharmacological specificity can yet be attained in corticosteroid drugs,
and highlight the importance of examining both genomic and nongenomic effects of candidate
steroids during the drug development process.
In conclusion, the HPA axis is clearly regulated on a rapid timescale by both glutamate
and glucocorticoids. Although this should not be surprising, given that the central nervous
system does rapidly react to stimuli in many, if not most cases, much of the work that has been
142 done, at least in the glucocorticoid field, has focused on slower events (usually those taking 2 hours or more to develop). This study of rapid HPA axis regulation has proven to be a rich field for investigation, and there is a great deal more work to be done in order to understand regulation by glucocorticoids and glutamate.
Figure Legends
Figure 1. Model for Homer-2-mediated interaction of GR and group I mGluR. One
explanation for how a membrane-bound form of GR could cause endocannabinoid synthesis and
release is that the protein Homer-2 could link GR with another protein. Using a group I
metabotropic glutamate receptor as an example, this is how this could occur. A) Glucocorticoid
binds to the membrane-bound GR. B) This binding induces a conformational change that is
transduced through Homer-2 to the metabotropic glutamate receptor. C) Activation of the
metabotropic glutamate receptor leads to activation of phospholipase C (PLC) through the Gq subunit associated with the glutamate receptor. D) PLC activation leads to synthesis of the endocannabinoid 2-arachidonyl glycerol (2-AG), which is then released and signals through the
CB1 receptor pathway.
143 Figure 5-1 Model for Homer-2 Mediation of Membrane GR Signaling
144 References
Abe T, Sugihara H, Nawa H, Shigemoto R, Mizuno N, Nakanishi S (1992) Molecular characterization of a novel metabotropic glutamate receptor mGluR5 coupled to inositol phosphate/Ca2+ signal transduction. J Biol Chem 267:13361-13368.
Abou-Samra AB, Catt KJ, Aguilera G (1986) Biphasic inhibition of adrenocorticotropin release by corticosterone in cultured anterior pituitary cells. Endocrinology 119:972-977.
Adams R (1941) Marihuana. Harvey Lect 37:168-197.
Adermark L, Lovinger DM (2007) Retrograde endocannabinoid signaling at striatal synapses requires a regulated postsynaptic release step. Proc Natl Acad Sci U S A 104:20564- 20569.
Akana SF, Chu A, Soriano L, Dallman MF (2001) Corticosterone exerts site-specific and state- dependent effects in prefrontal cortex and amygdala on regulation of adrenocorticotropic hormone, insulin and fat depots. J Neuroendocrinol 13:625-637.
Al-Majed HT, Jones PM, Persaud SJ, Sugden D, Huang GC, Amiel S, Whitehouse BJ (2004) ACTH stimulates insulin secretion from MIN6 cells and primary mouse and human islets of Langerhans. J Endocrinol 180:155-166.
Alger BE (2002) Retrograde signaling in the regulation of synaptic transmission: focus on endocannabinoids. Prog Neurobiol 68:247-286.
Anderson RG (1998) The caveolae membrane system. Annu Rev Biochem 67:199-225.
Anisman H, Zaharia MD, Meaney MJ, Merali Z (1998) Do early-life events permanently alter behavioral and hormonal responses to stressors? Int J Dev Neurosci 16:149-164.
Aroniadou-Anderjaska V, Qashu F, Braga MF (2007) Mechanisms regulating GABAergic inhibitory transmission in the basolateral amygdala: implications for epilepsy and anxiety disorders. Amino Acids 32:305-315.
Atkinson HC, Wood SA, Castrique ES, Kershaw YM, Wiles CC, Lightman SL (2008) Corticosteroids mediate fast feedback of the rat hypothalamic-pituitary-adrenal axis via the mineralocorticoid receptor. Am J Physiol Endocrinol Metab 294:E1011-1022.
Barna I, Zelena D, Arszovszki AC, Ledent C (2004) The role of endogenous cannabinoids in the hypothalamo-pituitary-adrenal axis regulation: in vivo and in vitro studies in CB1 receptor knockout mice. Life Sci 75:2959-2970.
Barnett V, Lewis T (1994) Outliers in statistical data, 3rd Edition. New York: John Wiley & Sons.
145 Bartis D, Boldizsar F, Szabo M, Palinkas L, Nemeth P, Berki T (2006) Dexamethasone induces rapid tyrosine-phosphorylation of ZAP-70 in Jurkat cells. J Steroid Biochem Mol Biol.
Bartis D, Boldizsar F, Kvell K, Szabo M, Palinkas L, Nemeth P, Monostori E, Berki T (2007) Intermolecular relations between the glucocorticoid receptor, ZAP-70 kinase, and Hsp- 90. Biochem Biophys Res Commun 354:253-258.
Benarroch EE (2008) Metabotropic glutamate receptors: synaptic modulators and therapeutic targets for neurologic disease. Neurology 70:964-968.
Binns KE, Turner JP, Salt TE (2003) Kainate receptor (GluR5)-mediated disinhibition of responses in rat ventrobasal thalamus allows a novel sensory processing mechanism. J Physiol 551:525-537.
Bisogno T, MacCarrone M, De Petrocellis L, Jarrahian A, Finazzi-Agro A, Hillard C, Di Marzo V (2001) The uptake by cells of 2-arachidonoylglycerol, an endogenous agonist of cannabinoid receptors. Eur J Biochem 268:1982-1989.
Bornstein SR, Engeland WC, Ehrhart-Bornstein M, Herman JP (2008) Dissociation of ACTH and glucocorticoids. Trends Endocrinol Metab 19:175-180.
Borski RJ (2000) Nongenomic membrane actions of glucocorticoids in vertebrates. Trends Endocrinol Metab 11:427-436.
Borski RJ, Hyde GN, Fruchtman S (2002) Signal transduction mechanisms mediating rapid, nongenomic effects of cortisol on prolactin release. Steroids 67:539-548.
Boston BA, Cone RD (1996) Characterization of melanocortin receptor subtype expression in murine adipose tissues and in the 3T3-L1 cell line. Endocrinology 137:2043-2050.
Boudaba C, Schrader LA, Tasker JG (1997) Physiological evidence for local excitatory synaptic circuits in the rat hypothalamus. J Neurophysiol 77:3396-3400.
Bradbury MJ, Giracello DR, Chapman DF, Holtz G, Schaffhauser H, Rao SP, Varney MA, Anderson JJ (2003) Metabotropic glutamate receptor 5 antagonist-induced stimulation of hypothalamic-pituitary-adrenal axis activity: interaction with serotonergic systems. Neuropharmacology 44:562-572.
Braga MF, Aroniadou-Anderjaska V, Xie J, Li H (2003) Bidirectional modulation of GABA release by presynaptic glutamate receptor 5 kainate receptors in the basolateral amygdala. J Neurosci 23:442-452.
Brakeman PR, Lanahan AA, O'Brien R, Roche K, Barnes CA, Huganir RL, Worley PF (1997) Homer: a protein that selectively binds metabotropic glutamate receptors. Nature 386:284-288.
146 Brann DW (1995) Glutamate: a major excitatory transmitter in neuroendocrine regulation. Neuroendocrinology 61:213-225.
Brasier DJ, Feldman DE (2008) Synapse-specific expression of functional presynaptic NMDA receptors in rat somatosensory cortex. J Neurosci 28:2199-2211.
Breivogel CS, Griffin G, Di Marzo V, Martin BR (2001) Evidence for a new G protein-coupled cannabinoid receptor in mouse brain. Mol Pharmacol 60:155-163.
Breuner CW, Greenberg AL, Wingfield JC (1998) Noninvasive corticosterone treatment rapidly increases activity in Gambel's white-crowned sparrows (Zonotrichia leucophrys gambelii). Gen Comp Endocrinol 111:386-394.
Buckingham JC (2006) Glucocorticoids: exemplars of multi-tasking. Br J Pharmacol 147 Suppl 1:S258-268.
Buckingham JC, John CD, Solito E, Tierney T, Flower RJ, Christian H, Morris J (2006) Annexin 1, glucocorticoids, and the neuroendocrine-immune interface. Ann N Y Acad Sci 1088:396-409.
Buttgereit F, Scheffold A (2002) Rapid glucocorticoid effects on immune cells. Steroids 67:529- 534.
Buttgereit F, Wehling M, Burmester GR (1998) A new hypothesis of modular glucocorticoid actions: steroid treatment of rheumatic diseases revisited. Arthritis Rheum 41:761-767.
Buttgereit F, Brand MD, Burmester GR (1999) Equivalent doses and relative drug potencies for non-genomic glucocorticoid effects: a novel glucocorticoid hierarchy. Biochem Pharmacol 58:363-368.
Campbell SL, Mathew SS, Hablitz JJ (2007) Pre- and postsynaptic effects of kainate on layer II/III pyramidal cells in rat neocortex. Neuropharmacology 53:37-47.
Cannon WB (1939) The wisdom of the body. New York: W.W. Norton & Company.
Carlson HE, Miglietta JT, Roginsky MS, Stegink LD (1989) Stimulation of pituitary hormone secretion by neurotransmitter amino acids in humans. Metabolism 38:1179-1182.
Castelli MP, Piras AP, Melis T, Succu S, Sanna F, Melis MR, Collu S, Grazia Ennas M, Diaz G, Mackie K, Argiolas A (2007) Cannabinoid CB1 receptors in the paraventricular nucleus and central control of penile erection: immunocytochemistry, autoradiography and behavioral studies. Neuroscience 147:197-206.
Chapman L, Nishimura A, Buckingham JC, Morris JF, Christian HC (2002) Externalization of annexin I from a folliculo-stellate-like cell line. Endocrinology 143:4330-4338.
147 Charalampopoulos I, Alexaki VI, Lazaridis I, Dermitzaki E, Avlonitis N, Tsatsanis C, Calogeropoulou T, Margioris AN, Castanas E, Gravanis A (2006) G protein-associated, specific membrane binding sites mediate the neuroprotective effect of dehydroepiandrosterone. Faseb J 20:577-579.
Chen F, Watson CS, Gametchu B (1999) Multiple glucocorticoid receptor transcripts in membrane glucocorticoid receptor-enriched S-49 mouse lymphoma cells. J Cell Biochem 74:418-429.
Chittajallu R, Braithwaite SP, Clarke VR, Henley JM (1999) Kainate receptors: subunits, synaptic localization and function. Trends Pharmacol Sci 20:26-35.
Choi DC, Furay AR, Evanson NK, Ostrander MM, Ulrich-Lai YM, Herman JP (2007) Bed nucleus of the stria terminalis subregions differentially regulate hypothalamic-pituitary- adrenal axis activity: implications for the integration of limbic inputs. J Neurosci 27:2025-2034.
Christian HC, Taylor AD, Flower RJ, Morris JF, Buckingham JC (1997) Characterization and localization of lipocortin 1-binding sites on rat anterior pituitary cells by fluorescence- activated cell analysis/sorting and electron microscopy. Endocrinology 138:5341-5351.
Clarke VR, Ballyk BA, Hoo KH, Mandelzys A, Pellizzari A, Bath CP, Thomas J, Sharpe EF, Davies CH, Ornstein PL, Schoepp DD, Kamboj RK, Collingridge GL, Lodge D, Bleakman D (1997) A hippocampal GluR5 kainate receptor regulating inhibitory synaptic transmission. Nature 389:599-603.
Coddington E, Lewis C, Rose JD, Moore FL (2007) Endocannabinoids mediate the effects of acute stress and corticosterone on sex behavior. Endocrinology 148:493-500.
Corchero J, Fuentes JA, Manzanares J (1999) Chronic treatment with CP-55,940 regulates corticotropin releasing factor and proopiomelanocortin gene expression in the hypothalamus and pituitary gland of the rat. Life Sci 64:905-911.
Croxtall JD, Choudhury Q, Flower RJ (2000) Glucocorticoids act within minutes to inhibit recruitment of signalling factors to activated EGF receptors through a receptor- dependent, transcription-independent mechanism. Br J Pharmacol 130:289-298.
Croxtall JD, van Hal PT, Choudhury Q, Gilroy DW, Flower RJ (2002) Different glucocorticoids vary in their genomic and non-genomic mechanism of action in A549 cells. Br J Pharmacol 135:511-519.
Dafny N, Phillips MI, Taylor AN, Gilman S (1973) Dose effects of cortisol on single unit activity in hypothalamus, reticular formation and hippocampus of freely behaving rats correlated with plasma steroid levels. Brain Res 59:257-272.
Danik M, Cassoly E, Manseau F, Sotty F, Mouginot D, Williams S (2005) Frequent coexpression
148 of the vesicular glutamate transporter 1 and 2 genes, as well as coexpression with genes for choline acetyltransferase or glutamic acid decarboxylase in neurons of rat brain. J Neurosci Res 81:506-521.
Darlington DN, Miyamoto M, Keil LC, Dallman MF (1989) Paraventricular stimulation with glutamate elicits bradycardia and pituitary responses. Am J Physiol 256:R112-119.
Daufeldt S, Lanz R, Allera A (2003) Membrane-initiated steroid signaling (MISS): genomic steroid action starts at the plasma membrane. J Steroid Biochem Mol Biol 85:9-23.
Daufeldt S, Klein R, Wildt L, Allera A (2006) Membrane initiated steroid signaling (MISS): Computational, in vitro and in vivo evidence for a plasma membrane protein initially involved in genomic steroid hormone effects. Mol Cell Endocrinol.
Dayanithi G, Antoni FA (1989) Rapid as well as delayed inhibitory effects of glucocorticoid hormones on pituitary adrenocorticotropic hormone release are mediated by type II glucocorticoid receptors and require newly synthesized messenger ribonucleic acid as well as protein. Endocrinology 125:308-313.
De Souza EB, Van Loon GR (1989) Rate-sensitive glucocorticoid feedback inhibition of adrenocorticotropin and beta-endorphin/beta-lipotropin secretion in rats. Endocrinology 125:2927-2934.
De Wied D, Jolles J (1982) Neuropeptides derived from pro-opiocortin: behavioral, physiological, and neurochemical effects. Physiol Rev 62:976-1059.
Deutsch DG, Glaser ST, Howell JM, Kunz JS, Puffenbarger RA, Hillard CJ, Abumrad N (2001) The cellular uptake of anandamide is coupled to its breakdown by fatty-acid amide hydrolase. J Biol Chem 276:6967-6973.
Devane WA, Dysarz FA, 3rd, Johnson MR, Melvin LS, Howlett AC (1988) Determination and characterization of a cannabinoid receptor in rat brain. Mol Pharmacol 34:605-613.
Devane WA, Hanus L, Breuer A, Pertwee RG, Stevenson LA, Griffin G, Gibson D, Mandelbaum A, Etinger A, Mechoulam R (1992) Isolation and structure of a brain constituent that binds to the cannabinoid receptor. Science 258:1946-1949.
Dhawan L, Liu B, Blaxall BC, Taubman MB (2007) A novel role for the glucocorticoid receptor in the regulation of monocyte chemoattractant protein-1 mRNA stability. J Biol Chem 282:10146-10152.
Di Marzo V, De Petrocellis L (2006) Non-CB1 non-CB2 receptors for endocannabinoids. Endocannabinoids: The brain and body's marijuana and beyond:151.
Di Marzo V, Melck D, Bisogno T, De Petrocellis L (1998) Endocannabinoids: endogenous cannabinoid receptor ligands with neuromodulatory action. Trends Neurosci 21:521-528.
149 Di Marzo V, Fontana A, Cadas H, Schinelli S, Cimino G, Schwartz JC, Piomelli D (1994) Formation and inactivation of endogenous cannabinoid anandamide in central neurons. Nature 372:686-691.
Di Marzo V, Breivogel CS, Tao Q, Bridgen DT, Razdan RK, Zimmer AM, Zimmer A, Martin BR (2000) Levels, metabolism, and pharmacological activity of anandamide in CB(1) cannabinoid receptor knockout mice: evidence for non-CB(1), non-CB(2) receptor- mediated actions of anandamide in mouse brain. J Neurochem 75:2434-2444.
Di S, Malcher-Lopes R, Halmos KC, Tasker JG (2003) Nongenomic glucocorticoid inhibition via endocannabinoid release in the hypothalamus: a fast feedback mechanism. J Neurosci 23:4850-4857.
Dingledine R, Borges K, Bowie D, Traynelis SF (1999) The glutamate receptor ion channels. Pharmacol Rev 51:7-61.
Doherty J, Dingledine R (2003) Functional interactions between cannabinoid and metabotropic glutamate receptors in the central nervous system. Curr Opin Pharmacol 3:46-53.
Edwardson JA, Bennett GW (1974) Modulation of corticotrophin-releasing factor release from hypothalamic synaptosomes. Nature 251:425-427.
Ehrengruber MU, Kato A, Inokuchi K, Hennou S (2004) Homer/Vesl proteins and their roles in CNS neurons. Mol Neurobiol 29:213-227.
Engeland WC, Miller P, Gann DS (1989) Dissociation between changes in plasma bioactive and immunoreactive adrenocorticotropin after hemorrhage in awake dogs. Endocrinology 124:2978-2985.
Evans SB, Wilkinson CW, Gronbeck P, Bennett JL, Zavosh A, Taborsky GJ, Jr., Figlewicz DP (2004) Inactivation of the DMH selectively inhibits the ACTH and corticosterone responses to hypoglycemia. Am J Physiol Regul Integr Comp Physiol 286:R123-128.
Evans SJ, Murray TF, Moore FL (2000) Partial purification and biochemical characterization of a membrane glucocorticoid receptor from an amphibian brain. J Steroid Biochem Mol Biol 72:209-221.
Eyigor O, Minbay Z, Cavusoglu I, Jennes L (2005) Localization of kainate receptor subunit GluR5-immunoreactive cells in the rat hypothalamus. Brain Res Mol Brain Res 136:38- 44.
Felder CC, Joyce KE, Briley EM, Mansouri J, Mackie K, Blond O, Lai Y, Ma AL, Mitchell RL (1995) Comparison of the pharmacology and signal transduction of the human cannabinoid CB1 and CB2 receptors. Mol Pharmacol 48:443-450.
Feldman S, Weidenfeld J (1997) Hypothalamic mechanisms mediating glutamate effects on the
150 hypothalamo-pituitary-adrenocortical axis. J Neural Transm 104:633-642.
Feldman S, Weidenfeld J (1999) Glucocorticoid receptor antagonists in the hippocampus modify the negative feedback following neural stimuli. Brain Res 821:33-37.
Francis AB, Pace TW, Ginsberg AB, Rubin BA, Spencer RL (2006) Limited brain diffusion of the glucocorticoid receptor agonist ru28362 following i.c.v. administration: Implications for i.c.v. drug delivery and glucocorticoid negative feedback in the hypothalamic- pituitary-adrenal axis. Neuroscience 141:1503-1515.
Fremeau RT, Jr., Burman J, Qureshi T, Tran CH, Proctor J, Johnson J, Zhang H, Sulzer D, Copenhagen DR, Storm-Mathisen J, Reimer RJ, Chaudhry FA, Edwards RH (2002) The identification of vesicular glutamate transporter 3 suggests novel modes of signaling by glutamate. Proc Natl Acad Sci U S A 99:14488-14493.
Freund TF, Katona I, Piomelli D (2003) Role of endogenous cannabinoids in synaptic signaling. Physiol Rev 83:1017-1066.
Gaab J, Baumann S, Budnoik A, Gmunder H, Hottinger N, Ehlert U (2005) Reduced reactivity and enhanced negative feedback sensitivity of the hypothalamus-pituitary-adrenal axis in chronic whiplash-associated disorder. Pain 119:219-224.
Galiegue S, Mary S, Marchand J, Dussossoy D, Carriere D, Carayon P, Bouaboula M, Shire D, Le Fur G, Casellas P (1995) Expression of central and peripheral cannabinoid receptors in human immune tissues and leukocyte subpopulations. Eur J Biochem 232:54-61.
Gametchu B (1987) Glucocorticoid receptor-like antigen in lymphoma cell membranes: correlation to cell lysis. Science 236:456-461.
Gametchu B, Chen F, Sackey F, Powell C, Watson CS (1999) Plasma membrane-resident glucocorticoid receptors in rodent lymphoma and human leukemia models. Steroids 64:107-119.
Gaoni Y, Mechoulam R (1964) Isolation, structure, and partial synthesis of an active constituent of hashish [8]. Journal of the American Chemical Society 86:1646-1647.
Garrett ER (1962) Prediction of stability in pharmaceutical preparations. X. Alkaline hydrolysis of hydrocortisone hemisuccinate. J Pharm Sci 51:445-450.
Gavrilova-Jordan LP, Price TM (2007) Actions of steroids in mitochondria. Semin Reprod Med 25:154-164.
Gay VL, Plant TM (1987) N-methyl-D,L-aspartate elicits hypothalamic gonadotropin-releasing hormone release in prepubertal male rhesus monkeys (Macaca mulatta). Endocrinology 120:2289-2296.
151 Ginsberg AB, Pecoraro NC, Van den Heuvel JK, Horneman HF, Warne JP, Dallman MF (2006) Cannabinoid receptor CB1 blockade rapidly enhances the hormonal response to restraint. In: Society for Neuroscience. Atlanta, Georgia.
Giuditta A, Kaplan BB, van Minnen J, Alvarez J, Koenig E (2002) Axonal and presynaptic protein synthesis: new insights into the biology of the neuron. Trends Neurosci 25:400- 404.
Gold T (1992) The deep, hot biosphere. Proc Natl Acad Sci U S A 89:6045-6049.
Gong JP, Onaivi ES, Ishiguro H, Liu QR, Tagliaferro PA, Brusco A, Uhl GR (2006) Cannabinoid CB2 receptors: immunohistochemical localization in rat brain. Brain Res 1071:10-23.
Greenwood SM, Connolly CN (2007) Dendritic and mitochondrial changes during glutamate excitotoxicity. Neuropharmacology 53:891-898.
Groner B, Hynes NE, Rahmsdorf U, Ponta H (1983) Transcription initiation of transfected mouse mammary tumor virus LTR DNA is regulated by glucocorticoid hormones. Nucleic Acids Res 11:4713-4725.
Guo Z, Chen YZ, Xu RB, Fu H (1995) Binding characteristics of glucocorticoid receptor in synaptic plasma membrane from rat brain. Funct Neurol 10:183-194.
Haller J, Mikics E, Makara GB (2008) The effects of non-genomic glucocorticoid mechanisms on bodily functions and the central neural system. A critical evaluation of findings. Front Neuroendocrinol 29:273-291.
Haller J, Varga B, Ledent C, Barna I, Freund TF (2004) Context-dependent effects of CB1 cannabinoid gene disruption on anxiety-like and social behaviour in mice. Eur J Neurosci 19:1906-1912.
Han JZ, Lin W, Lou SJ, Qiu J, Chen YZ (2002) A rapid, nongenomic action of glucocorticoids in rat B103 neuroblastoma cells. Biochim Biophys Acta 1591:21-27.
Hanus L, Abu-Lafi S, Fride E, Breuer A, Vogel Z, Shalev DE, Kustanovich I, Mechoulam R (2001) 2-arachidonyl glyceryl ether, an endogenous agonist of the cannabinoid CB1 receptor. Proc Natl Acad Sci U S A 98:3662-3665.
Hanus LO (2005) Cannabinoid chemistry: an overview. In: Cannabinoids as Therapeutics (Mechoulam R, ed), pp 23-45. Basel, Switzerland: Birkhauser Verlag.
Hedman E, Widen C, Asadi A, Dinnetz I, Schroder WP, Gustafsson JA, Wikstrom AC (2006) Proteomic identification of glucocorticoid receptor interacting proteins. Proteomics 6:3114-3126.
Heitzer MD, Wolf IM, Sanchez ER, Witchel SF, DeFranco DB (2007) Glucocorticoid receptor
152 physiology. Rev Endocr Metab Disord 8:321-330.
Hennessy MB (1997) Hypothalamic-pituitary-adrenal responses to brief social separation. Neurosci Biobehav Rev 21:11-29.
Herkenham M, Lynn AB, Johnson MR, Melvin LS, de Costa BR, Rice KC (1991) Characterization and localization of cannabinoid receptors in rat brain: a quantitative in vitro autoradiographic study. J Neurosci 11:563-583.
Herman JP, Mueller NK, Figueiredo H (2004) Role of GABA and glutamate circuitry in hypothalamo-pituitary-adrenocortical stress integration. Ann N Y Acad Sci 1018:35-45.
Herman JP, Eyigor O, Ziegler DR, Jennes L (2000) Expression of ionotropic glutamate receptor subunit mRNAs in the hypothalamic paraventricular nucleus of the rat. J Comp Neurol 422:352-362.
Herman JP, Cullinan WE, Ziegler DR, Tasker JG (2002) Role of the paraventricular nucleus microenvironment in stress integration. Eur J Neurosci 16:381-385.
Herman JP, Figueiredo H, Mueller NK, Ulrich-Lai Y, Ostrander MM, Choi DC, Cullinan WE (2003) Central mechanisms of stress integration: hierarchical circuitry controlling hypothalamo-pituitary-adrenocortical responsiveness. Front Neuroendocrinol 24:151- 180.
Hertz L (1979) Functional interactions between neurons and astrocytes I. Turnover and metabolism of putative amino acid transmitters. Prog Neurobiol 13:277-323.
Hertz L, Dringen R, Schousboe A, Robinson SR (1999) Astrocytes: glutamate producers for neurons. J Neurosci Res 57:417-428.
Hibbeln JR, Palmer JW, Davis JM (1989) Are disturbances in lipid-protein interactions by phospholipase-A2 a predisposing factor in affective illness? Biol Psychiatry 25:945-961.
Hinoi E, Takarada T, Ueshima T, Tsuchihashi Y, Yoneda Y (2004) Glutamate signaling in peripheral tissues. Eur J Biochem 271:1-13.
Hinz B, Hirschelmann R (2000) Rapid non-genomic feedback effects of glucocorticoids on CRF- induced ACTH secretion in rats. Pharm Res 17:1273-1277.
Hoffman AF, Lupica CR (2006) Man-made marijuana: Endocannabinoid modulation of synaptic transmission and implications for the regulation of synaptic plasticity. Endocannabinoids: The Brain and Body's Marijuana and Beyond:229.
Hokfelt T, Broberger C, Xu ZQ, Sergeyev V, Ubink R, Diez M (2000) Neuropeptides--an overview. Neuropharmacology 39:1337-1356.
153 Holsboer F (2001) Stress, hypercortisolism and corticosteroid receptors in depression: implications for therapy. J Affect Disord 62:77-91.
Houamed KM, Kuijper JL, Gilbert TL, Haldeman BA, O'Hara PJ, Mulvihill ER, Almers W, Hagen FS (1991) Cloning, expression, and gene structure of a G protein-coupled glutamate receptor from rat brain. Science 252:1318-1321.
Hrabovszky E, Wittmann G, Turi GF, Liposits Z, Fekete C (2005) Hypophysiotropic thyrotropin- releasing hormone and corticotropin-releasing hormone neurons of the rat contain vesicular glutamate transporter-2. Endocrinology 146:341-347.
Huettner JE (2001) Kainate receptors: knocking out plasticity. Trends Neurosci 24:365-366.
Huettner JE (2003) Kainate receptors and synaptic transmission. Prog Neurobiol 70:387-407.
Jaferi A, Bhatnagar S (2006) Corticosterone can act at the posterior paraventricular thalamus to inhibit hypothalamic-pituitary-adrenal activity in animals that habituate to repeated stress. Endocrinology 147:4917-4930.
Jain S, Li Y, Kumar A, Sehgal PB (2005) Transcriptional signaling from membrane raft- associated glucocorticoid receptor. Biochem Biophys Res Commun 336:3-8.
Jaskolski F, Coussen F, Mulle C (2005) Subcellular localization and trafficking of kainate receptors. Trends Pharmacol Sci 26:20-26.
Jasper MS, Engeland WC (1997) Splanchnicotomy increases adrenal sensitivity to ACTH in nonstressed rats. Am J Physiol 273:E363-368.
Jezova D (2005) Control of ACTH secretion by excitatory amino acids: functional significance and clinical implications. Endocrine 28:287-294.
Jezova D, Tokarev D, Rusnak M (1995) Endogenous excitatory amino acids are involved in stress-induced adrenocorticotropin and catecholamine release. Neuroendocrinology 62:326-332.
Joanny P, Steinberg J, Oliver C, Grino M (1997) Glutamate and N-methyl-D-aspartate stimulate rat hypothalamic corticotropin-releasing factor secretion in vitro. J Neuroendocrinol 9:93- 97.
Joels M, Karst H, Alfarez D, Heine VM, Qin Y, van Riel E, Verkuyl M, Lucassen PJ, Krugers HJ (2004) Effects of chronic stress on structure and cell function in rat hippocampus and hypothalamus. Stress 7:221-231.
John C, Cover P, Solito E, Morris J, Christian H, Flower R, Buckingham J (2002) Annexin 1- dependent actions of glucocorticoids in the anterior pituitary gland: roles of the N- terminal domain and protein kinase C. Endocrinology 143:3060-3070.
154 Johnson LR, Farb C, Morrison JH, McEwen BS, LeDoux JE (2005) Localization of glucocorticoid receptors at postsynaptic membranes in the lateral amygdala. Neuroscience 136:289-299.
Johnson MP, Kelly G, Chamberlain M (2001) Changes in rat serum corticosterone after treatment with metabotropic glutamate receptor agonists or antagonists. J Neuroendocrinol 13:670- 677.
Jones MT, Hillhouse EW (1976) Structure-activity relationship and the mode of action of corticosteroid feedback on the secretion of corticotrophin-releasing factor (corticoliberin). J Steroid Biochem 7:1189-1202.
Jones MT, Hillhouse EW, Burden JL (1977) Dynamics and mechanics of corticosteroid feedback at the hypothalamus and anterior pituitary gland. J Endocrinol 73:405-417.
Jonsson KO, Holt S, Fowler CJ (2006) The endocannabinoid system: current pharmacological research and therapeutic possibilities. Basic Clin Pharmacol Toxicol 98:124-134.
Jung KM, Mangieri R, Stapleton C, Kim J, Fegley D, Wallace M, Mackie K, Piomelli D (2005) Stimulation of endocannabinoid formation in brain slice cultures through activation of group I metabotropic glutamate receptors. Mol Pharmacol 68:1196-1202.
Kajantie E (2006) Fetal origins of stress-related adult disease. Ann N Y Acad Sci 1083:11-27.
Kaneko M, Hiroshige T (1978) Fast, rate-sensitive corticosteroid negative feedback during stress. Am J Physiol 234:R39-45.
Kaneko T, Fujiyama F, Hioki H (2002) Immunohistochemical localization of candidates for vesicular glutamate transporters in the rat brain. J Comp Neurol 444:39-62.
Karst H, Berger S, Turiault M, Tronche F, Schutz G, Joels M (2005) Mineralocorticoid receptors are indispensable for nongenomic modulation of hippocampal glutamate transmission by corticosterone. Proc Natl Acad Sci U S A 102:19204-19207.
Keller-Wood ME, Dallman MF (1984) Corticosteroid inhibition of ACTH secretion. Endocr Rev 5:1-24.
Khan MM, Hadman M, De Sevilla LM, Mahesh VB, Buccafusco J, Hill WD, Brann DW (2006) Cloning, distribution, and colocalization of MNAR/PELP1 with glucocorticoid receptors in primate and nonprimate brain. Neuroendocrinology 84:317-329.
Kino T, Tiulpakov A, Ichijo T, Chheng L, Kozasa T, Chrousos GP (2005) G protein beta interacts with the glucocorticoid receptor and suppresses its transcriptional activity in the nucleus. J Cell Biol 169:885-896.
Kino T, Souvatzoglou E, Charmandari E, Ichijo T, Driggers P, Mayers C, Alatsatianos A, Manoli
155 I, Westphal H, Chrousos GP, Segars JH (2006) RHO family guanine nucleotide exchange factor BRX couples extracellular signals to the glucocorticoid signaling system. J Biol Chem.
Koenig JI, Cho JY (2005) Provocation of kainic acid receptor mRNA changes in the rat paraventricular nucleus by insulin-induced hypoglycaemia. J Neuroendocrinol 17:111- 118.
Koukouritaki SB, Theodoropoulos PA, Margioris AN, Gravanis A, Stournaras C (1996) Dexamethasone alters rapidly actin polymerization dynamics in human endometrial cells: evidence for nongenomic actions involving cAMP turnover. J Cell Biochem 62:251-261.
Kristensen P (1993) Differential expression of AMPA glutamate receptor mRNAs in the rat adrenal gland. FEBS Lett 332:14-18.
Kubo M, Shimizu C, Kijima H, Nagai S, Koike T (2004) Alternate promoter and 5'-untranslated exon usage of the mouse adrenocorticotropin receptor gene in adipose tissue. Endocr J 51:25-30.
Landfield PW, Eldridge JC (1991) The glucocorticoid hypothesis of brain aging and neurodegeneration: recent modifications. Acta Endocrinol (Copenh) 125 Suppl 1:54-64.
Landys MM, Piersma T, Ramenofsky M, Wingfield JC (2004) Role of the low-affinity glucocorticoid receptor in the regulation of behavior and energy metabolism in the migratory red knot Calidris canutus islandica. Physiol Biochem Zool 77:658-668.
Lariviere WR, Melzack R (2000) The role of corticotropin-releasing factor in pain and analgesia. Pain 84:1-12.
Lees GJ (2000) Pharmacology of AMPA/kainate receptor ligands and their therapeutic potential in neurological and psychiatric disorders. Drugs 59:33-78.
Lerma J (1997) Kainate reveals its targets. Neuron 19:1155-1158.
Li HL, Lin H (1974) An archeological and historical account of cannabis in China. Econ Bot 28:437-448.
Liberman AC, Druker J, Perone MJ, Arzt E (2007) Glucocorticoids in the regulation of transcription factors that control cytokine synthesis. Cytokine Growth Factor Rev 18:45- 56.
Liposits Z, Bohn MC (1993) Association of glucocorticoid receptor immunoreactivity with cell membrane and transport vesicles in hippocampal and hypothalamic neurons of the rat. J Neurosci Res 35:14-19.
Lipworth BJ (2000) Therapeutic implications of non-genomic glucocorticoid activity. Lancet
156 356:87-89.
Liu C, Zhou J, Zhang LD, Wang YX, Kang ZM, Chen YZ, Jiang CL (2007) Rapid inhibitory effect of corticosterone on histamine release from rat peritoneal mast cells. Horm Metab Res 39:273-277.
Liu X, Chen YZ (1995) Membrane-mediated inhibition of corticosterone on the release of arginine vasopressin from rat hypothalamic slices. Brain Res 704:19-22.
Lopez FJ, Negro-Vilar A (1988) Estimation of endogenous adrenocorticotropin half-life using pulsatility patterns: a physiological approach to the evaluation of secretory episodes. Endocrinology 123:740-746.
Lowenberg M, Verhaar AP, van den Brink GR, Hommes DW (2007) Glucocorticoid signaling: a nongenomic mechanism for T-cell immunosuppression. Trends Mol Med 13:158-163.
Lowenberg M, Tuynman J, Bilderbeek J, Gaber T, Buttgereit F, van Deventer S, Peppelenbosch M, Hommes D (2005) Rapid immunosuppressive effects of glucocorticoids mediated through Lck and Fyn. Blood 106:1703-1710.
Loxley HD, Cowell AM, Flower RJ, Buckingham JC (1993) Modulation of the hypothalamo- pituitary-adrenocortical responses to cytokines in the rat by lipocortin 1 and glucocorticoids: a role for lipocortin 1 in the feedback inhibition of CRF-41 release? Neuroendocrinology 57:801-814.
Mackie K (2005) Cannabinoid receptor homo- and heterodimerization. Life Sci 77:1667-1673.
Madsen U, Brauner-Osborne H, Frydenvang K, Hvene L, Johansen TN, Nielsen B, Sanchez C, Stensbol TB, Bischoff F, Krogsgaard-Larsen P (2001) Synthesis and pharmacology of 3- isoxazolol amino acids as selective antagonists at group I metabotropic glutamic acid receptors. J Med Chem 44:1051-1059.
Maejima T, Hashimoto K, Yoshida T, Aiba A, Kano M (2001) Presynaptic inhibition caused by retrograde signal from metabotropic glutamate to cannabinoid receptors. Neuron 31:463- 475.
Mahesh VB, Zamorano P, De Sevilla L, Lewis D, Brann DW (1999) Characterization of ionotropic glutamate receptors in rat hypothalamus, pituitary and immortalized gonadotropin-releasing hormone (GnRH) neurons (GT1-7 cells). Neuroendocrinology 69:397-407.
Maier C, Runzler D, Schindelar J, Grabner G, Waldhausl W, Kohler G, Luger A (2005) G- protein-coupled glucocorticoid receptors on the pituitary cell membrane. J Cell Sci 118:3353-3361.
Makara GB, Stark E (1975) Effect of intraventricular glutamate on ACTH release.
157 Neuroendocrinology 18:213-216.
Makara GB, Haller J (2001) Non-genomic effects of glucocorticoids in the neural system. Evidence, mechanisms and implications. Prog Neurobiol 65:367-390.
Malcher-Lopes R, Franco A, Tasker JG (2008) Glucocorticoids shift arachidonic acid metabolism toward endocannabinoid synthesis: A non-genomic anti-inflammatory switch. Eur J Pharmacol 583:322-339.
Mandelbrod I, Feldman S, Werman R (1974) Inhibition of firing is the primary effect of microelectrophoresis of cortisol to units in the rat tuberal hypothalamus. Brain Res 80:303-315.
Mao L, Yang L, Tang Q, Samdani S, Zhang G, Wang JQ (2005) The scaffold protein Homer1b/c links metabotropic glutamate receptor 5 to extracellular signal-regulated protein kinase cascades in neurons. J Neurosci 25:2741-2752.
Marsicano G, Lutz B (1999) Expression of the cannabinoid receptor CB1 in distinct neuronal subpopulations in the adult mouse forebrain. Eur J Neurosci 11:4213-4225.
Martin BR (1986) Cellular effects of cannabinoids. Pharmacol Rev 38:45-74.
Massa EM, Morero RD, Bloj B, Farias RN (1975) Hormone action and membrane fluidity: effect of insulin and cortisol on the Hill coefficients of rat erythrocyte membrane-bound acetylcholinesterase and (Na+ + K+)-ATPase. Biochem Biophys Res Commun 66:115- 122.
Matsuda LA, Lolait SJ, Brownstein MJ, Young AC, Bonner TI (1990) Structure of a cannabinoid receptor and functional expression of the cloned cDNA. Nature 346:561-564.
Mayer ML, Armstrong N (2004) Structure and function of glutamate receptor ion channels. Annu Rev Physiol 66:161-181.
Mechoulam R (1970) Marihuana chemistry. Science 168:1159-1166.
Mechoulam R, Feigenbaum JJ (1987) Towards cannabinoid drugs. Prog Med Chem 24:159-207.
Mechoulam R, Feigenbaum JJ, Lander N, Segal M, Jarbe TU, Hiltunen AJ, Consroe P (1988) Enantiomeric cannabinoids: stereospecificity of psychotropic activity. Experientia 44:762-764.
Mechoulam R, Ben-Shabat S, Hanus L, Ligumsky M, Kaminski NE, Schatz AR, Gopher A, Almog S, Martin BR, Compton DR, et al. (1995) Identification of an endogenous 2- monoglyceride, present in canine gut, that binds to cannabinoid receptors. Biochem Pharmacol 50:83-90.
158 Meeker RB, Greenwood RS, Hayward JN (1994) Glutamate receptors in the rat hypothalamus and pituitary. Endocrinology 134:621-629.
Mehvar R, Dann RO, Hoganson DA (2000) Kinetics of hydrolysis of dextran- methylprednisolone succinate, a macromolecular prodrug of methylprednisolone, in rat blood and liver lysosomes. J Control Release 68:53-61.
Menkin V (1957) Direct effect of corticotropin (ACTH) on capillary permeability in inflammation. Am J Physiol 189:98-104.
Mikics E, Barsy B, Barsvari B, Haller J (2005) Behavioral specificity of non-genomic glucocorticoid effects in rats: effects on risk assessment in the elevated plus-maze and the open-field. Horm Behav 48:152-162.
Mikuriya TH (1969) Marijuana in medicine: past, present and future. Calif Med 110:34-40.
Mizoguchi K, Ishige A, Aburada M, Tabira T (2003) Chronic stress attenuates glucocorticoid negative feedback: involvement of the prefrontal cortex and hippocampus. Neuroscience 119:887-897.
Moldrich G, Wenger T (2000) Localization of the CB1 cannabinoid receptor in the rat brain. An immunohistochemical study. Peptides 21:1735-1742.
Moore FL, Orchinik M (1994) Membrane receptors for corticosterone: a mechanism for rapid behavioral responses in an amphibian. Horm Behav 28:512-519.
Mori H, Mishina M (1995) Structure and function of the NMDA receptor channel. Neuropharmacology 34:1219-1237.
Munro S, Thomas KL, Abu-Shaar M (1993) Molecular characterization of a peripheral receptor for cannabinoids. Nature 365:61-65.
Neckers L, Sze PY (1975) Regulation of 5-hydroxytryptamine metabolism in mouse brain by adrenal glucocorticoids. Brain Res 93:123-132.
Newton R, Seybold J, Kuitert LM, Bergmann M, Barnes PJ (1998) Repression of cyclooxygenase-2 and prostaglandin E2 release by dexamethasone occurs by transcriptional and post-transcriptional mechanisms involving loss of polyadenylated mRNA. J Biol Chem 273:32312-32321.
Nicholls D, Attwell D (1990) The release and uptake of excitatory amino acids. Trends Pharmacol Sci 11:462-468.
Noon LA, Bakmanidis A, Clark AJ, O'Shaughnessy PJ, King PJ (2006) Identification of a novel melanocortin 2 receptor splice variant in murine adipocytes: implications for post- transcriptional control of expression during adipogenesis. J Mol Endocrinol 37:415-420.
159 Nunez E, Benito C, Pazos MR, Barbachano A, Fajardo O, Gonzalez S, Tolon RM, Romero J (2004) Cannabinoid CB2 receptors are expressed by perivascular microglial cells in the human brain: an immunohistochemical study. Synapse 53:208-213.
O'Shaugnessy WB (1843) On the Cannabis indica or Indian hemp. Pharmacol J Trans 2:594-595.
Oelofsen W, Ramachandran J (1983) Studies of corticotropin receptors on rat adipocytes. Arch Biochem Biophys 225:414-421.
Oliet SH, Baimoukhametova DV, Piet R, Bains JS (2007) Retrograde regulation of GABA transmission by the tonic release of oxytocin and endocannabinoids governs postsynaptic firing. J Neurosci 27:1325-1333.
Onaivi ES, Ishiguro H, Gong JP, Patel S, Perchuk A, Meozzi PA, Myers L, Mora Z, Tagliaferro P, Gardner E, Brusco A, Akinshola BE, Liu QR, Hope B, Iwasaki S, Arinami T, Teasenfitz L, Uhl GR (2006) Discovery of the presence and functional expression of cannabinoid CB2 receptors in brain. Ann N Y Acad Sci 1074:514-536.
Orchinik M, Murray TF, Moore FL (1991) A corticosteroid receptor in neuronal membranes. Science 252:1848-1851.
Orchinik M, Murray TF, Moore FL (1994) Steroid modulation of GABAA receptors in an amphibian brain. Brain Res 646:258-266.
Orchinik M, Murray TF, Franklin PH, Moore FL (1992) Guanyl nucleotides modulate binding to steroid receptors in neuronal membranes. Proc Natl Acad Sci U S A 89:3830-3834.
Ostasov P, Bourova L, Hejnova L, Novotny J, Svoboda P (2007) Disruption of the plasma membrane integrity by cholesterol depletion impairs effectiveness of TRH receptor- mediated signal transduction via G(q)/G(11)alpha proteins. J Recept Signal Transduct Res 27:335-352.
Ostrander MM, Ulrich-Lai YM, Choi DC, Richtand NM, Herman JP (2006) Hypoactivity of the hypothalamo-pituitary-adrenocortical axis during recovery from chronic variable stress. Endocrinology 147:2008-2017.
Ozawa S, Kamiya H, Tsuzuki K (1998) Glutamate receptors in the mammalian central nervous system. Prog Neurobiol 54:581-618.
Paoletti P, Neyton J (2007) NMDA receptor subunits: function and pharmacology. Curr Opin Pharmacol 7:39-47.
Pappas TC, Gametchu B, Watson CS (1995) Membrane estrogen receptors identified by multiple antibody labeling and impeded-ligand binding. Faseb J 9:404-410.
Parker AJ, Wessely S, Cleare AJ (2001) The neuroendocrinology of chronic fatigue syndrome
160 and fibromyalgia. Psychol Med 31:1331-1345.
Patel S, Roelke CT, Rademacher DJ, Hillard CJ (2005) Inhibition of restraint stress-induced neural and behavioural activation by endogenous cannabinoid signalling. Eur J Neurosci 21:1057-1069.
Patel S, Roelke CT, Rademacher DJ, Cullinan WE, Hillard CJ (2004) Endocannabinoid signaling negatively modulates stress-induced activation of the hypothalamic-pituitary-adrenal axis. Endocrinology 145:5431-5438.
Pechnick RN, George R, Poland RE (1989) Characterization of the effects of the acute and repeated administration of MK-801 on the release of adrenocorticotropin, corticosterone and prolactin in the rat. Eur J Pharmacol 164:257-263.
Petralia RS, Wenthold RJ (1992) Light and electron immunocytochemical localization of AMPA- selective glutamate receptors in the rat brain. J Comp Neurol 318:329-354.
Petralia RS, Wang YX, Wenthold RJ (1994a) Histological and ultrastructural localization of the kainate receptor subunits, KA2 and GluR6/7, in the rat nervous system using selective antipeptide antibodies. J Comp Neurol 349:85-110.
Petralia RS, Yokotani N, Wenthold RJ (1994b) Light and electron microscope distribution of the NMDA receptor subunit NMDAR1 in the rat nervous system using a selective anti- peptide antibody. J Neurosci 14:667-696.
Pfaff DW, Silva MT, Weiss JM (1971) Telemetered recording of hormone effects on hippocampal neurons. Science 172:394-395.
Philip JG, Flower RJ, Buckingham JC (1995) Evidence for different molecular species of lipocortin 1 in the rat. Pharmacol Res 31:84.
Philip JG, Flower RJ, Buckingham JC (1997) Glucocorticoids modulate the cellular disposition of lipocortin 1 in the rat brain in vivo and in vitro. Neuroreport 8:1871-1876.
Pin JP, Duvoisin R (1995) The metabotropic glutamate receptors: structure and functions. Neuropharmacology 34:1-26.
Pinheiro PS, Rodrigues RJ, Rebola N, Xapelli S, Oliveira CR, Malva JO (2005) Presynaptic kainate receptors are localized close to release sites in rat hippocampal synapses. Neurochem Int 47:309-316.
Pistovcakova J, Makatsori A, Sulcova A, Jezova D (2005) Felbamate reduces hormone release and locomotor hypoactivity induced by repeated stress of social defeat in mice. Eur Neuropsychopharmacol 15:153-158.
Porter AC, Sauer JM, Knierman MD, Becker GW, Berna MJ, Bao J, Nomikos GG, Carter P,
161 Bymaster FP, Leese AB, Felder CC (2002) Characterization of a novel endocannabinoid, virodhamine, with antagonist activity at the CB1 receptor. J Pharmacol Exp Ther 301:1020-1024.
Poulin JF, Castonguay-Lebel Z, Laforest S, Drolet G (2008) Enkephalin co-expression with classic neurotransmitters in the amygdaloid complex of the rat. J Comp Neurol 506:943- 959.
Rasband W (1997-2007) ImageJ. In, 1.38x Edition. Bethesda, Maryland, USA: National Institutes of Health.
Rivier C, Vale W (1983) Interaction of corticotropin-releasing factor and arginine vasopressin on adrenocorticotropin secretion in vivo. Endocrinology 113:939-942.
Roland BL, Sawchenko PE (1993) Local origins of some GABAergic projections to the paraventricular and supraoptic nuclei of the hypothalamus in the rat. J Comp Neurol 332:123-143.
Ronesi JA, Huber KM (2008) Homer interactions are necessary for metabotropic glutamate receptor-induced long-term depression and translational activation. J Neurosci 28:543- 547.
Rosa ML, Guimaraes FS, Pearson RC, Del Bel EA (2002) Effects of single or repeated restraint stress on GluR1 and GluR2 flip and flop mRNA expression in the hippocampal formation. Brain Res Bull 59:117-124.
Rose JD, Moore FL, Orchinik M (1993) Rapid neurophysiological effects of corticosterone on medullary neurons: relationship to stress-induced suppression of courtship clasping in an amphibian. Neuroendocrinology 57:815-824.
Rosenmund C, Stern-Bach Y, Stevens CF (1998) The tetrameric structure of a glutamate receptor channel. Science 280:1596-1599.
Rouyer M (1810) Sur les medicaments usuels des Egyptiens. Bull Pharm 2:385-415.
Ruf K, Steiner FA (1967) Steroid-sensitive single neurons in rat hypothalamus and midbrain: identification by microelectrophoresis. Science 156:667-669.
Sagan S, Venance L, Torrens Y, Cordier J, Glowinski J, Giaume C (1999) Anandamide and WIN 55212-2 inhibit cyclic AMP formation through G-protein-coupled receptors distinct from CB1 cannabinoid receptors in cultured astrocytes. Eur J Neurosci 11:691-699.
Sajadi AA, Samaei SA, Rashidy-Pour A (2006) Intra-hippocampal microinjections of anisomycin did not block glucocorticoid-induced impairment of memory retrieval in rats: an evidence for non-genomic effects of glucocorticoids. Behav Brain Res 173:158-162.
162 Sakakura M, Saito Y, Takebe K, Ishii K (1976) Studies on fast feedback mechanisms by endogenous glucocorticoids. Endocrinology 98:954-957.
Sandi C, Venero C, Guaza C (1996a) Nitric oxide synthesis inhibitors prevent rapid behavioral effects of corticosterone in rats. Neuroendocrinology 63:446-453.
Sandi C, Venero C, Guaza C (1996b) Novelty-related rapid locomotor effects of corticosterone in rats. Eur J Neurosci 8:794-800.
Sapolsky RM, Romero LM, Munck AU (2000) How do glucocorticoids influence stress responses? Integrating permissive, suppressive, stimulatory, and preparative actions. Endocr Rev 21:55-89.
Sawchenko PE, Swanson LW (1983) The organization of forebrain afferents to the paraventricular and supraoptic nuclei of the rat. J Comp Neurol 218:121-144.
Sayers G, Sayers MA (1947) Regulation of pituitary adrenocorticotrophic activity during the response of the rat to acute stress. Endocrinology 40:265-273.
Scaccianoce S, Matrisciano F, Del Bianco P, Caricasole A, Di Giorgi Gerevini V, Cappuccio I, Melchiorri D, Battaglia G, Nicoletti F (2003) Endogenous activation of group-II metabotropic glutamate receptors inhibits the hypothalamic-pituitary-adrenocortical axis. Neuropharmacology 44:555-561.
Selye H (1936) A syndrome produced by diverse nocuous agents. Nature 138:32.
Selye H (1956) What is stress? Metabolism 5:525-530.
Serova LI, Gueorguiev V, Cheng SY, Sabban EL (2008) Adrenocorticotropic hormone elevates gene expression for catecholamine biosynthesis in rat superior cervical ganglia and locus coeruleus by an adrenal independent mechanism. Neuroscience 153:1380-1389.
Sheng WS, Hu S, Min X, Cabral GA, Lokensgard JR, Peterson PK (2005) Synthetic cannabinoid WIN55,212-2 inhibits generation of inflammatory mediators by IL-1beta-stimulated human astrocytes. Glia 49:211-219.
Shigeri Y, Seal RP, Shimamoto K (2004) Molecular pharmacology of glutamate transporters, EAATs and VGLUTs. Brain Res Brain Res Rev 45:250-265.
Shivaji S, Jagannadham MV (1992) Steroid-induced perturbations of membranes and its relevance to sperm acrosome reaction. Biochim Biophys Acta 1108:99-109.
Silverman AJ, Hoffman DL, Zimmerman EA (1981) The descending afferent connections of the paraventricular nucleus of the hypothalamus (PVN). Brain Res Bull 6:47-61.
Silverman AJ, Jhamandas J, Renaud LP (1987) Localization of luteinizing hormone-releasing
163 hormone (LHRH) neurons that project to the median eminence. J Neurosci 7:2312-2319.
Smith Y, Charara A, Hanson JE, Paquet M, Levey AI (2000) GABA(B) and group I metabotropic glutamate receptors in the striatopallidal complex in primates. J Anat 196 (Pt 4):555-576.
Solito E, Mulla A, Morris JF, Christian HC, Flower RJ, Buckingham JC (2003) Dexamethasone induces rapid serine-phosphorylation and membrane translocation of annexin 1 in a human folliculostellate cell line via a novel nongenomic mechanism involving the glucocorticoid receptor, protein kinase C, phosphatidylinositol 3-kinase, and mitogen- activated protein kinase. Endocrinology 144:1164-1174.
Sommer B, Keinanen K, Verdoorn TA, Wisden W, Burnashev N, Herb A, Kohler M, Takagi T, Sakmann B, Seeburg PH (1990) Flip and flop: a cell-specific functional switch in glutamate-operated channels of the CNS. Science 249:1580-1585.
Stallkamp I, Dowhan W, Altendorf K, Jung K (1999) Negatively charged phospholipids influence the activity of the sensor kinase KdpD of Escherichia coli. Arch Microbiol 172:295-302.
Strijbos PJ, Tilders FJ, Carey F, Forder R, Rothwell NJ (1991) Localization of immunoreactive lipocortin-1 in the brain and pituitary gland of the rat. Effects of adrenalectomy, dexamethasone and colchicine treatment. Brain Res 553:249-260.
Stromberg J, Backstrom T, Lundgren P (2005) Rapid non-genomic effect of glucocorticoid metabolites and neurosteroids on the gamma-aminobutyric acid-A receptor. Eur J Neurosci 21:2083-2088.
Stubbs CD, Smith AD (1984) The modification of mammalian membrane polyunsaturated fatty acid composition in relation to membrane fluidity and function. Biochim Biophys Acta 779:89-137.
Subczynski WK, Wisniewska A (2000) Physical properties of lipid bilayer membranes: relevance to membrane biological functions. Acta Biochim Pol 47:613-625.
Suemaru S, Darlington DN, Akana SF, Cascio CS, Dallman MF (1995) Ventromedial hypothalamic lesions inhibit corticosteroid feedback regulation of basal ACTH during the trough of the circadian rhythm. Neuroendocrinology 61:453-463.
Sugiura T, Waku K (2002) Cannabinoid receptors and their endogenous ligands. J Biochem (Tokyo) 132:7-12.
Sugiura T, Kobayashi Y, Oka S, Waku K (2002) Biosynthesis and degradation of anandamide and 2-arachidonoylglycerol and their possible physiological significance. Prostaglandins Leukot Essent Fatty Acids 66:173-192.
Sugiura T, Oka S, Ikeda S, Waku K (2006) Occurrence, biosynthesis, and metabolism of
164 endocannabinoid. Endocannabinoids: The brain and body's marijuana and beyond:177.
Sugiura T, Kondo S, Sukagawa A, Nakane S, Shinoda A, Itoh K, Yamashita A, Waku K (1995) 2- Arachidonoylglycerol: a possible endogenous cannabinoid receptor ligand in brain. Biochem Biophys Res Commun 215:89-97.
Sutherland ER, Ellison MC, Kraft M, Martin RJ (2003) Altered pituitary-adrenal interaction in nocturnal asthma. J Allergy Clin Immunol 112:52-57.
Tasker JG (2006) Rapid glucocorticoid actions in the hypothalamus as a mechanism of homeostatic integration. Obesity (Silver Spring) 14 Suppl 5:259S-265S.
Tatro JB (1990) Melanotropin receptors in the brain are differentially distributed and recognize both corticotropin and alpha-melanocyte stimulating hormone. Brain Res 536:124-132.
Tauchi M, Zhang R, D'Alessio DA, Seeley RJ, Herman JP (2008) Role of central glucagon-like peptide-1 in hypothalamo-pituitary-adrenocortical facilitation following chronic stress. Exp Neurol 210:458-466.
Taylor AD, Cowell AM, Flower J, Buckingham JC (1993) Lipocortin 1 mediates an early inhibitory action of glucocorticoids on the secretion of ACTH by the rat anterior pituitary gland in vitro. Neuroendocrinology 58:430-439.
Thrivikraman KV, Nemeroff CB, Plotsky PM (2000) Sensitivity to glucocorticoid-mediated fast- feedback regulation of the hypothalamic-pituitary-adrenal axis is dependent upon stressor specific neurocircuitry. Brain Res 870:87-101.
Tierney T, Christian HC, Morris JF, Solito E, Buckingham JC (2003) Evidence from studies on co-cultures of TtT/GF and AtT20 cells that Annexin 1 acts as a paracrine or juxtacrine mediator of the early inhibitory effects of glucocorticoids on ACTH release. J Neuroendocrinol 15:1134-1143.
Tillman TS, Cascio M (2003) Effects of membrane lipids on ion channel structure and function. Cell Biochem Biophys 38:161-190.
Todd AR (1946) Hashish. Experientia 2:55-60.
Tokarev D, Jezova D (1997) Effect of central administration of the non-NMDA receptor antagonist DNQX on ACTH and corticosterone release before and during immobilization stress. Methods Find Exp Clin Pharmacol 19:323-328.
Towle AC, Sze PY (1983) Steroid binding to synaptic plasma membrane: differential binding of glucocorticoids and gonadal steroids. J Steroid Biochem 18:135-143.
Tsou K, Brown S, Sanudo-Pena MC, Mackie K, Walker JM (1998) Immunohistochemical distribution of cannabinoid CB1 receptors in the rat central nervous system. Neuroscience
165 83:393-411.
Urbach V, Walsh DE, Mainprice B, Bousquet J, Harvey BJ (2002) Rapid non-genomic inhibition of ATP-induced Cl- secretion by dexamethasone in human bronchial epithelium. J Physiol 545:869-878.
Vahl TP, Ulrich-Lai YM, Ostrander MM, Dolgas CM, Elfers EE, Seeley RJ, D'Alessio DA, Herman JP (2005) Comparative analysis of ACTH and corticosterone sampling methods in rats. Am J Physiol Endocrinol Metab 289:E823-828.
Vale W, River C (1977) Substances modulating the secretion of ACTH by cultured anterior pituitary cells. Fed Proc 36:2094-2099.
Vale W, Spiess J, Rivier C, Rivier J (1981) Characterization of a 41-residue ovine hypothalamic peptide that stimulates secretion of corticotropin and beta-endorphin. Science 213:1394- 1397.
Van den Pol AN (1994) Metabotropic glutamate receptor mGluR1 distribution and ultrastructural localization in hypothalamus. J Comp Neurol 349:615-632. van den Pol AN, Romano C, Ghosh P (1995) Metabotropic glutamate receptor mGluR5 subcellular distribution and developmental expression in hypothalamus. J Comp Neurol 362:134-150.
Varma N, Carlson GC, Ledent C, Alger BE (2001) Metabotropic glutamate receptors drive the endocannabinoid system in hippocampus. J Neurosci 21:RC188.
Venero C, Borrell J (1999) Rapid glucocorticoid effects on excitatory amino acid levels in the hippocampus: a microdialysis study in freely moving rats. Eur J Neurosci 11:2465-2473.
Verriere VA, Hynes D, Faherty S, Devaney J, Bousquet J, Harvey BJ, Urbach V (2005) Rapid effects of dexamethasone on intracellular pH and Na+/H+ exchanger activity in human bronchial epithelial cells. J Biol Chem 280:35807-35814.
Wang D, Zhang H, Lang F, Yun CC (2007) Acute activation of NHE3 by dexamethasone correlates with activation of SGK1 and requires a functional glucocorticoid receptor. Am J Physiol Cell Physiol 292:C396-404.
Wang S, Lim G, Zeng Q, Sung B, Yang L, Mao J (2005) Central glucocorticoid receptors modulate the expression and function of spinal NMDA receptors after peripheral nerve injury. J Neurosci 25:488-495.
Watanabe M, Inoue Y, Sakimura K, Mishina M (1993) Distinct distributions of five N-methyl-D- aspartate receptor channel subunit mRNAs in the forebrain. J Comp Neurol 338:377-390.
Watson CS, Gametchu B (2001) Membrane estrogen and glucocorticoid receptors--implications
166 for hormonal control of immune function and autoimmunity. Int Immunopharmacol 1:1049-1063.
Whiting KP, Restall CJ, Brain PF (2000) Steroid hormone-induced effects on membrane fluidity and their potential roles in non-genomic mechanisms. Life Sci 67:743-757.
Widmaier EP, Dallman MF (1984) The effects of corticotropin-releasing factor on adrenocorticotropin secretion from perifused pituitaries in vitro: rapid inhibition by glucocorticoids. Endocrinology 115:2368-2374.
Wiegand SJ, Price JL (1980) Cells of origin of the afferent fibers to the median eminence in the rat. J Comp Neurol 192:1-19.
Wiegert O, Joels M, Krugers H (2006) Timing is essential for rapid effects of corticosterone on synaptic potentiation in the mouse hippocampus. Learn Mem 13:110-113.
Wieronska JM, Branski P, Szewczyk B, Palucha A, Papp M, Gruca P, Moryl E, Pilc A (2001) Changes in the expression of metabotropic glutamate receptor 5 (mGluR5) in the rat hippocampus in an animal model of depression. Pol J Pharmacol 53:659-662.
Wong CW, McNally C, Nickbarg E, Komm BS, Cheskis BJ (2002) Estrogen receptor-interacting protein that modulates its nongenomic activity-crosstalk with Src/Erk phosphorylation cascade. Proc Natl Acad Sci U S A 99:14783-14788.
Wood TB, Spivey WTN, Easterfield TH (1896) XL. - Charas. The resin of Indian hemp. Journal of the Chemical Society, Transactions 69:539-546.
Wu J, Gorenstein DG (1993) Structure and dynamics of cytochrome c in nonaqueous solvents by 2D NH-exchange NMR spectroscopy. J Am Chem Soc 115:6843-6850.
Wu LJ, Xu H, Ren M, Zhuo M (2007) Genetic and pharmacological studies of GluR5 modulation of inhibitory synaptic transmission in the anterior cingulate cortex of adult mice. Dev Neurobiol 67:146-157.
Xiao B, Tu JC, Petralia RS, Yuan JP, Doan A, Breder CD, Ruggiero A, Lanahan AA, Wenthold RJ, Worley PF (1998) Homer regulates the association of group 1 metabotropic glutamate receptors with multivalent complexes of homer-related, synaptic proteins. Neuron 21:707-716.
Xiao L, Qi A, Chen Y (2005) Cultured embryonic hippocampal neurons deficient in glucocorticoid (GC) receptor: a novel model for studying nongenomic effects of GC in the neural system. Endocrinology 146:4036-4041.
Xu H, Wu LJ, Zhao MG, Toyoda H, Vadakkan KI, Jia Y, Pinaud R, Zhuo M (2006) Presynaptic regulation of the inhibitory transmission by GluR5-containing kainate receptors in spinal substantia gelatinosa. Mol Pain 2:29.
167 Yates FE, Leeman SE, Glenister DW, Dallman MF (1961) Interaction between plasma corticosterone concentration and adrenocorticotropin-releasing stimuli in the rat: evidence for the reset of an endocrine feedback control. Endocrinology 69:67-80.
Yoneda Y, Ogita K (1989) Solubilization of stereospecific and quisqualate-sensitive activity of [3H]glutamate binding in the pituitary of the rat. Neuropharmacology 28:611-616.
Zelena D, Makara GB, Jezova D (1999) Simultaneous blockade of two glutamate receptor subtypes (NMDA and AMPA) results in stressor-specific inhibition of prolactin and corticotropin release. Neuroendocrinology 69:316-323.
Zelena D, Mergl Z, Makara GB (2005) Glutamate agonists activate the hypothalamic-pituitary- adrenal axis through hypothalamic paraventricular nucleus but not through vasopressinerg neurons. Brain Res 1031:185-193.
Zhu BG, Zhu DH, Chen YZ (1998) Rapid enhancement of high affinity glutamate uptake by glucocorticoids in rat cerebral cortex synaptosomes and human neuroblastoma clone SK- N-SH: possible involvement of G-protein. Biochem Biophys Res Commun 247:261-265.
Zhu Y, Bond J, Thomas P (2003) Identification, classification, and partial characterization of genes in humans and other vertebrates homologous to a fish membrane progestin receptor. Proc Natl Acad Sci U S A 100:2237-2242.
Ziegler DR, Herman JP (2000) Local integration of glutamate signaling in the hypothalamic paraventricular region: regulation of glucocorticoid stress responses. Endocrinology 141:4801-4804.
Ziegler DR, Cullinan WE, Herman JP (2002) Distribution of vesicular glutamate transporter mRNA in rat hypothalamus. J Comp Neurol 448:217-229.
Ziegler DR, Cullinan WE, Herman JP (2005) Organization and regulation of paraventricular nucleus glutamate signaling systems: N-methyl-D-aspartate receptors. J Comp Neurol 484:43-56.
Zuardi AW (2006) History of cannabis as a medicine: a review. Rev Bras Psiquiatr 28:153-157.
168