Copper handling machinery of the brain.

Abstract

Copper plays an indispensable role in the physiology of the human central nervous system (CNS). As a cofactor of dopamine-β-hydroxylase, peptidyl-α-monooxygenase, superoxide dismutases, and many other enzymes, copper is a critical contributor to catecholamine biosynthesis, activation of neuropeptides and hormones, protection against reactive oxygen species, respiration and other processes essential for normal CNS function. Copper content in the CNS is tightly regulated, and changes in copper levels in the brain are associated with a wide spectrum of pathologies. However, the mechanistic understanding of copper transport in the CNS is still in its infancy. Little is known about copper distribution among various cell types or cell-specific regulation of copper homeostasis, despite the fact that the molecules mediating copper transport and distribution in the brain (CTR1, Atox1, CCS, ScoI/II, ATP7A and ATP7B) have been identified and their importance in CNS function increasingly understood. In this review, we summarize current knowledge about copper levels and uses in the CNS and describe the molecules involved in maintaining copper homeostasis in the brain.

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596Metallomics, 2010, 2, 596–608This journal is c The Royal Society of Chemistry 2010
Copper handling machinery of the brain
Svetlana Lutsenko,*aAshima Bhattacharjeeaand Ann L. Hubbardb
Received 27th April 2010, Accepted 5th July 2010
DOI: 10.1039/c0mt00006j
Copper plays an indispensable role in the physiology of the human central nervous system (CNS).
As a cofactor of dopamine-b-hydroxylase, peptidyl-a-monooxygenase, superoxide dismutases, and
many other enzymes, copper is a critical contributor to catecholamine biosynthesis, activation of
neuropeptides and hormones, protection against reactive oxygen species, respiration and other
processes essential for normal CNS function. Copper content in the CNS is tightly regulated, and
changes in copper levels in the brain are associated with a wide spectrum of pathologies.
However, the mechanistic understanding of copper transport in the CNS is still in its infancy.
Little is known about copper distribution among various cell types or cell-specific regulation of
copper homeostasis, despite the fact that the molecules mediating copper transport and
distribution in the brain (CTR1, Atox1, CCS, ScoI/II, ATP7A and ATP7B) have been identified
and their importance in CNS function increasingly understood. In this review, we summarize
current knowledge about copper levels and uses in the CNS and describe the molecules involved
in maintaining copper homeostasis in the brain.
Copper content in the central nervous system
The human brain requires copper for its normal development
and function. Compared to other organs in the body, copper
concentration in the brain is one of the highest, second only to
the liver,1which is the major organ for copper uptake and
excretion. The estimated copper content of human brain
ranges from 2.9 to 10.7 mg Cu/g wet weight, while rat brains
appear to have a lower copper content-1–2.3 mg Cu/g wet
weight.2,3Fractionation of brain homogenates demonstrates
that copper is present in all major cell compartments, with
nuclei and mitochondria containing 1.12–1.3 mg Cu/g, micro-
somes-0.49–0.65 mg Cu/g, and the cytosol-2.56 ? 1.02 mg
Cu/g.4Disease states alter copper concentrations in the
CNS. Copper elevation (in the range of 7.8–37.8 mg/l) was
observed in patients with neurological symptoms related to
dementia;5in Alzheimer’s-like dementia CNS copper was
reported to be 2-fold higher than in age matched controls.6,7
Copper is not uniformly distributed in the CNS. Experiments
in adult rats (6–12 months old) using laser ablation inductively
coupled plasma mass spectrometry (LA-ICP-MS) have yielded
two-dimensional maps of copper distribution in brain sections.8
These maps show63Cu enrichment in the medial geniculate
nucleus (the center processing visual information), superior
colliculus (a component of the midbrain associated with motor
functions) and periaqueductal grey (the gray matter located
around the cerebral aqueduct of the midbrain that modulates
pain and defensive behavior). Copper is also high in the lateral
amygdala (groups of nuclei involved in memory and emotional
reactions) and in the dorsomedial aspect of the diencephalon
(the part of the forebrain containing thalamus, hypothalamus
and the posterior portion of the pituitary gland). It is notable
that copper distribution within the CNS varies between species.
For example, copper levels in rat hippocampus are moderate.
In contrast, the human hippocampus has the highest copper
aDepartment of Physiology, Johns Hopkins University,
725N Wolfe St., Baltimore, MD 21205, USA.
E-mail: lutsenko@jhmi.edu; Tel: +1 410-614-4661
bDepartment of Cell Biology, Johns Hopkins University,
725N Wolfe St., Baltimore, MD 21205, USA
Svetlana Lutsenko
Dr Svetlana Lutsenko is a
faculty member
Department of Physiology at
the Johns Hopkins Medical
School in Baltimore, USA.
Her laboratory is working on
dissecting the mechanisms of
ATP driven ion transport,
regulation of human copper
homeostasis,
developing tractable models
for human disorders of copper
metabolism.
inthe
aswellas
Ashima Bhattacharjee
Dr Ashima Bhattacharjee is a
postdoctoralfellow
Department of Physiology at
the Johns Hopkins Medical
School in Baltimore, USA.
Sheisinterested
pathophysiology
genetics disorders. Her current
research focuses on determining
the role of mutations and
polymorphisms in the Wilson’s
diseasegene
function, and the intracellular
behavior of copper-transporting
ATPase ATP7B.
inthe
in
human
the
of
onstability,
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This journal is c The Royal Society of Chemistry 2010Metallomics, 2010, 2, 596–608597
concentration (up to 15 mg g?1) compared to other regions.9
Since human brains are considerably larger than rodent
brains, the LA-ICP-MS methodology has only been applied
to certain regions, such as the insular, central and hippo-
campal areas.9,10Analysis of these areas further confirmed the
non-uniform distribution of copper. Thus, in the hippo-
campus, copper is highly enriched in the pyramidal and
lacunosum molecular layers of cornu ammonis.10
Higher copper concentrations in certain regions of the brain
most likely reflect higher metabolic demands for copper in
these regions. The metabolic demands for copper within the
CNS change during brain development. In humans and in rats,
copper concentration in the CNS increases with age.11,12
Detailed studies by Tarohda and colleagues12demonstrated
that the rat neonatal brain has low copper levels, which
rapidly increase during the first two weeks after birth
(Fig. 1). A particularly striking increase is observed between
the postnatal day 7 and 14, when the striatum, thalamus,
and superior colliculus show marked and preferential
accumulation of copper.12The proper copper delivery to the
brain during this developmental period can be literally a
matter of life and death. Severely copper-deficient mice, which
otherwise die, can be rescued by a single copper injection
administered before postnatal day 7.13–15
supplementation results in a significant restoration of
morphology and function of the CNS. The sufficiency of a
one-dose injection suggests that copper in the brain is largely
recyclable, and only a small additional amount of copper
is needed during an animal’s life. In humans, very early
treatment with copper-(histidine)2complex produces beneficial
effects in copper deficient Menkes disease patients and corrects
some neurological symptoms.16,17
This copper
Copper-requiring enzymes play essential roles in
brain development and function
In the CNS, copper is utilized for general cellular metabolism
such as respiration and radical defense, as well as for specia-
lized processes such as production of neuroendocrine peptides
and hormones. The mitochondria function and protection
against reactive oxygen species in the cytosol rely on the
activities of copper-dependent enzymes cytochrome C oxidase
(CCO) and Cu,Zn-dependent superoxide dismutase 1 (SOD1),
respectively. Superoxides at the cell surface are neutralized
by extracellularmembrane-bound
maintains cerebral vascular tone and regulates neurogenesis.18,19
Many peptide hormones and neurotransmitters such as
oxytocin, vasopressin, gastrin, corticotropin-releasing factor,
thyotropin-releasing hormone and others require the presence
of an a-amide moiety for their activity.20Amidation of neuro-
peptides and the production of norepinephrine is mediated
by twohomologous cuproenzymes:
monooxygenase (PAM) and dopamine-b-hydroxylase (DBH),
respectively. Another DBH-like monooxygenase MOXD1 is
expressed throughout the brain (olfactory bulb, pituitary,
cerebellum, parietal cortex), but its physiological role is
unclear.21
Recent studies indicate that several enzymes well known for
their roles in peripheral tissues are also present in the brain and
therefore contribute to normal brain function. For example, the
copper-requiring enzymes, tyrosinase and tyrosinase-related
protein, are best known as key players in melanin synthesis in
melanocytes. These two proteins have also been found in
developing and adult brain with particularly high expression
in the cortex, olfactory system, hippocampus, epithalamus and
substantia nigra.22–24The role of tyrosinase in the CNS has yet
to be fully understood; one suggestion is that it oxidizes excess
dopamine and thus prevents damage of catecholaminergic
neurons caused by dopamine auto-oxidation.25However,
increased susceptibility of neurons to dopamine in response
to tyrosinase overexpression has also been reported,24illustrating
the need for better understanding of the function of this
copper-dependent enzyme in the CNS. An interesting signaling
role for tyrosinase was proposed in studies on the development
of the visual system where tyrosinase expression was shown to
determine the pathway taken by ganglion cell axons.26
The lysyl oxidase (LOX) protein family, another class of
copper dependent enzymes with a well-characterized peripheral
function, is also found in the CNS. These proteins catalyze the
oxidation of the side chain of a lysine as the first step of a
cross-linking process that leads to the formation of collagen
and elastin. In the CNS of normal rats and mice, expression of
LOX was observed in the choroid plexus, blood vessel walls,
brain matrix, and neurons.27The LOX activity is needed for
the formation and remodeling of the extracellular matrix
during development and for maintaining normal structural
organization of tissue.28In pathological states, the involvement
of LOX is also apparent; recent data suggest that LOX is
upregulated in malignant astrocytes and its activity might be
necessary for the migratory behavior of these cells.29
SOD3,whichalso
peptidyl-a-amidating
Fig. 1
regions of rat brain (reproduced with modifications from Tarohda
et al., 2004, Springer permission #2414301072545) Pseudo-colors
indicate minimal amounts of copper (blue) and maximum amount of
copper (red). The scale (0–3.95 mg g?1tissue).
Time-dependent changes in copper concentration in different
Ann Hubbard
Dr Ann Hubbard is a professor
in the Department of Cell
Biology at the Johns Hopkins
Medical School in Baltimore,
USA. Her research has been
focused generally on pathways
and mechanisms of membrane
protein trafficking in polarized
epithelialcells,
hepatocytesand
cells. Currently, her lab is
determiningthe
signals that direct the copper-
dependent movements of the
copper-transporting ATPases,
ATP7B and ATP7A, within
epithelial cells.
specifically
intestinal
structural
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598Metallomics, 2010, 2, 596–608This journal is c The Royal Society of Chemistry 2010
The multicopper oxidase ceruloplasmin and the blood
coagulation factor fVIII share significant structural homology
but have very different properties and functions. Factor fVIII
is a component of a blood clotting cascade; copper binding to
fVIII is required for stable association of protein chains, i.e.
the major role for copper in fVIII is structural.30In contrast,
ceruloplasmin uses copper for electron transfer. By oxidizing
ferrous ion into ferric ion, ceruloplasmin facilitates iron efflux
from tissues.31The neuronal degeneration caused by the loss
of ceruloplasmin function in the CNS becomes particularly
apparent with age.32–34In brains of patients with acerulo-
plasminemia and in mice lacking ceruloplasmin, astrocytes show
a notable accumulation of iron associated with a significant
deformity and cell loss. The number of Purkinje neurons is also
reduced and their iron metabolism is altered. However, in
contrast to astrocytes, neurons of ceruloplasmin-deficient mice
appear iron-deprived.34Since the activity of ceruloplasmin
strictly depends on copper availability, an inadequate supply of
copper to CNS is likely to trigger cell-specific pathological
responses similar to those observed in aceruloplasminemia.
There is accumulating evidence linking changes in cellular
copper levels to changes in the biochemical and cell biological
properties of amyloid precursor proteins (APP and APLP),
which are associated with the pathology of Alzheimer’s disease
(AD).35,36The abnormal proteolytic processing of APP is
known to result in the accumulation of Ab fragment and
plaque formation in the brains of AD patients. Copper is
found in high concentration in these deposits; drugs that
decrease copper levels in the CNS decrease the rate of plaque
formation.37–39In vitro, complexes of Ab with copper oxidize
cholesterol producing 4-cholesten-3-one; coincidentally brain
tissue from AD subjects has almost 100% more 4-cholesten-3-
one than tissue from age-matched controls.40Copper was also
shown to bind to the full-length APP and regulate APP
endocytosis.41In animal models of AD, modulation of copper
levels significantly affects the course of the disease.42,43
Reciprocally, copper accumulation in cultured fibroblasts
from Menkes disease patients shows significant upregulation
of APP mRNA and protein, while copper depletion down-
regulates APP.44,45The connection between copper and the
aggregation state of a-synuclein (in Parkinson’sisease) and
prion protein (transmissible spongiform encephalopathy) has
also emerged (for recent detailed review see ref. 46).
Since many, if not all, cells in the CNS express more than
one copper-dependent enzyme, the complexity of cellular
responses to changing copper levels is bound to be very high.
The shift in distribution of copper between cuproenzymes and/
or between different cell compartments, as well as cell-specific
loss (or accumulation) of the metal may trigger/enhance CNS
pathology, even if total copper levels in the brain do not
change dramatically. The apparent involvement of copper in
the pathology of Alzheimer’s disease47–49and Parkinson’s
disease50–52is likely to reflect not only the direct interaction
of copper with APP53–55or its role in the catecholamine
production,56–58but an overall change in copper metabolism
in the aging brain. The next important frontier of copper
neurobiology is to provide mechanistic understanding of
copper metabolism in the CNS and its regulation in different
cell types as well as the entire brain.
Copper imbalance has severe consequences for CNS
function
Given numerous physiological processes that require copper
(see above), it is not surprising that insufficient copper supply
during brain development has long-lasting consequences.59
Severe copper deprivation is associated with marked develop-
mental defects in humans and animals. Inactivation of copper-
dependent enzymes can result in altered cell morphology,28
increased inflammation,60and even embryonic death. In
fish, genetic or dietary copper limitation is associated with
abnormal notochord development and impaired neurogenesis
affecting both the midbrain-hindbrain region as well as primary
motor neurons.61In rodents, copper deficiency is associated with
vacuolization of neurons, necrosis, edema in the cerebral cortex
and corpus striatum and convulsive seizures.3,62
Copper deficiency in humans can be caused by several
factors, such as genetic defects resulting in lower copper
absorption and abnormal copper distribution (as in Menkes
disease), or limited absorption due to an inadequate diet.
Neuropathology due to copper malabsorption has also been
reported as a complication of bariatric surgeries or other
operations affecting the gastrointestinal tract.63Overuse of
dietary supplements such as zinc and iron, and certain drugs
(such as D-penicillamine) may greatly decrease copper availability
and, if used in pregnancy, result in congenital malformations
in infants.64In adults, copper deficiency results in myeloneuro-
pathy, anemia65and motor neuron disease that can have
manifestations similar to those of amyotrophic lateral
sclerosis.66Either genetic or acquired copper deficiency leads
to demyelination and microcavitation of the neuropil in the
white matter of the spinal cord and brainstem.67There has
been a growing appreciation that neurological manifestations
caused by copper deficiency may represent a distinct syndrome;
early recognition of this syndrome may prevent neurologic
disability.65
Angiogenesis, an important physiological process of blood
vessel growth, is essential for formation of the circulatory
system. On the other hand, angiogenic blood vessels are the major
source of sustenance for neoplastic tumors and facilitators of
metastatic tumor migration. There is a large body of literature
indicating that copper sequestration diminishes angiogenesis
and that tumors are more significantly affected by such treatment
comparedto surroundingtissue.68,69
D-penicillamine and tetrathiomolybdate have been repeatedly
shown to decrease the microvascular density and tumor
volume in murine models.70–72However, the application of
D-penicillamine to treatment of glioblastoma in clinical trials
did not improve patient survival73illustrating the need for a
better mechanistic understanding of the role of copper in
angiogenesis and tumor development.
Another process in the CNS with a well-established link to
copper homeostasis is a myelination of neurons. Copper
deficiency is associated with the loss of myelin; this process
can be reversed by copper supplementation. Delayed myelination
has been observed in infants suffering from Menkes disease,74
a disorder associated with severe copper deficiency in the CNS.
Similar to angiogenesis, the specific role of copper in myelination
is not well understood; the reversible nature of copper effects
Copperchelators
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This journal is c The Royal Society of Chemistry 2010Metallomics, 2010, 2, 596–608599
suggests a regulatory role. One suggested mechanism by which
Cu deficiency might lead to a delayed or decreased rate of
myelination involves AMP-activated protein kinase (AMPK)
inhibiting acetyl-CoA carboxylase (ACC), the latter being
involved in fatty acid biosynthesis.75In addition, studies of
copper-dependent myelination in rats have demonstrated that
copper imbalance is associated with changes in mRNA levels
of proteins forming myelin sheath.76,77Similarly, the micro-
array experiments using mRNA from a post-mortem cerebral
cortex and cerebellum of a Menkes disease patient showed
preferential dysregulation of genes involved in myelination,
energy metabolism, and translation/ribosomal function.78
Thus, copper levels appear to influence transcription and/or
translation of protein components of myelin.
Two human genetic disorders illustrate harmful consequences
of either copper deficiency or copper overload. Menkes
disease, caused by mutations in the gene encoding copper-
transporting ATPase ATP7A, is an X-linked, fatal disorder of
copper metabolism. Impaired copper delivery to the brain in
patients with Menkes disease results in marked metabolic and
developmental changes, progressive neuro-degeneration and
death of a majority of patients in early childhood.74,79Patients
exhibit a loss of neurons in the granular layer of the cerebellum,
cortical atrophy, and extensive degeneration of grey matter.
Insufficient copper supply is a likely cause of low dopamine-b-
hydroxylase activity and a resultant abnormal ratio of DOPA
to catechol, which is used as a diagnostic test. Among
cuproenzymes in the CNS, lysyl oxidase appears particularly
sensitive to a diminished copper supply. In Menkes disease
patients, vascular tortuosity is common and is ascribed to the
loss of lysyl oxidase function.80,81
Wilson’s disease (WD) is an example of severe consequences
for CNS function caused by copper overload. WD is caused by
mutations inactivating the copper transporting ATPase
ATP7B.82In WD patients, copper accumulates in the liver,
brain and kidneys; the CSF copper concentration is also
elevated (76.25 ? 5.95 mg L?1).83A large proportion of WD
patients (about 40%) display neurological and/or psychiatric
symptoms84that are ascribed to copper imbalance in the CNS.
In advanced WD, several regions of the brain can be affected,
including the thalamus, subthalamic nuclei, brainstem, and
frontal cortex. The pathology manifests as atrophy, spongy
degeneration, increased ventricular size, cavitation and cyst
formation in the putamen.85
Copper transport in the CNS
The molecular mechanism through which copper is distributed
within the brain among different cell populations is not
clear. The concentration and form(s) in which copper
exists in the interstitial fluid bathing cells are also unknown.
Copper concentration in cerebrospinal fluid (CSF), largely
produced by the choroid plexus, was estimated to be
0.3–0.5 mM or 22.3 ? 2.23 mg L?1. These levels are
considerably lower than the total copper concentration in
serum (1129 ? 124 mg L?1,5,86,87), although the amounts of
exchangeable copper in the blood and CSF have not been
measured/compared. Nevertheless, the B50-fold difference
between copper in the blood and CSF suggests that the blood
brain barrier effectively limits copper’s access to the brain.
Analysis of CSF by gel filtration produced a complex profile of
copper-containing molecules with masses ranging from small
ligands to proteins with apparent molecular weights of
15–66 kDa.86It remains unknown whether these protein
components represent carriers of exchangeable copper or
copper-containing enzymes. The in vitro studies using
hypothalamic slices and the clinical data indicate that the
copper-His2 complex can be used by the cells of the
CNS;17,88,89whether such a complex exists endogenously is
unclear.
Recently, brain perfusion with64CuCl2,64Cu-albumin, and
64Cu-ceruloplasmin was employed to compare the rates of
copper uptake into brain capillaries, parenchyma, choroid
plexus, and CSF.90Very little protein-bound copper was
absorbed compared to ‘‘free’’ copper (from
major accumulation of
Comparisonoftheinitial
demonstrated that the entry of copper into cells of the choroid
plexus was 3–4 times faster than that into brain capillaries, and
9–10 times faster than copper entry into the brain parenchyma.
Copper entry into the CSF was negligible in these short-term
experiments.90These observations suggest that copper efflux
from the cells of brain barriers rather than uptake is the rate
limiting step for copper delivery to brain parenchyma. It was
suggested that choroid plexus may tightly regulate the movement
of Cu into the CSF.90The data also suggest that parenchymal
cells receive copper from the blood and not from CSF.90The
machinery involved in copper uptake, distribution and export
in the CNS is described in sections below.
At the individual cell level, the uptake of copper by astrocytes
was examined using enriched primary cultures. These studies
showed transport with an apparent Km of 9.7 ? 2.4 mM
(5–30 min).91This Km value is similar to the Km of copper
transporter hCTR1 (see below for details) heterologously
expressed in insect cells (8.9 mM)92and slightly higher than
that of hCTR1 expressed in HEK293 cells (2.56 ? 1.04 mM).93
Similar uptake rates and basal copper content of astocytes
(1.1 nmol copper per mg protein) and liver parenchymal cells
(1.4 nmolmg?1)94
or hepatoblastoma
(1.3 nmol mg?1)95suggest that copper handling in the CNS
and peripheral cells could be similar. At the same time, some
pharmacologic differences were observed between astrocytes
and HEK293 cells pointing to additional components
contributing to copper regulation in the CNS. For example,
a strong inhibition of copper accumulation by zinc was seen in
cultured astrocytes but not in HEK293 cells, suggesting that
additional zinc-sensitive copper transporter(s) may be present
in astrocytes. The molecular identity of these other transporters
is unknown; two other proteins with proposed roles in copper
uptake are DMT96and CTR2.97
64CuCl2); the
64Cu was in the choroid plexus.
ratesofcoppertransport
HepG2cells
Molecules regulating copper homeostasis in the
brain
Presently, our knowledge of copper handling machinery in
different cell types of the CNS is rudimentary. Nevertheless, it
is clear that all the key copper handling proteins mediating
copper homeostasis in peripheral tissues are also present in the
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600Metallomics, 2010, 2, 596–608This journal is c The Royal Society of Chemistry 2010
brain (Fig. 2). These proteins can be divided into three major
groups. Copper uptake transporters transfer copper into the
cytosol; copper chaperones facilitate copper distribution to
intracellular protein targets; whereas Cu-transporting ATPases
translocate copper from the cytosol to the lumen of the
secretory pathway and small vesicles for the delivery of copper
to newly synthesized cuproenzymes and export into extra-
cellular fluids, respectively (Fig. 2). Cu-ATPase may also
facilitate copper efflux into the synaptic cleft,98,99although
this has not been formally shown (the in vitro measurements
using synaptosomes and the copper-dependent quenching of
tetrakis-(4-sulfophenyl)porphine estimated copper concentration
in synaptic cleft as 2–4 mM100). The physiologic significance of
synaptic release of copper is not very clear; one suggestion is
that copper protects against excitotoxic cell death by regulating
the activities of extra-synaptic NMDA receptors.98,99Overall,
the complexity of CNS architecture and homeostasis implies
sophisticated regulation of copper metabolism. Mechanistic
understanding of this regulation on the molecular and cellular
levels is a wide-open area of study.
High affinity copper transporter CTR1
Copper transporter 1 (CTR1) is the major pathway for copper
entry into cells. CTR1 function is essential for organism
growth and development; the whole-body knockout of
CTR1 is associated with embryonic lethality in mice and
fish.101–103This phenotype is likely due to severe copper
deficiency in tissues, although a copper-independent role for
CTR1 in development has also been reported.104Experiments
in zebrafish revealed that neural tissue is particularly sensitive
to the loss of CTR1 function,103as indicated by marked
cell death in the brain and spinal cord in response to CTR1
down-regulation. In mice lacking CTR1, neuroepithelial
layering is aberrant, and the embryo exhibits impaired neural
tube closure.101,102
In mammals, CTR1 mRNA is uniformly expressed in the
brain and is higher in the choroid plexus.105The CTR1 protein
levels appear to parallel the mRNA levels with the most
intense immunostaining observed in the choroid plexus and
in the endothelial cells of small blood vessels.106Astrocytes
and neurons do not show marked differences in CTR1 levels,
at least in culture. For example, human CCF-STTG1
astrocytoma and SY5Y neuroblastoma cells express comparable
amounts of CTR1 mRNA and show similar uptake of copper.107
The level of CTR1 in the choroid plexus can be altered
(upregulated) by dietary copper deficiency.106
By analogy with other tissues, CTR1 is expected to be
present at the plasma membrane of cells forming the CNS
(Fig. 2). However, the polarity of CTR1’s distribution needs to
be determined, particularly for cells of brain barriers. In
peripheral tissues, the abundance of CTR1 at the plasma
membrane is regulated in response to changes in copper
levels.106,108,109High copper facilitates endocytosis of CTR1
to intracellular vesicles, whereas lowering copper recruits
CTR1 to the plasma membrane. Whether CTR1 is ‘‘silent’’
in the intracellular vesicles or is able to release the luminal
Fig. 2
(exchangeable copper) or to enzymes that use copper as a cofactor (cuproenzymes). Copper enters the cell via the high affinity copper transporter
CTR1, located at the plasma membrane. The levels of CTR1 at the membrane can be regulated via recycling mechanism. Copper binds to cytosolic
copper chaperones CCS and Atox1, which facilitate copper delivery to SOD1 and Cu-ATPases ATP7A and ATP7B, respectively. ATP7A and
ATP7B transfer copper into the lumen of the TGN for incorporation into secreted and plasma membrane-bound cuproenzymes. When Cu is
elevated or in response to other signals (such as activation of NMDA receptor), ATP7A moves from the TGN and facilitates copper excretion.
Whether or not ATP7B traffics in the CNS is presently unknown.
Copper distribution in a generalized cell in the CNS. In extracellular fluids copper (green balls) is bound to either specific copper carriers
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This journal is c The Royal Society of Chemistry 2010Metallomics, 2010, 2, 596–608601
copper content is presently unknown. Also, despite significant
effort, the biochemical mechanism of copper transport by
CTR1 remains largely enigmatic. Current data suggest that
CTR1-mediated permeation of copper does not require ATP
hydrolysis or co-transport of another ion.92,102,110Reducing
agents facilitate transport suggesting that CTR1 transports
Cu(I); this conclusion is further supported by the Ag+-mediated
inhibition of CTR1. Zinc and other divalent metals do not
affect the CTR1 transport function.102
The structure of human CTR1 has been solved by electron
microscopy with 6–7 A resolution.111CTR1 is a stable trimer
of identical monomers with a conducting pore for copper
located in the center of the molecule between the monomers.
Each monomer of CTR1 is a 190 amino-acid membrane
protein with three trans-membrane segments. The N-terminus
of CTR1 is oriented extracellularly and is N- and O-glycosylated;
the glycosylation protects the protein against proteolysis.112
Structural data in combination with extensive site-directed
mutagenesis92,93,113have demonstrated that the functionally
and structurally important residues are predominantly clustered
around the pore entry and exit or contribute to helix packing.113
At the same time, the vast majority of residues lining the
‘‘pore’’ are not critical for copper transport.92This observation
points to the lack of tight interaction between copper and the
walls of the permeation pathway during transport. The
conserved MxxxM motif at the pore entrance is a notable
exception; mutation of this motif abolishes copper transport.92,93
The structure of CTR1 suggests that it may act as a gated
channel in which copper binding to the extracellular surface
triggers opening of the cytosolic gate and allowing copper
release. Copper that exits CTR1 is thought to bind to cytosolic
copper chaperones that act as specific shuttles delivering
copper to various intracellular destinations (Fig. 2).
Copper chaperone CCS
Human Copper Chaperone for Superoxide dismutase 1 (CCS)
has two important functions: it facilitates the formation of
an essential disulfide bond in its target protein superoxide
dismutase 1 (SOD1) and delivers copper to the SOD1 catalytic
site. CCS is a soluble protein with a predominant localization
in the cytosol. The 274 amino-acid residues that compose CCS
are folded into three distinct domains (Fig. 3). Domain I is a
structural homologue of another copper chaperone, Atox1
(see below). This domain has a ferredoxin fold with a MxCxxC
copper binding motif in the exposed loop and is thought to
participate in copper binding, although the CxxC motif can be
mutated without a negative effect on copper transfer or
disulfide formation in SOD1. In rats, Domain I was shown
to link CCS to an aspartic protease BACE1 allowing
co-transport of the two proteins along the axon.114The
physiological consequences of this interesting partnership,
which can be disrupted by RNA aptamers,115is unknown.
Domain II of CCS is highly homologous to SOD1 and is
responsible for hetero-dimerization of CCS and SOD1 during
copper transfer. Domain III is small and has a large extended
loop with the highly conserved CXC motif (Fig. 3). This
domain is required for both functional activities of CCS: it
participates in the disulfide bond formation and in copper
transfer to SOD1. The proposed mechanism of CCS-mediated
activation has been described in a recent excellent review.116
Briefly, CCS interacts with SOD1 via Domain II; the hetero-
dimerization facilitates the formation of a disulfide bond
between Cys229of Domain III in CCS and Cys57of SOD1.
Under aerobic conditions this bond is rapidly converted into
the intramolecular disulfide in SOD1.117At which step Cu is
being inserted and the precise copper coordination during
transfer remain controversial.
In addition to its primary location and function in the
cytosol, a small fraction of CCS is found in mitochondria
and peroxisomes. Hypoxia facilitates the entry of SOD1, the
downstream target of CCS, into mitochondria. CCS is thought
to aid this process by incorporating copper and disulfide bond
into SOD1 that enters mitochondria, thus increasing retention
of the latter in the inter-membrane space. While the role of
CCS in the maturation of SOD1 is without dispute, the
function of CCS at the organism level appears to be critical
only under stress conditions. Genetic inactivation of CCS in
mice does not lead to an obvious phenotype. In contrast, the
overexpression of CCS markedly accelerates neurological
deficits and death of the transgenic mice bearing the
G93A-SOD1 (a mutant SOD1 associated with amyotropic
lateral sclerosis), apparently by facilitating mitochondria targeting
of mutant SOD1. The CCS/wild-type-SOD1 dual transgenic
mice are neurologically normal.118
Copper chaperone Atox1
Human Atox1 is a small (68 amino acids) cytosolic protein,
which is highly conserved (Z85% sequence identity) in
mammalian species and has orthologues in lower eukaryotes
and all other phyla. In solution, Atox1 exists as a monomer,
which likely represents the functional form. However, in the
presence of metals Atox1 can form dimers;119whether such
dimers have a distinct functional role in the cell is currently
unknown. The crystallographic and spectroscopic studies
Fig. 3
target SOD1 in the cytosol and in the inter-membrane space of
mitochondria. CCS has three structural domains (DI-DIII), which
have distinct function in binding of copper, docking to SOD1, and
incorporating copper and disulfide bond. The MxCxxC motif is shown
in red. The critical CxC motif is located in the extended part of DIII
(green).
Cytosolic copper chaperone CCS transfers copper to its
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602Metallomics, 2010, 2, 596–608This journal is c The Royal Society of Chemistry 2010
of Atox1 have demonstrated that the protein folds into a
compact ferredoxin-like structure (Fig. 4). Copper binds to
Atox1 within the surface-exposed loop that connects the first
b-strand and the first a-helix. The loop contains a highly
conserved motif MxCxxC, in which two cysteines coordinate
copper, whereas methionine is buried in the hydrophobic core
of Atox1 and stabilizes protein structure.120Copper binding to
Atox1 does not change protein structure but makes it more
rigid.121
In vitro, the binding of copper by Atox1 has been explored
in detail;121–123however the mechanism by which Atox1 and
other copper chaperones acquire copper in cells remains
poorly understood. Several possibilities exist that are not
mutually exclusive: (i) Atox1 may directly interact with
CTR1 and retrieve copper at the ‘‘exit gate’’ of the transporter;
(ii) it may directly or indirectly exchange copper with other
copper carrying proteins (such as copper chaperone CCS,
which has an Atox1-like domain, see above); and/or (iii)
Atox1 can bind copper from a copper-glutathione complex.
The last possibility has been shown to occur in vitro in
experiments where Cu-glutathione was used to produce a
functional Atox1-Cu(I) complex.124Glutathione also weakly
binds to Cu-Atox1 as a third copper-coordinating ligand with
a Kd of 10–25 mM.122In CNS, the intracellular levels of
glutathione are unlikely to reach sufficiently high concentrations
for it to form stable adducts with Atox1-Cu and interfere with
the subsequent transfer of copper, although this has not been
formally investigated. (The estimate for glutathione levels in
cultured cells lines ranges from 1–40 nmol mg?1protein, with
neurons containing less glutathione than astroglia.125,126)
The expression of Atox1 in a normal adult brain has been
characterized for humans and rats using Northern blotting
and in situ hybridization histochemistry.127,128Widespread
expression was observed,127with the highest levels of Atox1
mRNA found in the pyramidal neurons of cerebral cortex and
hippocampus, as well as in neurons of the locus coeruleus.128
Atox1 is moderately expressed throughout the olfactory bulb
and is low in cerebellum with the notable exception of Purkinje
neurons, where the expression was reported as very high. It is
thought that the function of Atox1 is common for all cell
types, including cells of the CNS, and involves primarily the
delivery of copper to the copper transporting ATPases
(Cu-ATPases) ATP7A and ATP7B (Fig. 4; for details on
Cu-ATPases see below). There is a strong evidence for direct
interaction between Atox1 and the N-terminal region of
Cu-ATPases as well as the transfer of copper from Atox1 to
the metal-binding sites of Cu-ATPases.124,129–132
mediated copper transfer is accompanied by upregulation of
the Cu-ATPase’s activity.124Reciprocally, apo-Atox1 can
retrieve copper from the ATPases, at least in vitro, and
down-regulate their activity.124Thus, in cells Atox1 may
function as a sensor that controls a proper delivery of copper
to the secretory pathway. In fact, recent studies demonstrate a
marked change in the intracellular distribution of copper in
cells lacking functional Atox1.133,134
In addition to a copper-transfer function, Atox1 was
proposed to play an antioxidant role, because its over-expression
in yeast cells lacking SOD1 was shown to suppress growth
defects caused by the loss of SOD1 activity.135The intra-
cellular levels of functional Atox1 have indeed significant
consequences for neuronal cell survival, although the mechanism
of this phenomenon remains unexplored. Experiments using
different neuronal cells (SKNMC, a neuroblastoma cell line;
NT-2, a teratocarcinoma cell line and GT-1, hypothalamic
neuronal cells) showed higher viability of cells over-expressing
Atox1 under conditions of serum deprivation.136Furthermore,
cells expressing the Atox1 variant with mutated cysteines in
the MxCxxC motif were less metabolically active and showed
decreased viability under either regular conditions or serum
deprivation.136It seems that the over-expressed active Atox1
may provide protection during stress, whereas the over-
expressed mutant may compete with the endogenous Atox1
for its docking sites at the N-terminus of ATP7A and/or
ATP7B and thus diminish copper delivery to the secretory
pathway. In addition, if Atox1 receives copper directly
through interactions with CTR1, then the excess of mutant
Atox1 may decrease normal copper uptake into cells.
Recent studies demonstrated the role of Atox1 in cell
growth and proliferation, and it was proposed that Atox1
may act as a transcription regulator.137It is also interesting
that the changes in the Atox1 levels appear to induce remodeling
of the entire copper-metabolic network. An increase in copper-
transporting ATPase ATP7A levels was observed in the
immortalized Atox1?/?cells,138whereas in a separate study,
the protein and mRNA levels of SOD3, a downstream target
of ATP7A, were dramatically decreased in cultured Atox1?/?
fibroblasts.139Complete Atox1 inactivation in mice has
Atox1-
Fig. 4
metal binding sites of Cu-transporting ATPases ATP7A and ATP7B.
The Cu-ATPases transport copper across membrane using the energy
of ATP hydrolysis. The structure of Atox1 (shown on the right) is
similar to the structure of individual metal binding domains of
Cu-ATPases (shown in blue and numbered 1–6). The copper-binding
loop MxCxxC is shown in red; copper is shown as a green ball.
Copper chaperone Atox1 transfers copper to the N-terminal
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grave consequences for their development and function. The
Atox1?/?mice display growth retardation and significant
mortality (up to 45%); in some cases death can be a result
of prolonged seizure activity.140Analysis of copper metabo-
lism revealed that newborn Atox1?/?mice have markedly
reduced levels of copper in the liver and brain due to decreased
placental copper transport.140They also exhibit low activity of
several copper-dependent enzymes including cytochrome C
oxidase (CCO). Some animals display severe congenital eye
defects and microphthalmia;140the precise molecular mechanisms
behind these pathologies remain unknown.
The cytochrome C oxidase assembly factors
Copper delivery to mitochondria is essential for cell survival,
because copper is required for the activity of CCO. CCO is a
terminal enzyme of the mitochondria respiratory chain; it
catalyzes electron transfer to molecular oxygen and contributes
to formation of the electrochemical potential that is used for
the synthesis of ATP. Human CCO consists of 13 subunits; the
assembly of the entire complex and copper delivery to the
catalytic sites of CCO is a fascinating and complex process,
which remains poorly understood. Studies in yeast have
suggested that the cytosolic protein Cox17 may play an
important role in copper delivery to mitochondria,141however
subsequent studies showed that tethering Cox17 to the
mitochondria inner membrane does not disrupt copper delivery
to CCO,142i.e. other cytosolic molecules must accept copper
from Ctr1 and deliver it to the organelle. Nevertheless, Cox17
plays an important role in copper transfer to CCO; genetic
inactivation of Cox17 in mice markedly reduces the CCO
activity and is embryonic lethal.143The siRNA knock-down
of COX17 in human cells has demonstrated that the down-
regulation of Cox17 is accompanied by abnormal formation of
a CCO supercomplex.144Human Cox17 is a 62-residue protein
with a hairpin-like structure (Fig. 5) and one copper(I) binding
site formed by two consecutive Cys residues.145Recent studies
also revealed the fascinating ability of Cox17 to transfer
copper to other CCO chaperones (Cox11 and SCOI) along
with the transfer of electrons.146It was suggested that such
coupled transfer may ensure the efficient delivery of the metal
to proteins when conditions are oxidizing.146
Other chaperones for CCO implicated in copper delivery
(Cox11, Sco1 and Sco2) are membrane proteins (Fig. 5). Cox
11 is required for the formation of Cu(B) site in CCO.147The
structure of the soluble domain of Cox11 from bacterial homo-
logue has been solved.148These studies revealed that Cox11 has a
novel b-immunoglobulin-like fold, shows propensity to dimerize
in the absence of reductants, and binds copper via Cys residues at
the dimerization interface.148Cox11p is needed in catalytic
amounts and is thought to act at the final step of CCO
assembly.147In contrast, Sco1 and Sco2 are more central to
CCO assembly, as evidenced by grave consequences of Sco1/2
inactivation. In humans, mutations in Sco2 are associated with
neonatal encephalocardiomyopathy,149,150while Sco1 mutations
cause neonatal hepatic failure
Analysis of Sco2 and Sco1 structures revealed that both
proteins have a thioredoxin fold, which led to the suggestion
that Scos function as ‘‘redox signaling molecules’’152rather
and encephalopathy.151
than copper carriers. The redox function for Sco2 has recently
been directly demonstrated by Leary and colleagues.153These
authors have found that Sco2 acts as a thiol–disulfide
oxidoreductaseto oxidize the
of Sco1 during maturation of the CuA site of CCO.153
Interestingly, mutations in Sco1 and Sco2 are also associated
with the reduction of cellular copper content, suggesting
that the assembly of CCO and/or mitochondria function
have a profound effect on overall cellular copper homeostasis.
The important role of mitochondria in regulating cellular
copper homeostasis is also suggested by the presence in the
mitochondria matrix of exchangeable pool of copper (for review
see ref. 154).
copper-binding cysteines
Copper-transporting ATPases ATP7A and ATP7B
(Cu-ATPases)
Human Cu-ATPases play a particularly important role in
CNS physiology. Genetic mutations in either ATP7A or
ATP7B are associated with neuronal degeneration, demyelination
of neurons, severe neurological, developmental and psychiatric
problems (see Menkes disease and Wilson’s disease, above).
Decreased expression of mouse ATP7A in Purkinje cells was
shown to be associated with impaired synaptogenesis and
dramatic cytoskeletal dysfunction.155Such severe consequences
of Cu-ATPase inactivation illustrate not only the functional
significance of these transporters but also the requirement for
precise balancing of copper in the CNS. Although ATP7A and
ATP7B are often co-expressed in same cells, permanently or
during a certain stage of development (as in Purkinje
neurons155–157), they do not fully compensate for each other’s
function when one is lost.155The precise reason for this
phenomenon is not clear, however differences in regulation
and trafficking properties of Cu-ATPases are likely to
contribute to the lack of complete functional complementation.
Fig. 5
CCO in mitochondria. Cox17 is a soluble protein found in the cytosol
and, preferentially, in the intermembarne space. Cox17 can transfer
copper (green ball) to Cox11, which is important for the CCO Cu(B)
site formation, and to Sco1. Sco2 acts as oxidoreductase for Sco1; the
activity of both Sco1 and Sco2 are required for the CCO assembly. In
addition to copper-binding proteins, mitochondria have labile copper
pool in the matrix.
Several proteins facilitate the maturation and assembly of
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604Metallomics, 2010, 2, 596–608This journal is c The Royal Society of Chemistry 2010
ATP7A and ATP7B are large (175 kDa and 160 kDa)
membrane proteins (Fig. 4). They belong to the diverse family
of P-type ATPases and use the energy of ATP hydrolysis to
transport copper from the cytosol into the lumen of the
secretory pathway where copper is incorporated as a co-factor
into various copper requiring enzymes (dopamine-b-hydroxylase,
PAM, ceruloplasmin, SOD3, lysyl oxidase, etc., see above). It
is thought that the biosynthetic incorporation of copper into
these enzymes occurs largely in the trans-Golgi network
(TGN). Consistent with this idea, ATP7A was found pre-
dominantly targeted to the TGN. Similarly, the presence of
ATP7B in the TGN was shown for Purkinje neurons in the
cerebellum. Interestingly, ATP7A was also found co-localized
with its target PAM in secretory vesicles. This localization
suggests that the function of ATP7A might be needed
downstream of the TGN to maintain the copper levels (and
thus the metallation state of PHM) in vesicles. In peripheral
cells, both Cu-ATPases were shown to traffic from the TGN in
response to changing copper levels and other stimuli.
Regulated trafficking was also observed in the CNS for
ATP7A (Fig. 2, see below section on trafficking).
ATP7A and ATP7B exhibit distinct expression patterns in
the CNS. ATP7A is expressed throughout the brain both in
embryonic and postnatal periods;105,158–160
suggests a housekeeping role for this copper transporter in
the CNS. In the early postnatal period, ATP7A expression is
most abundant in neocortex and cerebellum,158whereas in the
developing and adult brain, ATP7A levels are greatest in the
choroid plexus/ependymal cells of the lateral and third
ventricles.158ATP7A expression decreases in most neuronal
subpopulations from birth to adulthood, with the exception of
CA2 hippocampal pyramidal layer where the levels of ATP7A
increase.158An age-dependent increase in the staining of
ATP7A was also detected in Bergman glia;156and in cerebellar
Purkinje neurons, both up- and down-regulation were
reported.105,156,158–160The ATP7B mRNA and protein are
present at high levels in the postnatal cerebellum.156Tissue
blotting using rat brains suggested the presence of ATP7B in
neuronal cells of the hippocampus, olfactory bulbs, cerebellum,
cerebral cortex and nuclei in the brainstem in which high
amounts of dopamine-b-hydroxylase and SOD1 were also
detected.161
In recent years, there has been a significant progress in
describing the biochemical properties and domain structure of
Cu-ATPases (for reviews see ref. 162–164). The Cu-ATPases
show high sequence similarity, common domain structure and
both hydrolyze ATP in a copper dependent manner with the
formation of a transient phospho-intermediate. Unlike
ATP7B, ATP7A is glycosylated, which increases its mass
and allows separation of the two Cu-ATPases in gels. The
membrane portion of human Cu-ATPases consists of 8
transmembrane segments that form the copper translocation
pathway and copper-binding sites with a high selectivity for
Cu(I). The bulk of the Cu-ATPases is exposed to the cytosol
(Fig. 4). The cytosolic domains include: the N-terminal
domain with 6 copper binding sites; the nucleotide-binding
domain where ATP docks; the P (phosphorylation domain)
with an invariant Asp (the site of catalytic phosphorylation);
theA-domain that regulates
this pattern
thecatalyticcycleand
conformational transitions in the Cu-ATPases; and the
C-terminal tail that is needed for protein stability and traffick-
ing. In addition, both proteins have unique sequences that are
thought to be responsible for differences in their regulation.164
Regulation of copper balance
Available data indicate that copper balance in the CNS is
maintained under very tight control, and compensatory
mechanisms are activated to achieve copper balance in disease
situations. For example, in MoBr/ymice (an animal model of
Menkes disease) the decreased expression of ATP7A protein in
Purkinje cells is associated with impaired synaptogenesis and
marked decrease in overall copper levels.155These events are
accompanied by compensatory upregulation of ATP7A in
endothelial cells, as well as increased association of MoBr/y
astrocytes and microglia with the blood–brain barrier.
Interestingly, ATP7B is not upregulated.155
Very little is known about specific molecular mechanisms
that respond to changing copper levels in the CNS. The
available data indicate that copper handling machinery is
regulated in a cell-specific manner at many levels, including
transcriptional, translational, and posttranslational control.
For example, the mRNA and protein levels of copper chaperone
CCS are sensitive to intracellular copper levels, and the
response is cell-specific.165In mice, copper deficiency is
accompanied by an increase in the CCS levels in the cerebellum,
whereas CCS in choroid plexus is essentially unchanged.165In
another example, the levels of ATP7A mRNA and protein
change during development, as does the intracellular localization
of ATP7A.156,158,166In developing axons, ATP7A is initially
expressed in cell bodies, whereas later it moves to extending
axons, peaking in its amounts prior to synaptogenesis.
Similarly, injury-stimulated neurogenesis is associated with
the increase of ATP7A in neurons and axons prior to
synaptogenesis.166
Transcriptional control was reported for both ATP7A
and ATP7B. In neuroblastoma cells, ATP7A expression is
triggered by retinoic acid receptor b (RAR b2).167The factors
regulating expression of the full-length ATP7B in the brain are
unknown. However, the production of a shorter, pineal gland
specific form of ATP7B (PINA) appears to be under control of
a pineal/retina specific nuclear factor, cone rod homeobox
CRX,168and follows a circadian rhythm.52In cultured pineal
cells, expression of PINA can be stimulated by activating
the cAMP signal transduction pathway.52In addition to
transcriptional control, ATP7A and ATP7B mRNA are
subjects of alternative splicing, both in normal and disease
states.169–176The level of expression, developmental regulation,
and trafficking behavior of such variants, when different from
the full-length form, may have important effects on CNS
copper homeostasis.
Regulation of copper balance through trafficking of
Cu-ATPases
Although ATP7A and ATP7B are B65% identical and share
a common transport function in the secretory pathway of cells,
they exhibit several important differences.157,177,178Perhaps
most dramatic is the distinct trafficking behavior of each
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This journal is c The Royal Society of Chemistry 2010Metallomics, 2010, 2, 596–608605
protein when intracellular Cu(I) levels increase and the
enzymes initiate their Cu(I) export function. ATP7A disperses
from the TGN in small vesicles, which move toward/to the
basolateral region of polarized cells.178,179In contrast, ATP7B
in slightly larger vesicles moves toward the apical region of
polarized cells (e.g., liver hepatocytes180–182). Several molecular
signals necessary for the polarity of the trafficking have been
identified on each protein183–188but the recognition and
transport machinery are still unknown (although relevant
interacting partners such as dynactin p62189and the PDZ-domain
containing PICP/AIPP1190were identified). Recent interesting
study indicates that glutathionylation (and the removal of
glutathione by glutaredoxin) may regulate the activity and
trafficking of Cu-ATPases.191
While it is thought that Cu-ATPases in vesicles actively
sequester Cu(I) for export during vesicle fusion, the possibility
remains that they act as vehicles for delivery of the specific
Cu-ATPase to its appropriate plasma membrane destination.
The composition of the ATP7A and ATP7B carrying vesicles
needs to be determined to better understand the driving
forces behind cell specific regulation of ATP7A and ATP7B
(see below).
So far, work on Cu-ATPase trafficking has focused almost
exclusively on tissues/cells outside of the CNS. To our
knowledge, little is known of the Cu-responsive trafficking
behavior of either Cu-ATPase in any region of the brain. Since
in some tissues, where both Cu-ATPAses are expressed, there
is evidence for one or the other protein not responding to
changes in Cu levels,192–194it is formally possible that ATP7A
and ATP7B in the some cells of CNS function exclusively
attheTGN.However, Cu-independent
ATP7A clearly occurs in hippocampal neurons in response
to glutamate excitation of NMDA receptors.99The ATP7A-
mediated release of Cu appears to modulate the excitotoxicity
of the NMDA receptor through nitrosylation.99The signal(s)
recruiting ATP7A from the cell body to the periphery are not
known but are likely to be distinct from those responding to
Cu(I) levels. Such novel results emphasize the need for more
study of the CNS Cu-ATPases.
traffickingof
Conclusions
The above discussion has aimed to illustrate two major points.
First, the available data leave little doubt that copper
homeostasis plays a very important role in normal function
and development of the mammalian CNS. Second, despite its
importance, our knowledge of copper homeostasis on the
brain is rudimentary, particularly considering the complexity
of the CNS architecture and the abundance of regulatory
mechanisms. With the major components of copper handling
machinery now identified, the field is ripe for studies directed
on understanding the mechanisms that control copper levels
within the CNS, the role of different copper transporters and
chaperones in CNS development and their adaptation to
pathological changes. Such studies would contribute to better
understanding of neuropathology of many disorders including
Menkesdisease,Wilson’sdisease,Alzheimer’sdisease,Parkinson’s
disease, and cancer.
Acknowledgements
This work was supported by the NIH Program Project Grant
P01 GM067166.
References
1 T. Lech and J. K. Sadlik, Biol. Trace Elem. Res., 2007, 118(1),
10–15.
2 L. Rongzhu, W. Suhua, X. Guangwei, R. Chunlan, H. Fangan,
J. Junjie and M. Aschner, Biol. Trace Elem. Res., 2009, 130(1),
39–47.
3 A. A. Gybina, I. Tkac and J. R. Prohaska, Nutr. Neurosci., 2009,
12(3), 114–122.
4 D. J. Waggoner, B. Drisaldi, T. B. Bartnikas, R. L. Casareno,
J. R. Prohaska, J. D. Gitlin and D. A. Harris, J. Biol. Chem.,
2000, 275(11), 7455–7458.
5 V. Nischwitz, A. Berthele and B. Michalke, Anal. Chim. Acta,
2008, 627(2), 258–269.
6 C. O. Hershey, L. A. Hershey, A. Varnes, S. D. Vibhakar,
P. Lavin and W. H. Strain, Neurology, 1983, 33(10), 1350–1353.
7 H. Basun, L. G. Forssell, L. Wetterberg and B. Winblad,
J. Neural. Transm. Park. Dis. Dement. Sect., 1991, 3(4),
231–258.
8 B. Jackson, S. Harper, L. Smith and J. Flinn, Anal. Bioanal.
Chem., 2006, 384(4), 951–957.
9 J. Dobrowolska, M. Dehnhardt, A. Matusch, M. Zoriy,
N.Palomero-Gallagher,P.
J. S. Becker, Talanta, 2008, 74(4), 717–723.
10 J. S. Becker, M. V. Zoriy, C. Pickhardt, N. Palomero-Gallagher
and K. Zilles, Anal. Chem., 2005, 77(10), 3208–3216.
11 P. Zatta, D. Drago, P. Zambenedetti, S. Bolognin, E. Nogara,
A. Peruffo and B. Cozzi, J. Chem. Neuroanat., 2008, 36(1), 1–5.
12 T. Tarohda, Y. Ishida, K. Kawai, M. Yamamoto and R. Amano,
Anal. Bioanal. Chem., 2005, 383(2), 224–234.
13 Y. Nose, B. E. Kim and D. J. Thiele, Cell Metab., 2006, 4(3),
235–244.
14 T. Yamano, M. Shimada, A. Onaga, H. Kawasaki, S. Iwane,
K. Ono and M. Nishimura, Acta Neuropathol., 1988, 76(6),
574–580.
15 H. Kawasaki, T. Yamano, S. Iwane and M. Shimada, Acta
Neuropathol., 1988, 76(6), 606–612.
16 J. Kreuder, A. Otten, H. Fuder, Z. Tumer, T. Tonnesen, N. Horn
and D. Dralle, Eur. J. Pediatr., 1993, 152(10), 828–832.
17 Z. Tumer and L. B. Moller, Eur. J. Hum. Genet., 2010, 18(5),
511–518.
18 I. T. Demchenko, T. D. Oury, J. D. Crapo and C. A. Piantadosi,
Circ. Res., 2002, 91(11), 1031–1037.
19 R. Rola, Y. Zou, T. T. Huang, K. Fishman, J. Baure, S. Rosi,
H. Milliken, C. L. Limoli and J. R. Fike, Free Radical Biol. Med.,
2007, 42(8), 1133–1145; discussion 1131–1132.
20 B. A. Eipper, D. A. Stoffers and R. E. Mains, Annu. Rev.
Neurosci., 1992, 15, 57–85.
21 X. Xin, R. E. Mains and B. A. Eipper, J. Biol. Chem., 2004,
279(46), 48159–48167.
22 K. Tief, A. Schmidt and F. Beermann, Mol. Brain Res., 1998,
53(1–2), 307–310.
23 K. Tief, M. Hahne, A. Schmidt and F. Beermann, Eur. J.
Biochem., 1996, 241(1), 12–16.
24 E. Greggio, E. Bergantino, D. Carter, R. Ahmad, G. E. Costin,
V. J. Hearing, J. Clarimon, A. Singleton, J. Eerola, O. Hellstrom,
P. J. Tienari, D. W. Miller, A. Beilina, L. Bubacco and
M. R. Cookson, J. Neurochem., 2005, 93(1), 246–256.
25 Y. Higashi, M. Asanuma, I. Miyazaki and N. Ogawa,
J. Neurochem., 2000, 75(4), 1771–1774.
26 C. A. Cronin, A. B. Ryan, E. M. Talley and H. Scrable,
J. Neurosci., 2003, 23(37), 11692–11697.
27 P. A. Li, Q. He, T. Cao, G. Yong, K. M. Szauter, K. S. Fong,
J. Karlsson, M. F. Keep and K. Csiszar, Mol. Brain Res., 2004,
120(2), 115–122.
28 N. W. Bronson, J. S. Hamilton, M. Han, P. A. Li, I. Hornstra,
J. M. Horowitz and B. A. Horwitz, Neurosci. Lett., 2005, 390(2),
118–122.
Koscielniak,K.Zillesand
Downloaded on 10 October 2010
Published on 17 August 2010 on http://pubs.rsc.org | doi:10.1039/C0MT00006J
View Online

Page 11

606Metallomics, 2010, 2, 596–608This journal is c The Royal Society of Chemistry 2010
29 R. Laczko, K. M. Szauter, M. K. Jansen, P. Hollosi, M. Muranyi,
J. Molnar, K. S. Fong, A. Hinek and K. Csiszar, Neuropathol.
Appl. Neurobiol., 2007, 33(6), 631–643.
30 N. Bihoreau, S. Pin, A. M. de Kersabiec, F. Vidot and
M. P. Fontaine-Aupart, Eur. J. Biochem., 1994, 222(1), 41–48.
31 Z. L. Harris, A. P. Durley, T. K. Man and J. D. Gitlin, Proc. Natl.
Acad. Sci. U. S. A., 1999, 96(19), 10812–10817.
32 K. Kaneko, K. Yoshida, K. Arima, S. Ohara, H. Miyajima,
T. Kato, M. Ohta and S. I. Ikeda, J. Neuropathol. Exp. Neurol.,
2002, 61(12), 1069–1077.
33 H. Miyajima,S. Kono,
M. Sakamoto and N. Sakai, Neurology, 2001, 57(12), 2205–2210.
34 S. Y. Jeong and S. David, J. Neurosci., 2006, 26(38),
9810–9819.
35 N. C. Inestrosa, W. Cerpa and L. Varela-Nallar, IUBMB Life,
2005, 57(9), 645–650.
36 N. C. Inestrosa, A. E. Reyes, M. A. Chacon, W. Cerpa,
A. Villalon,J. Montiel,G.
F. Bozinovic and F. Aboitiz, Neurobiol. Aging, 2005, 26(7),
1023–1028.
37 C. S. Atwood, R. C. Scarpa, X. Huang, R. D. Moir, W. D. Jones,
D. P. Fairlie, R. E. Tanzi and A. I. Bush, J. Neurochem., 2000,
75(3), 1219–1233.
38 K. A. Price, P. J. Crouch and A. R. White, Recent Pat. CNS Drug
Discovery, 2007, 2(3), 180–187.
39 S. Schafer, F. G. Pajonk, G. Multhaup and T. A. Bayer, J. Mol.
Med., 2007, 85(4), 405–413.
40 L. Puglielli, A. L. Friedlich, K. D. Setchell, S. Nagano, C. Opazo,
R. A. Cherny, K. J. Barnham, J. D. Wade, S. Melov,
D. M. Kovacs and A. I. Bush, J. Clin. Invest., 2005, 115(9),
2556–2563.
41 Y. H. Hung, E. L. Robb, I. Volitakis, M. Ho, G. Evin, Q. X. Li,
J. G. Culvenor, C. L. Masters, R. A. Cherny and A. I. Bush,
J. Biol. Chem., 2009, 284(33), 21899–21907.
42 D. L. Sparks, J. Alzheimers. Dis., 2008, 15(4), 641–656.
43 A. L. Phinney, B. Drisaldi, S. D. Schmidt, S. Lugowski,
V. Coronado, Y. Liang, P. Horne, J. Yang, J. Sekoulidis,
J. Coomaraswamy, M. A. Chishti, D. W. Cox, P. M. Mathews,
R. A. Nixon, G. A. Carlson, P. St George-Hyslop and
D. Westaway, Proc. Natl. Acad. Sci. U. S. A., 2003, 100(24),
14193–14198.
44 A. D. Armendariz, M. Gonzalez, A. V. Loguinov and
C. D. Vulpe, Physiol. Genomics, 2004, 20(1), 45–54.
45 S. A. Bellingham, D. K. Lahiri, B. Maloney, S. La Fontaine,
G. Multhaup and J. Camakaris, J. Biol. Chem., 2004, 279(19),
20378–20386.
46 D. R. Brown, Metallomics, 2010, 2, 186–194.
47 Y. H. Hung, A. I. Bush and R. A. Cherny, JBIC, J. Biol. Inorg.
Chem., 2010, 15(1), 61–76.
48 D. R. Brown, Dalton Trans., 2009, (21), 4069–4076.
49 J. F. Quinn, S. Crane, C. Harris and T. L. Wadsworth, Expert
Rev. Neurother., 2009, 9(5), 631–637.
50 K. J. Barnham and A. I. Bush, Curr. Opin. Chem. Biol., 2008,
12(2), 222–228.
51 M. Alcaraz-Zubeldia, M. C. Boll-Woehrlen, S. Montes-Lopez,
F. Perez-Severiano, J. C. Martinez-Lazcano, A. Diaz-Ruiz and
C. Rios, Rev. Invest. Clin., 2009, 61(5), 405–411.
52 J. Borjigin, A. S. Payne, J. Deng, X. Li, M. M. Wang,
B. Ovodenko, J. D. Gitlin and S. H. Snyder, J. Neurosci., 1999,
19(3), 1018–1026.
53 M. Suazo, C. Hodar, C. Morgan, W. Cerpa, V. Cambiazo,
N. C. Inestrosa and M. Gonzalez, Biochem. Biophys. Res.
Commun., 2009, 382(4), 740–744.
54 G. K. Kong, L. A. Miles, G. A. Crespi, C. J. Morton, H. L. Ng,
K. J. Barnham, W. J. McKinstry, R. Cappai and M. W. Parker,
Eur. Biophys. J., 2008, 37(3), 269–279.
55 K. J. Barnham, W. J. McKinstry, G. Multhaup, D. Galatis,
C. J. Morton, C. C. Curtain, N. A. Williamson, A. R. White,
M. G. Hinds, R. S. Norton, K. Beyreuther, C. L. Masters,
M. W. Parker and R. Cappai, J. Biol. Chem., 2003, 278(19),
17401–17407.
56 D. S. Goldstein, C. S. Holmes and S. G. Kaler, Neurochem. Res.,
2009, 34(8), 1464–1468.
57 T. Saito, T. Nagao, M. Okabe and K. Saito, Neurosci. Lett., 1996,
216(3), 195–198.
Y. Takahashi,M. Sugimoto,
Merabachvili,R. Aldunate,
58 V. Gerbasi, S. Lutsenko and E. J. Lewis, Neurochem. Res., 2003,
28(6), 867–873.
59 J. R. Prohaska, Nutrition, 2000, 16(7–8), 502–504.
60 H. E. Lob, P. J. Marvar, T. J. Guzik, S. Sharma, L. A. McCann,
C. Weyand, F. J. Gordon and D. G. Harrison, Hypertension,
2010, 55(2), 277–283, 276p following 283.
61 E. C. Madsen and J. D. Gitlin, PLoS Genet., 2008, 4(11),
e1000261.
62 J. Y. Uriu-Adams, R. E. Scherr, L. Lanoue and C. L. Keen,
Biofactors, 36(2), 136–152.
63 K. Juhasz-Pocsine, S. A. Rudnicki, R. L. Archer and S. I. Harik,
Neurology, 2007, 68(21), 1843–1850.
64 M. L. Martinez-Frias, E. Rodriguez-Pinilla, E. Bermejo and
M. Blanco, Am. J. Med. Genet., 1998, 76(3), 274–275.
65 N. Kumar and P. A. Low, J. Neurol., 2004, 251(6), 747–749.
66 C. C. Weihl and G. Lopate, Muscle Nerve, 2006, 34(6), 789–793.
67 B. Schleper and H. J. Stuerenburg, J. Neurol., 2001, 248(8),
705–706.
68 X. Z. Wu, Neoplasma, 2008, 55(6), 472–481.
69 D. Yoshida, Y. Ikeda and S. Nakazawa, Neurosurgery, 1995,
37(2), 287–292, discussion 292–283.
70 C. Cox, S. D. Merajver, S. Yoo, R. D. Dick, G. J. Brewer,
J. S. Lee and T. N. Teknos, Arch. Otolaryngol., Head Neck Surg.,
2003, 129(7), 781–785.
71 S. S. Brem, D. Zagzag, A. M. Tsanaclis, S. Gately, M. P. Elkouby
and S. E. Brien, Am. J. Pathol., 1990, 137(5), 1121–1142.
72 B. Hassouneh, M. Islam, T. Nagel, Q. Pan, S. D. Merajver and
T. N. Teknos, Mol. Cancer Ther., 2007, 6(3), 1039–1045.
73 S. Brem, S. A. Grossman, K. A. Carson, P. New, S. Phuphanich,
J. B. Alavi, T. Mikkelsen and J. D. Fisher, Neuro-Oncology, 2005,
7(3), 246–253.
74 S. G. Kaler, C. S. Holmes, D. S. Goldstein, J. Tang,
S. C. Godwin, A. Donsante, C. J. Liew, S. Sato and
N. Patronas, N. Engl. J. Med., 2008, 358(6), 605–614.
75 A. A. Gybina and J. R. Prohaska, Brain Res., 2008, 1204, 69–76.
76 N. Franco-Pons, J. Tomas, B. Roig, C. Auladell, L. Martorell
and E. Vilella, J. Mol. Neurosci., 2009, 38(1), 2–11.
77 P. Morell, C. V. Barrett, J. L. Mason, A. D. Toews,
J. D. Hostettler, G. W. Knapp and G. K. Matsushima, Mol.
Cell. Neurosci., 1998, 12(4–5), 220–227.
78 P. C. Liu, Y. W. Chen, J. A. Centeno, M. Quezado, K. Lem and
S. G. Kaler, Mol. Genet. Metab., 2005, 85(4), 291–300.
79 L. B. Moller, M. Mogensen and N. Horn, Biochimie, 2009, 91(10),
1273–1277.
80 S. C. Godwin, T. Shawker, B. Chang and S. G. Kaler, J. Pediatr.,
2006, 149(3), 412–415.
81 P. M. Royce and B. Steinmann, Pediatr. Res., 1990, 28(2),
137–141.
82 S. K. Das and K. Ray, Nat. Clin. Pract. Neurol., 2006, 2(9),
482–493.
83 H. J. Stuerenburg, J. Neural Transm., 2000, 107(3), 321–329.
84 M. T. Lorincz, Ann. N. Y. Acad. Sci., 2010, 1184, 173–187.
85 S. Sinha, A. B. Taly, S. Ravishankar, L. K. Prashanth,
K. S. Venugopal, G. R. Arunodaya, M. K. Vasudev and
H. S. Swamy, Neuroradiology, 2006, 48(9), 613–621.
86 K. Gellein, P. M. Roos, L. Evje, O. Vesterberg, T. P. Flaten,
M. Nordberg and T. Syversen, Brain Res., 2007, 1174, 136–142.
87 G. Forte, B. Bocca, O. Senofonte, F. Petrucci, L. Brusa,
P. Stanzione, S. Zannino, N. Violante, A. Alimonti and
G. Sancesario, J. Neural Transm., 2004, 111(8), 1031–1040.
88 B. M. Katz and A. Barnea, J. Biol. Chem., 1990, 265(4),
2017–2021.
89 A. Barnea and B. M. Katz, J. Inorg. Biochem., 1990, 40(1), 81–93.
90 B. S. Choi and W. Zheng, Brain Res., 2009, 1248, 14–21.
91 I. F. Scheiber, J. F. Mercer and R. Dringen, Neurochem. Int.,
2010, 56(3), 451–460.
92 J. F. Eisses and J. H. Kaplan, J. Biol. Chem., 2005, 280(44),
37159–37168.
93 J. Lee, M. M. Pena, Y. Nose and D. J. Thiele, J. Biol. Chem.,
2002, 277(6), 4380–4387.
94 G. J. Van den Berg, J. J. de Goeij, I. Bock, M. J. Gijbels,
A. Brouwer, K. Y. Lei and H. F. Hendriks, J. Nutr., 1991,
121(8), 1228–1235.
95 L. Tapia, M. Suazo, C. Hodar, V. Cambiazo and M. Gonzalez,
BioMetals, 2003, 16(1), 169–174.
Downloaded on 10 October 2010
Published on 17 August 2010 on http://pubs.rsc.org | doi:10.1039/C0MT00006J
View Online

Page 12

This journal is c The Royal Society of Chemistry 2010Metallomics, 2010, 2, 596–608607
96 W. Zheng, N. Xin, Z. H. Chi, B. L. Zhao, J. Zhang, J. Y. Li and
Z. Y. Wang, FASEB J., 2009, 23(12), 4207–4217.
97 B. Zhou and J. Gitschier, Proc. Natl. Acad. Sci. U. S. A., 1997,
94(14), 7481–7486.
98 M. L. Schlief and J. D. Gitlin, Mol. Neurobiol., 2006, 33(2), 81–90.
99 M. L. Schlief, T. West, A. M. Craig, D. M. Holtzman and
J. D. Gitlin, Proc. Natl. Acad. Sci. U. S. A., 2006, 103(40),
14919–14924.
100 A. Hopt, S. Korte, H. Fink, U. Panne, R. Niessner, R. Jahn,
H. Kretzschmar and J. Herms, J. Neurosci. Methods, 2003,
128(1–2), 159–172.
101 J. Lee, J. R. Prohaska and D. J. Thiele, Proc. Natl. Acad. Sci.
U. S. A., 2001, 98(12), 6842–6847.
102 Y. M. Kuo, B. Zhou, D. Cosco and J. Gitschier, Proc. Natl. Acad.
Sci. U. S. A., 2001, 98(12), 6836–6841.
103 N. C. Mackenzie, M. Brito, A. E. Reyes and M. L. Allende, Gene,
2004, 328, 113–120.
104 T. Haremaki, S. T. Fraser, Y. M. Kuo, M. H. Baron and
D. C. Weinstein, Proc. Natl. Acad. Sci. U. S. A., 2007, 104(29),
12029–12034.
105 N. A. Platonova, S. A. Barabanova, R. G. Povalikhin,
N. V. Tsymbalenko, A. D. Nozdrachev and L. V. Puchkova,
Dokl. Biol. Sci., 2005, 401, 88–91.
106 Y. M. Kuo, A. A. Gybina, J. W. Pyatskowit, J. Gitschier and
J. R. Prohaska, J. Nutr., 2006, 136(1), 21–26.
107 Y. Qian, Y. Zheng, L. Abraham, K. S. Ramos and E. Tiffany-
Castiglioni, Mol. Brain Res., 2005, 134(2), 323–332.
108 S. A. Molloy and J. H. Kaplan, J. Biol. Chem., 2009, 284(43),
29704–29713.
109 Y. Guo, K. Smith, J. Lee, D. J. Thiele and M. J. Petris, J. Biol.
Chem., 2004, 279(17), 17428–17433.
110 J. F. Eisses and J. H. Kaplan, J. Biol. Chem., 2002, 277(32),
29162–29171.
111 C. J. De Feo, S. G. Aller, G. S. Siluvai, N. J. Blackburn and
V. M. Unger, Proc. Natl. Acad. Sci. U. S. A., 2009, 106(11),
4237–4242.
112 E. B. Maryon, S. A. Molloy and J. H. Kaplan, J. Biol. Chem.,
2007, 282(28), 20376–20387.
113 C. J. De Feo, S. Mootien and V. M. Unger, J. Membr. Biol.,
234(2), 113–123.
114 B. Angeletti, K. J. Waldron, K. B. Freeman, H. Bawagan,
I. Hussain, C. C. Miller, K. F. Lau, M. E. Tennant,
C. Dennison, N. J. Robinson and C. Dingwall, J. Biol. Chem.,
2004, 280(18), 17930–17937.
115 A. Rentmeister, A. Bill, T. Wahle, J. Walter and M. Famulok,
RNA, 2006, 12(9), 1650–1660.
116 J. M. Leitch, P. J. Yick and V. C. Culotta, J. Biol. Chem., 2009,
284(37), 24679–24683.
117 Y. Furukawa and T. V. O’Halloran, Antioxid. Redox Signaling,
2006, 8(5–6), 847–867.
118 M.Son,K. Puttaparthi,
P. J. Boyer, G. Manfredi and J. L. Elliott, Proc. Natl. Acad.
Sci. U. S. A., 2007, 104(14), 6072–6077.
119 V. Tanchou, F. Gas, A. Urvoas, F. Cougouluegne, S. Ruat,
O. Averseng and E. Quemeneur, Biochem. Biophys. Res. Com-
mun., 2004, 325(2), 388–394.
120 A. Rodriguez-Granillo and P. Wittung-Stafshede, Biochemistry,
2009, 48(5), 960–972.
121 I. Anastassopoulou, L. Banci, I. Bertini, F. Cantini, E. Katsari
and A. Rosato, Biochemistry, 2004, 43(41), 13046–13053.
122 M. Ralle, S. Lutsenko and N. J. Blackburn, J. Biol. Chem., 2003,
278(25), 23163–23170.
123 A. K. Wernimont, L. A. Yatsunyk and A. C. Rosenzweig, J. Biol.
Chem., 2003, 279(13), 12269–12276.
124 J. M. Walker, R. Tsivkovskii and S. Lutsenko, J. Biol. Chem.,
2002, 277(31), 27953–27959.
125 S. P. Raps, J. C. Lai, L. Hertz and A. J. Cooper, Brain Res., 1989,
493(2), 398–401.
126 E. Pileblad, P. S. Eriksson and E. Hansson, J. Neural Transm.,
1991, 86(1), 43–49.
127 L. W. Klomp, S. J. Lin, D. S. Yuan, R. D. Klausner, V. C. Culotta
and J. D. Gitlin, J. Biol. Chem., 1997, 272(14), 9221–9226.
128 G. S. Naeve, A. M. Vana, J. R. Eggold, G. S. Kelner, R. Maki,
E. B. Desouza and A. C. Foster, Neuroscience, 1999, 93(3),
1179–1187.
H.Kawamata,B.Rajendran,
129 F. Hussain, A. Rodriguez-Granillo and P. Wittung-Stafshede,
J. Am. Chem. Soc., 2009, 131(45), 16371–16373.
130 L. Banci, I. Bertini, F. Cantini, N. Della-Malva, M. Migliardi and
A. Rosato, J. Biol. Chem., 2007, 282(32), 23140–23146.
131 L. A. Yatsunyk and A. C. Rosenzweig, J. Biol. Chem., 2007,
282(12), 8622–8631.
132 M. Ralle, S. Lutsenko and N. J. Blackburn, J. Inorg. Biochem.,
2004, 98(5), 765–774.
133 T. Miyayama, K. T. Suzuki and Y. Ogra, Toxicol. Appl.
Pharmacol., 2009, 237(2), 205–213.
134 R. McRae, B. Lai and C. J. Fahrni, JBIC, J. Biol. Inorg. Chem.,
2010, 15(1), 99–105.
135 I. H. Hung, R. L. Casareno, G. Labesse, F. S. Mathews and
J. D. Gitlin, J. Biol. Chem., 1998, 273(3), 1749–1754.
136 G. S. Kelner, M. Lee, M. E. Clark, D. Maciejewski, D. McGrath,
S. Rabizadeh, T. Lyons, D. Bredesen, P. Jenner and R. A. Maki,
J. Biol. Chem., 2000, 275(1), 580–584.
137 S. Itoh, H. W. Kim, O. Nakagawa, K. Ozumi, S. M. Lessner,
H. Aoki, K. Akram, R. D. McKinney, M. Ushio-Fukai and
T. Fukai, J. Biol. Chem., 2008, 283(14), 9157–9167.
138 I. Hamza, J. Prohaska and J. D. Gitlin, Proc. Natl. Acad. Sci.
U. S. A., 2003, 100(3), 1215–1220.
139 S. Itoh, K. Ozumi, H. W. Kim, O. Nakagawa, R. D. McKinney,
R. J. Folz, I. N. Zelko, M. Ushio-Fukai and T. Fukai, Free
Radical Biol. Med., 2009, 46(1), 95–104.
140 I. Hamza, A. Faisst, J. Prohaska, J. Chen, P. Gruss and
J. D. Gitlin, Proc. Natl. Acad. Sci. U. S. A., 2001, 98(12),
6848–6852.
141 D. M. Glerum, A. Shtanko and A. Tzagoloff, J. Biol. Chem.,
1996, 271(24), 14504–14509.
142 A. B. Maxfield, D. N. Heaton and D. R. Winge, J. Biol. Chem.,
2003, 279(7), 5072–5080.
143 Y. Takahashi, K. Kako, S. Kashiwabara, A. Takehara, Y. Inada,
H. Arai, K. Nakada, H. Kodama, J. Hayashi, T. Baba and
E. Munekata, Mol. Cell. Biol., 2002, 22(21), 7614–7621.
144 C. Oswald, U. Krause-Buchholz and G. Rodel, J. Mol. Biol.,
2009, 389(3), 470–479.
145 L. Banci, I. Bertini, S. Ciofi-Baffoni, A. Janicka, M. Martinelli,
H. Kozlowski and P. Palumaa, J. Biol. Chem., 2008, 283(12),
7912–7920.
146 L. Banci, I. Bertini, S. Ciofi-Baffoni, T. Hadjiloi, M. Martinelli
and P. Palumaa, Proc. Natl. Acad. Sci. U. S. A., 2008, 105(19),
6803–6808.
147 L. Hiser, M. Di Valentin, A. G. Hamer and J. P. Hosler, J. Biol.
Chem., 2000, 275(1), 619–623.
148 L. Banci, I. Bertini, F. Cantini, S. Ciofi-Baffoni, L. Gonnelli and
S. Mangani, J. Biol. Chem., 2004, 279(33), 34833–34839.
149 B. C. Mobley, G. M. Enns, L. J. Wong and H. Vogel, Clin.
Neuropathol., 2009, 28(2), 143–149.
150 L. C. Papadopoulou, C. M. Sue, M. M. Davidson, K. Tanji,
I. Nishino, J. E. Sadlock, S. Krishna, W. Walker, J. Selby,
D. M. Glerum, R. V. Coster, G. Lyon, E. Scalais, R. Lebel,
P. Kaplan, S. Shanske, D. C. De Vivo, E. Bonilla, M. Hirano,
S. DiMauro and E. A. Schon, Nat. Genet., 1999, 23(3), 333–337.
151 I. Valnot, S. Osmond, N. Gigarel, B. Mehaye, J. Amiel,
V. Cormier-Daire, A. Munnich, J. P. Bonnefont, P. Rustin and
A. Rotig, Am. J. Hum. Genet., 2000, 67(5), 1104–1109.
152 J. C. Williams, C. Sue, G. S. Banting, H. Yang, D. M. Glerum,
W. A. Hendrickson and E. A. Schon, J. Biol. Chem., 2005,
280(15), 15202–15211.
153 S. C. Leary, F. Sasarman, T. Nishimura and E. A. Shoubridge,
Hum. Mol. Genet., 2009, 18(12), 2230–2240.
154 F. Pierrel, P. A. Cobine and D. R. Winge, BioMetals, 2007,
20(3–4), 675–682.
155 M. J. Niciu, X. M. Ma, R. El Meskini, J. S. Pachter, R. E. Mains
and B. A. Eipper, Neurobiol. Dis., 2007, 27(3), 278–291.
156 N. Barnes, R. Tsivkovskii, N. Tsivkovskaia and S. Lutsenko,
J. Biol. Chem., 2004, 280(10), 9640–9645.
157 R. Linz and S. Lutsenko, J. Bioenerg. Biomembr., 2007, 39(5–6),
403–407.
158 M. J. Niciu, X. M. Ma, R. El Meskini, G. V. Ronnett,
R. E. Mains and B. A. Eipper, Neuroscience, 2006, 139(3),
947–964.
159 E. Nishihara, T. Furuyama, S. Yamashita and N. Mori,
NeuroReport, 1998, 9(14), 3259–3263.
Downloaded on 10 October 2010
Published on 17 August 2010 on http://pubs.rsc.org | doi:10.1039/C0MT00006J
View Online

Page 13

608Metallomics, 2010, 2, 596–608 This journal is c The Royal Society of Chemistry 2010
160 Y. M. Kuo, J. Gitschier and S. Packman, Hum. Mol. Genet., 1997,
6(7), 1043–1049.
161 T. Saito, M. Okabe, T. Hosokawa, M. Kurasaki, A. Hata,
F. Endo, K. Nagano, I. Matsuda, K. Urakami and K. Saito,
Neurosci. Lett., 1999, 266(1), 13–16.
162 S. Lutsenko, N. L. Barnes, M. Y. Bartee and O. Y. Dmitriev,
Physiol. Rev., 2007, 87(3), 1011–1046.
163 S. Lutsenko, E. S. LeShane and U. Shinde, Arch. Biochem.
Biophys., 2007, 463(2), 134–148.
164 A. N. Barry, U. Shinde and S. Lutsenko, JBIC, J. Biol. Inorg.
Chem., 2010, 15(1), 47–59.
165 A. A. Gybina and J. R. Prohaska, Genes Nutr., 2006, 1(1), 51–59.
166 R. El Meskini, L. B. Cline, B. A. Eipper and G. V. Ronnett, Dev.
Neurosci., 2005, 27(5), 333–348.
167 A. Bohlken, B. B. Cheung, J. L. Bell, J. Koach, S. Smith,
E. Sekyere, W. Thomas, M. Norris, M. Haber, D. B. Lovejoy,
D. R. Richardson and G. M. Marshall, Br. J. Cancer, 2009,
100(1), 96–105.
168 X. Li, S. Chen, Q. Wang, D. J. Zack, S. H. Snyder and J. Borjigin,
Proc. Natl. Acad. Sci. U. S. A., 1998, 95(4), 1876–1881.
169 M. Lovicu, M. B. Lepori, S. Incollu, V. Dessi, A. Zappu, R. Iorio,
M. D’Ambrosi, M. T. Pellecchia, P. Barone, G. Maggiore, S. De
Virgiliis, A. Cao and G. Loudianos, Genetic Testing and
Molecular Biomarkers, 2009, 13(2), 185–191.
170 G. Loudianos, M. Lovicu, V. Dessi, M. Tzetis, E. Kanavakis,
L. Zancan, L. Zelante, C. Galvez-Galvez and A. Cao, Hum.
Mutat., 2002, 20(4), 260–266.
171 A. B. Shah, I. Chernov, H. T. Zhang, B. M. Ross, K. Das,
S. Lutsenko, E. Parano, L. Pavone, O. Evgrafov, I. A. Ivanova-
Smolenskaya, G. Anneren, K. Westermark, F. H. Urrutia,
G. K. Penchaszadeh, I. Sternlieb, I. H. Scheinberg, T. C. Gilliam
and K. Petrukhin, Am. J. Hum. Genet., 1997, 61(2), 317–328.
172 K. Petrukhin, S. Lutsenko, I. Chernov, B. M. Ross, J. H. Kaplan
and T. C. Gilliam, Hum. Mol. Genet., 1994, 3(9), 1647–1656.
173 M. C. Reddy, S. Majumdar and E. D. Harris, Biochem. J., 2000,
350(3), 855–863.
174 L. B. Moller, Z. Tumer, C. Lund, C. Petersen, T. Cole,
R. Hanusch, J. Seidel, L. R. Jensen and N. Horn, Am. J. Hum.
Genet., 2000, 66(4), 1211–1220.
175 M. Qi and P. H. Byers, Hum. Mol. Genet., 1998, 7(3), 465–469.
176 S. G. Kaler, L. K. Gallo, V. K. Proud, A. K. Percy, Y. Mark,
N. A. Segal, D. S. Goldstein, C. S. Holmes and W. A. Gahl, Nat.
Genet., 1994, 8(2), 195–202.
177 S. La Fontaine and J. F. Mercer, Arch. Biochem. Biophys., 2007,
463(2), 149–167.
178 B. Hardman, A. Michalczyk, M. Greenough, J. Camakaris,
J. Mercer and L. Ackland, Cell. Physiol. Biochem., 2007, 20(6),
1073–1084.
179 L. Nyasae, R. Bustos, L. Braiterman, B. Eipper and A. Hubbard,
Am. J. Physiol.: Gastrointest. Liver Physiol., 2006, 292(4),
G1181–1194.
180 M. Y. Bartee and S. Lutsenko, BioMetals, 2007, 20(3–4),
627–637.
181 Y. Guo, L. Nyasae, L. T. Braiterman and A. L. Hubbard,
Am. J. Physiol.: Gastrointest. Liver Physiol., 2005, 289(5),
G904–916.
182 H. Roelofsen, H. Wolters, M. J. Van Luyn, N. Miura, F. Kuipers
and R. J. Vonk, Gastroenterology, 2000, 119(3), 782–793.
183 L. Braiterman, L. Nyasae, Y. Guo, R. Bustos, S. Lutsenko and
A. Hubbard, Am. J. Physiol.: Gastrointest. Liver Physiol., 2008,
296(2), G433–444.
184 C. Lane, M. J. Petris, A. Benmerah, M. Greenough and
J. Camakaris, BioMetals, 2004, 17(1), 87–98.
185 C. Cobbold, J. Coventry, S. Ponnambalam and A. P. Monaco,
Hum. Mol. Genet., 2003, 12(13), 1523–1533.
186 M. J. Petris and J. F. Mercer, Hum. Mol. Genet., 1999, 8(11),
2107–2115.
187 M. J. Francis, E. E. Jones, E. R. Levy, R. L. Martin,
S. Ponnambalam and A. P. Monaco, J. Cell. Sci., 1999,
112(Pt 11), 1721–1732.
188 M. J. Petris, J. Camakaris, M. Greenough, S. LaFontaine and
J. F. Mercer, Hum. Mol. Genet., 1998, 7(13), 2063–2071.
189 C. M. Lim, M. A. Cater, J. F. Mercer and S. La Fontaine, J. Biol.
Chem., 2006, 281(20), 14006–14014.
190 S. E. Stephenson, D. Dubach, C. M. Lim, J. F. Mercer and S. La
Fontaine, J. Biol. Chem., 2005, 280(39), 33270–33279.
191 W. C. Singleton, K. T. McInnes, M. A. Cater, W. R. Winnall,
R. McKirdy, Y. Yu, P. E. Taylor, B. X. Ke, D. R. Richardson,
J. F. Mercer and S. La Fontaine, J. Biol. Chem..
192 A. Ibricevic, S. L. Brody, W. J. Youngs and C. L. Cannon,
Toxicol. Appl. Pharmacol., 2010, 243(3), 315–322.
193 N. Barnes, M. Y. Bartee, L. Braiterman, A. Gupta, V. Ustiyan,
V. Zuzel, J. H. Kaplan, A. L. Hubbard and S. Lutsenko, Traffic,
2009, 10(6), 767–779.
194 L. S. Mangala, V. Zuzel, R. Schmandt, E. S. Leshane,
J. B. Halder, G. N. Armaiz-Pena, W. A. Spannuth, T. Tanaka,
M. M. Shahzad, Y. G. Lin, A. M. Nick, C. G. Danes, J. W. Lee,
N. B. Jennings, P. E. Vivas-Mejia, J. K. Wolf, R. L. Coleman,
Z. H. Siddik, G. Lopez-Berestein, S. Lutsenko and A. K. Sood,
Clin. Cancer Res., 2009, 15(11), 3770–3780.
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