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PLATELETS AND THROMBOPOIESIS
Mechanism of platelet dense granule biogenesis: study of cargo transport and
function of Rab32 and Rab38 in a model system
Andrea L. Ambrosio,1 Judith A. Boyle,1 and Santiago M. Di Pietro1
1Department
of Biochemistry and Molecular Biology, Colorado State University, Fort Collins, CO
Dense granules are important in platelet
aggregation to form a hemostatic plug as
evidenced by the increased bleeding time
in mice and humans with dense granule
deficiency. Dense granules also are targeted by antiplatelet agents because of
their role in thrombus formation. Therefore, the molecular understanding of the
dense granule and its biogenesis is of
vital importance. In this work, we establish a human megakaryocytic cell line
(MEG-01) as a model system for the study
of dense granule biogenesis using a variety of cell biology and biochemical approaches. Using this model system, we
determine the late endocytic origin of
these organelles by colocalization of the
internalized fluid phase marker dextran
with both mepacrine and transmembrane
dense granule proteins. By mistargeting
of mutant dense granule proteins, we
demonstrate that sorting signals recognized by adaptor protein-3 are necessary
for normal transport to dense granules.
Furthermore, we show that tissue-specific
Rab32 and Rab38 are crucial for the fusion of vesicles containing dense granule
cargo with the maturing organelle. This
work sheds light on the biogenesis of
dense granules at the molecular level and
opens the possibility of using this powerful model system for the investigation of
new components of the biogenesis machinery. (Blood. 2012;120(19):4072-4081)
Introduction
Platelets contribute to normal hemostasis by releasing their ␣ granule (AG) and dense granule (DG) components at sites of vascular
injury. DGs concentrate small molecules such as serotonin, ADP,
and calcium, and their involvement in hemostasis is evident in
patients presenting with bleeding disorders because of deficiency
of these granules.1,2 In contrast, DG biogenesis and secretion have
been identified as targets for antithrombotic drugs.3-5 Despite the
importance of DGs for human health, very little is known about
their biogenesis. DGs are synthesized in the bone marrow by
megakaryocytes (MKs). These cells are difficult to isolate, culture,
and manipulate, which explains this knowledge gap. Thus, the lack
of convenient systems to study DG formation at the cellular and
molecular level has been a major limitation, leaving the mechanism
involved in biogenesis unclear.
Unlike most secretory granules produced in other cell types,
DGs may not originate from the trans-Golgi network. Instead, the
biogenesis of DGs may involve a specialized biosynthetic mechanism that connects the secretory and the endocytic pathways6,7
(Figure 1A). Consistent with that notion, one study suggested a
possible multivesicular body (MVB)/late endosome origin for DGs
using granulophysin as a DG marker.8 However, granulophysin was
later discovered to be the same molecule as CD63/LAMP3 that localizes
to several organelles in addition to DGs.9 Therefore, the MVB/late
endosome origin of dense granules remains an open question.
Platelet DGs belong to a group of lysosome-related organelles
(LROs) that also includes melanosomes in melanocytes. Defects in
genes and proteins involved in LRO biogenesis have been identified in Hermansky-Pudlak syndrome (HPS) patients and animal
models of the disease that present with a combination of albinism
and prolonged bleeding because of abnormal melanosomes and
DGs.1 Examples of HPS proteins are adaptor protein-3 (AP-3) and
Rab38. AP-3 operates at early endosome–associated tubules where
it recognizes both tyrosine-based and dileucine-based sorting
signals in the cytosolic tail of transmembrane protein cargo and
packages them into transport vesicles destined to lysosomes and
melanosomes.10-13 It is possible that similar sorting signals and
AP-3–dependent mechanism operate in a transport pathway from
early endosome–associated tubules to maturing DGs (Figure 1A).
This hypothesis has not yet been tested experimentally.
Rab proteins are key regulators of vesicular trafficking that
mediate vesicle motility, tethering, and fusion within the secretory
and endocytic pathways.14 Mutation of Rab38 in rodent disease
models cause DG and melanosome deficiency.15,16 Rab38 and its
very close homolog Rab32 cooperate in melanosome biogenesis, at
least in part working through the AP-3 pathway.17,18 It is possible
that Rab32 also participates in DG biogenesis and that both Rabs
cooperate with AP-3 in the delivery of transmembrane proteins to
the maturing DG. However, the cellular location and molecular
function of Rab32 and Rab38 in DG biogenesis are unknown.
Here, we show that the megakaryocytic cell line MEG-01 is a
very good model system that recapitulates what is known about
DGs and allows various experimental approaches to study DG
biogenesis. Using this system, we established the MVB/late
endosome origin for the organelle. We demonstrate that DG
transmembrane proteins depend on tyrosine-based and dileucinebased sorting signals for normal transport to DGs. We also show
that Rab32 and Rab38 likely define a biosynthetic transport
pathway from early endosome–associated tubules to maturing DGs
Submitted March 30, 2012; accepted August 12, 2012. Prepublished online as
Blood First Edition paper, August 27, 2012; DOI 10.1182/blood-2012-04-420745.
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 USC section 1734.
There is an Inside Blood commentary on this article in this issue.
The online version of the article contains a data supplement.
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© 2012 by The American Society of Hematology
BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
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BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
and that they have a key role in vesicle tethering, fusion with the
maturing organelle, or both.
Methods
Antibodies
See Tables 1 and 2 for lists of primary and secondary antibodies used in
this study.
Cloning of VMAT2 and LAMP2 cDNAs
The cDNAs for VMAT2 and LAMP2A were amplified from total RNA of
MEG-01 cells by reverse transcriptase-PCR and subsequently cloned
in-frame into pEGFP and pmCherry.
Cell culture
MEG-01 cells were obtained from ATCC and cultured in RPMI-1640
supplemented with 10% (v/v) fetal bovine serum, 100 U/mL penicillin,
100 ␮g/mL streptomycin, and 0.3 ␮g/mL glutamine. Mouse bone marrow
MKs were isolated by a 4-step BSA density gradient followed by a
continuous Ficoll velocity gradient as described previously.19
MEG-01 cells were transfected using the Nucleofector electroporation
system (Lonza). For siRNA treatments, 2 sequential transfections were
performed on days 1 and 4; cells were analyzed on day 7. Oligonucleotides
(Sigma) used for siRNA are as follows: negative control (SIC001-10
NMOL), Rab32 (SASI_Hs02_00342400), Rab38 (SASI_Hs01_00247037).
For dextran uptake experiments, cells were incubated for 16 hours at
37°C in medium containing 250 ␮g/mL dextran Alexa Fluor 647 or Oregon
Green 488 BAPTA-1 dextran (Invitrogen) followed by a 4-hour chase period in
medium lacking dextran. Mepacrine was added to cells to a final concentration of
10␮M followed by a 5-minute incubation at 37°C before imaging.
MECHANISM OF PLATELET DENSE GRANULE BIOGENESIS
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Table 1. List of primary antibodies used in the study
Target
Host
Type
Company/Laboratory
Rab32
Rabbit
Polyclonal
Di Pietro
Rab38
Rabbit
Polyclonal
Di Pietro
AP-3 ␦ (SA4)
Mouse
Monoclonal
Peden11
LAMP2
Mouse
Monoclonal
Santa Cruz Biotechnology
PF-4
Goat
Polyclonal
Santa Cruz Biotechnology
MRP4 (IF)
Rat
Monoclonal
Alexis Biochemicals
MRP4 (IB)
Mouse
Monoclonal
Abnova
Clathrin
Mouse
Monoclonal
Abcam
vWF
Rabbit
Polyclonal
Dako
Rab7a
Rabbit
Monoclonal
Cell Signaling Technology
␣-tubulin
Mouse
Monoclonal
Sigma
IF indicates immunofluorescence; and IB, immunoblotting.
cient21 (MOC) of dual-color images processed with both Gaussian and
Laplacian 2-dimensional filters. MOC was used to determine the level of
colocalization between marker pairs that are both membrane-bound or both
contained inside the organelles (luminal markers). For colocalization of a
membrane-bound marker with a luminal marker, the percentage of structures containing both markers was determined.
Subcellular fractionation
A postnuclear supernatant was prepared by homogenizing MEG-01 cells
with a dounce homogenizer in buffer H (20mM Hepes pH 7.4) containing
0.32M sucrose and protease inhibitors followed by centrifugation for
20 minutes at 800g at 4°C. The postnuclear supernatant (250 ␮L) was
loaded onto a 12-mL linear sucrose gradient (10%-60%) in buffer H. The
sample was centrifuged at 113 000g for 6 hours in a SW41Ti rotor in an
L8-70M ultracentrifuge (Beckman Coulter) at 4°C. Fractions of 1 mL
were collected and used for immunoblotting, immunoprecipitation, ADP
determination, and both Alexa Fluor 647 and Oregon Green 488 BAPTA-1
dextran fluorescence intensity reading.
Electron microscopy
Cells grown on Aclar (Ted Pella) were subsequently fixed in 2% electron
microscopy grade glutaraldehyde in phosphate-buffered saline (PBS) for
45 minutes, washed in PBS, postfixed in 1% aqueous osmium tetroxide for
15 minutes, and then washed in distilled water (dH2O). The cells were
dehydrated in an ethanol series and flat embedded in LR White resin.
Groups of cells were cut out, mounted, and 90-nm sections were collected,
stained with 2% aqueous uranyl acetate for 15 minutes, poststained in
Reynold’s lead stain, and viewed on a 2000 transmission electron microscope (JEOL). High-pressure freezing (HPF) was performed as described
previously.20 For immunogold labeling, 90-nm sections were collected on
nickel slot grids, blocked with 20% goat serum in PBS, blotted, and then
incubated in the primary antibodies overnight at 4°C. Grids were washed in
PBS-Tween 20 and incubated in the secondary antibodies for 1.5 hours.
Grids were washed first in PBS-Tween 20 and then in PBS, fixed in 0.5%
glutaraldehyde in PBS, washed in dH2O, and dried. The grids were stained
with 2% aqueous uranyl acetate in 70% methanol/30% water for 7 minutes,
rinsed in 70% methanol/30% water, dried, and then poststained in
Reynold’s lead stain, rinsed in dH2O, and dried. Images were taken on a
2000 transmission electron microscope (JEOL).
Confocal fluorescence microscopy
For the live cell imaging experiments, cells were plated in glass-bottomed
35-mm dishes, and phorbol 12-myristate 13-acetate was added to a final
concentration of 10nM 24 hours before imaging. The samples were imaged
using a temperature-controlled chamber at 37°C and 5% CO2 on an IX81
spinning-disk confocal fluorescence microscope (Olympus). Fixation and
immunofluorescence staining were performed as described previously.13
Immunofluorescence microscopy samples were examined using the same
microscope utilized for the live cell fluorescence imaging experiments.
Fixed and live cell sample images were acquired and analyzed in Slidebook
5.0 software (Intelligent Imaging Innovations). The colocalization module
with auto threshold was used to determine the Manders’ overlap coeffi-
Biochemical procedures
For immunoblotting, proteins were fractionated on precast 4% to 20%
gradient SDS/polyacrylamide gels (Invitrogen) and transferred by electroblotting to polyvinylidene difluoride membranes. Membranes were incubated sequentially with blocking buffer, primary antibody, and horseradish
peroxidase–conjugated secondary antibody as described previously.22
Bound antibodies were detected using ECL Prime Western blotting reagent
(GE Healthcare). Immunoprecipitations were carried out using protein G
magnetic beads (Millipore) and 2 ␮g of the appropriate antibody. ADP was
determined by bioluminescence using the Enzylight ADP assay kit (BioAssay Systems). The fluorescence intensity of dextran Alexa Fluor 647 and
Oregon Green 488 BAPTA-1 dextran in the sucrose gradient samples was
measured using a microplate reader Victor3V (PerkinElmer Life and
Analytical Sciences).
Results
MEG-01 cells provide a very good model system to study DG
biogenesis
MEG-01 cells display the typical markers of differentiated megakaryocytes, generate AGs and DGs, produce platelet-like particles,
Table 2. List of secondary antibodies used in the study
Reactivity
Conjugation
Rat, rabbit, mouse Alexa-488, -546, -647
Goat
Company
Invitrogen
HRP
Invitrogen
Rabbit
12-nm gold
Jackson ImmunoResearch Laboratories
Mouse
18-nm gold
Jackson ImmunoResearch Laboratories
HRP
GE Healthcare
Rabbit, mouse
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AMBROSIO et al
BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
Figure 1. MEG-01 cells have DGs that can be studied
by different microscopy techniques. (A) Model depicting the protein traffic to DG. DG transmembrane proteins
(gold) follow the secretory pathway through the Golgi
complex to early endosomes where they are selectively
targeted to maturing DGs originated from MVBs.
(B) Thin-section transmission electron microscopy image
of a MEG-01 cell subjected to HPF, fixed or embedded
with glutaraldehyde-uranyl acetate-Lowicryl HM20. Original magnification, ⫻3900 (bar represents 500 nm).
(C) Spinning-disk confocal fluorescence microscopy images of a MEG-01 cell fixed and immunostained with
LAMP2 and MRP4 antibodies to label DGs
(MOC ⫽ 0.63 ⫾ 0.06, n ⫽ 5 cells). Bar represents 10 ␮m.
(D) Spinning-disk confocal fluorescence microscopy images of a live MEG-01 cell expressing the DG markers
VMAT2-Cherry and LAMP2-GFP (MOC ⫽ 0.58 ⫾ 0.03,
n ⫽ 10 cells). Bar represents 5 ␮m. PM indicates plasma
membrane; and IDG, immature dense granule.
and seem to resemble primary megakaryocytes better than other
cell lines.23-27 We corroborated MEG-01 cells and primary megakaryocytes isolated from mouse bone marrow express surface
proteins such as CD41 to a similar extent (data not shown). We then
carried out several experiments to confirm the presence of DGs and
various markers in MEG-01 cells. First, the cells were subjected to
HPF and processed for thin-section electron microscopy. Of the
different approaches tested, samples fixed or embedded with
glutaraldehyde-uranyl acetate-Lowicryl HM20 worked best for
overall preservation of membrane-bound structures. Based on the
morphology and content of internal dense material, MEG-01 cells
showed the presence of mature DGs together with a large number
of immature DGs and MVBs (Figure 1B and supplemental Figure
1, available on the Blood Web site; see the Supplemental Materials
link at the top of the online article). Quantitative analysis of
electron micrographs of 19 MEG-01 cells showed they contain
0.6 ⫾ 0.1 mature DG/10 ␮m2, 1.8 ⫾ 0.3 immature DG/10 ␮m2,
and 1.6 ⫾ 0.2 MVB/10 ␮m2, whereas a similar analysis of 18 primary megakaryocytes isolated from mouse bone marrow showed
2.5 ⫾ 0.4 mature DG/10 ␮m2, 0.5 ⫾ 0.1 immature DG/10 ␮m2,
and 1.0 ⫾ 0.2 MVB/10 ␮m2 (970 organelles counted). Second, the
DG integral membrane protein markers MRP4 (ADP transporter)
and LAMP228,29 were detected by immunofluorescence microscopy in fixed MEG-01 cells (Figure 1C and supplemental Figure
2). MRP4 and LAMP2 display very good colocalization
(MOC ⫽ 0.63 ⫾ 0.06) consistent with previous findings in platelet
DGs. Third, LAMP2 and VMAT2 (the putative DG serotonin
transporter)30 were amplified from MEG-01 mRNA; cloned into
pEGFP and pmCherry, respectively; and transfected into MEG-01
cells. Live cell spinning-disk confocal fluorescence microscopy
show LAMP2-GFP and VMAT2-Cherry localize to the same
structures (MOC ⫽ 0.58 ⫾ 0.03), indicating the tagged proteins
are correctly targeted to DGs (Figure 1D and supplemental Video
1). These levels of colocalization were significantly higher than
those obtained in control experiments using LAMP2 and peroxisomal markers (MOC ⫽ 0.08 ⫾ 0.01 or 0.09 ⫾ 0.01) or MRP4 and
LAMP2 with the AG marker VWF (MOC ⫽ 0.12 ⫾ 0.01 and
0.18 ⫾ 0.01, respectively; supplemental Figure 3). Fourth, live
MEG-01 cells expressing VMAT2-Cherry were incubated with the
DG lumen marker mepacrine and imaged by confocal fluorescence
microscopy (Figure 2A). Mepacrine and VMAT2-Cherry largely
colocalize in MEG-01 cells (95% ⫾ 2% of the mepacrine structures contain both markers), corroborating both the presence of
DGs and VMAT2-Cherry targeting to these organelles. Similar
results were obtained when using LAMP2-Cherry and mepacrine
(supplemental Figure 4B). Furthermore, MEG-01 cells dense
granules labeled with LAMP2-GFP fused with the plasma membrane on stimulation with 1 U/mL thrombin, showing MEG-01
cells produce functional dense granules (supplemental Figure 5).
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MECHANISM OF PLATELET DENSE GRANULE BIOGENESIS
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Figure 2. DGs originate from late endocytic structures. (A) DGs were labeled with the green fluorescent
dye mepacrine in live MEG-01 cells expressing VMAT2Cherry and visualized by spinning-disk confocal microscopy. The inset shows examples of colocalization between the 2 DG markers; 94% ⫾ 2% of 171 mepacrine
structures (7 cells) also contain VMAT-Cherry. (B) Live
MEG-01 cells were allowed to internalize the fluid phase
marker dextran Alexa Fluor 647, and DGs were subsequently labeled with mepacrine. Cells were observed by
spinning-disk confocal fluorescence microscopy.
Examples of structures containing both markers are
presented in the magnified inset (MOC ⫽ 0.43 ⫾ 0.04,
n ⫽ 6 cells). (C) Live MEG-01 cells expressing the DG
marker VMAT2-GFP were labeled with dextran Alexa
Fluor 647 and imaged by spinning-disk confocal fluorescence microscopy. Inset: Examples of VMAT-GFP presence in the limiting membrane of organelles containing
dextran Alexa Fluor 647 in the lumen; 66% ⫾ 8% of
187 structures containing fluorescent dextran (7 cells)
also contain VMAT-GFP. Bars represent 5 ␮m.
DGs have a late endocityc origin
It has been proposed that DGs,8 similar to AGs,31 originate from
MVBs. To test this idea, we used a fluid phase marker taken up by
endocytosis, fluorescent dextran, to label late endocytic structures.
In nonspecialized cell types, this marker remains in the lumen of
endocytic organelles through their progress from early/recycling
endosomes to MVBs/late endosomes and it finally accumulates in
lysosomes, the terminal organelle of the endocytic pathway.
Pulse-chase experiments in MEG-01 cells show fluorescent dextran
follows a similar early/recycling endosome to MVBs/late endosomes path in these specialized cells (supplemental Figure 6). We
reasoned that if DGs originate from MVBs, internalized fluorescent
dextran should accumulate in DGs in megakaryocytic cells (Figure
1A). We allowed MEG-01 cells to internalize fluorescent dextran,
subsequently labeled DGs with mepacrine, and analyzed them by
live cell fluorescence microscopy (Figure 2B). The colocalization
of dextran and mepacrine (MOC ⫽ 0.43 ⫾ 0.04) supports the idea
of DGs originating from MVBs. Interestingly, although dextran
labels the lumen of the DG in a rather uniform manner, mepacrine
often seems more concentrated in an internal region of the DG,
perhaps labeling the dense core (Figure 2B inset and supplemental
Video 2). In a separate series of experiments, MEG-01 cells
expressing VMAT2-GFP were labeled with internalized fluorescent
dextran (Figure 2C). VMAT2-GFP shows green “doughnut”shaped structures—the membrane of DGs imaged at their median
plane—filled with red dextran (66% ⫾ 8% of the red dextran
structures contain both markers), consistent with an MVB origin of
DGs. Similar results were obtained with LAMP2-GFP and internalized fluorescent dextran (supplemental Figure 4C).
Normal transport of membrane proteins to the DG depends on
sorting signals bound by AP-3
DG integral membrane proteins may depend on tyrosine-based
sorting signals, dileucine-based sorting signals, or both, in their
cytosolic tails for vesicular transport to the organelle. Both LAMP2
and VMAT2 have sequences conforming to the tyrosine- and
dileucine-based signal consensus, respectively. We subjected the
putative signals to site-directed mutagenesis and analyzed the
cellular localization of the mutant proteins compared with the
wild-type proteins. Both LAMP2 and VMAT2 signal mutants were
mistargeted to the plasma membrane, indicating a severe transport
defect (Figure 3A-D). This result indicates transport to DGs is
mediated by similar sorting signals as transport to melanosomes
and lysosomes that are recognized by AP-3. These results are
consistent with a model in which AP-3 is a key adaptor that
packages proteins in vesicles at early endosome–associated tubules
for subsequent transport to the maturing DGs (Figure 1A). Mutant
cargo is unable to bind to AP-3, accumulates in early endosomal
membranes, and leaks into the recycling pathway to the plasma
membrane (Figures 1A, 3A-D, and 7J). An alternative explanation
would be that LAMP2 and VMAT2 normally traffic through the
plasma membrane on their way to the DG in an AP-3–independent
pathway, but their mutant forms fail to be internalized by the
endocytic adaptor AP-2. To test that possibility, wild-type LAMP2
and VMAT2 were expressed in MEG-01 cells subjected to AP-2
knockdown. Neither cargo was observed at the plasma membrane (data
not shown), arguing against biosynthetic traffic through the plasma
membrane en route to DGs, thus supporting the AP-3 model.
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AMBROSIO et al
BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
Figure 3. Sorting signals bound by AP-3 are crucial
for the correct targeting of DG proteins in MEG-01
cells. AP-3 partially colocalizes with Rab38 in MKs and
MEG-01 cells. (A-B) A single mutation in the LAMP2
cytosolic tail sorting signal (Y/A LAMP2: YEQF into
AEQF) is sufficient to mistarget Y/A LAMP2-GFP to the
plasma membrane in MEG-01 cells. Live cells were
visualized by spinning-disk confocal fluorescence microscopy. (C-D) Similarly, mutation of the VMAT2 cytosolic tail
sorting signal (IL/AA VMAT2: EEKMAIL into EEKMAAA)
causes mistrafficking of the mutant protein to the
plasma membrane in MEG-01 cells. Live cells were
visualized by spinning-disk confocal fluorescence microscopy. (E-E⬘) Primary MKs were fixed and immunostained with antibodies against AP-3, Rab38, and clathrin, and imaged by spinning-disk confocal fluorescence
microscopy. (E⬘) Close-up view of individual structures
allows observation of colocalization of AP-3 and Rab38
(merge panel, MOC ⫽ 0.34 ⫾ 0.01, n ⫽ 7 cells), whereas
clathrin is also present in many of these structures
(Rab38 and clathrin MOC ⫽ 0.33 ⫾ 0.02, n ⫽ 7 cells).
(F-F⬘) MEG-01 cells were fixed and immunostained with
antibodies against AP-3, Rab38, and clathrin and imaged
by spinning-disk confocal fluorescence microscopy.
(F⬘) Similarly to the results obtained with MKs, AP-3 and
Rab38 colocalize in structures that in many cases contain
clathrin (Rab38 and AP-3 MOC ⫽ 0.32 ⫾ 0.02,
n ⫽ 4 cells; Rab38 and clathrin MOC ⫽ 0.33 ⫾ 0.03,
n ⫽ 4 cells). Bars represent 5 ␮m.
Rab32 and Rab38 partially colocalize with AP-3 in MKs and
MEG-01 cells
Rab32 and Rab38 may cooperate with AP-3 in a pathway to deliver
transmembrane proteins to DGs. We tested for colocalization of
these proteins in both MEG-01 cells and primary MKs. Cells were
fixed or permeabilized, and simultaneously immunostained with
antibodies to Rab38, AP-3, and the vesicle coat protein clathrin. As
shown in Figure 3E and F, Rab38 partially colocalizes with AP-3 in
both MEG-01 cells and primary MKs (MOC ⫽ 0.32 ⫾ 0.04 and
0.34 ⫾ 0.03, respectively). This result indicates Rab38 is present at
exit sites in early endosome–associated tubules, transport vesicles
that have already pinched off and are en route to fuse with maturing
DGs, or both. The presence of clathrin in many of the Rab38/AP-3
structures (MOC ⫽ 0.33 ⫾ 0.05 and 0.33 ⫾ 0.04 for MEG-01 and
MKs, respectively) is consistent with Rab38 being recruited during
vesicle budding or soon after it pinched off from the donor
organelle. The fact that a similar result was obtained with MKs and
MEG-01 cells further supports this cell line as an appropriate
model system for studying DG biogenesis. Similar results were
obtained for Rab32-stained primary MKs and MEG-01 cells
(supplemental Figure 7). These data are consistent with a role of
Rab32 in the biogenesis of DGs similar to that of Rab38.
Rab32 and Rab38 are predominantly present in immature DGs
To test for the presence of Rab32 and Rab38 in DGs, we
determined the localization of both Cherry-Rab38 and CherryRab32 in live MEG-01 cells labeled with mepacrine. As shown in
Figure 4A, Cherry-Rab38 presents a high degree of localization to
mepacrine labeled structures. Interestingly, although some structures contain intermediate levels of both Cherry-Rab38 and mepacrine, others show the highest amount of Cherry-Rab38 and low
amounts of mepacrine and vice versa (see supplemental Figure 8B
for quantification). Similar results were obtained using CherryRab32 (supplemental Figure 8). Mepacrine accumulates in DG and
mature DGs display maximal mepacrine fluorescence; immature
granules stain less strongly.32 Thus, the inverse correlation observed between the fluorescence intensity of each Rab versus
mepacrine indicates the Rabs are enriched in immature DGs. A
close look at MEG-01 cells expressing Cherry-Rab38 also reveals
small structures (100-200 nm in diffraction limited puncta) that do
not contain mepacrine and likely represent transport vesicles
(Figure 4C and supplemental Video 3).
To corroborate the presence of Rab38 in DGs, particularly in
immature organelles, we investigated the localization of GFPRab38 relative to LAMP2-Cherry as a DG marker. Again, we
detected significant colocalization and structures that show the
highest amount of the DG marker contain less GFP-Rab38 and vice
versa (Figure 4D-F; see supplemental Figure 8B for quantification).
Similarly, a reverse correlation was observed for the labeling
intensity of Rab32 and LAMP2 in MEG-01 cells (supplemental
Figure 8C), suggesting that both Rabs are present primarily in
immature DGs.
Thin-section immunogold electron microscopy of MEG-01
cells stained for endogenous LAMP2 and Rab32 revealed the
presence of both proteins in the same organelle, many of them
immature DGs based on their low amount of internal dense
material (Figure 4G-L). Specifically, 73% of the organelles labeled
with Rab32 also contain LAMP2. Rab32 is restricted to the limiting
membrane of the organelle, whereas LAMP2 was found both in the
limiting membrane (Figure 4G,I,J) and inside of the organelle
(Figure 4H,K,L). In addition, we observed LAMP2 associated with
internal vesicles of MVBs and immature DGs (supplemental
Figure 9).
Rab5 is a well-known marker for early/recycling endosomes
and Rab7 for MVBs/late endosomes. Rab7 also has been found in
LROs such as melanosomes and lamellar bodies.33,34 We reasoned
that if MVBs/late endosomes are DG precursors, and Rab32 and
Rab38 localize primarily to immature DGs, these Rabs should
colocalize preferentially with Rab7 but only marginally with Rab5.
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BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
Figure 4. Rab32 and Rab38 are primarily present in immature DGs. (A-C) DGs
were labeled with mepacrine in live MEG-01 cells expressing Cherry-Rab38 and
visualized by spinning-disk confocal fluorescence microscopy; 95% ⫾ 2% of structures containing Cherry-Rab38 (40 cells) also contain mepacrine. (A) A structure
containing the highest amount of mepacrine and low Cherry-Rab38 levels is indicated
with a green arrowhead, a structure with high concentration of Cherry-Rab38 and low
mepacrine with a red arrowhead, and a structure with intermediate amounts of both
markers with a yellow arrowhead (bar represents 5 ␮m). (B) Both organelles and
vesicles are labeled with Cherry-Rab38, which is shown as an inset in panel C (bar
represents 5 ␮m). (C) Although mepacrine is only present in the organelles,
Cherry-Rab38 is also present in the vesicle (bar represents 500 nm). (D) Live
MEG-01 cell coexpressing LAMP2-Cherry, as a DG marker, and GFP-Rab38 imaged
by spinning-disk confocal fluorescence microscopy; 88 ⫾ 4% of structures containing
GFP-Rab38 (37 cells) also contain LAMP2-Cherry. Bar represents 5 ␮m. (E) A
close-up view of structures from panel D shows a reverse correlation between the
amount of Rab38 and the DG marker. (F) Fluorescence intensity line scan of the
structures shown in panel E (merge panel). A.U., arbitrary units. (G-L) Immunogold
electron microscopy images of immature DGs from MEG-01 cells subjected to HPF
using antibodies against LAMP2 (18 nm) and Rab32 (12 nm). LAMP2 is present in
73% of the organelles label with Rab32 (n ⫽ 84 organelles). Original magnifications
were ⫻15,000, ⫻20,000, ⫻15,000, ⫻20,000, ⫻25,000, and ⫻20,000, respectively.
Bars represent 200 nm.
Indeed, confocal fluorescence microscopy images and corresponding fluorescence intensity line scans showed significant colocalization between GFP-Rab7a and either Cherry-Rab32 or CherryRab38 (MOC ⫽ 0.52 ⫾ 0.02 and 0.42 ⫾ 0.03, respectively; Figure
5B,D). In contrast, colocalization between GFP-Rab5a and either
Cherry-Rab32 or Cherry-Rab38 was minimal (MOC ⫽ 0.16 ⫾ 0.02
and 0.14 ⫾ 0.01, respectively; Figure 5A,C). These results indicate
Rab32 and Rab38 localize primarily to immature DGs and that
these organelles are closely related to MVBs/late endosomes,
consistent with the dextran internalization results.
DGs can be studied biochemically in MEG-01 cells
To complement our microscopy studies, we carried out a subcellular fractionation of MEG-01 cells. A postnuclear supernatant
obtained from a MEG-01 total extract was subjected to a 10% to
60% linear sucrose gradient. Figure 6A shows an immunoblotting
analysis of each of the 11 fractions collected (Fs), with F1 being the
least dense. Markers for different cell compartments are as follow:
MECHANISM OF PLATELET DENSE GRANULE BIOGENESIS
4077
MRP4 and LAMP2 for immature and mature DGs, platelet factor-4
(PF-4) for AGs35; and Rab7a and Rab32 for cytosol, vesicles, and
immature DGs. The DG markers LAMP2 and MRP4 coelute in F6
to F9, suggesting these fractions contain an heterogeneous mixture
of DGs at different levels of maturation. LAMP2 is also present as a
faint band in F1 that possibly corresponds to the vesicular pool of
LAMP2. AGs eluted in F4 as detected by PF-4, consistent with
their lower density compared with DGs, and reproducing the
findings of other groups with platelet organelles.29 Rab32 and Rab7
presence in F1 and F2 corresponds to their cytosolic and vesicleassociated pools. Interestingly, Rab32 and Rab7 also coelute with
the DGs markers in F6 to F8, suggesting these fractions contain
immature DGs.
To identify the sucrose gradient fraction(s) that contained the
most mature DGs, the levels of ADP were determined for each
fraction (Figure 6B filled circles). A high amount of ADP was
measured in F1 to F3 that suggests the cytosolic pool of ADP (F1)
leaked into F2 and F3. Importantly, F9, which contains the DG
markers LAMP2 and MRP4, presented a small but reproducible
ADP peak. MRP4 has been suggested to be the ADP transporter of
the DG,28 so we enriched the sucrose gradient fractions in
MRP4-containing structures by immunoprecipitation using an
anti-MRP4 antibody. After immunoprecipitation, F9 showed an
increase of ⬃ 5 times in the amount of ADP, which coincided with
the concentration factor of the sample because of immunoprecipitation (Figure 6B open circles). This result indicates that F9 contains
the most mature DGs and that immature DGs present in F6 to F8
are not competent for ADP accumulation. The larger amount of
MRP4 and LAMP2 in fractions F6 to F8 containing immature DGs
compared with F9 representing mature DGs is also consistent with
the electron microscopy data showing more MVBs and immature
DGs than mature DGs in MEG-01 cells (Figure 1B).
The ability to fractionate the organelles biochemically provided
another angle to test the idea that DGs originate from MVBs. For
this purpose, MEG-01 cells were allowed to internalize fluorescent
dextran and a cellular extract fractionated in a sucrose gradient. The
fluorescence intensity of each fraction was measured and the results
presented in Figure 6B (dashed line). F9 shows the highest peak,
indicating DGs contain internalized fluid phase material, thus
supporting the idea of DGs originating from MVBs. The presence
of dextran in F4, the fraction containing the AGs labeled with PF-4,
likely reflects their published MVB origin.31 Taking advantage of
the high Ca2⫹ content of dense granules, we used the fluorescent
Oregon Green 488 BAPTA-1 calcium indicator conjugated to
dextran. MEG-01 cells were allowed to internalize this compound,
which increases its fluorescence intensity 14-fold upon Ca2⫹
binding. After density gradient fractionation, the fluorescence
intensity of each fraction was measured and the results presented in
Figure 6B (dotted line). The peak in F9 displays most of the fluorescence
indicating this fraction contains mature, Ca2⫹-containing DGs.
We found that in MEG-01 cells the DG marker LAMP2
colocalizes with Rab32 and Rab38 by confocal fluorescence
microscopy of overexpressed proteins and with Rab32 by electron
microscopy of endogenous proteins. To support those results with
biochemical data, sucrose gradient fractions were immunoprecipitated using an anti-Rab38 antibody. As shown in Figure 6C,
structures labeled by Rab38 also contain MRP4, predominantly in
F6 to F8, which most likely correspond to immature DGs. Although
very faint, MRP4 bands in F2 and F3 suggest Rab38 and MRP4
coexist in transport vesicles.
Finally, immunoprecipitation experiments were carried out to
verify that LAMP2 and MRP4 not only coelute in the same
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4078
AMBROSIO et al
BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
Figure 5. Rab32 and Rab38 colocalize with the late
endocytic marker Rab7a but not with the vacuolar
early endosome and recycling endosome marker
Rab5a. Cherry-Rab32 and Cherry-Rab38 were cotransfected with GFP-Rab5a or GFP-Rab7a in MEG-01
cells. Confocal fluorescence microscopy images of live
cells together with the corresponding fluorescence
intensity line scan graphs are shown for each experiment. The white lines in the merge panels indicate the
portions of the cells where fluorescence intensities for
both the red and green channels were measured.
(A,C) Neither Rab32 nor Rab38 colocalizes significantly with Rab5a, a marker of recycling vesicles/early
endosomes (MOC ⫽ 0.16 ⫾ 0.02, n ⫽ 13 cells and
MOC ⫽ 0.14 ⫾ 0.01, n ⫽ 7 cells, respectively).
(B,D) Consistent with Rab32 and Rab38 being present
in immature DGs, both proteins colocalize with Rab7a,
a marker of late endosomal compartments that is also
present in other LROs such as melanosomes and
lamellar bodies (MOC ⫽ 0.52 ⫾ 0.02, n ⫽ 7 cells and
MOC ⫽ 0.42 ⫾ 0.03, n ⫽ 7 cells, respectively).
fractions but that they are present in the same organelles. F8 and F9
were submitted to immunoprecipitation using either anti-MRP4 or
anti-LAMP2 antibodies. Immunoblotting analysis of the eluted
proteins indicated MRP4 is able to pull down structures containing
LAMP2 and vice versa (Figure 6D), indicating they are present in
the same organelles in MEG-01 cells as previously described for
platelets DGs.28,29 A control experiment using an irrelevant antibody demonstrates the specificity of the immunoprecipitation
procedure (supplemental Figure 10A). Immunoblotting for PF-4
shows AGs are not being nonspecifically isolated along with DGs
(supplemental Figure 10B).
Rab32 and Rab38 are involved either in tethering or fusion of
cargo-containing vesicles with the immature DG, or both
To further investigate the function of Rab32 and Rab38 in DG
biogenesis, MEG-01 cells were subjected to siRNA knockdown of
Figure 6. Biochemical study of MEG-01 dense granules. (A) Immunoblotting analysis of fractions obtained from MEG-01 postnuclear supernatants subjected to subcellular
fractionation with a 10% to 60% sucrose gradient. Markers for different cell compartments are as follow: MRP4 and LAMP2 for immature and mature DGs; PF-4 for ␣ granules;
and Rab7a and Rab32 for cytosol, vesicles, and DGs. (B) ADP (␮M, solid lines) and fluorescence intensity of both infrared fluorescent-dextran (A.U., dashed line) and the
fluorescent Ca2⫹ indicator Oregon Green BAPTA-1 dextran (A.U., dotted line) in fractions from panel A. The concentration of ADP was determined both in the untreated sucrose
gradient fractions (solid circles) and in sucrose gradient fractions enriched in MRP4 structures by immunoprecipitation using an MRP4 antibody (open circles). (C) Sucrose
gradient fractions were immunoprecipitated using a Rab38 antibody and the presence of MRP4 in the precipitated structures was determined by immunoblotting, confirming
the occurrence of the proteins in the same structures. (D) The coexistence of both LAMP2 and MRP4 in the same organelles was confirmed by coimmnunoprecipitation using a
MRP4 antibody (left) or a LAMP2 antibody (right) and immunoblotting analysis using an antibody against the other protein. SGF indicates sucrose gradient fraction; IP,
immunoprecipitation; IB, immunoblotting; and IDG, immature DG.
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BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
MECHANISM OF PLATELET DENSE GRANULE BIOGENESIS
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either Rab32 or Rab38 and expression of LAMP2-Cherry as a DG
protein cargo reporter. In control cells, LAMP2-Cherry labels
organelle-sized structures by live cell imaging (Figure 7A). In
contrast, in Rab32-deficient cells LAMP2-Cherry is present in
structures significantly smaller and more consistent with vesicles or
very small organelles (Figure 7C,G). Rab38-deficient cells show a
similar phenotype but to a lesser extent (Figure 7E,G). What is
more, in Rab-deficient cells most of the LAMP2 structures move
significantly faster than in the control cells, which is also consistent
with LAMP2 being present in vesicles (Figure 7H; also, compare
kymographs in Figure 7B with D and F and supplemental Videos
4-6). These results suggest Rab32 and Rab38 are crucial components in either tethering or fusion events (or both) between the
cargo-carrying vesicle and the maturing organelle. In addition, the
fact that deficiency of either Rab cannot be completely compensated by the presence of the other one indicates they are not fully
redundant at least for this function.
Subcellular fractions were obtained from extracts of control,
Rab32- and Rab38-deficient cells and the presence of endogenous
LAMP2 in F1 was studied by immunoblotting (Figure 7I). The
amount of LAMP2 in F1 from Rab32- and Rab38-deficient cells is
strikingly higher than the control. This result indicates that in cells
deficient for each Rab the amount of vesicular LAMP2 is higher
than in control cells, confirming the data obtained with LAMP2Cherry by fluorescence microscopy.
Discussion
The great level of difficulty in studying primary bone marrow MKs
is a key reason behind our poor understanding of DG biogenesis.
We demonstrate here that the megakaryocytic cell line MEG-01 is a
very good model system to study DG biogenesis. The presence of
mature DGs in MEG-01 cells was established by electron microscopy, mepacrine fluorescence microscopy in live cells, and ADP
accumulation in MRP4-containing high-density organelles. Consistently, known DG protein markers such as endogenous LAMP2 and
MRP4 or exogenously expressed LAMP2 and VMAT2 colocalize
in MEG-01 cells, indicating the DG biogenesis pathways are
mostly conserved. DG markers segregate from AG markers by both
microscopy and biochemical fractionation. MEG-01 cells present a
high proportion of MVBs and immature DGs, indicating these cells
represent relatively immature megakaryocytes. Despite the similarities between dense granules in MEG-01 and primary MKs,
potential limitations of using megakaryocytic leukemia cells should
be kept in mind.
Unlike most secretory organelles, AGs and DGs may not
originate from the trans-Golgi network. More than a decade ago,
MVBs/late endosomes were shown to be the precursors of AGs.31
One study proposed a similar MVB origin for DGs on the basis of
the localization of granulophysin, a marker later revealed as
CD63/LAMP3, which is also present in AGs and various other
cellular compartments.8,9 Our use of an internalized fluid phase
marker to label late endocytic compartments together with wellvalidated DG markers (mepacrine, LAMP2, ADP) shows unequivocally that DGs originate from endosomal compartments rather than
the trans-Golgi network. Consistently, the putative serotonin
transporter (VMAT2) also colocalize with internalized dextran.
Furthermore, the presence of the DG protein LAMP2 in the
limiting membrane and internal vesicles of MVBs was revealed by
immunogold electron microscopy. Rab7 is a classic marker of
MVBs/late endosomes; therefore, its colocalization by microscopy
Figure 7. Rab32 or Rab38 knock-down impairs normal fusion of vesicles
containing dense granule proteins with the organelle. (A-F) MEG-01 cells
were cotransfected with LAMP2-Cherry as a DG reporter and either control siRNA
shown in panels A and B, Rab32 siRNA in panels C and D, or Rab38 siRNA in
panels E and F (see supplemental Figure 11 for Rab32/38 siRNA knockdown
confirmation by immunoblotting). The kymographs presented in panels B, D, and
F were made by aligning on a time axis the pieces of images indicated with a white
rectangle in panels A, C, and E, respectively, from each of the 60 frames of the
corresponding movies (1 frame/second). (A) Control siRNA cells present diffractionlimited LAMP2 structures consistent in size with organelles. (B) The LAMP2
structures in Control siRNA cells present a limited range of motion. (C,E) Both
Rab32 and Rab38 siRNA cells present LAMP2 structures that are more consistent
in size with vesicles or small organelles. (D,F) The smaller LAMP2 structures in
Rab32 and Rab38 siRNA cells are more dynamic and move faster than structures
in Control siRNA cells. (G) The diameter of LAMP2 structures present in the
representative cells shown in panels A, C, and E was measured using the
Ruler function in Slidebook. (H) The average speed of LAMP2 structures
present in the representative cells shown in panels A, C, and E was measured
using the Manual Particle Tracking function in Slidebook. (I) Extracts from
control, Rab32, and Rab38 siRNA-treated cells were fractionated in sucrose
gradients. For each treatment, the amount of LAMP2 in the first fraction of the
gradient, which corresponds to the vesicular LAMP2, was analyzed by immunoblotting. The levels of tubulin in the same blot were used to confirm equal loading.
Bars represent 5 ␮m (*P ⬍ .05; **P ⬍ .001). (J) DG membrane proteins are
sorted in early endosomal compartments by adaptor protein complexes, such as
AP-3, which recognize sorting signals present in their cytosolic tails. Rab32 and
Rab38 are recruited to the nascent clathrin-coated vesicle and through interactions with so far unknown effectors target the vesicle to the maturing DG. The DG
precursor is a MVB that on receiving Rab32/Rab38 vesicles containing DG
proteins, such as the ADP transporter MRP4 or the serotonin transporter VMAT2,
matures into a DG.
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4080
BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
AMBROSIO et al
and cofractionation in density gradients with Rab32, Rab38, and DG
protein markers further supports a MVBs/late endosome origin for DGs.
In addition to endocytic material, DGs must receive newly
synthesized integral membrane proteins that normally function in
its limiting membrane. Our results show for the first time that
tyrosine- and dileucine-based sorting signals are required for
normal traffic to DGs of LAMP2 and VMAT2, respectively. These
types of signals are also used for AP-3–mediated transport to
melanosomes and lysosomes, suggesting the mechanism of biogenesis is highly conserved among different LROs. Our results are
most consistent with a model in which AP-3 carries out transport of
newly synthesized DG membrane proteins from early endosome–
associated tubules to maturing DGs, a pathway analogous to that
described for melanosomal proteins (Figure 7J).
Mutation of Rab38 causes deficiency of platelet DGs and other
LROs in rodent models of HPS.15-17,34 Rab38 and its very close
homolog Rab32 operate in a partially redundant manner in
melanosome biogenesis.17,18 We found that Rab32 and Rab38
partially colocalize with AP-3 and clathrin both in primary MKs
and MEG-01 cells. This result suggests Rab32 and Rab38 are
recruited to cargo-filled vesicles budding from early endosome–
associated tubules and likely destined for maturing DGs. Rab32
and Rab38 are also present predominantly in immature DGs
(Figures 4-6 and supplemental Figure 8). We also detect small
structures labeled by fluorescently tagged Rabs that likely represent
transport vesicles. Overall, these localization experiments fit very
well with a model in which Rab32 and Rab38 define a pathway
from early endosome–associated tubules to maturing dense granules (Figure 7J). In this context, the LAMP2-Cherry trafficking
phenotype observed on knock-down of either Rab32 or Rab38 are
indicative of a deficiency in the tethering or fusion of vesicles
containing DG cargo with the organelles. These results constitute
the first evidence at the molecular level of Rab32 and Rab38
involvement in DGs biogenesis.
Our model for the biogenesis of DGs is depicted in Figure 7J.
Newly synthesized DG proteins are sorted in specific early
endodomal tubules by the recognition of sorting signals present in
their cytosolic tails by adaptors such as AP-3. At this point or
quickly after pinching off, cytosolic Rab32 and Rab38 are recruited
to these vesicles and through interactions with still unknown
effectors regulate docking and fusion of the vesicles with the
MVBs/immature DGs.
This work represents an important step forward in our understanding of the molecular mechanism of platelet DG biogenesis.
Moreover, our data indicate MEG-01 cells are a valid and powerful
model system for the study of DG formation and the identification
of new players involved in these pathways.
Acknowledgments
The authors thank Andrew Peden for the AP-3 antibody and
Thomas Giddings for HPF.
This work was supported by American Heart Association award
09SDG2280525 and National Institutes of Health grant
1R01HL106186-01A1 (S.M.D.). Microscopes used in this work
are supported in part by the Colorado State University Microscope
Imaging Network core infrastructure grant.
Authorship
Contribution: A.L.A., J.A.B., and S.M.D. designed the study,
performed the experiments, analyzed the data, and wrote the
manuscript.
Conflict-of-interest disclosure: The authors declare no competing financial interests.
Correspondence: Santiago M. Di Pietro, Department of Biochemistry and Molecular Biology, 1870 Campus Delivery, Colorado State University, Fort Collins, CO 80523-1870; e-mail:
[email protected].
References
1. Huizing M, Helip-Wooley A, Westbroek W, GunayAygun M, Gahl WA. Disorders of lysosome-related
organelle biogenesis: clinical and molecular genetics. Annu Rev Genomics Hum Genet. 2008;9:359386.
2. McNicol A, Israels SJ. Platelet dense granules:
structure, function and implications for haemostasis. Thromb Res. 1999;95(1):1-18.
3. Kim S, Kunapuli SP. P2Y12 receptor in platelet
activation. Platelets. 2011;22(1):54-58.
4. Konopatskaya O, Matthews SA, Harper MT, et al.
Protein kinase C mediates platelet secretion and
thrombus formation through protein kinase D2.
Blood. 2011;118(2):416-424.
5. Graham GJ, Ren Q, Dilks JR, Blair P, Whiteheart SW,
Flaumenhaft R. Endobrevin/VAMP-8-dependent
dense granule release mediates thrombus formation in vivo. Blood. 2009;114(5):1083-1090.
6. Gunay-Aygun M, Huizing M, Gahl WA. Molecular
defects that affect platelet dense granules. Semin
Thromb Hemost. 2004;30(5):537-547.
7. King SM, Reed GL. Development of platelet secretory granules. Semin Cell Dev Biol. 2002;
13(4):293-302.
8. Youssefian T, Cramer EM. Megakaryocyte dense
granule components are sorted in multivesicular
bodies. Blood. 2000;95(12):4004-4007.
9. Pols MS, Klumperman J. Trafficking and function
of the tetraspanin CD63. Exp Cell Res. 2009;
315(9):1584-1592.
10. Bonifacino JS, Traub LM. Signals for sorting of
transmembrane proteins to endosomes and lysosomes. Annu Rev Biochem. 2003;72:395-447.
11. Peden AA, Oorschot V, Hesser BA, Austin CD,
Scheller RH, Klumperman J. Localization of the
AP-3 adaptor complex defines a novel endosomal exit site for lysosomal membrane proteins.
J Cell Biol. 2004;164(7):1065-1076.
12. Theos AC, Tenza D, Martina JA, et al. Functions
of adaptor protein (AP)-3 and AP-1 in tyrosinase
sorting from endosomes to melanosomes. Mol
Biol Cell. 2005;16(11):5356-5372.
13. Di Pietro SM, Falcon-Perez JM, Tenza D, et al.
BLOC-1 interacts with BLOC-2 and the AP-3
complex to facilitate protein trafficking on endosomes. Mol Biol Cell. 2006;17(9):4027-4038.
14. Grosshans BL, Ortiz D, Novick P. Rabs and their
effectors: achieving specificity in membrane traffic. Proc Natl Acad Sci U S A. 2006;103(32):
11821-11827.
15. Oiso N, Riddle SR, Serikawa T, Kuramoto T,
Spritz RA. The rat Ruby (R) locus is Rab38: identical mutations in Fawn-hooded and TesterMoriyama rats derived from an ancestral Long
Evans rat sub-strain. Mamm Genome. 2004;
15(4):307-314.
16. Ninkovic I, White JG, Rangel-Filho A, Datta YH.
The role of Rab38 in platelet dense granule defects. J Thromb Haemost. 2008;6(12):2143-2151.
17. Wasmeier C, Romao M, Plowright L, Bennett DC,
Raposo G, Seabra MC. Rab38 and Rab32 con-
trol post-Golgi trafficking of melanogenic enzymes. J Cell Biol. 2006;175(2):271-281.
18. Bultema JJ, Ambrosio AL, Burek CL, Di Pietro SM.
BLOC-2, AP-3, and AP-1 proteins function in concert with Rab38 and Rab32 proteins to mediate
protein trafficking to lysosome-related organelles.
J Biol Chem. 2012;287(23):19550-19563.
19. Leven RM. Isolation of primary megakaryocytes
and studies of proplatelet formation. Methods Mol
Biol. 2004;272:281-291.
20. Meehl JB, Giddings TH Jr, Winey M. High pressure freezing, electron microscopy, and immunoelectron microscopy of Tetrahymena thermophila
basal bodies. Methods Mol Biol. 2009;586:227241.
21. Manders EMM, Verbeek FJ, Aten JA. Measurement of colocalization of objects in dual-color
confocal images. J Microscopy. 1993;169:375382.
22. Nazarian R, Starcevic M, Spencer MJ, Dell’Angelica
EC. Reinvestigation of the dysbindin subunit of
BLOC-1 (biogenesis of lysosome-related organelles complex-1) as a dystrobrevin-binding protein. Biochem J. 2006;395(3):587-598.
23. Ogura M, Morishima Y, Ohno R, et al. Establishment of a novel human megakaryoblastic leukemia cell line, MEG-01, with positive Philadelphia
chromosome. Blood. 1985;66(6):1384-1392.
24. Ogura M, Morishima Y, Okumura M, et al. Functional and morphological differentiation induction
of a human megakaryoblastic leukemia cell line
From www.bloodjournal.org by guest on June 11, 2017. For personal use only.
BLOOD, 8 NOVEMBER 2012 䡠 VOLUME 120, NUMBER 19
(MEG-01s) by phorbol diesters. Blood. 1988;
72(1):49-60.
25. Schweinfurth N, Hohmann S, Deuschle M,
Lederbogen F, Schloss P. Valproic acid and all
trans retinoic acid differentially induce megakaryopoiesis and platelet-like particle formation
from the megakaryoblastic cell line MEG-01.
Platelets. 2010;21(8):648-657.
26. Giannaccini G, Betti L, Palego L, et al. Human
serotonin transporter expression during megakaryocytic differentiation of MEG-01 cells. Neurochem Res. 2010;35(4):628-635.
27. Dionisio N, Albarran L, Lopez JJ, et al. Acidic
NAADP-releasable Ca(2⫹) compartments in the
megakaryoblastic cell line MEG01. Biochim Biophys Acta. 2011;1813(8):1483-1494.
28. Jedlitschky G, Tirschmann K, Lubenow LE, et al.
The nucleotide transporter MRP4 (ABCC4) is
MECHANISM OF PLATELET DENSE GRANULE BIOGENESIS
highly expressed in human platelets and present
in dense granules, indicating a role in mediator
storage. Blood. 2004;104(12):3603-3610.
29. Niessen J, Jedlitschky G, Grube M, et al. Subfractionation and purification of intracellular
granule-structures of human platelets: an improved method based on magnetic sorting. J Immunol Methods. 2007;328(1-2):89-96.
4081
32. Reddington M, Novak EK, Hurley E, Medda C,
McGarry MP, Swank RT. Immature dense granules in platelets from mice with platelet storage
pool disease. Blood. 1987;69(5):1300-1306.
33. Gomez PF, Luo D, Hirosaki K, et al. Identification
of rab7 as a melanosome-associated protein involved in the intracellular transport of tyrosinaserelated protein 1. J Invest Dermatol. 2001;117(1):
81-90.
30. Höltje M, Winter S, Walther D, et al. The vesicular
monoamine content regulates VMAT2 activity
through Galphaq in mouse platelets. Evidence for
autoregulation of vesicular transmitter uptake.
J Biol Chem. 2003;278(18):15850-15858.
34. Zhang L, Yu K, Robert KW, et al. Rab38 targets to
lamellar bodies and normalizes their sizes in lung
alveolar type II epithelial cells. Am J Physiol Lung
Cell Mol Physiol. 2011;301(4):L461-477.
31. Heijnen HF, Debili N, Vainchencker W, BretonGorius J, Geuze HJ, Sixma JJ. Multivesicular
bodies are an intermediate stage in the formation
of platelet alpha-granules. Blood. 1998;91(7):
2313-2325.
35. Huang CL, Cheng JC, Stern A, Hsieh JT, Liao CH,
Tseng CP. Disabled-2 is a novel alphaIIb-integrinbinding protein that negatively regulates plateletfibrinogen interactions and platelet aggregation.
J Cell Sci. 2006;119(Pt 21):4420-4430.
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2012 120: 4072-4081
doi:10.1182/blood-2012-04-420745 originally published
online August 27, 2012
Mechanism of platelet dense granule biogenesis: study of cargo
transport and function of Rab32 and Rab38 in a model system
Andrea L. Ambrosio, Judith A. Boyle and Santiago M. Di Pietro
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