Regulation of tyrosinase trafficking and processing
by presenilins: Partial loss of function by familial
Alzheimer’s disease mutation
Runsheng Wang*, Phuong Tang*†, Pei Wang*‡§, Raymond E. Boissy¶, and Hui Zheng*†‡?
*Huffington Center on Aging and Departments of†Molecular and Human Genetics and‡Molecular and Cellular Biology, Baylor College of Medicine,
Houston, TX 77030; and¶Department of Dermatology, University of Cincinnati College of Medicine, Cincinnati, OH 45267
Communicated by Xiaodong Wang, University of Texas Southwestern Medical Center, Dallas, TX, November 10, 2005 (received for review October 25, 2005)
Presenilins (PS) are required for ?-secretase cleavage of multiple
type I membrane proteins including the amyloid precursor protein
and Notch and also have been implicated in regulating intracellular
protein trafficking and turnover. Using genetic and pharmacolog-
ical approaches, we reveal here a unique function of PS in the
pigmentation of retinal pigment epithelium and epidermal mela-
nocytes. PS deficiency leads to aberrant accumulation of tyrosinase
(Tyr)-containing 50-nm post-Golgi vesicles that are normally des-
tined to melanosomes. This trafficking is ?-secretase-dependent,
and abnormal localization of Tyr in the absence of PS is accompa-
nied by the simultaneous accumulation of its C-terminal fragment.
Furthermore, we show that the PS1M146V familial Alzheimer’s
disease mutation exhibits a partial loss-of-function in pigment
synthesis. Our results identify Tyr and related proteins as physio-
logical substrates of PS and link ?-secretase activity with intracel-
lular protein transport.
?-secretase ? pigmentation ? melanocyte ? knock-out ? knock-in
A?40 and A?42, peptides that constitute the principal components
of the ?-amyloid plaques characteristic of Alzheimer’s disease (1).
Mutations in PS1 and PS2 lead to autosomal dominant inheritance
of familial Alzheimer’s disease (FAD). More than 100 mutations
have been identified in PS1 and PS2, and they spread throughout
the entire molecules. These mutations are known to alter the
regulated intramembrane proteolysis (4), PS are required for
processing and signaling of Notch (5), and this pathway likely
contributes to various PS developmental activities (6).
The PS-dependent ?-secretase activity requires the formation of
a high-molecular-weight complex containing nicastrin, Aph1,
and Pen2 (6). The active complex is assembled in a sequential and
interdependent manner through the endoplasmic reticulum and
Golgi compartments and requires posttranslational modifications.
The ?-secretase cleavage is preceded by extracellular processing
and exhibits relaxed sequence specificity. Nicastrin recently was
shown to function as the receptor for the ?-secretase substrates (7).
Besides APP and Notch, PS has been implicated in the processing
of a growing list of type I membrane proteins (reviewed in ref. 8).
However, with the exception of Notch, the physiological signifi-
cance of these proteolytic events remains speculative.
In addition to its ?-secretase activity, PS has been documented
to regulate intracellular protein trafficking (9). PS1 deficiency has
been reported to result in aberrant trafficking and maturation of
APP (10, 11), Notch (12, 13), tyrosine kinase receptor TrkB (14),
et al. (20) and Wilson et al. (21) implicated a role of PS1 in the
turnover of telencephalin and ?- and ?-synucleins, respectively.
They showed that these molecules accumulate in degradative
ammalian presenilins (PS) consists of two homologous pro-
by ?-secretase inhibitor treatment (20, 21). Thus, this activity is
likely mediated through a ?-secretase independent mechanism.
Similar to APP and Notch, tyrosinase (Tyr) (monophenyl mo-
nooxidase, EC 126.96.36.199), along with two Tyr-related proteins,
Tyr-related protein-1 (Tyrp1) and 3,4-dihydroxyphenylalanine
(DOPA) chrome tautomerase?Tyr-related protein-2 (DCT?
Tyrp2), are type I membrane proteins specialized in pigment
synthesis (22). Tyr catalyzes the conversion of tyrosine to DOPA,
which is an essential and rate-limiting step in melanin synthesis.
and DCT?Tyrp2 result in the synthesis of eumelanin (23). These
reactions are conducted within the melanosomes of vertebrate
and epidermal melanocytes. Melanosomes are endosomal?
lysosomal-related organelles. They progress through a series of
coated endosomes lacking pigmentation, to stages II, III, and IV
melanosomes, which are striated with increased melanin contents
(24). Under normal conditions, Tyr is glycosylated through the
secretory pathway, budded off from the trans-Golgi network as
50-nm vesicles, and transported to melanosomes to undergo pig-
ment synthesis and further maturation (25, 26).
pigment synthesis. Mutations in tyrosinase (Tyr) and Tyrp1 result in
retention of immature Tyr in the endoplasmic reticulum and are
associated with oculocutaneous albinism type 1 and 3, respectively
(27). In addition to the melanocyte-specific proteins, numerous
other molecules, including both highly conserved adaptor proteins,
soluble N-ethylmaleimide sensitive factor attachment protein re-
ceptors, and small GTPases of the Ras superfamily, as well as
unique vertebrate-specific protein complexes termed biogenesis of
lysosome-related organelles complexes, are known to regulate Tyr
trafficking and pigment processes (reviewed in refs. 24 and 28). In
addition to melanosomes, these proteins also play important roles
in regulating other endosomal?lysosomal-related organelles that
are essential for immune or neuronal functions.
In this work, we reveal an indispensable role of PS in Tyr
trafficking and pigmentation by a ?-secretase-dependent mecha-
FAD mutation in vivo.
Materials and Methods
Abs and Inhibitors. Rabbit polyclonal antisera against Tyrp1
(?PEP1), DCT?Tyrp2 (?PEP8), and C- and N-terminal Tyr
Conflict of interest statement: No conflicts declared.
Abbreviations: PS, presenilin; APP, amyloid precursor protein; FAD, familial Alzheimer’s
disease; RPE, retinal pigment epithelium; DOPA, 3,4-dihydroxyphenylalanine; Tyr, tyrosi-
AP3, adaptor protein-3; HPS, Hermansky–Pudlak syndrome; CTF, C-terminal fragment;
DAPT, N-[N-(3,5-difluorophenacetyl-L-alanyl)]-S-phenylglycine t-butyl ester.
§Present address: Department of Developmental Biology, Stanford University, Stanford,
?To whom correspondence should be addressed. E-mail: email@example.com.
© 2005 by The National Academy of Sciences of the USA
January 10, 2006 ?
vol. 103 ?
no. 2 ?
?PEP-7 and ?PEP-5, respectively, were generously provided by
V. J. Hearing (National Institutes of Health, Bethesda). The PS1
was obtained from Dako. The ?-secretase inhibitor N-[N-(3,5-
difluorophenacetyl-L-alanyl)]-S-phenylglycine t-butyl ester
(DAPT) was purchased from Calbiochem. DOPA was available
Mice and Cell Culture. The PS1 and PS2 knock-out, PS1M146V
were backcrossed onto C57BL?6J background to yield animals of
black coat colors. Primary melanocytes were established from
neonatal skins of C57BL?6J mice. Skins were incubated in 0.25%
trypsin for 2 h at 37°C. The epidermis was separated from dermis
layer and vortexed for 30 sec to separate the melanocytes from the
epidermal cell layer. The melanocytes were plated in melanocyte
growth medium, which consisted of M154 basal medium (Cascade
Biologicals, Portland, OR) supplemented with 4% heat-inactivated
melanocyte growth supplement (Cascade Biologicals). All cultures
The ?-secretase inhibitor treatment began 1 d after plating of the
melanocytes, and the medium was changed every 3–4 d. Concen-
tration at 500 nM was used unless otherwise indicated. Immuno-
staining and Western blotting was carried out 2 d after treatment,
whereas EM and melanin content assay was performed after 2 wk
of treatment. DOPA histochemistry was done by incubating the
fixed cells with 0.1% solution of DOPA for 2 h twice at 37°C.
Melanin Assay. Cells were rinsed with PBS and lysed with an
extraction buffer (50 mM Tris, pH 7.5?2 mM EDTA?150 mM
NaCl?1% Triton X-100) containing protease inhibitor mixture at
4°C. Cell pellet was collected and assayed for melanin content by
rinsing twice with ethanol–ether (1:1), dissolving in 2 M NaOH
solution, and measuring for A at 490 nm.
Histology and Immunofluorescence. Histology and immunohisto-
chemical staining of mouse tissues were performed as described
in ref. 33. Immunohistochemical staining was performed on
paraffin-fixed sections. Sections were blocked with 5% goat
serum, incubated with primary anti-PS1 Ab AB14 (1:250 dilu-
tion) at 4°C overnight, washed in PBS, incubated with 1?1,000
Alexa Fluor-594-conjugated secondary Ab (Molecular Probes)
for 1 h at room temperature, washed in PBS, and mounted in
glycerol?PBS. Digital images were obtained with a Zeiss con-
focal microscope (Axioskop 2).
Mouse melanocyte monolayers were seeded onto gelatin-coated
six-well Lab-Tek chamber slides (Nunc). The cells were fixed for 10
min in 2% formaldehyde in PBS. Cells were incubated for 1 h at
room temperature in a mixture of mouse monoclonal and rabbit
polyclonal Abs (diluted in 5% goat serum in PBS). After the
times for 5 min each. Cells then were incubated for 30 min with
Oregon Green 488 goat anti-rabbit IgG, and images were captured
by the confocal microscope (Zeiss).
Western Blotting. Protein extracts were separated by SDS?
PAGE and electroblotted onto 0.2-?m nitrocellulose mem-
branes (Schleicher & Schuell). Nonspecific sites were blocked
by incubation in 5% (wt?vol) nonfat dry milk and 0.1% Tween
20. The membranes were incubated with primary antiserum
followed by horseradish peroxidase-conjugated secondary Abs
and visualized with the enhanced chemiluminescence (ECL)
system (Amersham Pharmacia Biotech).
EM. Melanocytes were seeded onto gelatin-coated eight-well
Lab-Tek chamber slides (Nunc). The cells were fixed in wells
with one-half-strength Karnovsky’s fixative in 0.2 M sodium
cacodylate buffer (pH 7.2) for 30 min at room temperature. Cells
then were postfixed with 1% osmium tetroxide containing 1.5%
potassium ferrocyanide for 30 min, washed, stained en bloc with
0.5% uranyl acetate for 30 min, dehydrated, and embedded in
Eponate 12. Cells were sectioned on an MT 6000-XL ultrami-
crotome (RMC, Tucson, AZ), stained with aqueous solutions of
uranyl acetate (2%) and lead citrate (0.3%) for 15 min each, and
then viewed and photographed with a Hitachi transmission
electron microscope. All tissue-processing supplies were pur-
chased from Ted Pella (Tustin, CA).
PS Are Required for Pigmentation of Retinal Pigment Epithelium and
Cutaneous Melanocytes. We developed a unique CNS-restricted PS
‘‘rescue’’ system in which the early lethal phenotype of the PS1 and
PS2 double-null embryos could be partially rescued by neuronal
expression of the human PS1 transgene (33). These rescued em-
bryos could be readily identified because their eyes lacked pigmen-
tation (Fig. 1A). Histological analysis of the eye at embryonic day
13.5 showed that all cellular structures, in particular the RPE cells,
could be identified, and the morphology appeared to be normal
(Fig. 1B). This result suggests that the defect lies specifically in
melanin synthesis within the RPE. Immunohistochemical staining
by using an anti-PS1 Ab documented that although PS1 protein
reduced eye pigmentation in a CNS-restricted PS rescue embryo (Mutant,
and PS mutant (b) eyes, showing virtually absent pigment granules in the PS
mutant RPE (marked by arrowheads). (C) Immunohistochemical staining by
using the anti-PS1 N-terminal Ab (AB14) documented the presence of PS1
(b). Embryonic stage 13.5 was used for all of the analysis. (Scale bar: 20 ?m.)
Loss of PS leads to defective pigmentation in RPE. (A) Dramatically
www.pnas.org?cgi?doi?10.1073?pnas.0509822102 Wang et al.
could be detected in the littermate control, it was greatly reduced
is caused by the loss of PS, and the mutant is termed PS-null in the
context of RPE.
In addition to RPE, neural-crest-derived pigment cells also
comprise the cutaneous melanocytes of the skin and hair.
Because both cell types share common melanin synthesis path-
way, we decided to investigate whether PS is also required for the
pigmentation of mouse cutaneous melanocytes by using specific
PS inhibitors (34, 35). We cultured primary melanocytes from
newborn WT C57BL?6J mouse skin and allowed the culture to
mature and become pigmented (Fig. 2Aa). Addition of the
?-secretase inhibitor DAPT potently blocked the pigmentation
(Fig. 2Ab). The same result was obtained by using a different
?-secretase inhibitor, Compound 1 (36) (data not shown). This
effect was dose-dependent, and 0.25 ?m of DAPT resulted in
almost complete blockage of pigment synthesis (Fig. 2B). These
results establish an essential role of PS in the pigmentation of
both RPE and melanocytes.
Of significance, histochemical staining by using the Tyr substrate
DOPA efficiently induced pigment reaction product in inhibitor-
treated cultures (Fig. 2Ad), demonstrating that catalytically func-
tional Tyr was present and that the overt cellular morphology and
viability was not affected by ?-secretase inhibition. Immunohisto-
DCT?Tyrp2, the three principal enzymes in the melanin synthesis
PS Inactivation Leads to Abnormal Accumulation of Tyr-Containing
Post-Golgi Vesicles. Ultrastructural analysis was performed to de-
termine the effect of PS on melanosome structures (Fig. 3).
Examination of RPE showed that, compared with the control in
which melanosomes were mostly present in mature forms (Fig.
3Aa), the PS-null melanosomes were predominately at stages I and
II (Fig. 3Ab). They were less pigmented and smaller in size.
Nevertheless, the structure of the melanosomes appeared normal
exhibiting melanofilament matrix formation. Similar to that of
RPE, whereas vehicle-treated cultured melanocytes contained pre-
dominantly mature melanosomes (Fig. 3Ba), the ?-secretase inhib-
itor-treated melanocytes contained largely early stage (I and II)
melanosomes with apparently normal morphology (Fig. 3Bb), the
latter exhibited melanin reaction product upon DOPA histochem-
istry (Fig. 4Ba, solid arrow). Immunostaining and Western blot
analysis of the melanosome structural protein Pmel 17 showed
similar patterns and levels of expression in both control and
DAPT-treated cells (data not shown), further supporting the
the overall melanosomal protein levels were not significantly
changed by ?-secretase inhibitor treatment (discussed in Fig. 5),
these data indicate a possible role of PS in targeting these proteins
Tyr is synthesized and matured through the secretory pathway
and buds off the trans-Golgi network in 50-nm coated vesicles,
presumably en route to melanosomes in the vicinity. DOPA reac-
tion followed by EM analysis allows identification of these Tyr-
organelles by DOPA histochemistry revealed that, in vehicle-
treated primary melanocytes, DOPA histochemistry was restricted
to Golgi areas and to post-Golgi vesicles in the vicinity of trans-
the cell body and dendrites (data not shown). In ?-secretase
inhibitor-treated culture, however, there were numerous DOPA-
positive, 50-nm vesicles present peripheral to the Golgi zone (Fig.
4Ba, arrowheads) and throughout the cell body and dendrites (Fig.
Control primary melanocytes became pigmented after 2 wk of culturing. (b)
Complete blockage of pigment synthesis by 500 nM of DAPT. (c and d)
Postfixation treatment of DOPA showed reaction product in both the control
(c) and DAPT-treated culture (d), demonstrating that Tyr remained active in
the presence of the inhibitor. (B) Quantification of melanin content (?g?mg
of protein) documented dose-dependent inhibition of melanin synthesis by
DAPT.*, P ? 0.001;**, P ? 0.0001 (Student’s t test). (Scale bars: 20 ?m.)
Effect of ?-secretase inhibitor treatment on melanin synthesis. (Aa)
Representative EM images of control (a) and PS-null mutant (b) RPE. (B)
Representative EM images of control (a) and DAPT-treated (b) melanocytes.
I–IV denote melanosomes of various maturation stages. Genetic ablation and
?-secretase inhibitor treatment both resulted in the formation of predomi-
nately immature (stages I and II) melanosomes as opposed to mature (stages
III and IV) melanosomes prevalent in control melanocytes. (Scale bar: 0.5 ?m.)
EM analysis of melanosome structures in RPE and melanocytes. (A)
Wang et al.
January 10, 2006 ?
vol. 103 ?
no. 2 ?
50-nm vesicles, which are normally destined to melanosomes, is
likely the primary cause for the impaired pigmentation.
Inactivation of PS Leads to the Accumulation of Tyr, Tyrp1, and
DCT?Tyrp2 C-Terminal Fragments (CTFs). PS have been shown to
cleave a large number of type I transmembrane proteins based
on the evidence that loss of PS results in the accumulation of
their CTFs. Western blot analysis using Abs against the C-
terminal end of Tyr, Tyrp1, and DCT?Tyrp2 documented that,
although the 70-kDa full-length Tyr, Tyrp1, and DCT?Tyrp2
were expressed in vehicle- and ?-secretase inhibitor-treated
cultures, an additional CTF of 7 kDa was detected for all three
proteins only in inhibitor-treated samples (Fig. 5A). The fact that
(i) this length represents a combination of the intracellular
domain (?28 aa), the transmembrane domain (?22 aa), and
15–20 aa beyond the transmembrane domain of each of these
proteins (27), (ii) an Ab against the N-terminal sequence of Tyr
failed to recognize this fragment (Fig. 5B), and (iii) transfection
of a Tyr expression vector resulted in the production of the
7-kDa fragment only in PS1?/?cells, but not in PS1?/?cells (Fig.
5C), support the notion that these fragments are bona fide CTFs
analogous to those of APP and other putative PS targets, and
that an enzyme(s) with the characteristic of membrane shedding
is responsible for extracellular processing of these molecules.
Time course studies showed that, while the full-length proteins
remained constant, the CTFs of Tyr, Tyrp1, and DCT?Tyrp2
could be detected after 1 h of DAPT treatment and accumulate
rapidly overtime (Fig. 5D).
histochemistry identified Golgi tubules (G) in control melanocytes. The 50-nm
vesicles were infrequent and confined to the Golgi area. (B) In DAPT-treated
bar: 0.5 ?m.)
Western blotting using anti-C-terminal Tyr (Tyr-C), Tyrp1, and DCT?Tyrp2
(Tyrp2) Abs in the presence (?) or absence (?) of DAPT. A 7-kDa CTF can be
detected with each of the three C-terminal Abs in inhibitor-treated (?)
samples but not in vehicle-treated controls (?). (B) Western blotting by using
an anti-N-terminal Tyr Ab (Tyr-N). Only the 70-kDa full-length protein was
present. (C) Transfection of a Tyr expression vector into WT (PS1?/?) mouse
embryonic fibroblasts led to the expression of Tyr full-length protein at 70
appearance of both the full-length protein and a 7-kDa CTF, demonstrating a
direct processing of Tyr by PS1. Empty-vector transfection (Tyr?) was used as
a negative control. Hybridization with an anti-tubulin Ab (Tubulin) was used
as loading controls. (D) Time course analysis of Tyr, Tryp1, and DCT?Tryp2
used as loading control.
Western blot analysis of Tyr, Tyrp1, and DCT?Tyrp2 proteins. (A)
www.pnas.org?cgi?doi?10.1073?pnas.0509822102Wang et al.
PS1 FAD Mutation Exhibits Partial Loss of Function in Melanin Syn-
thesis. Having established a definitive role of PS in regulating
melanosomal protein and pigment processes, we next investigated
the effect of PS1 FAD mutations by using the PS1M146V knock-in
expressed in a completely physiological environment. Our previous
studies documented impaired contextual fear learning and adult
only be detected on a sensitized genetic background, i.e., when
endogenous WT PS1 allele was removed (37). Based on this
on PS2-null (PS2?/?) background. Specifically, crossing of
PS1M146V heterozygous (PS1M146V/?) with PS1?/?mice gener-
ated animals that were PS1?/?(???), PS1?/?(???), or
PS1M146V/?(M146V??). Replacing the WT allele with the
PS1M146V mutant allele led to a dramatic reduction in the coat
between PS1?/?and PS1?/?mice could not be readily recognized
on the coat (data not shown), mild hypopigmentation could be
observed in PS1?/?(???) tail skin when compared with that of
PS1?/?(???), and it was further reduced in PS1M146V/?
(M146V??) samples (Fig. 6B). This assessment was corroborated
by measuring the melanin content in melanocyte cultures derived
from these animals (Fig. 6C), which revealed lowered melanin
production by either removing the WT allele (compare ??? with
??? with M146V??). This result is consistent with our inhibitor
studies documenting a dose-dependent inhibition of melanin syn-
thesis by the ?-secretase inhibitor (Fig. 2B). This finding thus
establishes a partial loss-of-function activity of the PS1M146V
mutation in melanin synthesis in vivo.
The role of PS in regulated intramembrane proteolysis has been
of PS in protein trafficking remains speculative. The melanosomes
offer an attractive system to study intracellular protein transport
because all organelles with catalytically active Tyr can be marked
through the DOPA reaction and visualized by EM. Using this
system and taking advantage of our PS rescue model and the
availability of highly potent PS ?-secretase inhibitors, we report
here a unique function of PS in pigment synthesis in both RPE and
cutaneous melanocytes. We demonstrate that absence of PS or
application of PS inhibitor leads to mislocalization of post-Golgi
Tyr-containing vesicles, thus establishing a functional role of PS in
mediating Tyr trafficking in vivo. Our work identifies Tyr, Tyrp1,
and DCT?Tyrp2 as physiological substrates for PS and links the
intracellular transport with ?-secretase activity because ?-secretase
inhibition results in simultaneous mistrafficking of Tyr and related
proteins and accumulation of their CTFs. Although current studies
do not address the cause–effect relationship between the secretase
activity and Tyr transport, mislocalization of Tyr in post-Golgi
vesicles most likely contributes directly to the pigment defect. We
further document that the PS1M146V FAD mutation exhibits a
partial loss of function in melanin synthesis and, by extension, Tyr
trafficking, in vivo.
Tyr, Tyrp1, and Dct?Tyrp2 are type I membrane glycoproteins.
They undergo synthesis and maturation through endoplasmic re-
ticulum and Golgi and are ultimately transported to premelano-
somes via 50-nm post-Golgi vesicles. Although the initial processes
are similar, they exhibit distinct differences in subsequent intracel-
lular regulations, likely due to the differences in their targeting
signals in the carboxyl domains of these molecules. Adaptor pro-
teins have been linked to these processes (reviewed in ref. 38).
Specifically, adaptor protein-3 (AP3) deletion leads to hypopig-
of Tyr with AP3 has been shown to be required for its targeting to
melanosomes (38). On the contrary, Tyrp1 may be transported by
an AP3-independent but AP1-dependent mechanism (39, 40).
Although both Tyr and Tyrp1 contain the dileucine motif, which is
required for correct routing of the two molecules (41, 42), there is
no dileucine sequence in DCT?Tyrp2 C terminus, and a tyrosine-
based signal may be used for its trafficking to melanosomes instead
(43). Because all three molecules are affected in PS-null melano-
cytes, PS appears to act on these substrates at a common step
independent of adapter proteins. Indeed, Western blot analysis
showed normal expression of AP3 subunits in PS-null melanocytes
(data not shown). The fact that the APP C terminus does not have
the dileucine motif and that PS was reported to affect APP
trafficking to plasma membrane or endocytosis not known to
mediate Tyr routing further corroborate with this notion (10, 11).
Further work is required to identify the mechanism where by PS
regulates the trafficking of these diverse type I membrane proteins.
Of interest, PS-null melanocytes resemble the pathology of
melanocytes in Hermansky–Pudlak syndrome-type 3 (HPS3),
where in the absence of the HPS3 gene product, 50-nm vesicles
carrying Tyr cargo are aberrantly distributed throughout the entire
cytoplasm and that all three Tyr-related enzymes are affected (44).
However, HPS3 protein was found to be normally expressed in
PS-null melanocytes, and immunohistochemical staining failed to
detect changes of PS in HPS3 melanocytes (not shown). Thus,
PS-mediated Tyr trafficking is likely independent of that of the
HPS3 gene product.
Our finding that PS inhibition leads to the accumulation of Tyr,
Tyrp1, and DCT?Tyrp2 CTFs identifies these molecules as endog-
enous substrates for PS. The question arises as to what could be the
physiological function of proteolytic processing of these enzymes.
This question becomes increasingly important in light of the large
number of putative PS substrates that have been identified. In this
regard, two competing views have been put forward for the
PS-dependent ?-secretase activity: Extending from Notch receptor
studies in which ?-secretase cleavage of Notch has been shown to
be required to liberate the Notch intracellular domain and to
activate its downstream signaling, other type I membranes have
were expressed on PS2-null background. (A) Reduced coat color in a repre-
sentative 2-mo-old PS1M146V/?(M146V??) mouse as compared with a litter-
mate PS1?/?(???) animal. (B and C) Pigment colors of the tail skin of ???,
???, and M146V?? mice (B), which correlates with the melanin levels mea-
sured from melanocyte cultures derived from these animals (C).*, P ? 0.001;
**, P ? 0.0001 (Student’s t test).
Reduced pigmentation by PS1M146V FAD mutation. All PS1 alleles
Wang et al.
January 10, 2006 ?
vol. 103 ?
no. 2 ?
been implicated to subject to similar PS-dependent intracellular
domain production and cellular signaling (reviewed in ref. 45).
Alternately, ?-secretase cleavage has been proposed to degrade
membrane proteins and possibly attenuate signaling pathways (8,
46). The well-established enzymatic activities of Tyr and related
proteins make them unlikely cell signaling molecules. Because the
C-terminal sequences of these proteins contain melanosomal tar-
geting signals (41–43), breakage of the CTFs by ?-secretase is
expected to disrupt their melanosomal localization and melanin
here to degrade these membrane proteins. As such, the accumu-
lation of CTFs of Tyr family of proteins in the absence of PS would
represent degradation intermediates, which otherwise may escape
detection due to PS-mediated proteolysis. Future studies are re-
quired to identify the cellular sites of CTF accumulation and to
Support for a function of PS in intracellular protein degrada-
tion also came from recent studies by Wilson et al. (21) and by
Esselens et al. (20), which documented delayed turnover and
subsequent accumulation of ?- and ?-synucleins and telencepha-
lin, respectively, in degradative organelles in PS1-null cells.
However, because these effects are not seen by ?-secretase
inhibitor treatment and because full-length proteins, not the
CTFs, accumulate, these are likely mediated through indepen-
In summary, we identify here a unique function of PS in melanin
synthesis and pigmentation. We present data that it is mediated by
its intracellular transport of post-Golgi Tyr-containing vesicles and
that this effect is ?-secretase dependent. We document a partial
loss-of-function activity by the PS1 M146V FAD mutation. These
findings raise the intriguing possibility that a compromised post-
Golgi vesicle transport may contribute to Alzheimer’s disease
We thank G. Martin and B. Sopher (University of Washington, Seattle)
for PS1M146V knock-in mice, R. Swank (Roswell Park Cancer Institute,
Buffalo, NY) for HPS mouse tissues, and A. Houghton (Memorial
Sloan–Kettering Cancer Center, New York) for the Tyr expression
vector. We thank V. J. Hearing (National Institutes of Health, Bethesda)
for the generous gifts of anti-Tyr, Tyrp1, and DCT?Tyrp2 and Pmel17
Abs; and H. Xu and P. Greenguard (The Rockefeller University, New
York) for the AB14 Ab. We thank Claire Haueter for expert EM
assistance. This work was supported by National Institutes of Health
Grants NS40039 (to H.Z.), AG20670 (to H.Z.), and AR45429 (to
R.E.B.) and by the Ellison Medical Foundation (H.Z.). R.W. and P.T.
are trainees of National Institutes of Health Training Grant T32
AG000183. H.Z. is a Zenith Award recipient from the Alzheimer’s
1. Annaert, W. & De Strooper, B. (2002) Annu. Rev. Cell Dev. Biol. 18, 25–51.
2. Siman, R., Reaume, A. G., Savage, M. J., Trusko, S., Lin, Y. G., Scott, R. W.
& Flood, D. G. (2000) J. Neurosci. 20, 8717–8726.
3. Moehlmann, T., Winkler, E., Xia, X., Edbauer, D., Murrell, J., Capell, A.,
Kaether, C., Zheng, H., Ghetti, B., Haass, C. & Steiner, H. (2002) Proc. Natl.
Acad. Sci. USA 99, 8025–8030.
4. Brown, M. S., Ye, J., Rawson, R. B. & Goldstein, J. L. (2000) Cell 100, 391–398.
5. De Strooper, B., Annaert, W., Cupers, P., Saftig, P., Craessaerts, K., Mumm,
J. S., Schroeter, E. H., Schrijvers, V., Wolfe, M. S., Ray, W. J., et al. (1999)
Nature 398, 518–522.
6. Selkoe, D. & Kopan, R. (2003) Annu. Rev. Neurosci. 26, 565–597.
C. E., III, Sudhof, T. & Yu, G. (2005) Cell 122, 435–447.
8. Kopan, R. & Ilagan, M. X. (2004) Nat. Rev. Mol. Cell Biol. 5, 499–504.
9. Sisodia, S. S. & St. George-Hyslop, P. H. (2002) Nat. Rev. Neurosci. 3, 281–290.
10. Kaether, C., Lammich, S., Edbauer, D., Ertl, M., Rietdorf, J., Capell, A.,
Steiner, H. & Haass, C. (2002) J. Cell Biol. 158, 551–561.
11. Cai, D., Leem, J. Y., Greenfield, J. P., Wang, P., Kim, B. S., Wang, R., Lopes,
K. O., Kim, S. H., Zheng, H., Greengard, P., et al. (2003) J. Biol. Chem. 278,
12. Levitan, D. & Greenwald, I. (1998) Development (Cambridge, U.K.) 125,
13. Guo, Y., Livne-Bar, I., Zhou, L. & Boulianne, G. L. (1999) J. Neurosci. 19,
14. Naruse, S., Thinakaran, G., Luo, J. J., Kusiak, J. W., Tomita, T., Iwatsubo, T.,
Qian, X., Ginty, D. D., Price, D. L., Borchelt, D. R., et al. (1998) Neuron 21,
15. Noll, E., Medina, M., Hartley, D., Zhou, J., Perrimon, N. & Kosik, K. S. (2000)
Dev. Biol. 227, 450–464.
P., Holmes, E., Liang, Y., Kawarai, T., et al. (1999) Nat. Med. 5, 164–169.
17. Chen, F., Tandon, A., Sanjo, N., Gu, Y. J., Hasegawa, H., Arawaka, S., Lee,
F. J., Ruan, X., Mastrangelo, P., Erdebil, S., et al. (2003) J. Biol. Chem. 278,
18. Herreman, A., Van Gassen, G., Bentahir, M., Nyabi, O., Craessaerts, K.,
Mueller, U., Annaert, W. & De Strooper, B. (2003) J. Cell Sci. 116, 1127–1136.
19. Leem, J. Y., Vijayan, S., Han, P., Cai, D., Machura, M., Lopes, K. O., Veselits,
M. L., Xu, H. & Thinakaran, G. (2002) J. Biol. Chem. 277, 19236–19240.
20. Esselens, C., Oorschot, V., Baert, V., Raemaekers, T., Spittaels, K., Serneels,
L., Zheng, H., Saftig, P., De Strooper, B., Klumperman, J., et al. (2004) J. Cell
Biol. 166, 1041–1054.
21. Wilson, C. A., Murphy, D. D., Giasson, B. I., Zhang, B., Trojanowski, J. Q. &
Lee, V. M. (2004) J. Cell Biol. 165, 335–346.
22. del Marmol, V. & Beermann, F. (1996) FEBS Lett. 381, 165–168.
23. Sturm, R. A., Teasdale, R. D. & Box, N. F. (2001) Gene 277, 49–62.
24. Marks, M. S. & Seabra, M. C. (2001) Nat. Rev. Mol. Cell Biol. 2, 738–748.
25. Maul, G. G. (1969) J. Ultrastruct. Res. 26, 163–176.
26. Maul, G. G. & Brumbaugh, J. A. (1971) J. Cell Biol. 48, 41–48.
27. Oetting, W. S. & King, R. A. (1999) Hum. Mutat. 13, 99–115.
28. Huizing, M., Boissy, R. E. & Gahl, W. A. (2002) Pigm. Cell. Res. 15, 405–419.
29. Xia, X., Qian, S., Soriano, S., Wu, Y., Fletcher, A. M., Wang, X.-J., Koo, E. H.,
Wu, X. & Zheng, H. (2001) Proc. Natl. Acad. Sci. USA 98, 10863–10868.
30. Wong, P. C., Zheng, H., Chen, H., Becher, M. W., Sirinathsinghji, D. J.,
Trumbauer, M. E., Chen, H. Y., Price, D. L., Van der Ploeg, L. H. & Sisodia,
S. S. (1997) Nature 387, 288–292.
31. Donoviel, D. B., Hadjantonakis, A. K., Ikeda, M., Zheng, H., St. George-
Hyslop, P. & Bernstein, A. (1999) Genes Dev. 13, 2801–2810.
32. Guo, Q., Fu, W., Sopher, B. L., Miller, M. W., Ware, C. B., Martin, G. M. &
Mattson, M. P. (1999) Nat. Med. 5, 101–106.
33. Wang, P., Pereira, F. A., Beasley, D. & Zheng, H. (2003) Development
(Cambridge, U.K.) 130, 5019–5029.
34. Li, Y. M., Xu, M., Lai, M. T., Huang, Q., Castro, J. L., DiMuzio-Mower, J.,
Harrison, T., Lellis, C., Nadin, A., Neduvelil, J. G., et al. (2000) Nature 405,
Development (Cambridge, U.K.) 130, 5031–5042.
36. Qyang, Y., Chambers, S. M., Wang, P., Xia, X., Chen, X., Goodell, M. A. &
Zheng, H. (2004) Biochemistry 43, 5352–5359.
37. Wang, R., Dineley, K. T., Sweatt, J. D. & Zheng, H. (2004) Neuroscience 126,
38. Robinson, M. S. & Bonifacino, J. S. (2001) Curr. Opin. Cell Biol. 13, 444–453.
39. Huizing, M., Sarangarajan, R., Strovel, E., Zhao, Y., Gahl, W. A. & Boissy,
R. E. (2001) Mol. Biol. Cell 12, 2075–2085.
40. Raposo, G., Tenza, D., Murphy, D. M., Berson, J. F. & Marks, M. S. (2001)
J. Cell Biol. 152, 809–824.
41. Vijayasaradhi, S., Xu, Y., Bouchard, B. & Houghton, A. N. (1995) J. Cell Biol.
42. Calvo, P. A., Frank, D. W., Bieler, B. M., Berson, J. F. & Marks, M. S. (1999)
J. Biol. Chem. 274, 12780–12789.
43. Simmen, T., Schmidt, A., Hunziker, W. & Beermann, F. (1999) J. Cell Sci. 112,
44. Boissy, R. E., Richmond, B., Huizing, M., Helip-Wooley, A., Zhao, Y.,
Koshoffer, A. & Gahl, W. A. (2005) Am. J. Pathol. 166, 231–240.
45. Fortini, M. E. (2002) Nat. Rev. Mol. Cell Biol. 3, 673–684.
46. Parent, A. T., Barnes, N. Y., Taniguchi, Y., Thinakaran, G. & Sisodia, S. S.
(2005) J. Neurosci. 25, 1540–1549.
www.pnas.org?cgi?doi?10.1073?pnas.0509822102 Wang et al.