contributes to the regulation of energy balance
TRANSCRIPT
CaMKK2 CONTRIBUTES TO THE REGULATION OF ENERGY BALANCE
by
Fumin Lin
Department of Pharmacology and Cancer Biology Duke University
Date:_______________________ Approved:
___________________________ Anthony R. Means, Supervisor
___________________________
Christopher B. Newgard
___________________________ Donald P. McDonnell
___________________________
Deborah M. Muoio
Dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of
Pharmacology and Cancer Biology in the Graduate School of Duke University
2011
ABSTRACT
CaMKK2 CONTRIBUTES TO THE REGULATION OF ENERGY BALANCE
by
Fumin Lin
Department of Pharmacology and Cancer Biology Duke University
Date:_______________________ Approved:
___________________________ Anthony R. Means, Supervisor
___________________________
Christopher B. Newgard
___________________________ Donald P. McDonnell
___________________________
Deborah M. Muoio
An abstract of a dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Department of
Pharmacology and Cancer Biology in the Graduate School of Duke University
2011
Copyright by Fumin Lin
2011
iv
Abstract The incidence of obesity and associated diseases such as type 2‐diabetes and
hypertension has reached epidemic portions worldwide and attracted increased interest
to understand the mechanisms that are responsible for these diseases. Obesity can result
from excessive energy intake, and increasing evidence has emphasized the role of the
central nervous system, especially the hypothalamus, in regulating food intake. White
adipose, as a direct target of obesity and an important endocrine organ, also has long
been a subject of scientific inquiry. AMPK, a conserved energy sensor, has been shown
to play important roles in both the hypothalamus and adipose. Recently, CaMKK2 was
shown to function as an AMPK kinase. I used intracerebroventricular cannulation as a
means to acutely inhibit hypothalamic CaMKK2 with STO‐609 and characterize the
appetite change associated with loss of CaMKK2 function. Infusion of STO‐609 in wild‐
type mice, but not CaMKK2‐null mice, inhibited appetite and promoted weight loss
consistent with reduced NPY and AgRP mRNA. Furthermore, intraperitoneal injection
of ghrelin increased food intake in wild‐type but not CaMKK2‐null mice, and 2‐DG
increased appetite in both types of mice, indicating that CaMKK2 functions downstream
of ghrelin to activate AMPK and upregulate appetite. As CaMKK2‐null mice were
protected from high‐fat diet‐induced obesity and diabetes, I performed a pair feeding
experiment using a high‐fat diet and demonstrated that protection of CaMKK2‐null mice
v
did not require reduced food consumption. Analysis of brown adipose tissue and
metabolic analysis indicated that CaMKK2‐null mice did not expend more energy than
WT mice. Interestingly, we were surprised to find that CaMKK2‐null mice had more
adipose than wild‐type mice when fed standard chow (5001). By real‐time PCR and
immunoblot, I identified CaMKK2 expression in preadipocytes and showed that it
decreased during adipogenesis. I used STO‐609 or shRNA to block CaMKK2 activity in
preadipocytes, which resulted in enhanced adipogenesis and increased mRNA of
adipogenic genes. I also identified AMPK as the relevant downstream target of CaMKK2
involved in inhibiting adipogenesis via a pathway that maintained Pref‐1 mRNA.
Consistent with the in vitro data, we further demonstrated that CaMKK2‐null mice have
more adipocytes but fewer preadipocytes, which supports our hypothesis that loss of
CaMKK2 enhances adipogenesis by depleting the preadipocyte pool. Together the data
presented herein contribute to our understanding of distinct mechanisms by which
CaMKK2 contributes to feeding behavior and adipogenesis.
vi
Contents
Abstract .........................................................................................................................................iv
List of Tables.............................................................................................................................. viii
List of Figures ...............................................................................................................................ix
List of Abbreviations ...................................................................................................................xi
Acknowledgements ...................................................................................................................xiv
1. Introduction ............................................................................................................................... 1
1.1 Regulation of Food Intake in Hypothalamus ............................................................... 1
1.2 Adipogenesis..................................................................................................................... 8
1.3 Calcium/camodulin‐dependent Protein Kinase Cascades ....................................... 13
1.3.1 Structures of CaMKK, CaMKI and CaMKIV......................................................... 15
1.3.2 CaMKK ....................................................................................................................... 17
1.4 AMPK............................................................................................................................... 19
1.4.1 Structure and Regulation of AMPK........................................................................ 20
1.4.2 Regulation of Energy Balance by AMPK in the Hypothalamus......................... 21
1.4.3 Regulation of Energy Balance by AMPK in Peripheral Tissues ......................... 25
1.4.4 Regulation of Adipogenesis by AMPK .................................................................. 26
2. Materials and Methods........................................................................................................... 29
2.1 Animal Experiment ........................................................................................................ 29
2.2 Adipose Tissue Analysis ............................................................................................... 33
2.3 Cell Isolation, Cell Culture and Induction of Adipogenesis .................................... 35
vii
2.4 RNA and Protein Analysis............................................................................................ 40
2.5 RNAi, Transfections and Virus Infections .................................................................. 46
3. CaMKK2 Contributes to the Hypothalamic Regulation of Energy Balance................... 48
3.1 Introduction..................................................................................................................... 48
3.2 Results .............................................................................................................................. 50
3.3 Discussion........................................................................................................................ 65
4. CaMKK2 Regulation of Food Intake and Energy Balance Can Be Uncoupled .............. 67
4.1 Introduction..................................................................................................................... 67
4.2 Results .............................................................................................................................. 69
4.3 Discussion........................................................................................................................ 78
5. CaMKK2 Inhibits Preadipocyte Differentiation ................................................................. 80
5.1 Introduction..................................................................................................................... 80
5.2 Results .............................................................................................................................. 82
5.3 Discussion...................................................................................................................... 103
6. Future Directions .................................................................................................................. 110
References .................................................................................................................................. 120
Biography................................................................................................................................... 142
viii
List of Tables Table 1: Formula of diets............................................................................................................ 32
Table 2: Antibodies ..................................................................................................................... 42
Table 3: RT‐PCR primers............................................................................................................ 45
Table 4: siRNA targeting AMPKα ............................................................................................ 47
ix
List of Figures Figure 1: Transcriptional regulation of adipogenesis ............................................................ 12
Figure 2: Pref‐1 inhibition of adipogenesis.............................................................................. 13
Figure 3: CaMK cascades ........................................................................................................... 15
Figure 4: Schematic of CaMKs domains .................................................................................. 16
Figure 5: Domain structures of α, β and γ subunits of AMPK............................................. 22
Figure 6: CaMKK2 is expressed in the hypothalamus and regulates NPY ........................ 53
Figure 7: Ca2+‐dependent induction of NPY expression in N38 cells is blocked by STO‐609 ................................................................................................................................................. 56
Figure 8: Deletion or inhibition of CaMKK2 inhibits food intake........................................ 59
Figure 9: CaMKK2‐null mice consume less food and are resistant to high‐fat diet‐induced adiposity, glucose intolence and insulin resistence ............................................................... 63
Figure 10: CaMKK2‐null mice have the same food intake but less body weight gain in high‐fat diet pair feeding ........................................................................................................... 70
Figure 11: CaMKK2‐null mice gained less adiposity and remained glucose tolerant and insulin sensive compared to pair‐fed WT mice ...................................................................... 71
Figure 12: CaMKK2‐null mice have increased BAT and enhanced cold‐induced thermogenesis on standard chow............................................................................................. 73
Figure 13: CaMKK2‐null mice have less BAT and produce less heat when fed high‐fat diet................................................................................................................................................. 75
Figure 14: Adipocyte analysis of WAT from WT and CaMKK2‐null mice fed different diets............................................................................................................................................... 76
Figure 15: Adiposity analysis on retroperitoneal and epididymal fat pads in WT and CaMKK2‐null mice fed different diets...................................................................................... 77
Figure 16: Expression of CaMKK2 in preadipocytes ............................................................. 83
x
Figure 17: CaMKK2 mRNA and protein expression decreases after adipogenesis ........... 86
Figure 18: Loss of CaMKK2 activity enhances adipogenesis................................................ 89
Figure 19: CaMKK2‐null MEFs and preadipocytes exhibit no growth defect .................... 91
Figure 20: CaMKK2 works through AMPK to inhibit adipogenesis ................................... 93
Figure 21: CaMKK2 phosphorylates AMPK in preadipocytes............................................. 96
Figure 22: Loss of CaMKK2 activity increases adipogenic gene transcripts during adipogenesis ................................................................................................................................ 99
Figure 23: CaMKK2 maintains Pref‐1 mRNA........................................................................ 102
Figure 24: AMPK maintains Pref‐1 mRNA............................................................................ 105
Figure 25: CaMKK2‐null mice have fewer preadipocytes and more adipocytes in gonadal WAT............................................................................................................................................ 107
xi
List of Abbreviations
ACC acetyl CoA carboxylase
AgRP agouti‐related
AICAR 5‐aminoimidazole‐4‐carboxamide ribonucleoside
AMPK AMP‐activated protein kinase
aP2 adipocyte protein 2
ARC arcuate nucleus
BAT brown adipose tissue
C/EBP CCAAT/enhancer binding protein
CaM calmodulin
CaMKI Ca2+/CaM‐dependent protein kinase I
CaMKII Ca2+/CaM‐dependent protein kinase II
CaMKIV Ca2+/CaM‐dependent protein kinase IV
CaMKK Ca2+/CaM‐dependent protein kinase kinase
CART cocaine‐ and amphetamine‐regulated transcript
CNS central nervous system
CPT1 carnitine palmitoyl‐CoA transferase‐1
DEXA dual energy X‐ray absorptiometry
xii
ER endoplasmic reticulum
ERK extracellular signal‐regulated kinase
Fas fatty acid synthase
GHSR growth hormone secretagogue receptor
Glut4 glucose transporter type 4
i.c.v. intracerebroventricular
i.p. intraperitoneal
IRS‐1 insulin receptor substrate 1
JAK2 janus kinase 2
MCH melanin‐concentrating hormone
MEF mouse embryo fibroblast
mTOR mammalian target of rapamycin
NPY neuropeptide Y
PCR Polymerase chain reaction
PKA protein kinase A
PKB protein kinase B
POMC pro‐opiomelanocortin
PPARγ peroxisome proliferator‐activated receptor γ
xiii
Pref‐1 preadipocyte factor 1
Ser serine
shRNA short hairpin RNA
siRNA small interfering RNA
SNS sympathetic nervous system
Sox9 SRY (sex determining region Y)‐box 9
STAT3 signal transducer and activator of transcription 3
TACE tumor necrosis factor‐α converting enzyme
Thr threonine
UCP1 uncoupling protein 1
VMH ventromedial hypothalamus
WAT white adipose tissue
WT wild type
xiv
Acknowledgements
I would first and foremost like to express my gratitude to my thesis advisor, Dr.
Anthony Means for the freedom and encouragement he has given me in pursuing my
project. He has demonstrated a breadth of knowledge and incredible enthusiasm in
scientific research. His skill in logical thinking and identifying the key of a question is
what I could only wish one day to have. I am also grateful to my thesis committee which
includes Christopher Newgard, Donald McDonnell, Deborah Muoio and Xiao Fan Wang
for their brilliant advice and technique support from their labs. I also want to thank all
members in Means lab for providing me such a friendly and helpful environment
during graduate school training. I especially thank Tom Ribar for teaching me various
lab techniques and keeping me happy in lab. Most significantly I thank my parents,
Gongyang Lin and Liping Chen, for all of their sacrifices which allowed me to pursue
the best education. Finally, I deeply appreciate my wife, Daoyi Lin. Without her love,
patience and endless encourage, my graduate study would have been much more lonely
and difficult.
1
1. Introduction
1.1 Regulation of Food Intake in Hypothalamus
Energy balance is achieved by matching energy intake (food consumption) and energy
expenditure. Food intake disorders are related to a number of clinical conditions. For
example, excessive food intake has been long known to contribute to obesity, which can
trigger type 2‐diabetes, hypertension and cardiovascular disorders (Must et al., 1999).
On the other hand, anorexia is associated with neoplasia, infectious diseases and aging
(Broberger and Hokfelt, 2001). Therefore, understanding the mechanisms involved in
regulating food intake will be very helpful in developing pharmacological treatments.
Numerous studies have emphasized the role of the central nervous system (CNS),
especially the hypothalamus in integrating nutrient and hormone signaling to regulate
food intake. Initially the hypothalamus was realized to be critical to mediate appetite
because variations of food intake were observed in many species in response to
hypothalamic lesions. For example, lesions in the medial hypothalamus caused
hyperphagia which led to obesity (Brobeck et al., 1943). On the other hand, other lesions
resulted in decreased food intake or even complete inhibition of appetite (Anand and
Brobeck, 1951).
The arcuate nucleus (ARC), located in the mediobasal hypothalamus, is now
known to play a pivotal role in integrating various signal inputs which mediate appetite
2
(Wynne et al., 2005; Woods and D’Alessio, 2008; Lopaschuk et al., 2010). The ARC is
quite accessible to blood‐borne signaling molecules because the median eminence
beneath the ARC is not as well protected by the brain‐blood barrier as other brain areas
(Broadwell and Brightman, 1976). The ARC contains two main populations of neurons
which express different neuropeptides having opposite effects on food intake. One
population of neurons expresses neuropeptide Y (NPY), and agouti‐related peptide
(AgRP) (Chronwall et al., 1985; Hahn et al., 1998). Activation of these neurons results in
increased food intake (Stanley et al., 2005; Coll et al., 2007). The other population
expresses pro‐opiomelanocortin (POMC) and cocaine‐ and amphetamine‐regulated
transcript (CART) (Elias et al., 1998; Kristensen et al., 1998; Valassi et al., 2008).
Activation of POMC neurons leads to a decrease in food intake. Inside the ARC, POMC
can be cleaved into α‐melanocyte‐stimulating hormone (α‐MSH), which functions as a
ligand/agonist of melanocortin‐3 receptor (MC3R) and MC4R (Harrold et al., 1999).
MC4R plays a clear role in decreasing food intake, because MC4R agonists effectively
depressed appetite whereas its antagonists stimulated appetite (Fan W, Boston et al.,
1997; Benoit et al., 2000). Yaswen and colleagues also indicated that loss of a‐MSH in
POMC knockout mice resulted in hyperphagia and obesity (Yaswen et al., 1999). All
these phenotypes are consistent with an inhibitory role for POMC neurons on appetite.
Actually, mRNA of POMC was significantly reduced during fasting and restored by
refeeding (Swart et al., 2002). Similar to POMC, the expression of CART was also
3
suppressed by fasting (Robson et al., 2002). In addition, intracerebroventricular (i.c.v.)
administration of CART to rats decreased food intake (Kristensen et al., 1998). In
contrast to the POMC neurons, NPY/AgRP neurons stimulate food intake and both NPY
and AgRP have orexigenic effects in the hypothalamus. AgRP has been shown to be a
potent antagonist of MC3R and MC4R, and transgenic expression of AgRP effectively
increased food intake (Ollmann et al., 1997). In addition, i.c.v. administration of AgRP or
its C‐terminal fragment was capable of triggering hyperphagia that lasted for quite a
long time (Rossi et al., 1998; Hagan et al., 2000). As expected for an orexigenic peptide,
AgRP increased during fasting and remained elevated for at least 6 hr after refeeding
(Swart et al., 2002). NPY is another well‐known orexigenic peptide. Its expression and
release increased during fasting and decreased after refeeding (Swart et al., 2002). On the
other hand, the i.c.v. administration of NPY increased food intake (Clark et al., 1984;
Levine and Morley, 1984).
NPY binds to a series of G‐protein coupled receptors termed the Y receptors
(Lindner et al., 2008). Mashiko and colleagues demonstrated that administration of a
selective antagonist of the Y5 receptors reduced food intake (Mashiko et al., 2007). This
result confirmed the previous report that NPY Y5 receptor antisense oligonucleotides
inhibited food intake (Schaffhauser et al., 1997). Interestingly, when investigators tried to
delete either NPY or AgRP or both genes together, they did not observe the strong
inhibition of food intake that was expected. For example, mice deficient for NPY had
4
normal food intake and body weight, and they also responded normally to fasting
(Erickson et al., 1996). AgRP‐deficient mice or AgRP/NPY double‐knockout mice
exhibited no obvious feeding or body weight deficits, and responded normally to fasting
(Qian et al., 2002). On the other hand, acute ablation of NPY/AgRP neurons led to
profound anorexia in adult mice but not in neonates, which suggests these neurons are
essential for feeding but their loss can be compensated for under certain conditions
(Luquet et al., 2005; Gropp et al., 2005).
The hypothalamus can sense nutrient signals so as to regulate appetite. For
example, administration of glucose into the third ventricle of the brain (central injection)
caused a reduction of food intake, which partially contributed to the observed weight
loss (Davis et al., 1981). Recently, increased hypothalamic lactate (one of the end
products of glycolysis) level was also shown to reduce food intake. Intraperitoneal (i.p.)
injection of glucose or lactate effectively increased POMC expression and depressed
NPY expression (Wolfgang et al., 2007; Cha and Lane, 2009). Long chain fatty acids have
also been reported to have an orexigenic effect by acting on the hypothalamus. Central
injection of oleic acid led to reduction of food intake via decreased expression of NPY
and AgRP in the hypothalamus (Obici et al., 2002). Citrate, a common intermediate
product during carbohydrate and fatty acid metabolism, also has an anorexigenic effect
by acting on the hypothalamus, where it decreased NPY expression and increased
POMC mRNA (Stoppa et al., 2008).
5
The hypothalamus also responds to various hormone signals. Leptin and ghrelin
are two of the most well‐known hormones regulating feeding behaviors. Leptin is a
peptide hormone mainly secreted from adipocytes (Zhang et al., 1994). It has been
reported that the serum leptin level is correlated to fat content, and its concentration in
the CNS is proportional to its serum level. The serum level of leptin was suppressed by
restricting food intake, while restored by refeeding (Frederich et al., 1995). Leptin exerts
its anorexigenic effect through the ARC. The absence of circulating leptin due to a
mutation in the leptin gene resulted in hyperphagia and obesity in mice, and these
phenotypes could be overcome by supplying exogenous leptin to the mice (Campfield et
al., 1995; Halaas et al., 1995). Long term leptin administration in mice led to decreased
food intake, loss of body weight and fat content (Halaas et al., 1995). However, if the
ARC was destroyed before i.c.v. injection of leptin, then the administration of leptin
could not reduce food intake (Tang‐Christensen et al., 1999). Subsequently, leptin
receptors were shown to be expressed in the hypothalamus and localized to both
NPY/AgRP neurons and POMC neurons (Fei et al., 1997; Elmquist et al., 1998). As
predicted, leptin inhibited NPY/AgRP neurons and NPY expression, whereas low
circulating leptin resulted in the activation of these neurons (Elias et al., 1999; Morton et
al., 2003). On the other hand, leptin activated POMC neurons and POMC expression
(Schwartz et al., 1997), and MC4R antagonists impaired the anorexigenic effect of leptin,
indicating the importance of the melanocortin pathway in leptin signaling (Seeley et al.,
6
1997).
In contrast to leptin, ghrelin is a potent orexigenic hormone, which is mainly
secreted from the gastrointestinal tract (Date et al., 2000). Consistent with its orexigenic
role, the circulating level of ghrelin increased during fasting, but decreased after eating
(Cummings et al., 2001). Ghrelin was first identified as an endogenous ligand of the
growth hormone secretagogue receptor (GHSR), whose mRNA is co‐localized with NPY
mRNA in the ARC (Willesen et al., 1999). The orexigenic effect of ghrelin was
demonstrated by the fact that i.c.v. administration of ghrelin strongly increased food
intake whereas anti‐ghrelin antibody robustly suppressed feeding (Nakazato et al., 2001).
Nakazato and colleagues also demonstrated that the orexigenic effect of ghrelin was
mediated through NPY/AgRP neurons and was dependent on NPY and AgRP.
Administration of antibodies against NPY or AgRP eliminated the food intake
stimulated by ghrelin; so did the administration of antagonists of Y1 and Y5 receptors.
GHSR is also critical for ghrelin function, as ghrelin treatment of Ghsr‐null mice did not
stimulate food intake (Sun et al., 2004). Further research on GSHR revealed that it was a
7 transmembrane receptor coupled to Gq and its activation by ghrelin resulted in a rise
in intracellular Ca2+ in NPY neurons thus suggesting the importance of a Ca2+ pathway in
ghrelin signaling (Kohno et al., 2003).
Great efforts have been made to understand signal transduction pathways
downstream of leptin and ghrelin. Leptin binds to its receptor which has a long
7
intracellular domain, and this domain can bind to Janus kinase 2 (JAK2), which
phosphorylates and activates signal transducer and activator of transcription 3 (STAT3)
(Vaisse et al., 1996). Phosphorylated STAT3 forms dimmers which subsequently enter
the nucleus to promote POMC transcription or inhibit AgRP transcription (Bjorbaek et
al., 1997; Morton et al., 2006). On the other hand, leptin has also been shown to stimulate
the phosphorylation and activation of insulin receptor substrate 1 (IRS‐1) via JAK2. IRS‐
1, in turn, activates phophatidylinositol‐3‐OH kinase, which activates protein kinase B
(PKB/Akt). PKB can phosphorylate and inhibit the transcription factor FOXO1. FOXO1
increases both NPY and AgRP transcription in the hypothalamus, thereby stimulating
food intake (Kim et al., 2006). The mammalian target of rapamycin (mTOR) was also
shown to regulate leptin signaling. Central injection of leptin increased hypothalamic
mTOR signaling and decreased food intake, while inhibition of mTOR by rapamycin
blocked leptin’s anorexigenic effect (Cota et al., 2006). AMPK is also important for leptin
signaling, as leptin inhibited hypothalamic AMPK activity and constitutively active
AMPK blunted leptin effect (Minokoshi et al., 2004). Compared to leptin signaling, the
pathway by which ghrelin regulated NPY/AgRP expression was poorly understood. In
2004, Andersson and colleagues demonstrated that the hyperphagic effect of ghrelin is
correlated with increased AMPK activity (Andersson et al., 2004). Combined with the
fact that GHSR bound by ghrelin increases intracellular Ca2+, and CaMKK2 can function
as an AMPKK, we hypothesized that CaMKK2 could be the Ca2+‐activated target
8
downstream of ghrelin required to activate AMPK. Our evaluation of this hypothesis is
described in chapter 3.
1.2 Adipogenesis
Adipose tissue is the largest energy reservoir in the body. Adipocytes, which are the
majority cells in adipose tissue store surplus energy in the form of triglycerides in lipid
droplets. Recently, a number of studies have provided compelling evidence that
adipocytes actually serve as endocrine cells (Fruhbeck et al., 2001). The molecules that
are secreted from white adipose tissue (WAT) play an important role in immune
response, vascular homeostasis, glucose/lipid metabolism and appetite regulation.
Briefly, there are two ways to expand the adipose tissue. One is storing more fat content
into existing adipocyte cells leading to increased cell size, which is known as
hypertrophy. The other one is to increase the number of adipocytes by differentiating
more precursor cells, which is termed as hyperplasia. The process of adipocyte
precursor cells (preadipocytes) differentiating into adipocytes is known as adipogenesis,
and a number of studies have revealed that this process follows a highly ordered
temporal sequence of transcriptional events that is regulated by well‐defined
transcription factors (reviewed in Mandrup and Lane, 1997; Rosen et al., 2000; Farmer,
2006).
9
The most important transcription factor for adipogenesis is peroxisome
proliferator‐activated receptor γ (PPARγ). Two isoforms of PPARγ were generated from
alternative splicing and alternate translation initiation. Although PPARγ2 has been
shown to be the dominant isoform in fat cells, no functional differences are detected
between the two isoforms. As a member of nuclear hormone receptor subfamily, PPARγ
needs to heterodimerize with the retinoid X receptor (RXR) to bind DNA. The
heterodimer regulates transcription by recruiting different coactivators or corepressors.
Transcription of several important adipocyte‐specific genes is under the control of
PPARγ, including adipocyte protein 2 (aP2), lipoprotein lipase, acyl‐CoA synthase and
others (Desvergne and Wahli, 1999). Also PPARγ can be activated by ligand binding.
Synthetic compounds named thiazolidinediones (TZDs) bind PPARγ with high affinity.
TZD treatment in preadipocytes greatly enhances adipogenesis, suggesting the role of
PPARγ in this process (Kletzien et al., 1992; Sandouk et al., 1993). In 1994, Tontonoz and
colleagues showed that expression of PPARγ alone in nonadipogenic fibroblasts was
able to initiate the whole adipogenic program and generate mature adipocytes
(Tontonoz et al., 1994). Actually, PPARγ is potent enough to induce the
transdifferentiation of myoblasts into adipocytes (Hu et al., 1995). In contrast,
preadipocytes deficient of PPARγ could not differentiate into adipocytes (Rosen et al.,
1999). In vivo, specific knockout of PPARγ in WAT resulted in lipodystrophy, which
confirms its role in adipogenesis (Koutnikova et al., 2003).
10
The other important regulator of adipogenesis is CCAAT/enhancer binding
protein α (C/EBPα). C/EBPα belongs to the basic‐leucine zipper class of transcription
factors. Other family members include C/EBPβ and C/EBPδ. They can form homodimers
or heterodimers to regulate transcription. Ectopic expression of C/EBPα promoted
adipogenic program in a variety of mouse fibroblastic cells (Freytag et al., 1994). C/EBPα
has also been shown to be sufficient to stimulate adipogenesis in 3T3‐L1 preadipocytes
without any hormone stimuli (Lin and Lane, 1994). In contrast, knockdown of C/EBPα
by antisense RNA in 3T3‐L1 preadipocytes resulted in inhibition of adipogenesis (Lin
and Lane, 1992). In vivo, knockout of C/EBPα in mice made them fail to form WAT
(Wang et al., 1995). One important function of C/EBPα is to keep cells in terminally
differentiated state by maintaining the expression of PPARγ, and PPARγ is able to
stimulate the expression of C/EBPα (Wu et al., 1999).
During the process of searching for upstream regulators of PPARγ and C/EBPα,
C/EBPβ and C/EBPδ were identified. Yeh and colleagues demonstrated that expression
of C/EBPβ was sufficient to induce adipogenesis of 3T3‐L1 preadipocytes without any
hormone stimuli (Yeh et al., 1995). They also showed that expression of C/EBPδ
accelerated adipogenesis, and C/EBPβ and C/EBPδ were able to induce C/EBPα
expression. Furthermore, C/EBPβ and C/EBPδ stimulated PPARγ expression in NIH‐3T3
fibroblasts and made them commit to adipose lineage (Wu et al., 1996). Interestingly,
mice lacking both C/EBPβ and C/EBPδ showed defect in WAT development, and MEFs
11
from these mice could not express PPARγ and C/EBPα and therefore could not
differentiate into adipocytes either (Tanaka et al., 1997). Besides its function to induce
PPARγ and C/EBPα, C/EBPβ was also suggested to affect PPARγ ligand production for
terminal adipogenesis, because attenuation of C/EBPβ activity blocked adipogenesis
unless extra PPARγ ligand was added (Hamm et al., 2001).
Therefore, the classic transcriptional regulation of adipogenesis is summarized in
Fig. 1. Among the early events that follow pro‐adipogenic hormonal stimulation is the
expression of C/EBPβ and C/EBPδ. C/EBPβ and C/EBPδ then trigger the expression of
PPARγ, which in turn activates C/EBPα, which exerts positive feedback on PPARγ to
maintain the differentiated state. Finally PPARγ and C/EBPα stimulate expression of
adipocyte‐specific genes such as aP2, fatty acid synthase (Fas) and glucose transporter
type 4 (Glut4).
Additional studies have shown that adipogenesis is influenced by a variety of
extrinsic factors and intracellular signaling pathways (Rosen and Spiegelman, 2000;
MacDougald and Mandrup, 2002). One of them is preadipocyte Factor 1 (Pref‐1; variants
in other tissues include DLK1, fetal antigen‐1 and pG2) (reviewed in Sul, 2009). As
shown in Fig. 2, Pref‐1 is an EGF‐repeat containing transmembrane protein, and it needs
to be cleaved by tumor necrosis factor‐α converting enzyme (TACE) to
12
Figure 1: Transcriptional regulation of adipogenesis
Adapted from Rosen et al., Genes Dev. 2000 14: 1293‐1307
generate the biologically active soluble form (Wang and Sul, 2006). The soluble form of
Pref‐1 binds an unknown receptor and activates extracellular signal‐regulated kinase
(ERK) to maintain the expression of Sox9, which inhibits adipogenesis by repressing
C/EBPβ and C/EBPδ transcription (Smas and Sul, 1996; Wang et al., 2006; Kim et al., 2007;
Wang and Sul, 2009).
13
Figure 2: Pref‐1 inhibition of adipogenesis
Adapted from Sul HS, Mol Endocrinol, November 2009, 23(11):1717–1725
1.3 Calcium/calmodulin-dependent Protein Kinase Cascades
Calcium is one of the most frequently used second messenger molecules in the
cell, and it plays important roles in many cellular processes such as proliferation,
development and secretion. Cells are capable of reacting differentially to intracellular
Ca2+ level changes according to their source, duration, amplitude, and subcellular
location (Macrez and Mironneau, 2004; Thomas et al., 1996; Chow and Means, 2007). In
14
order to mediate so many different responses, mammalian cells possess a good many
proteins which bind intracellular Ca2+ with various affinities to act as buffers,
transducers or effectors. One of the most important Ca2+ transducers is calmodulin
(CaM). Binding of Ca2+ to CaM induces a conformational change to expose its
hydrophobic residues, and in turn promotes interaction of Ca2+/CaM complex with a
great number of proteins. Among them, a subfamily of Ser/Thr protein kinases is known
as Ca2+/CaM‐dependent protein kinase cascade (CaMK cascade).
At the beginning, CaMK cascade was described as consisting of three members:
Ca2+/CaM‐dependent protein kinase kinase (CaMKK), CaMKI and CaMKIV (Soderling,
1999; Means, 2000). Their activities all need the binding of Ca2+/CaM complex. CaMKKs
activate CaMKI and CaMKIV by phosphorylating their activation loop (Thr177 on
CaMKI and Thr196 on CaMKIV) (Haribabu et al., 1995; Selbert et al., 1995). Recently,
AMP‐activated protein kinase (AMPK) was reported to be activated though
phosphorylation of Thr172 by CaMKK2 (Hawley et al., 2005; Hurley et al., 2005; Woods
et al., 2005). Different from CaMKI and CaMKIV, AMPK is not a direct target of CaM.
The pathways are summarized in Fig. 3.
Pharmacological inhibitors have been utilized as an important approach to
understand CaMK functions. KN‐62 and KN‐93 are competitive to CaM, and they are
able to inhibit CaMKI, CaMKII and CaMKIV similarly (Tokumitsu et al., 1990; Sumi et
al., 1991; Enslen et al., 1994; Mochizuki et al., 1993). As shown in Fig. 3, KN‐62 and KN‐
15
93 inhibit CaMKI and CaMKIV but not CaMKKs. STO‐609 is an ATP‐competitive
inhibitor, which inhibits CaMKKs potently and selectively (CaMKK1, IC50=120ng/ml;
CaMKK2, IC50=40ng/ml) and does not inhibit CaMKI, CaMKIV or CaMKII
(IC50>=10μg/ml) (Tokumitsu et al., 2002). As for AMPK, there is also one selective
inhibitor, compound C, which is an ATP‐competitive inhibitor. Other than inhibitors,
overexpression of different mutant forms of kinases or knockdown by small interference
RNA are also often employed to understand the detail of this cascade.
Figure 3: CaMK cascades
Modified from Means, 2008 Mol Endocrinol 22:2759‐2765
1.3.1 Structures of CaMKK, CaMKI and CaMKIV
CaMKKs, CaMKI and CaMKIV have similar domain structures. Each of them has a well‐
conserved N‐terminal catalytic domain and a central regulatory domain containing
16
overlapping autoinhibitory domain (AID) and Ca2+/CaM binding domain (CBD) (Fig. 4).
CaMKKs have a unique Arg‐Pro (RP)‐rich insert in their catalytic domain, which does
not exist in CaMKI, CaMKIV or other kinases. Deletion of RP‐insert results in impaired
ability to phosphorylate and activate CaMKI or CaMKIV, but does not affect catalytic
activity (Tokumitsu et al., 1999). AIDs mimic the substrates; they contain substrate
recognition determinants but do not have a phosphorylatable Ser/Thr. In the absence of
Ca2+/CaM, AID interacts with and inhibits the catalytic domain. The binding of Ca2+/CaM
to CBD, which partially overlaps the AID, leads to conformational changes which
disrupt the interaction between AID and catalytic domain and increase kinase activity.
Interestingly, CaMKKs are much more sensitive to the activation of CaM binding
compared to CaMKI and CaMKIV (Matsushita and Nairn, 1998).
Figure 4: Schematic of CaMKs domains
The domains are shown as follows: catalytic domains, dark green; autoinhibitory domains, red; Ca2+/CaM‐binding domains, yellow. Modified from Soderling and Stull, 2001 Chem. Rev. 101: 2341‐2351
17
1.3.2 CaMKK
The identification of CaMKK starts from the discovery that in order to achieve full
activity of CaMKIV, some activating factor around 68kD is needed with ATP in addition
to Ca2+/CaM (Okuno and Fujisawa, 1993; Tokumitsu et al., 1994; Okuno et al., 1994).
After that, CaMKK1 as a 68 kD protein and CaMKK2 as an approximately 70 kD protein
were cloned by different groups (Tokumitsu et al., 1995; Kitani et al., 1997; Anderson et
al., 1998). CaMKK1 and CaMKK2 share 65% identical (80% similar) catalytic and
Ca2+/CaM binding domains, and they function similarly in vitro (Anderson et al., 1998).
Generally, CaMKK1 and CaMKK2 distribute similarly through different tissues with
abundant expression in brain. CaMKK2 has also been shown to be slightly expressed in
thymus, testis and spleen; however, only messenger RNA of CaMKK1 could be detected
in these tissues (Anderson et al., 1998; Tokumitsu et al., 1995). Existence of CaMKKs in
many other tissues still remains to be determined because of lack of robust antibodies.
At the cellular level, both CaMKK1 and CaMKK2 are localized dominantly in cytoplasm
but not nuclei, which was proved by immunohistochemistry of rat brain and GFP‐
CaMKK fusion protein overexpression in NG108‐15 cells (Sakagami et al., 2000)
CaMKK1 activity needs Ca2+/CaM binding; however, CaMKK2 has some
Ca2+/CaM‐independent activity (Tokumitsu and Soderling, 1996; Anderson et al., 1998).
CaMKKs are also subject to regulation by phosphorylation. Both CaMKK1 and CaMKK2
have been shown to autophosphorylate themselves, but autophosphorylation seems to
18
have nearly no effect on catalytic activity at least for CaMKK2 (Anderson et al., 1998;
Tokumitsu et al., 1999). There are at least three regulatory sites on CaMKK1 shown to be
phosphorylated by cAMP‐dependent protein kinase (PKA) and these phosphorylations
all inhibit CaMKK1 activity (Wayman et al., 1997; Matsushita and Nairn, 1999; Davare et
al., 2004). However, regulation of CaMKK2 by PKA has never been reported. Instead,
CaMKK2 can be sequentially phosphorylated by cyclin‐dependent kinase 5 and
glycogen synthae kinase 3 at N‐terminus. This results in an increase in the Ca2+/CaM
independent activity of CaMKK2 and changes its half‐life (Green et al., 2011).
CaMKKs play an important role in learning and memory. For example, CaMKK1‐
null mice showed impaired contextual fear memory formation (Mizuno et al., 2006;
Blaeser et al., 2006). CaMKK2‐null mice showed impaired spatial fear memory formation
and long‐term memory (Peters et al, 2003). The CaMKK‐CaMKI cascade is involved in
long‐term potentiation, a cellular model of learning and memory. CaMKK/CaMKI can
phosphorylate extracellular‐regulated kinase (ERK), which is necessary for LTP process;
STO‐609 or dominant‐negative CaMKI effectively blocked LTP‐induced ERK
phosphorylation (Schmitt et al., 2005; Fortin et al., 2010). CaMKKs are also critical in
neurite outgrowth. For example, in neuroblastoma NG108 cells, STO‐609 inhibited
neurite outgrowth but constitutively active CaMKKI or CaMKI promoted it (Schmitt et
al, 2004). The CaMKK2‐CaMKIV‐CREB pathway is important for brain‐derived
neurotrophic factor (BDNF) expression, which was required for normal development of
19
cerebellar granule neurons (Kokubo et al., 2009). In addition, CaMKKs may participate
in cell cycle regulation. Two kinases CMKC and CMKB in Aspergillus nidulams, which
share high sequence identity with CaMKK and CaMKI/V respectively, are important for
cell cycle progression especially G1/S transition (Joseph and Means, 2000). Kahl and
Means also demonstrated that in WI‐38 human fibroblasts, KN‐93 or overexpression of
dominant negative CaMKI prevented cyclin D/cdk4 activation and arrested cells in late
G1 (Kahl and Means, 2004). Most CaMKK2 functions so far identified are related to
CaMKI or CaMKIV; however, its function as an AMPK kinase is gradually being
revealed.
1.4 AMPK
AMPK is a serine/threonine kinase which is evolutionarily conserved from single cell
eukaryotes such as yeast to mammals (Hardie et al., 2003; Carling, 2004). AMPK is well
known as a key sensor of cellular energy status, because an elevated AMP:ATP ratio,
which indicates cellular energy status is compromised, triggers the activation of AMPK.
Thus, AMPK will be switched on by conditions interfering with ATP production
including hypoxia and glucose deprivation, or by processes that consume ATP such as
muscle contraction (Hardie, 2003). Once activated, AMPK turns on catabolic pathways
which increase ATP production, and turns off anabolic processes which consume ATP.
It is believed that AMPK functions via direct phosphorylation of corresponding
20
metabolic enzymes or altering long term gene expression. Moreover, recent studies
demonstrate that AMPK not only works as a master regulator of metabolism at the
cellular level, but also controls energy balance at the whole body level. Generally,
AMPK integrates nutrient and hormone signals to regulate energy balance in
hypothalamus (to increase food intake) and peripheral tissues (to increase free fatty acid
oxidation and glucose uptake and reduce gluconeogenesis) (Xue and Kahn, 2006).
1.4.1 Structure and Regulation of AMPK
Since its purification in 1994, AMPK has been revealed to be a heterotrimeric complex
consisting of an α catalytic subunit and regulatory β and γ subunits (Davies et al., 1994;
Mitchelhill et al., 1994). In mammals, each subunit is encoded by multiple genes (α1, α2,
β1, β2, γ1, γ2, γ3) (Carling, 2004). The structure of each subunit remains similar in
different eukaryotes as shown in Figure 5 (Hardie et al., 2006). The α subunit contains a
serine/threonine kinase domain in the N‐terminal region and has a threonine (Thr‐172)
in the activation loop which needs to be phosphorylated in order to activate AMPK
(Hawley et al., 1996). The α subunit also has a β/γ subunit binding region in its C‐
terminus. The β subunit has its α/γ binding region in the C‐terminus. It also has a central
region which binds glycogen to the AMPK complex, however, the physiological role of
this region is still unknown (Hudson et al., 2003; Polekhina et al., 2003). In the C‐
21
terminal region of the γ subunit, there are four CBS motifs which work in pairs to bind
AMP or ATP in a mutually exclusive manner (Cheung et al., 2000; Hardie et al., 2006).
As mentioned, activation of AMPK requires phosphorylation of T172 in the α
subunit. So far, three upstream kinases have been indentified for this event. They are
LKB1, CaMKK2 and TGF‐β activated kinase (TAK1) (Hawley et al., 2003; Woods et al.
2003; Shaw et al., 2004; Hawley et al., 2005; Hurley et al., 2005; Woods et al., 2005;
Momcilovic et al., 2006). LKB1 is constitutively active and phosphorylates AMPK when
the AMP concentration rises in the cell and binds to the γ subunit, which transforms
AMPK into a more suitable substrate for LKB1. CaMKK2 phosphorylates AMPK kinase
independently of AMP, instead requiring a change of intracellular Ca2+ concentration.
However, T172 is not the only phosphorylation site in AMPK. Further studies indicate
that Ser485 (Ser491 in α2) can be phosphorylated by protein kinase B/Akt or PKA or
AMPK itself, which attenuates the phosphorylation of T172 and AMPK activity (Hurley
et al., 2006; Horman et al., 2006; Soltys et al., 2006).
1.4.2 Regulation of Energy Balance by AMPK in the Hypothalamus
AMPK activity has been found in both central nervous system and peripheral tissues
such as muscle, liver and adipose tissue. A number of studies reported that
hypothalamic AMPK responded to different nutrient or hormone signals to regulate
food intake. Most obviously, fasting increased AMPK activity in the hypothalamus and
22
Figure 5: Domain structures of α, β and γ subunits of AMPK
Adapted from Hardie et al., 2006 J Physiol 574.1 pp 7–15
refeeding depressed it (Minokoshi et al., 2004; Kim et al., 2004). Furthermore, i.p.
injection of glucose decreased AMPKα2 activity in the hypothalamus, and i.c.v. injection,
which does not change serum glucose and insulin levels, also inhibited AMPKα2 in the
hypothalamus (Minokoshi et al., 2004; Perrin et al., 2004; Kim et al., 2004). Besides,
hypoglycemia caused by insulin injection was reported to increase AMPKα2 activity in
the hypothalamus (Han et al., 2005). 2‐deoxyglucose (2‐DG), a nonmetabolizable glucose
analog that inhibits glucose utilization, increased AMPK phosphorylation in neuronal
cell lines and ex vivo hypothalamus culture (Lee et al., 2005). Hypothalamic AMPK
activity is also regulated by hormone signals. Generally, orexigenic hormones stimulate
23
AMPK activity whereas anorexigenic hormones inhibit it. Cannabinoid and ghrelin are
both orexigenic hormones increasing food intake through interactions in the
hypothalamus (Nakazato et al., 2001; Jamshidi and Taylor, 2001). Either i.p. or i.c.v.
injection of either ghrelin or cannabinoid significantly elevated hypothalamic AMPK
phosphorylation and activity (Andersson et al., 2004; Kola et al., 2005). Anorexigenic
molecules such as leptin, insulin and MT‐II (melanocortin receptor agonist) inhibited
hypothalamic AMPK activity (Minokoshi et al., 2004; Andersson et al., 2004). It is more
important that alteration of AMPK activity in the hypothalamus itself is enough to
mediate food intake, body weight and expression of orexigenic neuropeptides. For
example, Minokoshi and colleagues exhibited that overexpression of dominant negative
(DN) AMPK in the medial hypothalamus decreased food intake and body weight gain
of mice, and they also showed that mice with DN‐AMPK had lower mRNA levels of
NPY and AgRP in the ARC. In contrast, mice with constitutively active (CA) AMPK
expressed in the medial hypothalamus had increased food intake and body weight gain
with elevated expression of NPY and AgRP during fasting. The ability of leptin to
reduce food intake and body weight on CA‐AMPK‐expressed mice was impaired,
because leptin was no longer able to decrease hypothalamic AMPK activity. 5‐
aminoimidazole‐4‐carboxamide ribonucleoside (AICAR) is an AMPK activator; it can be
taken into cells and phosphorylated into ZMP, which mimics the effect of AMP on
AMPK activation (Sabina et al., 1985; Sullivan et al., 1994). The i.c.v. injection of AICAR
24
effectively increased food intake whereas compound C, a selective AMPK inhibitor,
triggered downregulation of food intake and body weight (Kim et al., 2004; Andersson
et al., 2004). Although the role of AMPK in hypothalamic regulation of appetite was well
established, the corresponding upstream kinase of AMPK still remained unclear.
Ghrelin, which activates AMPK in the hypothalamus, binds to its receptor GHSR to
activate phospholipase C and generate inositol trisphosphate, which triggers Ca2+ release
from endoplasmic reticulum (ER) (Wells, 2009). This fact hints that CaMKK2 might be
the upstream AMPK kinase. The abundance of CaMKK2 in the hypothalamus also
supports this hypothesis. Actually our lab has presented evidence to support the idea
that CaMKK2 functions as an AMPK kinase downstream of ghrelin in the hypothalamus
to regulate NPY/AgRP expression (Anderson et al., 2008), which is described in chapter
3.
Hypothalamic AMPK may also regulate energy expenditure in brown adipose
tissue (BAT), which is under the control of the sympathetic nervous system (SNS)
(Lowell & Spiegelman 2000). Lopez and colleagues suggested that thyroid hormone
depressed hypothalamic AMPK activity so as to upregulate de novo lipogenesis, which
resulted in enhanced SNS activity and upregulated thermogenesis in BAT (Lopez et al.,
2010). In 2003, another group reported that AMPKα2 null mice had increased daily
urinary epinephrine, norepinephrine and dopamine excretion, which also suggested that
loss of hypothalamic AMPK activity enhanced SNS activity (Viollet et al., 2003). The
25
i.c.v. injection of ghrelin suppressed the electrophysiological activity of the sympathetic
nerves innervating BAT in rats (Yasuda et al., 2003). Central administration of ghrelin
also decreased norepinephrine release in BAT, which is an important stimulator for BAT
activity (Mano‐Otagiri et al., 2009). In contrast, leptin, which inhibits AMPK activity in
the hypothalamus, increased sympathetic outflow to BAT and stimulated thermogenesis
(Collins et al., 1996; Haynes et al., 1999). Leptin mediated thermogenesis through the
hypothalamic melanocortin (MC) system (Satoh et al., 1998). AgRP, a potent antagonist
of MC3R and MC4R, negatively regulated sympathetic activation caused by leptin (Fong
et al., 1997; Ebihara et al., 1999). NPY is also reported to affect thermogenesis in BAT.
Recently, Chao and colleagues knocked down NPY expression in the dorsomedial
hypothalamus in rats and observed increased BAT development and energy
expenditure (Chao et al., 2011). Given that AMPK activity in the hypothalamus controls
NPY/AgRP expression, it would be reasonable to think hypothalamic AMPK also
regulates BAT activity.
1.4.3 Regulation of Energy Balance by AMPK in Peripheral Tissues
AMPK regulates metabolic pathways in most peripheral tissues to maintain normal ATP
levels (Xue and Kahn, 2006). In heart and skeletal muscle, AMPK stimulates fatty acid
uptake and oxidation as well as glucose uptake (Xue and Kahn, 2006; Russell et al., 1999;
Merrill et al. 1997). AMPK also stimulates mitochondrial biogenesis in skeletal muscle
26
and glycolysis in heart (Winder et al. 2000; Marsin et al. 2000). AMPK inhibits insulin
secretion from pancreatic β cells (Rutter et al., 2003). AMPK inhibits protein synthesis in
both skeletal muscle and liver (Bolster et al., 2002; Viollet et al., 2006); it also inhibits
fatty acid and cholesterol synthesis and gluconeogenesis in liver (Corton et al., 1995;
Henin et al., 1995). In adipose tissue, AMPK has been shown to inhibit fatty acid
synthesis and lipolysis but increase fatty acid oxidation (Daval et al., 2006).
The central metabolic pathway that AMPK utilizes to regulate fatty acid
oxidation is through acetyl CoA carboxylase (ACC), which converts acetyl CoA to
malonyl CoA (Merrill et al., 1997). Phosphorylation of ACC by AMPK inhibits its
activity, which leads to a decrease of malonyl CoA. Malonyl CoA inhibits carnitine
palmitoyl‐CoA transferase‐1 (CPT1), an enzyme transporting long‐chain fatty acid into
the mitochondria for oxidation. Thus, decrease of malonyl CoA will increase CPT1
activity and increase fatty acid oxidation.
1.4.4 Regulation of Adipogenesis by AMPK
There is evidence to suggest that AMPK is involved in preadipocyte differentiation.
AICAR inhibited adipogenesis in 3T3‐L1 or F442A preadipocytes (Habinowski et al.,
2001; Giri et al., 2006; Dagon et al., 2006; Lee et al., 2011). Giri and colleagues showed
that the expression of late adipogenesis markers such as aP2 and adipsin was reduced
by AICAR treatment, as was the expression of adipogenic transcription factor C/EBPα
27
and PPARγ. However, the expression of Pref‐1, which usually significantly decreases
during adipogenesis, was restored by AICAR treatment (Giri et al., 2006). Most groups
used AICAR stimulation to study the effect of AMPK on adipogenesis; however, at least
one group reported that inhibition of AMPK by compound C enhanced PPARγ
expression (Tong et al., 2008). Besides, metformin, a drug frequently used to treat type
2‐diabetes, inhibited lipid accumulation in 3T3‐L1 cells via a pathway that involves
activation of AMPK (Alexandre et al., 2008). However, the detailed knowledge about
interactions upstream and downstream of AMPK in this process still needs exploration.
In this thesis, I tried to understand the function of CaMKK2 in the hypothalamus
and white adipose tissue using genetically altered mice and cell lines. At first, we found
that CaMKK2 was expressed in NPY/AgRP neurons within the ARC of the
hypothalamus and CaMKK2‐null mice displayed a reduced hypothalamic AMPK
activity as well as a decreased level of NPY/AgRP. This fact led us to hypothesize that
CaMKK2 might work as an AMPK kinase in the hypothalamus to control appetite. In
order to prove this idea, I compared the feeding responses of WT and mutant mice to i.p.
injection of ghrelin and 2‐DG. The role of CaMKK2 in hypothalamus was also confirmed
by i.c.v. infusion of its inhibitor, STO‐609, which reduced food intake and body weight
in WT mice but not in CaMKK2‐null mice. Later on, I demonstrated that CaMKK2‐null
mice were resistant to high‐fat diet‐induced obesity, glucose intolerance and insulin
28
resistance, and it could be partially explained by reduced food intake during high‐fat
diet feeding. To clarify whether reduced appetite in CaMKK2‐null mice was the primary
contributor to their resistance to high‐fat diet effects, I performed a pair feeding
experiment between WT and CaMKK2‐null mice. Since WT mice were still more obese,
glucose intolerant and insulin resistant compared to CaMKK2‐null mice, I hypothesized
that loss of CaMKK2 enhanced energy expenditure via BAT, because increased BAT and
cold‐induced thermogenesis were observed in mutant mice under standard chow.
However, knockout mice possessed smaller BAT and decreased heat production when
fed high‐fat diet, which does not support this hypothesis. Finally, I turned my focus to
WAT. I demonstrated that CaMKK2 inhibited adipogenesis in several preadipocyte cell
lines by activating AMPK and maintaining Pref‐1 mRNA. These results are consistent
with the observation that CaMKK2‐null mice had more adiposity and fewer
preadipocytes on standard chow. Therefore, here I demonstrated that the absence of
CaMKK2 reduced appetite via the hypothalamus and enhanced adipogenesis in WAT.
29
2. Materials and Methods
2.1 Animal Experiment
Animal care
All mice were bred and maintained in either the Duke University LSRC or MSRBII
animal facility under a 12 hr light (0600‐1800), 12 hr dark cycle (1800‐0600). Food and
water were provided ad libitum and all care was given in compliance within NIH and
institutional guidelines on the care and use of laboratory and experimental animals.
Cannulation and infusion of STO‐609 into third ventricle
Insertion of cannulae (Plastics One) and subsequent infusion of solutions into the third
ventricle of adult WT or CaMKK2‐null mice using Alzet osmotic minipumps (DURECT
Corporation) were performed as follows. A stock solution of STO‐609 (20mM in 100mM
NaOH) was diluted 1:1000 in 0.9% sterile saline to obtain a 20uM working solution
(pH≈7.0). One hundred microliters of STO‐609 or carrier was loaded into individual
osmotic minipumps (Alzet model 1007D) 12 hr prior to surgical implantation and
allowed to equilibrate in 0.9% sterile saline at 37 degree celsius to ensure that the pumps
began to work immediately after implantation at the maximal flow rate of 0.5 μl/hr.
Prior to surgery, mice were administered ketamine 80 mg/kg; xylazine 10 mg/kg i.p.
After the anesthetic had taken effect, mice were secured in a stereotaxic apparatus (Kopf
30
Instruments), and an incision was made down the midline of the head to expose the
skull. After swabbing the exposed skull with 95% EtOH to remove any contaminants,
the bregma line was located and marked. A small access hole was drilled at bregma ‐1.94,
and the cannula was inserted to a depth of 5.8 mm and secured with epoxy bone cement
(3M ESPE). After the bone cement dried, a small incision was made in the skin between
the scapulae. A small pocket was formed in the subcutaneous tissue, and the pump was
implanted and connected to the cannula via a small sterile plastic tube. The incision was
then closed, and the animal was placed on a heating pad to recover. Mice were
monitored daily for feeding behavior over a 7 day period, at which time they were
sacrificed, and the hypothalamus was removed for RNA extraction.
Intraperitoneal injection of ghrelin or 2‐DG
WT and CaMKK2‐null mice were given intraperitoneal (i.p.) injection of saline control or
ghrelin (3nmol/100μl per mouse, Sigma G8903) 1hr before the onset of dark cycle, and
food intake was quantified at 1hr and overnight. For 2‐DG (Calbiochem #25972), a dose
of 15mg/100ul per mouse was given and food intake was also quantified at 1hr and
overnight.
Glucose and insulin tolerance assays
Glucose tolerance assays were performed on mice that were fasted overnight (>12 hr). At
31
0900, mice were measured for baseline glucose by collecting a small drop of blood from
the tail vein for analysis using a handheld Bayer Ascensia Contour glucometer. After the
baseline glucose values were established, each mouse was weighted and given an i.p.
injection of 2 mg glucose/g body weight (Fisher Scientific, D14‐212). Glucose was
dissolved in sterile water in a concentration of 400mg/ml. Glucose injections and blood
glucose sampling were timed to take approximately the same amount of time per animal,
so that the sample times were accurate for each animal. Blood glucose was quantified as
a function of time until glucose levels returned to near baseline values. Insulin tolerance
test was conducted using the same glucometer, also at 0900. Mice were fasted for 3 hr
prior to starting the procedure. After the baseline glucose values were established, mice
were given recombinant human insulin (1 U/kg i.p. Eli Lilly). Clearance of plasma
glucose was subsequently monitored at 15, 30, 45, 60, 90, 120 and 150 minutes
postinjection.
Feeding mice with different diets
For high‐fat diet experiment, twenty male WT and CaMKK2‐null mice were housed five
per cage at the age of weaning (three weeks) and fed with low‐fat (D12328) or high‐fat
(D12330) diet from Research Diets for 31 weeks. After 31 weeks, DEXA scan, glucose
tolerance test and insulin tolerance test were performed and serum was collected for
hormone analysis. Measurements of food intake and weight gain were performed
32
weekly. Information for three different diets is displayed in Table 1.
Table 1: Formula of diets
Standard (5001) Low Fat(CCO) High Fat (HCO)
Producer Lab Diet Research Diets Research Diets
Total kcal/gram 3.36 4.07 5.56
Fat% 13.4% 10.5% 58.0%
Protein% 28.5% 16.4% 16.4%
Carbohydrate% 58.0% 73.1% 25.5%
Pair feeding in high fat diet
For pair‐feeding experiment, fourteen WT and CaMKK2‐null mice around three‐month‐
old were housed individually. Initial body weight and adiposity were measured and
mice were subjected to 5 days adaptation to high‐fat diet. Food intake and body weights
were monitored every day. CaMKK2‐null mice had free access to the food. WT mice,
which would eat more if food was provided ad libitum, received the average amount of
the food that was consumed by CaMKK2‐null mice one day before. At the same time,
both types of mice had free access to water. Pair‐feeding was carried out for 30 weeks;
DEXA scan, glucose tolerance test and insulin tolerance test were performed around 10
weeks after the start of pair‐feeding and at the end. White adipose tissue, brown fat
tissue, liver, kidney and serum were collected for future study at the end of experiment.
33
Thermogenic response of WT and CaMKK2‐null mice
Ten WT and CaMKK2‐null mice were housed individually with food and water
provided ad libitum. All mice were fasted for 1 hr after dark cycle started, and then
moved into 8 degree celsius cold room for following 8 hr. Rectal temperature was
measured hourly by mouse rectal temperature probe (ADinstruments, MLT1404).
2.2 Adipose Tissue Analysis
Dual energy X‐ray absorptiometry (DEXA)
Mice were anesthetized (ketamine 100 mg/kg; zylaxine 10 mg/kg) and scanned 4 times
(with no repositioning between scans) using a Lunar PIXImus II densitometer (software
version 2.0; GE Lunar Corp, Madison, WI). All mice had access to food and water ad
libitum before the DEXA measurements which were typically performed between 8:00
and 11:30 AM. Anesthetized mice were then placed on the imaging positioning tray in a
prostrate fashion with the front and back legs extended away from the body. Heads
were excluded from all analyses by placing an exclusion region of interest (ROI) over the
head. Thus, all body composition data exclude the head. Sizes and locations of ROI were
modified to evaluate the amount of scapula brown adipose tissue, epididymal fat pad
and retroperitoneal fat pad.
34
White Fat analysis
White fat analysis from mice was performed on euthanized mice. After euthanasia,
gonadal fat pads were removed and were fixed in 10% neutral buffered formalin
overnight. Fixed samples were dehydrated in graded ethanol, cleared in xylene and
embedded in paraffin. Five‐micron serial sections were cut, mounted on poly‐L‐lysine‐
coated glass slides (Histology Control System, Glenn Head, NY), and dried at 55 degree
Celsius for 1hr . Sections were deparaffinized in Hemo‐D (Fisher Scientific, Pittsburg,
PA) and rehydrated through graded ethanol to PBS. Cell dimension measurements were
performed on 5u H&E‐stained sections using a Zeiss AxioImager D1 microscope (Carl
Zeiss, Thornwood, NJ) and Axiovision v.4.4 software.
Adipocyte number analysis
Isolation of mature adipocytes from WAT was performed according to a previously
described method by Etherton et al (Etherton et al., 1977) with some modification.
Briefly, one gonadal fat pad was isolated from freshly sacrificed WT and CaMKK2‐null
mice and rinsed twice with 0.154M NaCl at 37 degree Celsius to get rid of free lipid. The
rinsed WAT was fixed with 2% osmium tetroxide (Electron Microscopy Sciences, #19100)
in collidine‐HCl buffer, pH 7.4 (Electron Microscopy Sciences, #11500). Adipose tissue
was left in the fixative for 72 hr at room temperature in a ventilated fume hood. After
fixation, osmium tetroxide was removed and tissue was rinsed with 0.154M NaCl for 24
35
hr. After 24 hr, NaCl was removed, and 8 M urea in 0.154M NaCl was added. Following
the addition of 8 M urea, samples were occasionally swirled by hand and left at room
temperature for 48 hr. After observing the suspension of free adipocytes devoid of
connective tissue debris, the sample was passed through a sterile 230μm stainless steel
tissue sieve (Bellco Glass, #1985‐00069); connective tissue debris and undigested tissue
were left on screen. The fixed and urea‐isolated adipocytes were collected by screening
through a sterile 25μm stainless steel tissue sieve (Bellco Glass, #1985‐00500). The
adipocytes were washed three times with 0.154M NaCl, pH 10.0, containing 0.01%
Triton X‐100 (Sigma, T‐9284). The adipocytes were resuspended in 20ml of 0.154M NaCl,
and then 100ul of adipocyte suspension was diluted to 10ml with balanced electrolyte
solution (Beckman Coulter, 624994‐B ) and counted using a Coulter Counter (Beckman
Coulter, Z1).
2.3 Cell Isolation, Cell Culture and Induction of Adipogenesis
Isolation and growth of preadipocyte
Isolation of preadipocytes was performed according to previously described method by
Permana et al (Permana et al., 2004) with some modification. Briefly, gonadal fat pad
was isolated from freshly sacrificed WT and CaMKK2‐null mice. Isolated mouse adipose
tissue was minced in DMEM containing 5.5 mM glucose, 5% fatty acid‐free BSA (Sigma,
A9647), and then centrifuged at 500g for 5 minutes to get rid of blood cells. Supernatant
36
was transferred to a new tube and digested with Collagenase P (Roche, #11249002001) at
a concentration of 1mg/ml dissolved in DMEM. Tissues were incubated at 37 degree
Celsius for 45 minutes and filtered through a sterile 230μm stainless steel tissue sieve
(Bellco Glass, #1985‐00069). After centrifugation at 500g for 10 minutes, the pelleted cells
containing preadipocytes were resuspended in DMEM and strained through a sterile
25μm stainless steel tissue sieve (Bellco Glass, #1985‐00500). The filtrate was transferred
to a 60mm culture plate and maintained in an incubator at 37 degree Celsius in 5% CO2.
Cells were allowed to attach, and the next day floating red blood cells were removed by
aspiration, and the culture medium was replenished. Medium was changed every other
day.
For growth rate measurement, preadipocytes were isolated from four WT and
CaMKK2‐null mice, and kept growing for three days. At day three, preadipocytes were
detached by trypsin treatment and counted in Coulter counter. The preadipocytes from
individual mice were plated at a concentration of 0.25X105 /ml into a 6 well plate. Cells
were allowed to grow, and 12, 24, 36, 48, 72 and 96 hr after seeding, preadipocytes from
one well were detached by trypsin and counted in Coulter counter.
Isolation of mouse embryonic fibroblasts
Isolation of mouse embryonic fibroblasts was performed according to the protocol
modified from Jacks Lab at David H. Koch Institute for Integrative Cancer Research of
37
MIT. The pregnancy of female mice was checked and female mice were scarified at day
13.5 after onset of pregnancy. The abdomen of freshly sacrificed mice was soaked in 70%
EtOH and opened by dissection scissors. Uterus was exposed and two strings of
embryos were removed and transferred into 10cm culture dish containing 10ml HBSS
with 1X pen/strep. Embryos were carefully dissected out of placenta and transferred
individually into a new 60mm dish with HBSS. Rostral portion of the embryo’s head
(above the eyes) was cut off and saved for PCR analysis for genotyping. Hematopoietic
tissue was removed as much as possible. Each embryo was transferred into a new 60mm
dish with 1 ml trypsin‐EDTA and minced into small pieces. The dishes then were
incubated at 37 degree Celsius for 45 minutes. 4 ml of DMEM supplemented with 15%
heat‐inactivated FBS was added into each dish to stop the digestion. Each trypsinized
embryo was pipette vigorously to separate cells and then transferred into a labeled T75
flask with 15 ml DMEM supplemented with 15% hiFBS. Flasks were placed in an
incubator at 37 degree Celsius with 5% CO2, and cells were allowed to grow 72 hr until
they reached over‐confluent. At the same time, genotyping PCR analysis was carried out
to identify the genotype for each embryo. After 72 hr, each T75 flask was rinsed with
HBSS and treated with 2 ml of trypsin‐EDTA. 8 ml of DMEM with 15% hiFBS was
added to stop the digestion and after pipetting, 5 ml of cells were transferred into T175
flask with 50 ml DMEM supplemented with 15% hiFBS. Cells were incubated at 37
degree Celsius with 5% CO2 for 72 hr until they reached confluence. The confluent cells
38
were detached by treating each T175 flask with 3 ml of trypsin‐EDTA and resuspended
in 25 ml DMEM supplemented with 15% hiFBS. Cells from same embryo were combined
and spun down at 1000 RPM, room temperature for 3 minutes. Cells were resuspended
in 32 ml freezing media per embryo (25% hiFBS, 10% TC‐grade DMSO, 65% DMEM). 1
ml of resuspended cells (around 3X106) was pipetted into each cryotube (nunc, 375418).
All vials were transferred into a Styrofoam rack and frozen overnight at ‐80 degree
Celsius. On the next day, cryotubes were transferred into a liquid nitrogen cryo
biological safety system.
Cell culture and induction of adipogenesis
MEF were isolated as described above. One vial of frozen MEFs was thawed and seeded
in 10cm culture dish with DMEM (Cellgro, #17‐205‐CV) supplemented with 10% heat‐
inactivated FBS (Atlanta Biologicals, #S11150), 2mM L‐glutamine (Cellgro, #25‐005‐CL),
100 units/ml penicillin and 100ug/ml streptomycin (Gibco, #15140). Culture medium was
refreshed every other day. For growth rate measurement, WT and CaMKK2‐null MEFs
were detached by trypsin treatment, counted in Coulter counter and plated at a
concentration of 0.5X105 /ml into a 6 well plate. Cells were allowed to grow, and 12, 24,
36, 48, 72 and 96 hr after seeding, cells were detached from one well by trypsin and
counted in Coulter counter.
Mouse 3T3L1 cells from ATCC were cultured in DMEM with 10% Bovine Serum
39
(Gibco, #16170‐078), 2mM L‐glutamine, 100 units/ml penicillin and 100ug/ml
streptomycin. Culture medium was refreshed every other day. Cells were split 1:10
before they reached 70% confluence.
C3H10T1/2 cells from ATCC were cultured in α–MEM (Gibco, #12561‐056) with
10% heat‐inactivated FBS, 2mM L‐glutamine, 100 units/ml penicillin and 100ug/ml
streptomycin. Culture medium was refreshed every other day. Cells were split 1:10
before they reach 70% confluence.
For induction of adipogenesis, cells were seeded in six well plate with 106 cells
per well. On day 0 (2 days after cells reach confluence), adipogenesis was induced by
changing from growth medium to MDI medium which is DMEM with 10% FBS, 1μg/ml
(0.2μg/ml for 3T3L1) insulin（Gibco, #12585‐014） , 0.5mM isobutylmethylxanthine
(Sigma, #I7018), and 1μM Dexamethasone (Sigma, #D4902). Forty‐eight hours later,
culture medium was changed to insulin medium which is DMEM with 10% FBS
containing 1μg/ml insulin (0.2μg/ml for 3T3L1) for 4 days. On day 6, cells were refreshed
with DMEM with 10%FBS. Cells were left in DMEM for at least 4 days. After
differentiation, adipocytes were either stained by Oil Red O or used to extract mRNA by
TRIzol. In order to test effects of different compounds on adipogenesis, 2μM STO‐609
(TOCRIS, 1551), 0.25mM or 0.5mM AICAR (Calbiochem, #123040), or 0.5μM KN‐93
(Calbiochem, #422711) were added into medium 2 days before confluence (day ‐4) and
refreshed every other day. Compound C (Calbiochem, #171261) was used at
40
concentration of 2μM for 1 hr in WT MEFs.
Oil Red O staining
After differentiation, medium was removed by aspiration. Cells were rinsed once with
HBSS (Cellgro, #21‐022‐CV) and fixed in 10% buffered formalin for 10 minutes. Then,
cells were incubated in propylene glycol (Sigma, 398039) for 2 minutes and switched to
0.5% Oil Red O solution (0.5g Oil Red O in 100ml propylene glycol) for 16 hr. After the
staining, Oil Red O solution was removed by aspiration and remaining Oil Red O was
removed by treating cells with 85% propylene glycol for 1 minute. Stained cells were
replenished with HBSS and pictures were taken using an Olympus DP20 system
connected to microscope. In order to evaluate the amount of neutral fat droplet stained
by Oil Red O, stained cells were washed by isopropyl alcohol which will elute Oil Red O
from the cells and the optical density of eluate was monitored spectrophotometrically at
520nm.
2.4 RNA and Protein Analysis
Immunoblotting
Cells or tissue were lysed in RIPA buffer which consists of 20 mM Tris‐HCl (pH 7.5), 150
mM NaCl, 1 mM Na2EDTA, 1 mM EGTA, 1% NP‐40, 1% sodium deoxycholate, 2.5 mM
sodium pyrophosphate, 1 mM beta‐glycerophosphate, 1 mM Na3VO4, 1 μg/ml leupeptin,
41
0.1uM okadaic acid, 20mM NaF and protease inhibitor cocktail set III (Calbiochem
#535140). After incubation on ice for 30 minutes, samples were centrifuged at 14,000 rpm
for 15 minutes in a cold centrifuge and supernatant was saved. Samples were
fractionated by SDS‐PAGE and transferred to Immobilon‐P membranes (Millipore
Corp.). Membranes were blocked for 1 hr in TBS containing 5% nonfat milk and then
membranes were incubated with primary antibody in TBS containing 5% BSA or milk.
Membranes were washed 4 times with 10 minutes each time in TBS with 0.5% Tween 20.
Horseradish peroxidase‐conjugated secondary antibodies were from Jackson
ImmunoResearch Laboratories, Inc. and were used at 1:5000 in TBS containing 5%
nonfat milk. Membranes were incubated in secondary antibody at room temperature for
1 to 2 hr and washed 4 times with 10 minutes each time in TBS with 0.5% Tween 20.
Exposure was done with the ECL kit from Amersham Biosciences (RPN2106). Protein
amount is quantified with ImageJ software. Antibodies used are listed in Table 2.
42
Table 2: Antibodies
Catalog No. Company Species Dilution
Anti‐phospho‐ AMPKα (Thr172)
#2535 Cell Signaling Rabbit 1:1000
Anti‐AMPKα #2532 Cell Signaling Rabbit 1:1000
Anti‐phospho‐ACC (S79)
#3661 Cell Signaling Rabbit 1:1000
Anti CaMKK #610545 BD Biosciences Mouse 1:1000
Anti‐β‐actin #A5441 Sigma Mouse 1:2000
Anti‐phospho‐p44/42 MAPK (Erk1/2) (Thr
202/Tyr 204) #9106S Cell Signaling Mouse 1:1000
Anti‐p44/42 MAPK (Erk1/2)
#9107 Cell Signaling Mouse 1:1000
Peroxidase‐conjugated affinipure goat anti‐rabbit IgG (H+L)
111‐035‐003Jackson
ImmunoResearchGoat 1:5000
Peroxidase‐conjugated affinipure goat anti‐mouse IgG (H+L)
115‐035‐003Jackson
ImmunoResearchGoat 1:5000
In Situ hybridization
Brains were removed from 3‐month‐old mice, frozen in powdered dry ice, and stored at
‐80 degree Celsius. Ten micrometer coronal sections, cut serially on a cryostat, were
collected at the midregion of the hypothalamus (bregma ‐1.7), thaw mounted, and
stored at ‐80 degree Celsius until use. The slides were then processed for in situ
43
hybridization by transferring directly into ice‐cold 4% paraformaldehyde and fixing for
10 min at room temperature. After a 10 min wash in 2 X SSC, sections were crosslinked
for 5 min with an ultraviolet light placed 30 cm from the tissue. Prehybridization,
hybridization, and washing steps were then performed as described previously
(Anderson et al., 1998). RNA probes were synthesized as follows: cDNAs encoding PCR‐
generated fragments of NPY, MCH, and CaMKK2 coding regions were subcloned into
the polylinker of the pCR2 (Invitrogen) vector. The PCR primer pairs were forward
primer 131–151 and reverse primer 665–646 for CaMKK2, 5’‐
CTCCGCTCTGCGACACCTAC‐3’ (forward) and 5’‐AATCAGTGTCTCAGGGCT‐3’
(reverse) for NPY, and 5’‐ATTCAAAGAACACAGGCTCCAAAC‐3’ (forward) and 5’‐
CGGATCCTTTCAGAGCGAG‐3’ (reverse) for MCH. Runoff [S35] UTP‐labeled sense and
antisense transcripts corresponding to base pairs ‐10–524 of CaMKK2, base pairs 220–294
of NPY, and base pairs 307–399 of MCH were generated using SP6 or T7 RNA
polymerase according to the manufacturer’s protocol (Promega). Unincorporated
nucleotides were removed by washing the EtOH‐precipitated RNA pellets with 70%
EtOH.
Immunocytochemistry to detect NPY protein in N38 cells
N38 cells were cultured on glass coverslips. Following treatment with STO‐609,
ionomycin, or 2‐DG, the cells were fixed for 10 min in 1% paraformaldehyde/PBS and
44
washed and permeabilized for 10 min in 0.05% Triton X‐100, 20mM Tris (pH 7.4), 50 mM
NaCl, 300 mM sucrose, and 3 mM MgCl2. After permeabilization, the cells were washed
and blocked for 20 min in 5% normal sheep serum/PBS and incubated overnight at 4
degree Celsius with rabbit anti‐NPY (Sigma, N9528) diluted 1:4000 in 5% sheep
serum/PBS. The cells were washed and incubated in the dark for 1 hr with anti‐rabbit
Cy3 (Jackson ImmunoResearch Laboratories, Inc.) diluted 1:200 in 5% sheep serum/PBS,
washed again, and mounted in hard‐set mounting medium (Vectashield) with DAPI. All
washing steps were performed in triplicate for 5 minutes with PBS, and the procedure
was performed at room temperature except for the 4 degree Celsius incubation with
primary antibody.
RNA isolation, reverse transcription, PCR and RT‐PCR
Total RNA from cells and tissues was extracted using TRIzol (Invitrogen, #15596‐026)
and quantified spectrophotometrically. Single‐stranded cDNA was synthesized using
SuperScript II Reverse Transcriptase (Invitrogen, #18064‐022) according to the
manufacturer’s directions. PCR was carried out with Taq DNA Polymerase (Apex)
according to manufacturer’s directions and primers for CaMKK2: 5’‐
TCCACTTGGGCATGGAATCCT‐3’ (forward) and 5’‐TCCTGGTACACTTGCTCGATG‐
3’ (reverse). PCR protocol is an initial denaturing step of five minutes at 95oC followed
by 35 cycles of 95 oC for 30 seconds, 60 oC for 30 seconds and 72 oC for 1 minute and then
45
a final extension at 72 oC for 10 minutes. Real‐time PCR was carried out using iCycler
(Bio‐Rad) with the IQ SYBR Green Supermix (Bio‐Rad, #170‐8882) and the primers for
different genes are shown in Table 3. After deriving the relative amount of each
transcript from a standard curve, transcript levels were normalized to 18S ribosomal
RNA.
Table 3: RT‐PCR primers
Gene Forward Reverse
CaMKK2 5’‐CATGAATGGACGCTGC‐3’ 5’‐TGACAACGCCATAGGAGCC‐3’
18S 5’‐AGGGTTCGATTCCGGAGAGG‐3’ 5’‐CAACTTTAATATACGCTATTGG‐3’
C/EBPβ 5’‐CGCCTTTAGACCCATGGAAG‐3’ 5’‐CCCGTAGGCCAGGCAGT‐3’
C/EBPδ 5’‐GACTCCTGCCATGTACGA‐3’ 5’‐GGTTGCTGTTGAAGAGGT‐3’
C/EBPα 5’‐TATAGACATCAGCGCCTACATCGA‐3’ 5’‐GTCGGCTGTGCTGGAAGAG‐3’
PPARγ 5’‐GGCCATCCGAATTTTTCAAG‐3’ 5’‐GGGATATTTTTGGCATACTCTGTGA‐3’
aP2 5’‐AAGAAGTGGGAGTGGGCTTTG‐3’ 5’‐CTCTTCACCTTCCTGTCGTCTG‐3’
Glut4 5’‐CATTCCCTGGTTCATTGTGG‐3’ 5’‐GAAGACGTAAGGACCCATAGC‐3’
Fas 5’‐CCGAAGCCACAAAGCT‐3’ 5’‐GTCAAGGTTCAGGGTGC‐3’
Pref‐1 5’‐AATAGACGTTCGGGCTTGCA‐3’ 5’‐TCCAGGTCCACGCAAGTTCCATTGTT‐3’
Sox9 5’‐AGGTTTCAGATGCAGTGAGGAGCA‐3’ 5’‐ACATACAGTCCAGGCAGACCCAAA‐3’
46
2.5 RNAi, Transfections and Virus Infections
shRNA against CaMKK2 was obtained from Open Biosystems, including empty pLKO1
vector, shRNA3143 (TRCN0000028761) and shRNA3145 (TRCN0000028776). These
vectors were transfected into 293T cells with lentiviral packaging plasmids to make viral
particles by FuGENE (Roche, 11 814 443 001) according to manufacturer’s
recommendations. 24 and 48 hr after transfection, cell medium was harvested, filtered
and centrifuged at 25000 rpm/4 degree Celsius for 3 hr to collect virus. 3T3L1
preadipocytes were infected with virus containing pLKO1 vector, 3143 or 3145 shRNA
vector using polybrene according to the manufacturer’s recommendations; the next day,
infected cells were screened by treating with 2μg/ml puromycin. Selected stable cells
were maintained in media with 1μg/ml puromycin.
Three siRNAs against AMPKα1 and three against AMPKα2 were obtained from
Integrated DNA Technologies, and their sequences are shown in Table 4. Control
siRNA or AMPKα siRNA was transfected into WT MEFs using Lipofectamine 2000
reagent (Invitrogen, #11668‐019) according to the manufacturer’s recommendations.
Protein and RNA were harvested 4 days after transfection.
Ad‐CMV‐GFP and Ad‐Cre‐IRES‐GFP viruses were obtained from Vector Biolabs.
Flox‐CaMKK2 MEFs were plated in 6‐wells plates and infected with 4X109 Pfu/well
viruses using polybrene. Protein and RNA were extracted 6 days after infection.
47
Table 4: siRNA targeting AMPKα
siRNA targeting AMPKα1 siRNA targeting AMPKα2
5’‐GUGGAGUAGCAGUCCCUGAUUUGGCUU‐3’3’‐CACCUCAUCGUCAGGGACUAAACCG‐3’
5’‐UUUGAAUUACCCACCCUGCUUGUUCAA‐3’ 3’‐AAACUUAAUGGGUGGGACGAACAAG‐5’
5’‐GCUUGAUUGCUCUACAAACUUCUGCCA‐3’3’‐CGAACUAACGAGAUGUUUGAAGACG‐5’
5’‐ACAGAAAGCCACUGCUCUCUUACUCUU‐3’ 3’‐TGUCUUUCGGUGACGAGAGAAUGAG‐5’
5’‐UAUAUAUACCUUAACAUCUCAGUGGUU‐3’3’‐ATAUAUAUGGAAUUGUAGAGUCACC‐5’
5’‐CAGCACAUUCUCUGGCUUCAGGUCCCU‐3’ 3’‐GTCGUGUAGAGACCGAAGUCCAGG‐5’
Statistical analysis
Statistical significance was tested with unpaired student’s t test or ANOVA (GraphPad).
P values < 0.05 were considered as significant.
48
3. CaMKK2 contributes to the hypothalamic regulation of energy balance
3.1 Introduction
Energy balance is achieved by precisely matching anabolic and catabolic processes. If
energy expenditure is exceeded by energy intake, the unused energy will be stored in
the form of fat and can result in obesity. The incidence of obesity and associated diseases
such as type 2‐diabetes, hypertension and cardiovascular disorders, has reached
epidemic proportions world‐wide (Mokdad et al., 2003; Must et al., 1999; Zamboni et al.,
2005). Extensive studies have been done to understand the mechanisms responsible for
food intake. The CNS, especially the hypothalamus, has been emphasized in integrating
hormonal and nutrient signals from the periphery to modulate food intake and body
weight (Morton et al., 2006). One especially important orexigenic hormone functioning
in the CNS is ghrelin, which is produced by the stomach and stimulates food intake via
the hypothalamus. Ghrelin mediates its effect on food intake by upregulation of NPY
and AgRP, the most potent physiological appetite transducers known in mammals
(Kalra and Kalra, 2004). At the molecular level, ghrelin binds the Gq‐coupled receptor
expressed on NPY neurons and leads to an increase of intracellular Ca2+ which is
required for transcriptional activation of the NPY and AgRP. Blocking NPY action or
selectively ablating NPY neurons in rodents suppresses feeding and reduces obesity and
49
fasting‐induced responses (Broberger and Hokfelt, 2001; Gropp et al., 2005; Luquet et al.,
2005). These observations have promoted considerable effort to design antagonists
against NPY or its receptors as therapeutics. However, poor bioavailability and brain
penetration and non‐selective nature of action have thus far confounded development of
an effective therapeutic and have underscored the need to identify additional signaling
components in the pathway (Kalra and Kalra, 2004).
Around 2005, AMPK was identified as one component downstream of ghrelin
and upstream of NPY/AgRP. The i.p. or i.c.v. injection of ghrelin elevated the
phosphorylation and activation of AMPK in the hypothalamus; while the i.c.v.
administration of AICAR induced food intake and body weight gain. In addition,
overexpression of dominant negative or constitutively active AMPK revealed a
correlation between AMPK activity and expression of NPY (Minokoshi et al., 2004).
However, how the ghrelin signal reaches AMPK still remains unclear, as we know
AMPK needs to be phosphorylated and activated by an upstream kinase. CaMKK2 was
shown to function as a physiologically relevant AMPK kinase in cells, responding to
elevated intracellular Ca2+. The abundant expression of CaMKK2 in brain led us to
predict that Ca2+/CaM/CaMKK2 may regulate the phosphorylation of AMPK in NPY
neurons of the hypothalamus (Witters et al., 2006).
Here we show that one primary defect in CaMKK2‐null mice is reduced
hypothalamic AMPK activity and downregulation of NPY and AgRP gene expression in
50
NPY neurons. Acute i.c.v. injection of a CaMKK inhibitor to WT mice results in
decreased food intake that correlates with decreased hypothalamic NPY and AgRP
mRNAs, but not in CaMKK2‐null mice. Intriguingly, the absence of CaMKK2 also
protects mice from diet‐induced weight gain, glucose intolerance and insulin resistance.
Taken together, our data suggest CaMKK2 functions as an AMPK kinase to regulate
food intake in the hypothalamus.
3.2 Results
CaMKK2 regulates NPY in the hypothalamus
Immunoblotting of hypothalamic extracts from wild type (WT) and CaMKK2‐null (KO)
mice revealed that CaMKK2 protein is present in this part of the brain appearing as a
doublet (Fig. 6A). To identify special regions where CaMKK2 might co‐localize with
AMPK and thus function as an AMPKK, its expression pattern was examined by in situ
hybridization. NPY and melanin‐concentrating hormone (MCH) are neuropeptides
expressed within neurons present only in the ARC or the VMH respectively, and serve
as positive controls. Consistent with previous studies in rat brain, CaMKK2 signal is
strong in the hippocampus and cortex and is shown here also present in the
hypothalamus, especially the ARC (Fig. 6B). ARC punches also enriched CaMKK2, NPY,
AgRP and POMC mRNA relative to the whole hypothalamus (Fig. 6C). In contrast,
51
MCH, the control for VMH, was barely detectable. To evaluate the possibility of a
functional correlation between CaMKK2 and NPY, we compared hypothalamic NPY,
AgRP, and POMC mRNA levels in WT and CaMKK2‐null mice. Fig. 6D shows decreased
NPY and AgRP levels in the mutant mice, but no significant difference in POMC. These
results implicate CaMKK2 in a pathway that regulates NPY and AgRP in NPY neurons.
Since CaMKK2 can function as an AMPK kinase in cells, we questioned whether the
absence of CaMKK2 alters hypothalamic AMPK activity. AMPK was
immunoprecipitated from protein extracts prepared from WT and CaMKK2‐null mice
and assayed in vitro. A statistically significant decrease in AMPK activity was observed
in the CaMKK2‐null samples (Fig. 6E), consistent with diminished AMPK‐NPY signaling.
We suspect that the modest decrease in total hypothalamic AMPK activity reflects the
observation that CaMKK2 expression is restricted to a small region of the hypothalamus
and is considerably enriched in punch biopsies containing the ARC (Fig. 6B and 6C),
whereas AMPK activity is detected throughout the hypothalamus (Minokoshi et al.,
2004).
Ghrelin can bind to GHSR present on NPY neurons, which leads to an increase in
intracellular Ca2+ and activation of the AMPK‐NPY pathway (Cummings et al., 2005).
This signaling pathway was further evaluated by systemic administration of ghrelin to
WT and CaMKK2‐null mice. Ghrelin increased food intake of WT animals at 1 hr and
overnight but had no effect on food intake of CaMKK2‐null mice (Fig. 6F). The ghrelin
52
53
Figure 61: CaMKK2 is expressed in the hypothalamus and regulates NPY.
A. CaMKK2 protein is expressed in the hypothalamus. Protein extracts of hypothalamus from wild‐type (WT) and CaMKK2‐null (KO) mice were immunoblotted for CaMKK2. CaMKK2 protein appears as a doublet in the WT sample and is absent from the KO sample. B. CaMKK2 mRNA is expressed in the hypothalamus. Coronal mouse brain sections collected at the midregion of the hypothalamus were incubated as indicated with antisense riboprobes against NPY and MCH, which serve as positive controls for the arcuate nucleus (ARC) and ventromedial hypothalamus (VMH), respectively, and against CaMKK2. C. CaMKK2, NPY, AgRP, and POMC mRNAs are enriched in ARC punch biopsies from hypothalamus of WT mice. n = 3. D. NPY and AgRP mRNA levels are decreased in hypothalamus of CaMKK2‐null mice. Total RNA was isolated from hypothalamus of WT and CaMKK2‐null mice that had been fasted overnight, and NPY, AgRP, and POMC mRNA levels were quantified by RT‐PCR. n = 8; *p < 0.05. E. AMPK activity is decreased in hypothalamus of CaMKK2‐null mice. Following immunoprecipitation of AMPK from hypothalamic extracts of fasted WT and CaMKK2‐null mice, AMPK activity was determined in vitro with SAMS peptide as substrate. A representative experiment is shown. n = 4; *p < 0.05. F. Food intake of CaMKK2‐null mice is unaffected by exogenously administered ghrelin. WT and CaMKK2‐null mice were given intraperitoneal (i.p.) injections of saline or ghrelin (3 nmol/100 ml per mouse) 1 hr before onset of dark cycle, and food intake was quantified at 1 hr and overnight. n = 11 (WT), n = 12 (CaMKK2‐null); *p < 0.05. G. Food intake of CaMKK2‐null and WT mice is affected similarly by 2‐DG. In the control experiment, mice were given i.p. injections of 2‐deoxyglucose (2‐DG) at a dose of 15 mg/100 ml per mouse, and food intake was monitored as in (F). n = 11 (WT), n = 12 (CaMKK2‐null); *p < 0.05. Data are presented as mean ± SEM.
1 Figure 6A‐E were kindly contributed by Dr. Kristin Anderson.
54
signal can be bypassed by 2‐DG, which causes AMP/LKB1‐dependent activation of
AMPK due to depletion of cellular energy stores and a rise in the AMP/ATP ratio. As
shown in Fig. 6G, systemic administration of 2‐DG stimulated the food intake of both
WT and CaMKK2‐null mice to the same extent. Collectively, our data indicate a signaling
defect in the NPY neurons of CaMKK2‐null mice lying downstream of ghrelin and
upstream of AMPK that affects the expression of NPY and AgRP mRNAs.
STO-609 Blocks Ca2+-Dependent Induction of NPY
In order to provide more direct evidence linking CaMKK2, AMPK, and NPY
production, we employed N38 cells, an immortalized cell line derived from mouse
hypothalamus that have been shown to express NPY. These cells express CaMKK2
(Fig.7A). In these cells, AMPK signaling can be activated by treating the cells with
ionomycin, which increases intracellular Ca2+ leading to activation of CaMKK2 (Fig. 7B).
2‐DG, which raises the AMP/ATP ratio, also activates AMPK in N38 cells indicated by
increased levels of phosphorylated AMPK and phosphorylated ACC. The selective
CaMKK2 inhibitor STO‐609 blocks the ionomycin‐induced increases in AMPK and
ACC phosphorylation, but not the activation of AMPK caused by 2‐DG (Fig. 7B). We
used immunocytochemistry to show that ionomycin increases the amount of NPY in
N38 cells and that this increase is prevented by the selective CaMKK inhibitor STO‐609
(Fig. 7C and 7D). These data confirmed in vitro that a rise in Ca2+ results in activation of
55
56
Figure 72: Ca2+‐dependent induction of NPY expression in N38 cells is blocked by STO‐609
A. CaMKK2 protein is detected in N38 cell protein extracts by immunoblotting. B. N38 cells were incubated for 1 hr with vehicle or with 10 mM STO‐609, followed by stimulation with 1 mM ionomycin for 5 min or 50 mM 2‐DG for 15 min. Cell extracts were prepared and immunoblotted for phosphorylated AMPK (p‐AMPK), total AMPK, and p‐ACC. STO‐609 selectively blocks the ionomycin‐induced increase in p‐AMPK and p‐ACC. One representative experiment is shown. C. N38 cells were incubated for 1 hr with vehicle or with 10 mM STO‐609, followed by stimulation with 1 mM ionomycin or with 50 mM 2‐deoxy glucose (2‐DG) for 6 hr. The cells were then fixed, and the NPY protein signal (red) was visualized by immunocytochemistry. The cells were co‐stained with the DAPI nuclear marker (blue). One representative experiment is shown. n = 3. D. NPY signal intensity in (C) was quantified using MetaMorph version 7.1 software. Forty cells were analyzed for each condition. Shown are mean ± SEM. n = 40; *p < 0.0002.
2 Figure 7 was kindly contributed by Dr. Kristin Anderson.
57
CaMKK2, which in turn phosphorylates AMPK, leading to an increase in NPY.
Depletion or Inhibition of CaMKK2 Inhibits Food Intake
NPY is a well‐established potent orexigenic factor, and depleting NPY signaling in mice
reduces refeeding after a fast. This phenotype is most severe during the first hour of the
refeeding period and gradually returns to normal by 24 hr (Segal‐Lieberman et al., 2003).
When refeeding behavior was examined in CaMKK2‐null animals, they were found to
exhibit a defect similar to that previously reported for NPY‐depleted mice (Fig. 8A).
After a 48 hr fast, CaMKK2‐null mice ate less than WT mice during a 6 hr refeeding
period, with the greatest decrease in food intake occurring at the early time points, but
by 48 hr the difference was abrogated (Fig. 8A). Fig. 8B shows that the food intake of
nonfasted WT and CaMKK2‐null animals over a 48 hr period is the same. i.c.v.
administration of NPY to rats led to robust induction of food intake, while chronic
infusion resulted in increased adiposity and insulin resistance (Clark et al., 1984), and
ablation of NPY neurons in adult mice induced life‐threatening anorexia (Gropp et al.,
2005; Luquet et al., 2005). The hypothesis that CaMKK2 controls appetite by regulating
NPY gene expression in the hypothalamus predicts that acute inhibition of CaMKK2 in
WT animals should decrease appetite but not in CaMKK2‐null mice. To test this idea, the
selective CaMKK inhibitor STO‐609 was administered i.c.v. to adult male WT mice, and
58
59
Figure 83: Deletion or inhibition of CaMKK2 inhibits food intake
A. CaMKK2‐null mice display reduced refeeding after a fast. Food intake was quantified hourly during a 6 hr refeeding period and at 48 hr, following a 48 hr fast. n = 7; *p < 0.02. B. In the control experiment, the food intake of nonfasted animals was measured. C and D. Intracerebroventricular (i.c.v.) administration of STO‐609 to WT mice, but not to CaMKK2‐null mice, decreases food intake. STO‐609 was administered i.c.v. to WT and CaMKK2‐null mice continuously for 6 days at a concentration of 20 mM and a rate of 0.5 ml/hr, during which time food intake was measured daily. Cumulative food intake is shown. n = 9; *p < 0.05 (C); n = 10 (D). E and F. Change in body weight by the end of the 6 day experiment is plotted as a percentage of the starting value obtained before the cannulation surgery. n = 7. G. At the end of the experiment, WT animals were sacrificed, and hypothalamic NPY and AgRP mRNAs were quantified by real‐time PCR. n = 7; *p < 0.05. Data are presented as mean ± SEM.
3 Figure 8A and 8B were kindly contributed by Thomas Ribar.
60
the effect on appetite was assessed by quantifying food intake. Compared to animals
receiving saline, the WT STO‐609 group ate significantly less food during the 6 days
over which food intake was monitored (Fig. 8C) and also lost body weight (Fig. 8E). We
repeated the same experiment using adult male CaMKK2‐null mice. Whereas the mutant
mice receiving vehicle consumed a similar amount of food as vehicle treated WT mice,
STO‐609 did not change the feeding behavior or decrease the body weight of CaMKK2‐
null mice (Fig. 8D and 8F). In addition, STO‐609 also decreased hypothalamic NPY and
AgRP mRNAs in the WT mice (Fig. 8G). The effects of STO‐609 on food intake and NPY
and AgRP mRNA expression in WT mice, but not CaMKK2‐null mice, provide
compelling independent evidence that CaMKK2 signaling is required for appetite
control.
CaMKK2-null mice are resistant to high-fat diet-induced obesity, glucose intolerance and insulin resistance
Impaired NPY signaling in CaMKK2‐null mice indicates these mice will consume less
food and accumulate less body weight. However, we did not observe predicted
phenotype when mice were fed standard chow. Therefore we utilized the Surwit diet; a
nutritionally paired low‐fat/high‐fat diet that is commonly used for the diet‐induced
obese model and that was developed for C57BL/6J mice, the predominant genetic
background of our mice (Petro et al., 2004; Surwit et al., 1988). WT and CaMKK2‐null
mice were fed either a low‐fat control diet (D12328 CCO) or the high‐fat equivalent of
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this diet (D12330 HCO) from weaning (3 weeks of age) to 34 weeks of age. The body
weight gain of CaMKK2‐null mice was significantly decreased compared to WT control
animals fed the low‐fat diet (Fig. 9A), and this difference was even bigger for animals
maintained on the high‐fat diet (Fig. 9B). As predicted, CaMKK2‐null mice displayed a
reduced average daily food intake over the course of the experiment (Fig. 9C) and they
also displayed a reduction in adiposity measured after 31 weeks on either diet (Fig. 9D).
Glucose tolerance testing revealed that WT mice on the high‐fat diet had become less
tolerant compared to WT animals maintained on low‐fat diet (Fig. 9E and 9F). CaMKK2‐
null mice fed the low‐fat diet responded to glucose almost the same as WT mice on the
same diet. However, CaMKK2‐null mice fed the high‐fat diet managed to remain glucose
tolerant as well as mutant mice kept on low‐fat diet (Fig. 9E and 9F). After 1 week of
recovery, the same group of animals was tested for insulin sensitivity. Consistent with
the results from the glucose tolerance test, WT mice fed high‐fat diet had developed the
predicted insulin resistance, whereas CaMKK2‐null animals were protected from diet‐
induced changes in insulin resistance (Fig. 9G and 9H). After an additional several
weeks of recovery, the animals were sacrificed and serum levels of ghrelin and leptin
were quantified. As shown in Fig. 9I, CaMKK2‐null mice did not exhibit a decrease in
ghrelin levels, indicating their ability to synthesize and secrete ghrelin. In fact, ghrelin
levels were increased in CaMKK2‐null mice fed high‐fat diet, suggesting the possibility
that these mice attempt to compensate for a defect in the ghrelin signaling pathway by
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Figure 94: CaMKK2‐null mice consume less food and are resistant to high‐fat diet‐induced adiposity, glucose intolerance and insulin resistance
Twenty male WT and CaMKK2‐null mice were housed five per cage at the age of weaning and fed control (D12328) or high‐fat (D12330) diets from Research Diets for 31 weeks. Measurements of food intake and weight gain were performed weekly. A and B. Growth curves of the two groups on both diets show that the CaMKK2‐null mice gain significantly less weight than the WT mice. *p < 0.01 by one‐way ANOVA. C. Average daily food intake of CaMKK2‐null mice is reduced compared to control animals. *p < 0.02. D. Dual energy X‐ray absorptiometry (DEXA) scans of CaMKK2‐null mice indicate a significant decrease in adiposity compared to WT animals after 31weekson either control or high‐fat diet. *p<0.02. E‐H. On the high‐fat diet, CaMKK2‐null mice also remain glucose tolerant (E and F) and insulin sensitive (G and H) relative to WT mice. *p < 0.01. I and J. After 31 weeks on low‐fat (n = 5) or high‐fat (n = 10) diet, WT and CaMKK2‐null mice were sacrificed, and total serum ghrelin (I) and leptin (J) levels were quantified. *p < 0.05. K. After 31 weeks on high‐fat chow, WT and CaMKK2‐null mice were either fasted overnight or fed ad libitum. The animals were then sacrificed, and hypothalamic NPY mRNA levels were quantified by real‐time PCR. n = 4; *p < 0.02. Data are presented as mean ± SEM.
4 Figure 9 was kindly contributed by Thomas Ribar.
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increasing its production. Serum leptin was decreased in CaMKK2‐null mice fed high‐fat
diet compared to WT mice (Fig. 9J). This may result from lower adiposity in CaMKK2‐
null mice. It is also possible that mutant animals attempted to compensate for the defect
in the ghrelin pathway by depressing the opposite leptin signaling. Hypothalamic NPY
mRNA was decreased in the CaMKK2‐null mice fed the high‐fat diet relative to WT mice,
and the decrease was accentuated upon fasting (Fig. 9K). Together, these data are
consistent with our observations in Fig. 6 that CaMKK2‐null mice cannot respond to
ghrelin with the appropriate increase in feeding and that this may be partially due to
decreased production of NPY.
3.3 Discussion
In this chapter, we provide evidence that hypothalamic CaMKK2 mediates ghrelin‐
induced feeding. In the absence of CaMKK2, mice ate less and accumulated less body
weight and fat stores when maintained on either low‐fat or high‐fat diet. The CaMKK2‐
null animals expressed reduced levels of hypothalamic NPY and AgRP mRNAs and
were unresponsive to the orexigenic effects of exogenously administered ghrelin.
Furthermore, i.c.v. infusion of the selective CaMKK2 inhibitor STO‐609 in adult WT mice
resulted in the acute suppression of NPY expression and food intake. The resistance of
CaMKK2‐null mice to suppression of food intake by STO‐609 infusion provides
compelling evidence that CaMKK2 is the direct target of the drug and that the specific
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inhibition of CaMKK2 in NPY neurons is likely responsible for the decreased level of
NPY and AgRP mRNAs and feeding behavior.
We hypothesize that CaMKK2 stimulates NPY expression by functioning as the
AMPK kinase linking ghrelin‐dependent Ca2+ signaling to AMPK activation. The fact
that CaMKK2‐null mice exhibited reduced hypothalamic AMPK activity supports this
idea. According to this model, we would predict that CaMKK2 and NPY null mice
should share a similar metabolic phenotype. Indeed, CaMKK2‐null mice share reduced
refeeding behavior with NPY‐null mice. However, other than that, CaMKK2‐null mice
act quite normally when fed standard chow ad libitum. They do not show decreased
appetite and reduced body weight. In fact, CaMKK2‐null mice gain more adiposity,
which is described in chapter 4. Similar phenotypes were observed in NPY/NPY‐Y‐null
mice and people have suggested it results from certain compensatory mechanisms. Once
CaMKK2‐null mice were switched to the Surwit diet, they began to eat less and
accumulated less body weight and fat on either low‐fat or high‐fat diet. Our data
suggest that, like NPY‐null animals, CaMKK2‐null mice develop pathways to
compensate for diminished NPY signaling, and these pathways function to prevent
eating disorders as long as the animals are fed standard chow. Once on the low‐ or high‐
fat Surwit diets, the compensatory pathways acquired by CaMKK2‐null mice are
apparently no longer effective, and the feeding defects predicted from low NPY are
exposed.
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An intriguing phenotypic consequence of depleting CaMKK2 was observed in
animals maintained on the high‐fat diet, as these mice were protected from the glucose
intolerance and insulin resistance that developed in WT controls over the course of
several months. Although the mechanism responsible for the improved glucose
handling is unknown, there are several possibilities. Since obesity represents the leading
risk factor for the development of type 2‐diabetes, the improvements observed in
CaMKK2‐null mice may result directly from their accumulating less fat stores compared
to WT mice prior to glucose and insulin testing (Golay and Ybarra, 2005). Improved
glucose tolerance is also consistent with enhanced leptin signaling (Elmquist et al., 2005),
implicated in CaMKK2‐null mice by decreased hypothalamic AMPK activity (Minokoshi
et al., 2004). In fact, it is possible that enhanced leptin signaling in CaMKK2‐null mice
may contribute as well to the observed decreases in food intake and body weight gain.
We also cannot rule out the possibility that CaMKK2 regulates glucose homeostasis
through action in energy expenditure. NPY and AgRP are both shown to suppress
energy expenditure by inhibiting thermogenesis in brown adipose tissue (Billington et
al., 1991; Small et al., 2003). Absence of CaMKK2 in the hypothalamus may result in
increased energy expenditure through reducing NPY/AgRP signaling. Anyway, the
behavior of CaMKK2‐null mice on the high‐fat diet underscores the value of targeting
CaMKK2 as a possible therapeutic locus in the treatment of obesity and diabetes.
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4. CaMKK2 Regulation of Food Intake and Energy Balance Can Be Uncoupled
4.1 Introduction
As we showed in the last chapter, when fed high‐fat diet, CaMKK2‐null mice gained
much less adiposity compared to WT animals. At the same time CaMKK2‐null mice were
much more resistant to high‐fat diet‐induced glucose intolerance and insulin resistance.
Although we have revealed that these phenotypes could be explained by the reduction
of food intake resulting from decreased NPY/AgRP expression in the ARC of the
hypothalamus in CaMKK2‐null mice, we could not rule out the possibility that other
pathway(s) affected by loss of CaMKK2 also participate. In order to understand whether
the reduction of food intake is the only primary factor changed in CaMKK2‐null mice, a
pair feeding experiment using high‐fat diet was arranged.
Regulation of energy expenditure is also important to maintain energy balance.
One of the most significant energy dissipating mechanisms in mammals takes place in
BAT. BAT is critical for respiratory uncoupling and nonshivering thermogenesis in
rodents (Heldmaier and Buchberger, 1985). In BAT, uncoupling protein 1 (UCP1), a
proton carrier residing in the inner mitochondrial membrane, allows protons to leak
through the membrane bypassing ATP synthase and turning the energy directly into
heat (Nubel and Ricquier, 2006). BAT is regulated by the hypothalamus through the
sympathetic nervous system (SNS). It has been shown that inactivation of AMPK in the
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hypothalamus increased SNS activity and upregulated thermogenic markers expression
in BAT (Lopez et al., 2010). AgRP is also revealed to regulate BAT activity. Chronic
central nervous system administration of AgRP significantly decreased the amount of
UCP1 in rat (Small et al., 2001). In another pathway, NPY released from the ARC
interacted with Y5 receptors in the PVN, which leads to the inhibition of the VMH cells
and BAT activity (Billington et al., 1991, 1994). The decrease of AMPK activity and
reduced NPY/AgRP expression in the hypothalamus of CaMKK2‐null mice infers BAT
might also contribute to metabolic phenotype of knockout mice.
In this chapter, we show that even with same calorie intake on high‐fat diet,
CaMKK2‐null mice still gained less body weight and adiposity compared to WT controls
and knockout mice also displayed better glucose and insulin tolerance. Although
restriction on food intake did decrease the body weight gain in WT mice, similar
increase in adiposity, glucose intolerance and insulin resistance suggests that something
other than food intake is altered due to loss of CaMKK2. We also found that under 5001
standard chow, CaMKK2‐null mice possessed larger interscapular brown adipose tissue
and displayed stronger thermogenesis when exposed to a cold environment. However,
further DEXA scan analysis indicated that CaMKK2‐null mice had equal amount of BAT
on low‐fat diet or even less BAT on high‐fat diet compared to WT animals. Metabolic
chamber analysis also showed that pair fed CaMKK2‐null mice produced less heat on
high‐fat diet. We also presented that under 5001 standard chow, CaMKK2‐null mice
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have more adiposity and larger adipocyte size. When switched to low‐fat or high‐fat diet,
WT mice had a significant increase in adipocyte size but the increase in CaMKK2‐null
mice was much smaller. Taken together, our data indicate that CaMKK2 regulation of
food intake can be uncoupled from its effect on energy balance.
4.2 Results
WT mice pair-fed with CaMKK2-null mice still display high-fat diet-induced adiposity, glucose intolerance and insulin resistance
To test whether food intake is the primary reason for protection of CaMKK2‐null mice
from high‐fat diet‐induced obesity and diabetes, we employed a pair feeding experiment
to compare metabolic phenotypes between WT and knockout mice under high fat diet.
Young adult mice (3 months old) were housed individually and food intake of WT mice
was restricted to the amount that CaMKK2‐null mice consumed. After 30 weeks, WT and
CaMKK2‐null mice shared the same food intake, but WT mice still gained much more
weight compared to mutant mice (Fig. 10A and 10B). However, body weights of WT
mice were less compared to those when fed high‐fat diet ad libitum (Fig. 9B). DEXA scan
showed that WT mice still gained more adiposity compared to CaMKK2‐null mice (Fig.
11A). In glucose tolerance test, WT mice showed intolerance to glucose whereas
CaMKK2‐null mice remained tolerant (Fig. 11B). After a week of recovery, same groups
of mice were tested for insulin sensitivity. WT mice developed insulin resistance while
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Figure 10: CaMKK2‐null mice have the same food intake but less body weight gain in high‐fat diet pair feeding
Fourteen male 3 month‐old WT and CaMKK2‐null mice were housed individually and fed high‐fat (D12330) diets from Research Diets for 30 weeks. Measurements of food intake and weight gain were performed daily. A. Cumulative food intake curves show that CaMKK2‐null mice consumed same amount of food compared to WT animals. B. Growth curves of the two groups show that the CaMKK2‐null mice gain significantly less weight than the WT mice. *p < 0.01 by one‐way ANOVA.
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Figure 11: CaMKK2‐null mice gained less adiposity and remained glucose tolerant and insulin sensitive compared to pair‐fed WT mice
A. DEXA scans of CaMKK2‐null mice indicate a significant decrease in adiposity compared to WT animals after 30 weeks’ pair feeding on high‐fat diet. *p<0.05. B and C. CaMKK2‐null mice also remain glucose tolerant (B) and insulin sensitive (C) relative to WT mice. *p < 0.01. n=12 for WT mice; n=14 for CaMKK2‐null mice.
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CaMKK2‐null animals remained insulin‐sensitive (Fig. 11C). These data infer that food
intake is not the primary factor contributing to protection of CaMKK2‐null mice from
high‐fat diet‐induced obesity and diabetes.
CaMKK2-null mice have increased brown adipose tissue and thermogenic response to a cold environment on standard chow
Since normalizing food intake did not eliminate the metabolic difference between WT
and knockout mice, we tried to pursue the answer from the other direction. BAT is an
important organ for energy expenditure under control of SNS activity. Inactivation of
AMPK stimulates the activity of SNS and enhances BAT activity (Lopez et al., 2010).
Hypothalamic AgRP and NPY were both reported to inhibit BAT activity (Small et al.,
2001; Chao et al., 2011). We have shown that CaMKK2‐null mice had decreased AMPK
activity and reduced AgRP/NPY expression. Oury and colleagues also presented that
CaMKK2/CaMKIV signaling pathway in VMH phosphorylates and activates CREB,
which ultimately leads to inhibition of SNS (Oury et al., 2010). It is reasonable to predict
that activities of SNS and BAT would be enhanced in CaMKK2‐null mice. Therefore, we
checked BAT size in both types of mice. Indeed, there is more BAT in CaMKK2‐null mice
compared to WT animals when fed standard chow (Fig. 12A). We also tested their
thermogenic responses in a cold environment. Consistent with the observation of larger
BAT, CaMKK2‐null mice also displayed a stronger thermogenic ability to maintain their
body temperature in an 8 hr 8 degree Celsius exposure (Fig. 12B).
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Figure 12: CaMKK2‐null mice have increased BAT and enhanced cold‐induced thermogenesis on standard chow
A. The interscapular brown adipose tissue from 14 WT and CaMKK2‐null mice fed 5001 standard diet was quantified by DEXA scan. n=14, *p < 0.01. B. The thermogenic response of fasted WT and CaMKK2‐null mice was compared by housing animals individually at 8°C for 8 hr, during which time rectal temperature was monitored hourly. n = 10; *p < 0.006.
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CaMKK2-null mice have smaller BAT and produced less heat on high-fat diet
As we observed more BAT and enhanced thermogenesis capability in CaMKK2‐null
mice on standard chow, we wanted to know whether enhanced thermogenesis in BAT
protected knockout mice from high‐fat diet‐induced obesity. Analysis on DEXA scans of
mice fed low‐fat or high‐fat diet revealed the opposite of what we predicted. As shown
in Fig. 13A, CaMKK2‐null mice have an equal amount of interscapular BAT compared to
WT animal when fed low‐fat diet. WT mice had a large increase in BAT on high‐fat diet
but CaMKK2‐null mice only displayed a slight increase. We also directly isolated BAT
from mice pair‐fed on high‐fat diet. Again, we found much larger BAT in WT mice (Fig.
13B). Consistent with what we observed in BAT, metabolic chamber analyses revealed
that CaMKK2‐null mice produced significantly less heat compared to WT controls pair‐
fed on high‐fat diet. These data infer that BAT may not play an important role in
protecting CaMKK2‐null mice from high‐fat diet‐induced obesity and diabetes.
High-fat diet affects adiposity and adipocyte size differently in WT and CaMKK2-null mice
Although CaMKK2‐null mice had less adiposity compared to WT mice when fed low‐fat
or high‐fat diet, we found that adult mutant mice have significantly more adipose tissue
under standard diet, and this difference was not age‐dependent as it was also evident
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Figure 13: CaMKK2‐null mice have less BAT and produce less heat when fed high‐fat diet
A. DEXA scans on interscapular area of mice fed ad libitum on CCO or HCO diet. WT mice have more BAT under HCO. *p < 0.05. n=17 for WT CCO; n=18 for KO CCO; n=13 for WT HCO; n=16 for KO HCO. B. Half of interscapular BAT was isolated from mice pair fed on HCO diet. C. WT and CaMKK2‐null mice pair‐fed on high‐fat diet were housed in individual metabolic chamber, and heat production was monitored for 48 hr. *p < 0.001, n=12.
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Figure 14: Adipocyte analysis of WAT from WT and CaMKK2‐null mice fed different diets
A. DEXA scans of CaMKK2‐null (KO) mice indicate a significant increase in adiposity compared to WT animals at weaning (3 weeks, n=10, *p<0.02) or after 3 months on standard 5001 chow. n=10; * P<0.005. B. H&E staining of gonadal fat pad sections. C. Size of adipocytes in B was quantified by ImageJ software. n=50, *p< 0.001.
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Figure 15: Adiposity analysis on retroperitoneal and epididymal fat pads in WT and CaMKK2‐null mice fed different diets
A and B. Fat in the region corresponding to retroperitoneal fat (A) and epididymal fat (B) was quantified from DEXA scan. Shown are mean+/‐SEM. n=14 for WT or KO 5001; n=17 for WT CCO; n=18 for KO CCO; n=13 for WT HCO; n=16 for KO HCO. *p<0.05, **p<0.01, ***p<0.001.
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in 3‐week‐old mice at the time of weaning (Fig. 14A). Furthermore, H&E staining of
gonadal fat pad sections revealed that the size of adipocytes in adult CaMKK2‐null mice
was much larger than WT controls when fed standard chow. Interestingly, consistent
with the fact that WT mice had more adiposity on low‐fat or high‐fat diet (Fig. 9D),
adipocyte size of WT mice on low‐fat diet significantly increased but there was no clear
change for CaMKK2‐null mice. When animals were fed high‐fat diet, adipocyte size
increased even more in WT mice and increase was considerably smaller in CaMKK2‐null
mice (Fig. 14B and 14C). We also reanalyzed the DEXA scans from WT and CaMKK2‐
null mice fed on 3 different diets for specific regions corresponding to retroperitoneal
and epididymal fat pads, and similar changes were found (Fig. 15A and 15B). These
findings make us wonder whether CaMKK2 may play a role in the development of
adiposity.
4.3 Discussion
In this study, we demonstrated that reduction of appetite cannot entirely explain the
protection of CaMKK2‐null mice from high‐fat diet‐induced obesity and diabetes. In
order to evaluate the importance of food intake reduction, we carried out a pair feeding
experiment. WT mice which were restricted to the same amount of food consumed by
CaMKK2‐null mice still exhibited more adiposity accumulation, glucose intolerance and
insulin resistance.
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We also tried to understand this phenotype from the aspect of energy
expenditure. We found larger BAT and stronger thermogenic capability in CaMKK2‐null
mice on standard chow, which matches the hypothesis that knockout mice have
enhanced BAT activity due to inhibited AMPK/NPY/AgRP signaling. However,
significantly more adiposity was also discovered in these knockout animals. What is
more, when fed low‐fat or high‐fat diet, WT mice possessed equal or even more BAT
compared to CaMKK2‐null mice. This finding together with the fact that knockout
animals actually produced less heat on high‐fat diet informed us that resistance of
CaMKK2‐null mice to high‐fat diet did not result from enhanced energy expenditure.
These facts also indicated that signaling which enhanced BAT activity on standard chow
might be overcome by other means on high‐fat diet.
Finally, similar to the adiposity change among the 3 different diets, CaMKK2‐null
mice have more retroperitoneal and epididymal fat on standard diet but a considerably
smaller increase compared to WT mice when switched to low‐fat or high‐fat diet. The
sizes of adipocytes from gonadal fat pad also showed a similar trend. These facts
suggest CaMKK2 plays a role in development of adiposity, which will be studied in next
chapter.
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5. CaMKK2 Inhibits Preadipocyte Differentiation
5.1 Introduction
Obesity can arise due to excessive lipid accumulation in adipocytes resulting in
increased cell size coupled with the recruitment of adipocyte progenitor cells
(preadipocytes) to white adipose tissue (WAT) that differentiate into additional mature
adipocytes. WAT is not only an energy storage depot that helps to maintain lipid
homeostasis (Spiegelman and Flier, 2001), but also functions as a dynamic endocrine
organ. Adipocytes, which comprise the bulk of WAT mass, secrete hormones
(adipokines) that play central roles in regulating energy balance, insulin sensitivity,
immunological responses and vascular disease (Fruhbeck et al., 2001). Because
adipocytes play such a critical role, elucidating molecular pathways that control
adipocyte differentiation may help to develop strategies for the prevention and
treatment of obesity and diabetes.
Adipogenesis is regulated by calcium signaling pathways, as increases of
intracellular Ca2+ in preadipocytes during the early phase of differentiation could inhibit
adipogenesis (Ntambi and Takova, 1996; Miller et al., 1996; Shi et al., 2000) as could
exposure of preadipocytes to high levels of extracellular Ca2+ (Jensen et al., 2004).
Although the mechanism by which Ca2+ represses adipogenesis is not fully understood, a
logical one to explore is the CaM kinase cascade. In this cascade a rise in Ca2+ results in
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activation of CaM which, in turn, activates CaMKK2. CaMKK2 then phosphorylates
downstream kinases including CaMKI, CaMKIV (Anderson et al., 1998; Haribabu et al.,
1995; Tokumitsu et al., 1994; Lee and Edelman, 1994; Matsushita and Nairn, 1998;
Edelman et al., 1996; Selbert et al., 1995) and AMPK (Hawley et al., 2005; Hurley et al.,
2005; Woods et al., 2005). Of these three CaMKK2 substrates AMPK seems to be the most
relevant to adipogenesis for several reasons. First, it has been shown to inhibit fatty acid
synthesis and increase fatty acid oxidation in WAT (Daval et al., 2006; Xue and Kahn,
2006). Second, AICAR, an activator of AMPK, inhibits adipogenesis in 3T3L1 or F442A
preadipocytes (Habinowski and Witters, 2001; Giri et al., 2006; Dagon et al., 2006). Third,
Metformin, a drug frequently used to treat type 2‐diabetes, inhibits lipid accumulation
in 3T3L1 cells via a pathway that involves activation of AMPK (Alexandre et al., 2008).
Therefore, since the mechanistic details of how Ca2+ and AMPK participate in
adipogenesis remain unresolved we questioned if CaMKK2 was involved.
Our early studies on CaMKK2‐null mice resulted in the unexpected finding that
they have more adipose tissue than wild type mice when fed standard chow (5001).
Here we show that CaMKK2 is present in preadipocytes but significantly reduced in
adipocytes. Deletion or inhibition of CaMKK2 results in enhanced adipogenesis when
preadipocytes are exposed to medium that initiates their differentiation, but does not
affect cell proliferation. We further show that the effects of CaMKK2 on adipogenesis are
due to its ability to phosphorylate AMPK (rather than its other two substrates CaMKI or
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CaMKIV) in preadipocytes. Finally, in tracing the adipogenic pathway backwards we
show that loss of CaMKK2 or AMPK activity decreases Pref‐1 mRNA. Our results
suggest that a CaMKK2‐AMPK signaling pathway functions to maintain expression of
Pref‐1 in preadipocytes, which functions to inhibit adipogenesis.
5.2 Results
CaMKK2 is expressed in preadipocytes but barely detectable in WAT
Previously, CaMKK2 was mainly known to be abundantly expressed throughout the
brain especially in the hypothalamus (Anderson et al., 1998). We wanted to know
whether CaMKK2 exists in white adipose tissue (WAT). Immunoblot of gonadal fat
extracts could not detect CaMKK2 protein but reverse transcription PCR showed a weak
signal for CaMKK2 mRNA from whole WAT sample (Fig. 16A, B). We then tested
CaMKK2 expression in isolated primary preadipocytes. Reverse transcription PCR
showed that mRNA of CaMKK2 existed in WT primary preadipocytes and a smaller
mRNA form was also can be found in CaMKK2‐null primary preadipocytes (Fig. 16C);
which is consistent with exon 2 to exon 4 truncation in CaMKK2‐null mice. Also
immunoblot clearly showed that CaMKK2 protein existed in WT preadipocytes but not
CaMKK2‐null preadipocytes (Fig 16D). Several cell lines are able to differentiate into
adipocytes. We confirmed that CaMKK2 is also expressed in these preadipocytes.
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Figure 16: Expression of CaMKK2 in preadipocytes
A and B. CaMKK2 expression is barely detectable in WAT by immunoblot or reverse‐transcription PCR. C and D. Reverse‐transcription PCR (C) and immunoblot (D) show expression of CaMKK2 in preadipocytes; brain samples work as controls.
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Reverse transcription PCR again showed CaMKK2 mRNA in MEF, 3T3‐L1 and
C3H10T1/2 cells (Fig. 16C) and CaMKK2 protein can also be detected by western blot
(Fig. 16D). These data show that CaMKK2 is expressed in preadipocytes and could
therefore play a direct role.
CaMKK2 expression decreases after adipogenesis.
The fact that CaMKK2 is expressed in primary preadipocytes but barely detectable in
WAT suggested to us that CaMKK2 expression may decrease during adipogenesis. To
address this we used real‐time PCR to compare CaMKK2 mRNA before and after
adipogenesis with aP2 as a diagnostic marker for mature adipocytes. Primary
preadipocytes had high amounts of CaMKK2 mRNA and protein but little aP2 mRNA;
however, this was reversed in mature adipocytes as aP2 mRNA greatly increased while
CaMKK2 mRNA and protein greatly decreased (Fig. 17A, 17B). This reciprocal
relationship between CaMKK2 and aP2 was also observed in three other cell lines that
can be induced to differentiate into adipocytes namely, 3T3L1, MEF, and C3H10T1/2
cells (Fig 17A, 17B). These data show that CaMKK2 is present in 4 types of preadipocytes
but mature adipocytes differentiated from these cells contain markedly decreased
amounts of CaMKK2 mRNA and protein. These data suggested to us that CaMKK2
might be inhibitory to adipogenesis.
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Figure 17: CaMKK2 mRNA and protein expression decreases after adipogenesis
A. CaMKK2 mRNA decreases after adipogenesis. Total mRNA was isolated from primary‐cultured preadipocytes, adipocytes and from undifferentiated (UD, day 0) or differentiated (FD, day 10) MEFs, 3T3L1 and C3H10T1/2 cells. CaMKK2 mRNA level was quantified by RT‐PCR (top panel). aP2 is included as a marker of adipogenesis (bottom panel). Data are presented as mean +/‐ SEM. n=3; *p<0.05. B. CaMKK2 protein decreases after adipogenesis. Protein was extracted from primary‐cultured preadipocytes, adipocytes and from undifferentiated (UD, day 0) or differentiated (FD, day 10) MEFs, 3T3L1 and C3H10T1/2 cells. Samples were western blotted for panCaMKK. One representative blot (upper) and quantification (lower) of three independent experiments are shown. The amount of CaMKK2 protein in each type of preadipocyte is normalized to 1.
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CaMKK2 inhibits adipogenesis via phosphorylation of AMPK
As aP2 is a late marker of adipogenesis we questioned if depletion or inhibition of
CaMKK2 resulted in accelerated fat accumulation when preadipocytes were exposed to
agents that stimulate adipogenesis. As shown in Fig 18A, MEFs from CaMKK2‐null mice
formed more triglyceride (based on Oil Red O staining) containing cells than did WT
MEFs. Similarly, the presence of the CaMKK2 selective inhibitor STO‐609 accelerated
adipogenesis of 3T3L1 and C3H10T1/2 cells (Fig. 18 B, 18C). Finally, the depletion of
CaMKK2 from 3T3L1 cells using a shRNA strategy similarly accelerated adipogenesis
(Fig 18D).
These data support our contention that CaMKK2 functions in a signaling
pathway that prevents differentiation of preadipocytes in several cell models of
adipogenesis. However, CaM kinase pathways have been shown to support cell
proliferation (Kahl and Means, 2003). Since cells must exit from the proliferative cycle in
order to differentiate we questioned if the absence of CaMKK2 altered the doubling time
of preadipocytes isolated from CaMKK2‐null mice or MEFs in culture. We found that the
absence of CaMKK2 did not alter cell cycle progression of preadipocytes or MEFs (Fig.
19A, 19B) providing additional support that CaMKK2 primarily functioned to inhibit
adipogenesis.
CaMKK2 has been shown to have three direct substrates, CaMKI, CaMKIV and
AMPK, so we next questioned which of these substrates may lie downstream of
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Figure 18: Loss of CaMKK2 activity enhances adipogenesis
A. WT and CaMKK2‐null (KO) MEFs were induced to differentiate as outlined in the methods. After 10 days of differentiation, intracellular lipid was stained with Oil Red O (left). To quantify the amount of Oil Red O, the dye was eluted with isopropyl alcohol and its optical density was monitored spectrophotometrically at 520nm (right). B and C. Inhibition of CaMKK2 in 3T3L1 or C3H10T1/2 preadipocytes enhances adipogenesis. 3T3L1 or C3H10T1/2 preadipocytes were induced to differentiate in the presence of vehicle or 2μM STO‐609. Amount of adipogenesis was determined by Oil Red O staining (left) which was quantified as in (A) shown on the right. D. Deletion of CaMKK2 in 3T3L1 preadipocytes enhances adipogenesis. 3T3L1 preadipocytes were infected with control or one of two different shRNA viruses against CaMKK2 (3143, 3145). The efficiency of CaMKK2 knockdown was determined by western blot. Cells were induced to differentiate and analyzed as in A. Quantification of Oil Red O is presented as mean +/‐ SEM, *p<0.05. Data shown are representative of at least 3 independent experiments; and each was done in triplicate.
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CaMKK2 in the signaling pathway that prevents adipogenesis. Since we had previously
generated mice null for CaMKIV we compared CaMKIV‐null MEFs with WT and
CaMKK2‐null MEFs in the triglyceride accumulation assay. The absence of CaMKIV did
not accelerate adipogenesis (Fig. 20A). CaMKI is essential for cell proliferation and
functions predominantly in G1 of the cell cycle (Kahl and Means, 2003; Kahl and Means,
2004). Since the absence of CaMKK2 did not slow cell proliferation, we surmised that
CaMKI was not likely to be the relevant substrate supporting that anti‐adipogenic role
for CaMKK2. To confirm this idea we cultured 3T3L1 cells with KN‐93 which inhibits
CaMKI and CaMKIV with similar potency (Tokumitsu et al., 1990; Sumi et al., 1991;
Enslen et al., 1994; Mochizuki et al., 1993). As KN‐93 did not accelerate adipogenesis (Fig.
20B) we reasoned that neither CaMKI nor CaMKIV was the relevant target of CaMKK2.
To test the third substrate, AMPK, we cultured 3T3L1 cells from which CaMKK2 had
been depleted by shRNA and CaMKK2‐null MEFs with the AMPK activator, AICAR.
AICAR inhibited the ability of CaMKK2‐depleted cells to support adipogenesis in a
dose‐dependent manner suggesting that activation of AMPK was negatively correlated
with adipogenesis and could be the relevant substrate by which CaMKK2 inhibits
adipogenesis (Fig. 20C and 20D).
We wanted to confirm that CaMKK2 is a functional AMPK kinase in
preadipocytes. As has been shown in the hypothalamus (Anderson et al., 2008; Thornton
et al., 2008), western blots of primary preadipocyte extracts showed that basal AMPK
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Figure 19: CaMKK2‐null MEFs and preadipocytes exhibit no growth defect
A. WT or CaMKK2‐null MEFs (KO) were plated at the same concentration in 6 well plates. At indicated time points, cells were detached by trypsin treatment and cell numbers were determined using a particle counter. There was no significant difference in proliferation between WT and KO MEFs. n=3. B. Isolated WT or CaMKK2‐null preadipocytes (KO) were allowed to grow in culture for 4 days. Preadipocytes were then detached by trypsin and plated at the same concentration in 6 well plates. At the indicated time points, cells were detached by trypsin treatment and cell number determined using a particle counter. There was no significant difference in proliferation between WT and KO preadipocytes. n=4.
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Figure 20: CaMKK2 works through AMPK to inhibit adipogenesis
A. WT, CaMKIV null and CaMKK2‐null MEFs were induced to differentiate. Loss of CaMKIV did not alter adipogenesis in MEFs as determined by Oil Red O staining. B. 3T3L1 preadipocytes were induced to differentiate in the presence of vehicle or 5μM KN‐93; adipogenesis was not affected as determined by Oil Red O staining. C. 3T3L1 preadipocytes infected with CaMKK2 shRNA (3145) were induced to differentiate in the presence of 0, 0.25 or 0.5mM AICAR. Adipogenesis was inhibited by AICAR as determined by Oil Red O staining. *p<0.05 as compared to 0mM AICAR. D. CaMKK2‐null MEFs (KO) were induced to differentiate in the presence of 0, 0.25 or 0.5mM AICAR. Adipogenesis was inhibited by AICAR as determined by Oil Red O staining. *p<0.05 as compared to 0mM AICAR. Quantification of Oil Red O is presented as mean +/‐ SEM. Data shown are representative of 3 independent experiments; and each was done in triplicate.
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phosphorylation was markedly decreased in the absence of CaMKK2 (Fig. 21A). In WT
MEFs, ionomycin, which increases intracellular Ca2+ leading to CaMKK2 activation, up‐
regulated AMPK phosphorylation, and its effect was blocked by preincubation of cells
with the CaMKK2 inhibitor, STO‐609; On the other hand, not only was basal AMPK
phosphorylation reduced in CaMKK2‐null MEFs, but it failed to respond to ionomycin
or STO‐609 (Fig. 21B). Additionally, deletion of CaMKK2 from 3T3L1 preadipocytes by
shRNA resulted in a large decrease in basal AMPK phosphorylation (Fig. 21C). As found
with WT MEFs, treatment of normal 3T3L1 preadipocytes with ionomycin increased
AMPK phosphorylation and was blocked by STO‐609 (Fig. 21D). Ionomycin and STO‐
609 had similar effects in normal C3H10T1/2 preadipocytes to those in MEFs (Fig. 21E).
We conclude from these data that CaMKK2 functions as an AMPK kinase in several
types of preadipocytes.
Loss of CaMKK2 activity increases mRNA of adipogenic transcription factors and markers
The temporal sequence of transcriptional events that lead to adipogenesis is well known.
In response to agents that stimulate adipogenesis a sequence of events occur as
summarized in Fig. 24D. Early in the process C/EBPβ and C/EBPδ are transcriptionally
induced and, in turn, these transcriptional activators trigger transcription from the
PPARγ and C/EBPα genes. PPARγ and C/EBPα then induce transcription of genes that
are required in adipocytes such as aP2, Fas and Glut4. In order to determine whether
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Figure 21: CaMKK2 phosphorylates AMPK in preadipocytes
A. Basal AMPK phosphorylation is down‐regulated in CaMKK2‐null preadipocytes. Protein extract from WT and CaMKK2‐null primary cultured preadipocytes were immunoblotted for T172 phosphorylated AMPK (pAMPK) and total AMPK. *p<0.05 compared to WT. B. WT and CaMKK2‐null MEFs were incubated for 1hr with vehicle or 2μM STO‐609, followed by stimulation with 1μM ionomycin for 5min. Cell extracts were immunoblotted for pAMPK and total AMPK. Ionomycin increased AMPK phosphorylation and was blocked by STO‐609 in WT but not CaMKK2‐null MEFs. *p<0.05 compared to control WT MEF. C. CaMKK2 was knocked down by shRNA in 3T3L1 preadipocytes. Basal AMPK phosphorylation was down‐regulated in CaMKK2‐null cells. *p<0.05 compared to control 3T3L1 preadipocytes. D and E. Ionomycin increased AMPK phosphorylation and was blocked by STO‐609 in 3T3L1 or C3H10T1/2 preadipocytes. *p<0.05 compared to control. For A‐E, one representative blot and quantification of 3 independent experiments are shown.
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CaMKK2 acts at any of these steps that lead from preadipocyte to adipocyte, we utilized
real‐time PCR to quantify the various adipogenic gene transcripts shown in Fig. 24D in
wild type and CaMKK2‐null MEFs after exposure to adipogenic stimulation. We found
that adipocytes derived from CaMKK2‐null MEFs express much more mRNA encoding
all 7 adipogenic genes than adipocytes derived from WT MEFs (Fig. 22A). This was also
true for 3T3L1 adipocytes derived from cells treated with CaMKK2 shRNA or cultured
in STO‐609 relative to those cells subjected to a nonsense shRNA or cultured in the
absence of the inhibitor, respectively (Fig. 22B and 22C). The presence of STO‐609 during
adipogenic stimulation of C3H10T1/2 cells also resulted in enhanced expression of all 7
adipogenic mRNAs examined (Fig. 22D). These results suggest that CaMKK2 must
function very early in adipogenesis or in a pathway that inhibits adipogenesis in
preadipocytes.
CaMKK2 and AMPK maintain Pref-1 signaling
Pref‐1 is expressed in preadipocytes where it inhibits adipogenesis by regulating a
signaling pathway that leads to the phosphorylation and activation of ERK and
subsequently to the transcriptional upregulation of Sox9, which functions to
transcriptionally repress the C/EBPβ and C/EBPδ genes (Smas and Sul, 1996; Wang et al.,
2006; Kim et al., 2007; Wang and Sul, 2009). To examine if CaMKK2 might function in
the Pref‐1 signaling pathway we quantified Pref‐1 and Sox9 mRNAs in primary
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Figure 22: Loss of CaMKK2 activity increases adipogenic gene transcripts during adipogenesis
A. mRNA levels of C/EBPβ, C/EBPδ, PPARγ, C/EBPα, aP2, Fas and Glut4 are increased in CaMKK2‐null MEF adipocytes compared to WT. Total RNA was isolated from WT or CaMKK2‐null MEFs after adipogenesis. mRNA levels of adipogenic genes were determined by RT‐PCR. B. 3T3L1 preadipocytes infected with control or CaMKK2 shRNA were induced to differentiate and total RNA was isolated after adipogenesis. RT‐PCR indicated mRNA levels of PPARγ, C/EBPα, aP2, Fas and Glut4 are increased in 3T3L1 adipocytes with CaMKK2 knocked down. C and D. mRNA levels of C/EBPβ, C/EBPδ, PPARγ, C/EBPα, aP2, Fas and Glut4 are increased in 3T3L1 or C3H10T1/2 adipocytes treated with 2μM STO‐609 during differentiation. For A‐D, data are presented as mean +/‐ SEM; *p<0.05. Data shown are representative of 3 independent experiments; and each was done in triplicate.
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preadipocytes and MEFs from WT and CaMKK2‐null mice. These data revealed that the
absence of CaMKK2 resulted in considerably reduced amounts of Pref‐1 and Sox9
mRNAs in both cell types (Fig. 23A and 23B). We also used immunoblot analysis to
quantify the phosphorylation of ERK1/2 in MEFs prepared from WT mice compared to
MEFs isolated from mice heterozygous (Het) or null (KO) for CaMKK2. As shown in Fig.
23C, ERK phosphorylation was decreased in both Het and KO cells relative to WT. To
confirm that CaMKK2 plays a role in regulating the Pref‐1 pathway, we cultured WT
MEFs in the presence of STO‐609 and found that the CaMKK2 inhibitor also reduced the
amount of Pref‐1 and Sox9 mRNAs (Fig. 23D). To determine if reduction of Pref‐1 and
Sox9 resulted from acute depletion of CaMKK2, we examined the effect of cre‐mediated
deletion of CaMKK2 in MEFs isolated from CaMKK2‐loxP mice and found indeed that
both mRNAs were reduced (Fig. 23E). Finally, shRNA‐mediated depletion of CaMKK2
from 3T3L1 cells also markedly reduced Pref‐1 and Sox9 mRNAs (Fig. 23F). Therefore,
CaMKK2 is important for maintaining the level of Pref‐1 and consequently the Pref‐1
signaling pathway in multiple preadipocyte cell types.
We have presented evidence that AMPK serves as the primary CaMKK2
substrate to regulate the pathway by which CaMKK2 inhibits adipogenesis (Fig. 21). It
has been reported that AICAR, a compound which can activate AMPK, also reduces
adipogenic gene transcripts and restores Pref‐1 mRNA during the differentiation of
3T3L1 cells (Giri et al., 2006). Thus, we questioned whether AMPK was involved in
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Figure 23: CaMKK2 maintains Pref‐1 mRNA
A and B. Pref‐1 and Sox9 mRNA levels are decreased in CaMKK2‐null primary preadipocytes and MEFs. Total RNA was isolated from WT and CaMKK2‐null (KO) primary preadipocytes or MEFs; Pref‐1 and Sox9 mRNA levels were quantified by RT‐PCR. C. Protein extracts from WT, CaMKK2‐/+ and CaMKK2‐null MEFs were immunoblotted for phosphorylated ERK (pERK) and total ERK (left). Phosphorylation of ERK is reduced in CaMKK2‐null MEFs as shown in quantification of blot (right). Each lane represents MEFs derived from an individual embryo. D. Total RNA was isolated from WT MEFs incubated with vehicle or 2μM STO‐609 for 2hr. Pref‐1 and Sox9 mRNA levels were determined by RT‐PCR. E. MEFs from CaMKK2 loxP mice were infected with Ad‐CMV‐GFP or Ad‐Cre‐IRES‐GFP viruses. Cells were harvested 6 days after infection. Protein extract was immunoblotted for CaMKK2 and pAMPK (left). Ad‐Cre‐IRES‐GFP virus effectively depleted CaMKK2 and decreased AMPK phosphorylation. Pref‐1 and Sox9 mRNA levels were reduced as determined by RT‐PCR (right). F. Total RNA was extracted from 3T3L1 preadipocytes infected with control or CaMKK2 shRNA. Pref‐1 and Sox9 mRNA levels were decreased in shRNA infected cells compared to control, as determined by RT‐PCR. Data are presented as mean +/‐ SEM. Data shown in A, B, D, E and F are representative of at least 2 independent experiments, and each was done in triplicate.
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regulating Pref‐1 in MEFs. Knocking down the AMPKα subunits by siRNA decreased
Pref‐1 and Sox9 mRNA in WT MEFs (Fig. 24A). Furthermore, Compound C, an inhibitor
of AMPK, reduced Pref‐1 and Sox9 mRNA in WT MEFs (Fig. 24B), but treatment with
KN‐93, a compound that inhibits both CaMKI and CaMKIV did not (data not shown).
Finally, AICAR, an activator of AMPK, increased Pref‐1 and Sox9 mRNA in CaMKK2‐
null MEFs (Fig. 24C). These data reveal that CaMKK2 functions in preadipocytes to
activate AMPK which results in up‐regulation of Pref‐1 mRNA leading to maintenance
of the Pref‐1 signaling pathway and therefore inhibition of adipogenesis (Fig. 24D).
5.3 Discussion
CaMKK2‐null mice have been utilized to demonstrate a number of roles for this protein
kinase in the brain (Anderson et al., 2008; Kokubo et al., 2009; Thornton et al., 2011;
Antunes‐Martins et al., 2007). However, these mice have not been employed to identify
cell autonomous effects of CaMKK2 in peripheral cells. Here we show CaMKK2 to be
expressed in preadipocytes and that germ line depletion of this gene results in increased
mass of WAT coupled with an elevated adipocyte numbers and a decreased number of
preadipocytes that can be isolated from WAT (Fig. 25) when mice are fed standard lab
chow (5001). When exposed to adipogenic stimuli primary preadipocytes (in vivo) or
MEFs (in culture) from CaMKK2‐null mice differentiate into adipocytes to a greater
extent than do cells isolated from WT mice, and the increase in molecular markers of the
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Figure 24: AMPK maintains Pref‐1 mRNA
A. WT MEFs were transfected with control or AMPKα siRNA. Protein and total RNA were extracted 4 days after transfection. Both phosphorylated AMPK and total AMPK were effectively decreased, as determined by immunoblot (left). Pref‐1 and Sox9 mRNA levels were decreased, as determined by RT‐PCR (right). *p<0.05. Data shown are from one experiment that is representative of 3 independent experiments which were each done in triplicate. B. Total RNA was extracted from 3T3L1 preadipocytes incubated with vehicle or 2μM Compound C (Calbiochem) for 1hr. Pref‐1 and Sox9 mRNA levels were decreased after Compound C treatment as determined by RT‐PCR. *p<0.05. Data shown are from one experiment that is representative of 3 independent experiments which were each done in triplicate. C. Total RNA was extracted from CaMKK2‐null MEFs treated with vehicle or 0.25mM AICAR for 1hr. Pref‐1 and Sox9 mRNA levels were increased in AICAR treated MEFs as determined by RT‐PCR. *p<0.05, n=6. Data shown are from one experiment that is representative of 2 independent experiments. D. CaMKK2 inhibits adipogenesis by maintaining Pref‐1 mRNA via AMPK. In response to increased intracellular Ca2+ concentration, CaMKK2 phosphorylates and activates AMPK. CaMKK2 and AMPK maintain mRNA level of Pref‐1 through an unknown mechanism. Pref‐1 stimulates phosphorylation and activation of ERK1/2, which maintains expression of Sox9. Sox9 inhibits transcription of the early adipogenic transcription factors C/EBPβ and C/EBPδ, which reduces expression of PPARγ and C/EBPα and, in turn, adipocyte‐specific genes such as aP2, Glut4 and Fas.
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mature fat cell is inversely correlated with the disappearance of CaMKK2. That this is
cell autonomous rather than mediated via the nervous system or developmentally is
revealed by the fact that virtually identical changes result when CaMKK2 activity is
acutely depressed by pharmacological inhibition or shRNA‐mediated knock‐down in
3T3L1 and CH310T1/2 cells. We demonstrate that the inhibitory effect of CaMKK2 on
adipogenesis is due to a role of CaMKK2 and its substrate AMPK in maintaining the
level of Pref‐1 mRNA. The Pref‐1 protein functions via the activation of MAP kinase to
enhance the amount of SOX9 which, in turn, acts as a transcriptional repressor to
prevent the expression of C/EBPβ and C/EBPδ, two early genes that positively regulate
the differentiation of preadipocytes to adipocytes (Smas and Sul, 1996; Wang et al., 2006;
Kim et al., 2007; Wang and Sul, 2009). Thus, in response to initiating a series of events
that lead to adipocyte formation and fat production by these cells, adipogenic
stimulation also results, by a yet to be discovered mechanism, in down‐regulation of
CaMKK2 mRNA and protein. In this context, CaMKK2 and AMPK participate in a
signaling pathway to maintain the preadipocyte rather than to stimulate the production
of mature adipocytes from these progenitor cells (Fig. 24D).
We have previously reported that CaMKK2‐null mice accumulate less body
weight when fed a high fat diet which is due to reduced expansion of WAT (Anderson
et al., 2008). As shown in Fig. 14 of chapter 4, the average size of mature adipocytes is
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Figure 25: CaMKK2‐null mice have fewer preadipocytes and more adipocytes in gonadal WAT.
A. Gonadal fat pads of equal weight were isolated from WT or CaMKK2‐null mice and digested with collagenase. Adipocytes were allowed to float by gravity. After centrifugation, pelleted preadipocytes were resuspended and allowed to attach and proliferate for 4 days. Red blood cells and other debris were removed by refreshing media. Compared to WT, CaMKK2‐null mice yielded fewer preadipocytes. B. Preadipocytes shown in A were detached by trypsin and quantified using a particle counter. n=4, *p<0.05. C. Adipocyte number in gonadal fat pads was analyzed as described in method. CaMKK2‐null mice have more adipocytes compared to WT mice. Shown are mean+/‐SEM. n=5, *p<0.01.
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greater in WAT isolated from CaMKK2‐null mice than it is in WT mice fed standard lab
chow (5001). In the study by Anderson et al (Anderson et al., 2008) the pregnant females
were also maintained on standard lab chow and remained on this diet while nursing
their pups. At the time of weaning the pups were placed on solid food for the first time
and fed either a high‐fat diet or its “control” low‐fat diet for 30 weeks. We carried out
histology of the WAT and found that the size of adipocytes of WT mice increased
markedly between standard chow and low‐fat diets and increased again, in a
statistically significant manner, between WT mice fed low‐fat and high fat diet (Fig. 14).
However, although adipocyte size was greater in WAT from CaMKK2‐null mice than
WT mice fed standard chow, there was no increase in adipocyte size in WAT from the
CaMKK2‐null mice between standard chow and low‐fat diets and only a very small
increase in size between low‐fat and high fat diet. These data are consistent with our
data showing depletion of preadipocytes in CaMKK2‐null mice fed standard chow and
suggest that preadipocyte depletion is likely an important factor contributing to the
resistance to diet‐induced obesity exhibited by the CaMKK2‐null mice. It is possible that
diet‐induced fat accumulation is also reduced in CaMKK2‐null mature adipocytes but, if
that occurs, it is due to a different and yet to be explored mechanism.
It came as a considerable surprise to us that preadipocytes were depleted in mice
with a germ line deletion of the CaMKK2 gene as CaMKK2 had not previously been
shown to be expressed in these cells. Our novel data reveal that AMPK serves as the
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primary CaMKK2 substrate in the signaling pathway by which CaMKK2 delays (or
prevents) adipogenesis and that a relevant consequence of this pathway is regulation of
Pref‐1 mRNA. Intriguingly, forced expression of Pref‐1 in 3T3L1 cells makes them
resistant to differentiation (Smas and Sul, 1996; Kim et al., 2007) and reduced expression
of Pref‐1 is required for differentiation of these cells into adipocytes. The same is true for
forced expression or inhibition of CaMKK2 or AMPK in 3T3L1 cells. This sequence of
events suggests that the down‐regulation of CaMKK2 in response to adipogenic stimuli
may be an even earlier event than a reduction in Pref‐1 and be required to down‐
regulate Pref‐1 mRNA. Regulation of Pref‐1 transcription is poorly understood but
studies have reported that HIF1α, HIF2α and Foxa‐2 promote Pref‐1 transcription
whereas Smad1/4 and Hes‐1 inhibit it (Kim et al., 2009; Ross et al., 2004; Wolfrum et al.,
2003; Zhang et al., 2010). This information may help to inform future efforts designed to
identify the primary AMPK target in the CaMKK2‐mediated pathway that inhibits
adipogenesis. Elucidating the relevant stimuli and resultant signaling events both
upstream and downstream of CaMKK2/AMPK may lead to a more complete
understanding of how adipogenesis is gated physiologically. It would also support
CaMKK2 as a novel and potentially attractive target to develop therapeutics to combat
diet‐induced obesity.
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6. Future Directions
CaMKK2‐AMPK pathway in hypothalamus
As we already showed in chapter 3, loss of CaMKK2 in the hypothalamus results in
decreased AMPK activity, which leads to reduced NPY and AgRP production. We
hypothesized that CaMKK2 functioned downstream of ghrelin and provided evidence
that CaMKK2‐null mice did not respond to i.p. injection of ghrelin. However, more
experiments need to be done to prove the relationship between ghrelin and CaMKK2. If
CaMKK2 indeed is activated by increased intracellular Ca2+ triggered by GHSR,
CaMKK2 in WT hypothalamus is expected to elevate AMPK phosphorylation and
activity after ghrelin i.c.v. administration. And this activation should be inhibited by
pre‐administration of CaMKK2 selective inhibitor STO‐609. A similar result would be
expected in CaMKK2‐null mice; that is, i.c.v. injection of ghrelin in mutant mice should
fail to activate AMPK and downstream NPY and AgRP expression. N38 cells or other
neuronal cells derived from hypothalamus with GHSR would be good in vitro models.
Ghrelin treatment of these cells should result in increased AMPK activity and elevated
NPY and AgRP expression. STO‐609 and/or small interfering RNA against CaMKK2 is
expected to blunt the effect of ghrelin.
On the other hand, it would be interesting to know whether decreased
hypothalamic AMPK activity in CaMKK2‐null mice enhances leptin activity. It has been
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shown that leptin inhibits AMPK activity in hypothalamus and down‐regulation of
AMPK activity is necessary for its anorexigenic effect because expression of a
constitutively active form of AMPK significantly blocks leptin function. Therefore, I
would like to compare the responses to leptin i.c.v. administration between WT and
CaMKK2‐null mice. For example, the food intake can be measured in WT and mutant
mice after leptin injection. Besides, mTOR has been shown to be an important regulator
downstream of leptin (Cota et al., 2006). Activation of mTOR signaling decreases food
intake and body weight, and inhibition of mTOR signaling blocks leptin’s effect. Inoki
and colleagues have shown that AMPK can phosphorylate TSC2 and enhance its activity,
which leads to the inhibition of mTOR (Inoki et al., 2003). As predicted from this report,
loss of CaMKK2 will impair the phosphorylation of TSC2 by AMPK and mTOR activity
would not be inhibited by TSC2, which will result in decreased food intake. In order to
test whether activation of mTOR signaling contributes to decreased food intake in
CaMKK2‐null mice, phosphorylation of TSC2, S6K and S6 would be examined in fasted
WT and mutant mice. CaMKK2‐null mice are expected to have decreased TSC2
phosphorylation but increased S6K and S6 phosphorylation. If so, rapamycin, the
inhibitor of mTOR can be i.c.v. administered to WT mice along with STO‐609, and we
should expect rapamycin can reverse the anorexigenic effect of STO‐609.
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High‐fat diet feeding
We also observed an intriguing phenotype when mice were fed a high‐fat diet. CaMKK2‐
null mice consumed less food, accumulated less adiposity and maintained glucose
tolerance and insulin sensitivity compared to WT control animals. Decreased feeding
partially explained the resistance to high‐fat diet. But as shown in chapter 4, it is not the
only reason benefitting CaMKK2‐null mice. Although CaMKK2 was shown to mainly
exist in brain, its expression was gradually discovered in some peripheral tissues. In this
study, all the CaMKK2‐null mice used were global knockout mice. It is really hard to rule
out the possible contributions from peripheral tissues. For example, we know that before
high‐fat diet feeding, mutant mice have higher serum leptin which may contribute to
negative regulation of energy balance. Also our lab has shown that islets isolated from
knockout mice secret more insulin when stimulated with high concentration of glucose.
Later on, members of our lab confirmed that CaMKK2 is expressed in β cells, which
implicates CaMKK2 in pancreas function. In order to understand more specifically the
function of CaMKK2 in NPY/AgRP neurons, neuron specific knockout mice should be
generated. Actually, this project is currently underway in our lab. We are trying to cross
CaMKK2 loxP mice with mice possessing CRE gene controlled by the promoter of AgRP,
which is mainly expressed in NPY/AgRP neurons. With this type of knockout mouse,
we would be able to perform the high‐fat diet feeding experiment again and determine
the specific contribution of CaMKK2 in NPY neurons to the resistance to high‐fat diet. In
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case the neuron‐specific knockout mice have developed similar compensation to that of
global knockout mice, we could also bypass this issue by performing tamoxifen i.c.v.
administration in CaMKK2 loxP mice.
In the high‐fat diet pair feeding experiment, CaMKK2‐null mice consumed
similar amounts of food and water as WT animal controls but they still gained less body
weight and adiposity. By comparing the heat production, we found mutant mice even
had lower energy consumption. Therefore, CaMKK2‐null mice had same energy intake
but lower energy storage and expenditure. We may wonder where the extra energy goes.
The primary test would be collecting urine and feces of WT and CaMKK2‐null mice
during high‐fat diet pair feeding and analyzing their nutrient composition. This result
should tell us whether mutant mice could not absorb nutrients as well as WT controls or
if they just lost nutrients in urine.
CaMKK2‐AMPK pathway in adipogenesis
Increased intracellular Ca2+ concentration has been shown to inhibit adipogenesis by
blocking DNA synthesis and clonal expansion (Ntambi and Takova, 1996). Activation of
AMPK by AICAR during adipogenesis also inhibited clonal expansion (Habinowskit
and Witters, 2001; Giri et al., 2006). Although CaMKK2‐null MEFs and preadipocytes
displayed no defect in growth, more adipocytes were observed during adipogenesis
when CaMKK2 activity was depressed. Therefore, it is reasonable to speculate that loss
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of CaMKK2 activity may also affect clonal expansion, which could be an additional way
in which CaMKK2 controls adipogenesis. In order to examine this, WT and CaMKK2‐
null MEFs would be induced to differentiate in MDI media for 2 day. After 2 days, DNA
content and cell number would be determined. Besides, DNA replication can be
measured by the rate of 3H incorporation. If clonal expansion is inhibited by CaMKK2 as
predicted, CaMKK2‐null MEFs are expected to have more DNA content and cells as well
as higher 3H incorporation rate during two days’ differentiation. STO‐609 and shRNA
against CaMKK2 would also be used to confirm the effect is indeed a result of loss of
CaMKK2 activity. In order to know whether CaMKK2 works via AMPK, I would check
whether AICAR treatment inhibits clonal expansion in CaMKK2‐null MEFs.
As I presented in chapter 5, AMPK mediates the inhibitory effect of CaMKK2 on
adipogenesis. However, the direct downstream target of AMPK was not identified
during this study. As we mentioned before, AMPK can phosphorylate and activate TSC2
so as to inhibit mTOR activity. The important role of mTOR in adipogenesis has been
well established by using its inhibitor, rapamycin. It has been demonstrated that
rapamycin treatment strongly inhibits preadipocyte differentiation and thus
maintenance of adipocytes (Kim and Chen, 2004). Therefore, decreased AMPK activity
caused by depletion or inhibition of CaMKK2 in preadipocytes may result in increased
mTOR activity, which is required for further adipogenesis. However, the relationship
between AMPK and mTOR has not been studied in preadipocytes. In order to confirm
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this interaction, I would examine the mTOR signaling in both WT and CaMKK2‐null
MEFs. Phosphorylation of S6K and TSC2 would be immunoblotted and increased
phosphorylation on S6K as well as decreased phosphorylation on TSC2 would be
expected in CaMKK2‐null MEFs. Then I would inhibit or knock down CaMKK2 by either
STO‐609 or shRNA in 3T3‐L1 preadipocytes and check S6K or S6 phosphorylation.
Overexpression of CaMKK2 in preadipocytes is expected to inhibit mTOR activity. If
mTOR is indeed regulated by CaMKK2‐AMPK pathway in preadipocytes, I would
knock down TSC2 in 3T3‐L1 cells and try to differentiate these cells with or without
STO‐609. The ability of STO‐609 to enhance adipogenesis should be blocked or greatly
inhibited in TSC2 knockout cells because STO‐609 would not be able to increase mTOR
activity in these cells. At the same time, we would expect that AICAR could not inhibit
adipogenesis in these cells either.
In this study, I also demonstrated that the CaMKK2‐AMPK pathway functions to
maintain Pref‐1 mRNA level in preadipocytes. The link between AMPK and Pref‐1 is
missing. The regulation of Pref‐1 transcription is not clearly defined. So far, people have
suggested that HIF1α, HIF2α and Foxa2 activate its transcription, whereas Smad1/4 and
Hes1 inhibit it. Among these transcription factors, HIF1α is considered the most likely
candidate downstream of AMPK. Under normoxia, HIF1α is rapidly degraded by
ubiquitination and proteasome pathways. Under hypoxia or treatment of deferoxamine,
a hypoxia‐mimicking compound, HIF1α is stabilized and enhances Pref‐1 transcription
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(Kim et al., 2009). It has been reported that AMPK activity is important for the
expression, stabilization and transcriptional activity of HIF1α (Hwang et al., 2004; Jeong
et al., 2011; Lee et al., 2003). Therefore, I would check whether HIF1α can stimulate Pref‐
1 expression in preadipocytes. WT MEFs or 3T3‐L1 preadipocytes would be treated with
normoxia, hypoxia (1%O2) or deferoxamine. HIF1α level would be determined by
immunoblot and Pref‐1 mRNA would be examined by RT‐PCR. One well‐known HIF1α
target gene, vascular endothelial growth factor, could be used as a positive control. If
HIF1α indeed enhances Pref‐1 transcription in preadipocytes, the amount of HIF1α
protein in WT and CaMKK2‐null MEFs would be compared. It will be also interesting to
know whether AICAR treatment in CaMKK2‐null MEFs elevates HIF1α protein level.
Also I would check whether overexpression of HIF1α in CaMKK2‐null MEFs rescues
Pref‐1 mRNA level and inhibits adipogenesis.
CaMKK2‐AMPK pathway in multipotent mesenchymal cells
As I showed in chapter 5, the CaMKK2‐AMPK pathway maintains Pref‐1 and Sox9
mRNA levels in preadipocytes. Pref‐1 and Sox9 are also important in determining the
fate of multipotent mesenchymal cells (Wang and Sul, 2009). Pref‐1 can direct
mesenchymal cells into chondrogenic induction; however Pref‐1 prevents maturation of
chondrocyte, adipogenesis and osteogenesis. Since CaMKK2‐null MEFs have been
shown to have less Pref‐1 and Sox9 mRNA, it is reasonable to predict that osteogenesis
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will be enhanced in CaMKK2‐null MEFs. In order to examine the role of CaMKK2 in
osteogenesis, the simplest experiment is to induce osteogenesis in WT and mutant MEFs
by adding β‐glycerophosphate, ascorbic acid and BMP2 for 21 days. After the induction,
osteogenesis would be evaluated by Von Kossa staining and alkaline phosphatase
(ALPase) activity assay in both types of MEFs. I would expect CaMKK2‐null MEFs to
display stronger Von Kossa staining and ALPase activity. Also the mRNA level of
osteogenesis markers such as ALPase, osteocalcin and Runx2 would be expected to be
higher in CaMKK2‐null MEFs. In addition, the expression of CaMKK2 before and after
osteogenesis would be compared in WT MEFs. The effect of CaMKK2 could be
confirmed by inducing C3H10T1/2 cells into osteogenesis with or without acute
inhibition of CaMKK2 activity. I expect that osteogenesis would be enhanced in
C3H10T1/2 cells when CaMKK2 is inhibited by STO‐609 or deleted by shRNA.
I have shown that fewer preadipocytes could be isolated from the gonadal fat
pads of CaMKK2‐null mice fed standard chow and at the same time they have more
adipocytes. I hypothesized that depletion of preadipocytes from knockout mice by
enhanced adipogenesis contributes to the smaller increase on adiposity when they are
switched to high‐fat diet. The first step in confirming this hypothesis is to examine the
adipocyte numbers from both WT and CaMKK2‐null mice after high‐fat diet feeding. I
would expect that more adipocytes will be isolated from WT mice than from CaMKK2‐
null mice after high‐fat diet. Usually, people think new adipocytes come from resident
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preadipocytes in WAT; however, recent evidence suggests that progenitor cells from
other tissues, especially bone marrow, also contribute to the hyperplasia of adipocytes
(Crossno et al., 2006). As we know, mesenchymal stem cells (MSCs) from bone marrow
could be differentiated into adipocytes, chondrocytes and osteocytes (Tondreau et al.,
2004). It makes me wonder whether the maintenance and differentiation of MSCs are
also changed in CaMKK2‐null mice. Actually, members of our lab have observed that
CaMKK2‐null mice have more bone mass and increased fat content in isolated bone
marrow. These facts are consistent with the hypothesis that CaMKK2‐null MSCs may
have enhanced adipogenesis and osteogenesis because of deficient Pref‐1/Sox9 signaling.
Therefore, I would like to isolate MSCs from WT and CaMKK2‐null mice by specific
marker SH2, SH3 or Stro‐1 and confirm the expression of CaMKK2. The number of
MSCs isolated from both types of mice would also be compared. It would be expected
that fewer MSCs could be recovered from knockout mice. The potential of MSCs to
differentiate into osteocytes and adipocytes would be compared between WT and
CaMKK2‐null mice. In order to test the importance of MSCs in adiposity increase during
high‐fat diet feeding, whole bone marrow transplantation would be carried out. WT
mice receiving WT bone marrow would still be expected to be the most obese after high‐
fat diet treatment. WT mice with CaMKK2‐null bone marrow should gain less adiposity
and fewer adipocytes. In contrast, the CaMKK2‐null mice receiving WT bone marrow
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would gain more adiposity and more adipocytes compared to the knockout mice with
CaMKK2‐null bone marrow.
120
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Biography
Fumin Lin was born in Fuzhou, Fujian Province of China, on December 8, 1982,
and grew up in the same city. In 2001, Fumin graduated from Fuzhou No.3 high school,
and attended Tsinghua Univeristy in Beijing, China. In 2005, he earned a Bachelor of
Science degree in the major of biological science and sought his graduate study aboard.
In the fall of the same year, Fumin was admitted by Cell and Molecular Biology Program
in Duke University. After one year’s rotation, he joined laboratory of Dr. Anthony
Means in the Department of Pharmacology to pursue his Ph.D. degree. Fumin and his
wife, Daoyi Lin, who married him in March, 2009, live in Durham, NC now. Fumin is
finishing his first first‐author paper. During his time in Duke, Fumin contributed to two
papers and attended two annual Endocrine Society meetings, at which he gave poster
presentations. Upon completing his graduate school degree he plans to receive a
postdoctoral training.