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RED CELLS, IRON, AND ERYTHROPOIESIS
The ribosome-related protein, SBDS, is critical for normal erythropoiesis
Saswati Sen,1 Hanming Wang,1 Chi Lan Nghiem,1 Kim Zhou,1 Janice Yau,1 Chetankumar S. Tailor,1 Meredith S. Irwin,1 and
Yigal Dror1
1Cell Biology Program, Research Institute, and Division of Haematology/Oncology, Department of Paediatrics, Hospital for Sick Children and Institute of Medical
Sciences, University of Toronto, Toronto, ON
Although anemia is common in ShwachmanDiamond syndrome (SDS), the underlying
mechanism remains unclear. We asked
whether SBDS, which is mutated in most
SDS patients, is critical for erythroid development. We found that SBDS expression is high early during erythroid differentiation. Inhibition of SBDS in CD34ⴙ
hematopoietic stem cells and early progenitors (HSC/Ps) and K562 cells led to
slow cell expansion during erythroid differentiation. Induction of erythroid differentiation resulted in markedly acceler-
ated apoptosis in the knockdown cells;
however, proliferation was only mildly
reduced. The percentage of cells entering
differentiation was not reduced. Differentiation also increased the oxidative stress
in SBDS-knockdown K562 cells, and antioxidants enhanced the expansion capability of differentiating SBDS-knockdown
K562 cells and colony production of SDS
patient HSC/Ps. Erythroid differentiation
also resulted in reduction of all ribosomal
subunits and global translation. Furthermore, stimulation of global translation
with leucine improved the erythroid cell
expansion of SBDS-knockdown cells and
colony production of SDS patient HSC/
Ps. Leucine did not reduce the oxidative
stress in SBDS-deficient K562 cells. These
results demonstrate that SBDS is critical
for normal erythropoiesis. Erythropoietic
failure caused by SBDS deficiency is at
least in part related to elevated ROS
levels and translation insufficiency because antioxidants and leucine improved
cell expansion. (Blood. 2011;118(24):
6407-6417)
Introduction
Shwachman-Diamond syndrome (SDS) is a rare multisystem
disorder, primarily affecting the bone marrow, pancreas, and
skeletal systems.1,2 Hematologic abnormalities are a major cause
for morbidity and mortality and include cytopenia, myelodysplastic
syndromes, and leukemia. Neutropenia is observed in almost all
SDS patients. Approximately 85% of the patients have hypomorphic mutations in the Shwachman Bodian Diamond syndrome
gene, SBDS.3,4 Knockdown of the SBDS homolog in murine
myeloid 32Dcl3 cells resulted in normal neutrophil maturation but
reduced viability of granulocyte precursors.5 These studies provide
evidence that SBDS is critical for normal granulopoiesis. The role
of SBDS in erythroid development has not been characterized.
Anemia is observed in ⬃ 60% of SDS patients. In addition, 60% of
the patients have high erythrocyte mean corpuscular volume, and
70% have high fetal hemoglobin blood levels.4,6,7 Despite these
erythrocyte abnormalities, SDS has frequently been classified as an
inherited neutropenia disorder.8,9 The underlying mechanism of
anemia and the degree to which it can be attributed to erythropoietic failure, nutritional deficiencies, and/or recurrent infection have
not been studied.
Most SDS bone marrows are hypocellular with reduced hematopoietic progenitors, including erythroid cells. Nevertheless, the
defects mediated by SBDS deficiency that are responsible for this
phenotype are unclear. For example, it is unknown whether the
reduction in SDS hematopoietic progenitors is the result of an
impaired ability to differentiate or to proliferate or because of a
defect to survive throughout the process of maturation. Previous
studies support SBDS roles in several cellular pathways, including
cell survival.10,11 Studies on SDS marrow cells similarly showed
reduced hematopoietic stem cells/early progenitors (HSC/Ps) and
colony formation.12 In both SDS marrow cells and SBDSknockdown HeLa cells, the slow cell expansion was in part the
result of Fas-mediated apoptosis.13 Importantly, inhibition of
apoptosis in HeLa cells rescued the slow cell expansion phenotype.
Elevated levels of reactive oxygen species (ROS) may affect
erythroid cell survival and can cause cytopenia.14,15 We have
previously found that knockdown of SBDS in myeloid and
nonmyeloid cell lines causes elevated ROS levels, and balancing
ROS levels improved cell survival and expansion.16 Although these
studies shed light on the role of SBDS in cell survival, they were not
performed on differentiating cells and may not accurately represent
the SBDS role during maturation of hematopoietic cells.
A role of SBDS in promoting ribosome biogenesis has also been
suggested.17-20 The yeast homolog has been shown to play a role in
maturation of the 60S ribosome subunit.18 However, the dynamics
of ribosome biogenesis during hematopoiesis and how these
defects relate to the hematologic abnormalities seen in SDS are
unknown. Because the developing erythroid cells are highly
proliferative and require increased ribosome synthesis for the
increased demand of hemoglobin synthesis, it is possible that these
immature cells are more dependent on SBDS than mature cells.
In this study, we asked whether, in addition to its role in
granulocytic cell development, SBDS is also critical for the
expansion and differentiation of developing erythroid progenitors.
Because expansion capability was reduced, we studied whether
apoptosis and proliferation of differentiating erythroid cells are
Submitted February 6, 2011; accepted July 18, 2011. Prepublished online as Blood
First Edition paper, September 30, 2011; DOI 10.1182/blood-2011-02-335190.
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.
The online version of this article contains a data supplement.
© 2011 by The American Society of Hematology
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SEN et al
affected when SBDS is depleted. We found that apoptosis was
predominantly affected and was accelerated. We then asked
whether the cell expansion phenotype is dependent on defects in
global translation and ROS levels. In this study, we focused on
the erythroid lineage model, to ask whether the anemia in
SBDS-deficient cells is the result of hematopoietic failure and to
perform an in-depth analysis of the underlying defects leading to
cytopenia in SDS.
Methods
Subjects
All SDS patients had characteristic hematologic and pancreatic manifestations,7 biallelic SBDS mutations, and no signs of leukemic transformation or
clonal marrow cytogenetic abnormalities. Healthy control subjects were
donors for bone marrow transplantation. Cord blood was obtained from
women after labor. Informed written consent was obtained from all patients
and controls or their legal guardians in accordance with the Declaration of
Helsinki. The studies were approved by the Hospital for Sick Children,
University of Toronto Research Ethics Board.
Cell culture and erythroid differentiation
CD34⫹ HSC/Ps from marrow mononuclear cells were enriched using
immunomagnetic beads.11,12 CD133⫹ HSC/Ps from cord blood samples
were sorted using an anti-CD133/2–PE antibody (Miltenyi Biotec). HSC/Ps
were plated in serum-free granulocyte-erythrocyte-monocyte-megakaryocyte colony-forming unit assays.11 In other experiments, HSC/Ps were
plated in differentiation cultures containing IMDM, 20% FBS, 2 U/mL
erythropoietin, 50 ng/mL SCF, and 20 ␮U/mL insulin, followed by addition
of erythropoietin every 2 days.
K562 cells were from ATCC. To induce erythroid differentiation, K562
cells were seeded at a density of 2 ⫻ 105 cells/mL in IMDM (Invitrogen)
with FBS and 25 ␮m hemin solution (Sigma-Aldrich).21
shRNA expression cassettes and generation of
SBDS-knockdown cells
SBDS short hairpin RNA (shRNA) and a scrambled (SCR) RNA control
sequence were cloned into a pSEC/Neo plasmid (Ambion) as previously
described.10 K562 cells were transfected with the plasmids and selected by
plating in methylcellose Cell Selection medium (StemCell Technologies)
containing geneticin (Invitrogen), and individual colonies were isolated
after 14 days. Two knockdown lines (K562/shSBDS-1 and K562/
shSBDS-3) and 1 control line (K562/shSCR) were established.
Viral vectors production and transduction
We cloned SBDS in frame with YPet on a pLNCX/YPet vector (courtesy of
Dr Patrick Daugherty, University of California, Santa Barbara, CA). Viral
particles were produced by transfecting FLYRD18 retroviral packaging
cells22 with pLNCX retroviral vectors expressing SBDS-YPET and control
YPET. K562/shSBDS-3 cells were transduced in the presence of polybrene.
K562/shSBDS-3 cells express an shRNA targeting nucleotides 137 to 156 3⬘
to the open reading frame; therefore, the reintroduced SBDS open reading
frame was not inhibited by the shRNA.
The shRNA oligonucleotides, which were used to target SBDS in K562
cells, were also cloned into the YFP vector, pFCYsi (courtesy of Dr Dan
Link, Washington University). To knock down SBDS in human HSC/Ps,
marrow CD34⫹ cells from healthy control subjects were incubated in
StemSpan medium (StemCell Technologies) with FMS-like tyrosine kinase
3 ligand, SCF, and thrombopoietin for 24 hours. Cells were then transduced
3 times with viral supernatant in the presence of polybrene.23
Morphologic analysis by benzidine and Wright-Giemsa staining
Benzidine staining was used to detect pseudo-peroxidase activity of
hemoglobin.21 Cells were cytospun on glass slides and incubated with
3,3⬘-dimethoxybenzidine (Sigma-Aldrich) and hydrogen peroxide. Benzidine-positive cells were enumerated by light microscopy. Cytologic examination of bone marrow was done after Wright-Giemsa staining.24 The
proportion of the various marrow cell populations was determined by
counting at least 500 cells under light microscope.
Cell expansion assays
K562 cells were seeded at 2.0 ⫻ 104 cells per 0.1 mL in 96-well plates or
2.0 ⫻ 105 cells/mL in 35-mm dishes depending on the specific experiment.
At the indicated time points after plating, viable cell counts were
determined using either trypan blue dye exclusion or MTT assay (ATCC).11
Absorbance was measured at 570 nm in a microplate reader (Bio-Tek
Instruments).
RNA quantitative PCR
Expression of SBDS, GATA-1, and TFR2 was measured by RNA quantitative real-time PCR using SYBR Green technology.25 The specific primer
pairs for the studied genes and the internal control ACTB gene are described
in supplemental Table 1 (available on the Blood Web site; see the
Supplemental Materials link at the top of the online article). In each
experiment, no-DNA template control was also used. Relative expression
levels were calculated using the relative standard curve method.26
Western blotting
Western blot analysis was performed as previously described.11 Chicken
anti–human SBDS antibody was generated as previously described,10
Mouse anti–human ␤-actin antibody was from Sigma-Aldrich; mouse
anti–human p53 antibody was from Santa Cruz Biotechnology; mouse
anti-p63 (A4A) was from Santa Cruz Biotechnology; mouse anti-TAp73
(BL-906) was from Bethyl Laboratories; a rabbit anti–human cleaved
poly(ADP-ribose) polymerase (Asp214) antibody was from Cell Signaling
Technology.
Flow cytometry
Propidium iodide and Ki-67 costaining was performed as previously
described10 with minor modifications. In brief, 5 ⫻ 105 cells were washed
with PBS and fixed with 70% ethanol. Cells were rewashed and incubated
with mouse anti–Ki-67 (Dako Canada) at 4°C for 30 minutes in the dark,
followed by incubation with FITC-conjugated IgM antibody (GE Healthcare) for 30 minutes. Cells were rewashed and incubated 1 hour with
propidium iodide, RNaseA, and Triton X-100. DNA content was analyzed
by flow cytometry, and the sub-G1 cell populations were quantified,
whereas Ki-67-FITC expression was analyzed in viable cells.
For evaluation of intracellular ROS level, cells (5 ⫻ 105) were washed
and incubated for 30 minutes with 50M 2,7-dichlorodihydrofluorescein
diacetate (Cayman Chemical). Cells were washed with HBSS and analyzed
by flow cytometry within 15 minutes.
Ribosome profile analysis by sucrose gradient density
ultracentrifugation
Cells were incubated at 37°C for 10 minutes after treating with cycloheximide, washed in cold PBS containing cycloheximide, and resuspended in
TMK100 lysis buffer (Tris, MgCl2, KCl, HEPES, cycloheximide, Triton
X-100, and RNAseOUT).27 Nuclei were removed by centrifugation at
12 000g for 5 minutes. Lysates were layered on a 5% to 45% (weight/
volume) sucrose gradient in sucrose gradient buffer containing KCl, MgCl2,
and HEPES and centrifuged at 197 000g for 2.5 hours. To evaluate the ratio
of 40S to 60S, cells were lysed with EDTA instead of MgCl2 to dissociate
polysomes and mRNA. Gradients were centrifuged for 3 hours at 197 000g
and fractionated using the Brandel gradient fractionator system (Brandel).
Absorbance was monitored at 254 nm with an ISCO UA-6 flow cell
(Teledyne ISCO).
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BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
SBDS ROLE IN ERYTHROPOIESIS
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Evaluation of global translation by incorporation of 35S
methionine/cysteine
Metabolic labeling of K562 cells with [35S]-methionine was performed as
previously published27 with minor modifications. Cells were incubated with
[35S]methionine and placed for 30 minutes at 37°C. The labeling medium
was removed, and the cells were washed and lysed. [35S]methionine
incorporation was determined by trichloroacetic acid precipitation of the
supernatant aliquots. Counts per minute were obtained using a scintillation
counter (Beckman Coulter LS 6500).
Oligonucleotide microarray
Ficoll-extracted marrow mononuclear cells from 9 SDS patients and
7 healthy subjects were analyzed by oligonucleotide microarray using the
HG_U133_Plus 2.0 GeneChip (Affymetrix).17 Data analysis and generation
of moderated T-statistics value were performed as previously described.17
Genes with an average probe T-statistics value of either higher than 1.7 or
lower than ⫺1.7 were considered candidate ROS-regulating genes in SDS.
The average percentages of the various bone marrow cell lineages: myeloid,
erythroid, lymphoid, and other cells were in the normal range (⫾ SD):
46.4% ⫾ 6.8%, 31.1% ⫾ 7.3%, 19.5% ⫾ 5.5%, and 3.0% ⫾ 1.3%,
respectively. Differential expression of genes involved in leukemia, apoptosis, and ribosome biogenesis were reported elsewhere.10,17,25 All microarray
data are available on the Gene Expression Omnibus under accession
number GSE32057.
Statistics
Student t test was used to determine the statistical significance of
differences between 2 means. A 1-way ANOVA followed by Dunnett test or
Tukey test of multiple means was performed to determine statistical
significance of differences between multiple means.
Results
SBDS is expressed early during erythroid differentiation
To determine whether SBDS is expressed at a particular stage of
erythroid development, we analyzed its expression by quantitative
RNA-PCR. Early CD133⫹/HSC/Ps showed high expression of
SBDS, followed by a decline in expression as erythroid differentiation progressed (Figure 1A). We also used K562 human erythroleukemia cells as a model for erythroid development because they are
multipotent hematopoietic progenitors, which can differentiate to
normoblasts over ⬃ 5 days using agents, such as hemin.28 SBDS
was up-regulated 24 hours after induction of differentiation.
Expression returned to basal levels for the subsequent 4 days
(Figure 1B). Increased SBDS protein levels in K562 were detected
one day after the mRNA levels peaked (Figure 1C). Morphologic
examination of undifferentiated and differentiating cells is shown
in supplemental Figure 1A and B.
SBDS deficiency results in markedly reduced expansion
capability of differentiating erythroid cells
We studied the effect of SBDS deficiency on expansion of primary
human erythroid cells by plating SBDS-knockdown marrow CD34⫹
cells in erythroid differentiation medium. SBDS-knockdown erythroid cells showed impaired expansion compared with controls
(Figure 2A). Effective protein depletion is shown in supplemental
Figure 2A. Because these experiments apply transient transduction
of primary cells with limited life span, the viability and expansion
of the scrambled RNA control are slightly compromised compared
with nontransduced wild-type cells. We also studied BFU-E colony
Figure 1. SBDS expression during erythroid differentiation. (A) Real-time RNA
quantitative PCR was used to analyze daily SBDS mRNA expression in CD133⫹
HSC/Ps after induction of erythroid differentiation. The experiment was performed in
triplicate. The ACTB gene was used as an internal control. (B) SBDS mRNA
expression by RNA quantitative PCR was also determined in wild-type K562 cells
after induction of erythroid differentiation by hemin. Three separate experiments were
conducted in triplicate. (C) SBDS protein expression in hemin-induced K562 cells by
Western blotting (representative blot of 3 experiments).
growth in clonogenic assays. CD34⫹ cells from SDS patients
produced less BFU-E colonies than normal controls (SDS, median
⫾ SEM ⫽ 7 ⫾ 3 colonies/103 CD34⫹ cells, n ⫽ 3; and normal,
17 ⫾ 4 colonies, n ⫽ 5, P ⫽ .04).
Similarly, we studied the expansion of SBDS-knockdown K562
cells (supplemental Figure 2B). Western blotting confirmed approximately 5% residual protein in K562/shSBDS-3 cells and approximately 12% in K562/shSBDS-1 cells (supplemental Figure 2C).
After induction of erythroid differentiation, the expansion of the
SBDS-knockdown cells was markedly reduced compared with
control cells (P ⬍ .01 for K562/shSBDS-1 and P ⬍ .001 for
K562/shSBDS-3). To determine whether the reduced cell expansion phenotype was a direct consequence of SBDS deficiency,
SBDS (that is not targeted by the shRNA expressed in K562/
shSBDS-3) was reintroduced into K562/shSBDS-3 cells. SBDS
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BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
SEN et al
Figure 2. Cell expansion of SBDS-deficient cells during erythroid differentiation. (A) SBDS-knockdown and control CD34⫹ HSC/Ps were induced to undergo erythroid
differentiation by a combination of SCF and erythropoietin and cell counts were monitored daily by trypan blue exclusion. (B) SBDS-knockdown K562/shSBDS-3 cells were
transduced with a pLNCX retrovirus expressing YPet and SBDS open reading frame (which is not targeted by the shRNA expressed). YPet-positive cells were sorted and
plated at a concentration of 2 ⫻ 105 cells/mL and treated with 25 ␮m hemin for 5 days. Cell expansion was assessed by trypan blue exclusion. Data are the mean ⫾ SE
of 3 independent experiments. *P ⬍ .05, statistically significant results using 1-way ANOVA and Tukey test for multiple means against both S-3 ⫹ YPET and S-3 cells.
expression completely rescued cell expansion (Figure 2C). The
higher numbers of cells after reintroduction of SBDS to the
knockdown cells on day 3 and 4 might be related to higher SBDS
expression levels during these days. Transduction efficiency is
shown in supplemental Figure 2D and E.
SBDS-deficient cells retain the ability to enter into a
differentiation program
Because reduction of SDS marrow cells can be the result of early
arrest in differentiation, we asked whether SBDS deficiency
impairs the entry into an erythroid differentiation program. First,
we analyzed the relative percentage of different hematopoietic cells
in marrows from SDS patients (Table 1). Although the patients had
reduced erythropoietic cells because of the hypoplastic bone
marrows, the percentages of erythroid marrow population and
normoblast subpopulation (basophilic, polychromatic, and orthocromatic) were in the normal range. The percentage of proerythroblasts in most patients was mildly elevated. Next, we
examined the ability of SBDS-deficient K562 cells to undergo
hemoglobinization after induction of differentiation. The percentages of hemoglobinized (benzidine-positive) SBDS-knockdown
cells were higher than the control cells (supplemental Figure 3A-B)
with a significant difference between K562/shSBDS-S3 and K562/
shSCR cells on day 5. The expression of the transcription factors
GATA1 and TFR2, which are critical for erythroid development in
SBDS-knockdown cells, was also increased in the SBDSknockdown cells (supplemental Figure 3C).
Ablation of SBDS results in markedly increased apoptosis
during erythroid differentiation
The underlying mechanisms responsible for reduced cell expansion
can be either accelerated cell death and/or reduced proliferation.
We first studied the viability of differentiating SBDS-knockdown
by the MTT assay. The MTT yellow tetrazole dye is reduced to
purple formazan in living cells. We found a dramatic reduction of
viability in the SBDS-knockdown cells compared with controls
during differentiation (supplemental Figure 4A). Next, sub-G0/G1
apoptotic cells were determined using a propidium iodide binding
assay.10,29 Apoptosis was slightly increased in the nondifferentiating SBDS-knockdown cells (Figure 3A-B). For example, the
percentage of apoptotic K562/shSBDS-3 cells was 1.46-fold higher
than that of K562/shSCR cells (P ⬍ .05). After induction of
differentiation, apoptosis was markedly increased in both knockdown cell lines. For example, the percentage of apoptotic K562/
shSBDS-3 cells was 2.68-fold higher than that of K562/shSCR
cells (P ⬍ .01). Caspase-3 activation as determined by semiquantitative Western blotting of cleaved poly(ADP-ribose) polymerase
was also enhanced in SBDS-knockdown K562 cells compared with
controls in both undifferentiated and differentiated cells (supplemental Figure 4B). In addition, we transduced CD34⫹ cells with either
Table 1. Bone marrow differential of patients with SDS
Promyelocytes,
myelocytes,
metamyelocytes,
%)
Bands,
neutrophils,
%
Peripheral blood Hg,
g/L; reticulocytes,
ⴛ 109/L; neutrophils,
ⴛ 109/L
Patient
(mutation)*
Age, y
1 (A/A ⫹ B)
21.5
5
28
29
6
6
2.5
33-66
72;28;1.1
2 (A/A ⫹ B)
13.5
7
31.5
37.5
6
15.5
2.5
33-66
90;ND;1.05
3 (A/B)
20
2
21.5
44.5
7.5
23
2.5
Indeterminate
91;ND;2.4
4 (A/C)
20
2
24
40
5
28
1
33-66
123;ND;1.2
5 (A/D)
22.5
7.5
30
31.5
10
17.5
2.5
⬍ 33
114;44;0.56
4
5
21
33.5
18.5
18
2.5
⬎ 66
113;1;0.47
7 (A/B)
13.5
1
23
40
21
14
2
Indeterminate
164;4;3.0
8 (A/B)
31.5
2
39
31
15.5
15
2
33-66
0.2-1.3
18.8-36.2
19.6-34.2
13.5-27.3
11.1-23.2
0.2-1.5
6 (A/A ⫹ B)
Normal54
⬎ 12
Proerythroblasts,
%
Normoblasts
(basophilic,
polychromatic,
orthochromatic),
%
Lymphocytes,
%
Myeloblasts,
%
Cellularity,†
%
120;ND;0.9
*A indicates 258 ⫹ 2T ⬎ C; B, 183-184TA ⬎ CT; C, 260T ⬎ G; and D, 203A ⬎ G.
†The cellularity was determined as the percentage of areas with hematopoietic cells on bone marrow biopsies. In cases where no biopsy was available, the percentage of
areas of hematopoietic cells on aspirate fragments was determined. In case of no fragments seen on the aspirates, the cellularity was deemed “indeterminate.”
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Figure 3. Apoptosis and proliferation levels in differentiating SBDS-deficient cells. (A) Control and knockdown K562 cells were induced to undergo erythroid
differentiation for 5 days. Cells were fixed in 70% ethanol and stained with propidium iodide for DNA content analysis. A representative diagram of 4 independent experiments is
shown. Pre-G0/G1 cells represent apoptotic cells. Representative distributions of the cell cycle phases are depicted on day 0 of wild-type cells. (B) Cells were costained for
propidium iodide to evaluate pre-G0/G1 apoptotic cells and for Ki-67 expression on nonapoptotic cells to evaluate proliferating cells. Cells were then analyzed by flow cytometry.
Comparison of the mean fold increase in apoptosis versus mean fold decrease in proliferation of 4 independent experiments is shown. The values for apoptosis and
proliferation were normalized to the value obtained for each day in control K562 cells, and assigned a value of 1.0 for proliferating cells and apoptotic cells. (C) Mobilized CD34⫹
cells from 2 donors for hematopoietic stem cell transplantation were purified, transduced, and induced to undergo erythroid differentiation as described in “Cell culture and
erythroid differentiation.” After 5 days, cells were harvested and evaluated for apoptosis rates using annexin V staining. For each donor cell (N1 and N2), annexin V staining of
cells transduced with scrambled RNA controls (SCR) or shRNA against SBDS (S-3) are shown. (D) Results of inhibition of SBDS mRNA in CD34⫹ cell samples are shown. The
faster running SBDS band corresponds to a previously published shorter isoform. (Cell numbers did not allow corresponding testing of SBDS protein levels.)
an shSBDS-3 or shSCR cassette, induced erythroid differentiation,
and evaluated apoptosis by annexin V staining 5 days after plating.
CD34⫹ cells from 2 different healthy persons manifested 6- and
3-fold higher rates of apoptosis when shSBDS was transduced
compared with transduction with shSCR (Figure 3C-D).
To assess the effect of SBDS deficiency on proliferation during
erythroid differentiation, cells were costained with propidium
iodide and Ki-67.30 Viable (non–sub-G0/G1) cells were assessed for
Ki-67 expression. Proliferation of SBDS-knockdown cells tended
to be lower than that of the controls only in the cells with more
severe SBDS-knockdown (K562/hSBDS-3; P ⫽ .05) compared
with controls. Generally, the fold change in apoptosis was markedly higher than that in proliferation (Figure 3B). For example, on
day 5 of differentiation, the increase in apoptosis in K562/
shSBDS-3 cells was 3.6-fold compared with only a 1.29-fold
decrease in proliferation. Because the percentage of cells at
different stages of the cell cycle did not indicate accumulation
at a specific phase (supplemental Figure 5), it is unlikely that
SBDS-knockdown erythroid cells are arrested at a specific cellcycle phase.
SBDS deficiency leads to increased levels of ROS during
erythroid differentiation
We previously showed that SBDS-deficient HeLa and TF-1 cells
have elevated ROS levels.16 It is unknown whether increased ROS
levels have a detrimental role during differentiation of SBDSdeficient hematopoietic cells. Using 2⬘,7⬘-dichlorofluoresceindiacetate, which is oxidized to the fluorescent compound, dichlorofluorescein by intracellular H2O2, we found higher ROS levels in
nondifferentiating SBDS-knockdown cells compared with controls.
Importantly, the difference in ROS levels between SBDSknockdown to control cells increased by 25% to 50% during differentiation (Figure 4A-C). In addition, we analyzed ROS levels in SBDSknockdown CD34⫹ cells from 2 donors after 5 days in erythroid
differentiation cultures. The SBDS-knockdown cells showed higher
ROS levels than mock-transduced cells (Figure 4D-F).
Antioxidants improve the SBDS-deficient cell expansion capacity
To ask whether elevated ROS levels impair the expansion of
SBDS-deficient cells, we performed rescue experiments with
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BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
Figure 4. Level of ROS. (A) For evaluation of intracellular ROS level, K562 cells (5 ⫻ 105) were washed and incubated for 30 minutes with 50M 2⬘,7⬘-dichlorofluoresceindiacetate (DCFH-DA). Cells and analyzed by flow cytometry. Duplicate cells were analyzed before inducing to undergo erythroid differentiation (a representative graph
of 2 independent experiments). (B) Cells were analyzed 3 days after induction of erythroid differentiation (a representative graph of 2 independent experiments). (C) Calculation
of the fold increase in the dichlorofluorescein fluorescence relative to the control wild-type cells. (D-E) Purified CD34⫹ from 2 donors were transduced with lentivectors
containing either shSBDS-3 or shSCR control, sorted for expressing the YFP reporter gene, and induced erythroid differentiation. The cells were analyzed on day 5 for ROS
levels by DCFH-DA. (F) Calculation of the fold increase in the dichlorofluorescein fluorescence relative to the scrambled RNA expressing cells.
antioxidants. Addition of N-acetylcysteine to duplicate clonogenic
assays of 3 SDS patients increased BFU-E colony numbers by
283%, 10%, and 40%, respectively (average 111%, P ⫽ .01).
Addition of catalase to the 3 patients’ cultures increased BFU-E
colony numbers by 167%, 63%, and 27%, respectively (average
86%, P ⫽ .04; Figure 5A). The average increase in BFU-E colony
production from CD34⫹ from 5 normal controls by N-acetylcysteine and catalase was 11% (P ⫽ .41) and 30% (P ⫽ .24), respectively. Addition of N-acetylcysteine and catalase to the SBDSknockdown K562 cells, but not to controls, improved cell expansion
(Figure 5B).
Ribosome biogenesis is more severely affected in
differentiating erythroid cells
The apoptosis and proliferation data in Figure 3 support the notion
that loss of SBDS during erythroid differentiation affects erythrocyte expansion, mainly by enhancing cell death. It is unknown,
however, what events trigger apoptosis during differentiation of
SBDS-deficient cells. Because SBDS plays a role in ribosome
biogenesis18,31 and expanding cells require substantial amounts of
energy, we hypothesized that defects in ribosome biogenesis would
be exacerbated during erythroid cell development. We found that
nondifferentiating SBDS-knockdown cells displayed slightly reduced amounts of 80S subunits compared with controls (Figure
6A). By contrast, during erythroid differentiation, SBDSknockdown cells showed markedly impaired ribosome profiles
with substantially reduced polysomes (Figure 6B). K562/shSBDS-3 cells, which express ⬃ 5% of the normal SBDS protein
concentration, also showed global reduction in all ribosome
subunits, including 80S, 40S, and 60S subunits. Importantly,
reintroduction of SBDS into SBDS-knockdown cells normalized the
ribosome profile (Figure 6B), indicating that these alterations in
ribosome profile are specifically the result of inhibition of SBDS. It
is noteworthy that, after removal of dead cells, SBDS-knockdown
cells still manifested general reduction of 80S subunits, indicating
that this defect was not the result of accumulation of dead cells
(supplemental Figure 6).
Because previous studies provided conflicting data on the ratio
of 60S to 40S ribosome subunits in SBDS-deficient cells, we
assayed the ribosome profile in the presence of EDTA. Normal
levels and ratio of 40S and 60S subunits were detected in
nondifferentiating SBDS-knockdown cells (Figure 6C). On differentiation, we detected a general reduction in both 40S and 60S
subunits; however, the 40S/60S ratio was not affected (Figure 6D).
The impaired global translation of SBDS-deficient K562 is
heightened during erythroid differentiation
Next, we investigated whether the ribosome abnormalities found in
SBDS-deficient cells resulted in reduced global translation. Global
protein synthesis of nondifferentiating K562/shSBDS-3 cells was
significantly reduced to 72% of K562/SCR cells (Figure 7A). After
48 hours of erythroid differentiation, the reduction of protein
synthesis became more prominent as translation in K562/
shSBDS-3 cells was reduced to 43% of K562/SCR cells (Figure
7A). Reintroduction of SBDS restored global translation during
erythroid differentiation (Figure 7B). These results show that
SBDS is necessary for normal protein translation, particularly
during erythroid differentiation.
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BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
SBDS ROLE IN ERYTHROPOIESIS
6413
levels. Leucine did not reduce ROS levels as determined by the
DCFH-DA assay at both 6 hours and 24 hours after administration
on day 0, day 3, or day 4 of differentiation (data not shown). Thus,
we analyzed the expression of genes involved in ROS regulation in
SDS marrow mononuclear cells compared with controls. We found
dysregulation of several genes, which may act together to increase
ROS levels in SDS cells (supplemental Table 2). Nox4, which
encodes the superoxide-generating enzyme NADPH-oxidase 4,
was up-regulated in SDS cells. In addition, 3 genes that encode
proteins that reduce ROS levels were down-regulated: EPX, CYGB,
and FoxM1. EPX encodes eosinophil peroxidase, which catalyzes
the conversion of H2O2 and bromide to hypobromite and water.38
CYGB encodes cytoglobin, which scavenges various ROS.39 FoxM1
encodes a transcription factor that up-regulates the expression of
ROS scavenger genes.40
The p53 family members were not up-regulated in
SBDS-deficient cells
Figure 5. Rescue of SBDS-knockdown slow cell growth with antioxidants.
(A) SDS patients and healthy control bone marrow CD34⫹ cells were plated in
duplicates at a density of 1 ⫻ 103 cells/1 mL dish with serum-free containing
methylcellulose and a cytokine cocktail (SCF, GM-CSF, IL-3, IL-6, G-CSF, and
erythropoietin). Additional duplicate cultures were plated in the same conditions, but
with N-acetylcysteine or catalase. BFU-E colonies containing 50 or more cells were
scored after 14 days under an inverted microscope. (B) SBDS-knockdown and
control K562 cells were induced to undergo erythroid differentiation by plating
2 ⫻ 104 cells/35-mm dishes with hemin. Replicate cultures were either not treated or
treated with N-acetylcysteine 500␮M or treated with catalase 500 U/mL and assessed daily for cell numbers by trypan blue exclusion. The cell growth of the
SBDS-knockdown K562 cells after 5 days was significantly improved with N-acetylcysteine (P ⬍ .05) and catalase (P ⬍ .05) treatment.
Leucine improves translation and cell expansion capacity of
SBDS-deficient cells
To ask whether translation insufficiency has a detrimental effect on
the expansion of SBDS-deficient cells, we performed rescue
experiments in which a translation stimulator was added to marrow
CD34⫹ cell cultures from 3 SDS patients. The addition of leucine,
which activates the mTOR pathway and stimulates protein synthesis (Figure 8A),32-37 resulted in improved BFU-E colony growth in
2 of 3 patients, by 52% and 83%, respectively; whereas a third
patient did not respond (Figure 8B). Addition of leucine to
clonogenic assays of CD34⫹ cells from 5 normal controls resulted
in an increase by an average of 28% (P ⫽ .25). In addition, leucine
improved SBDS-deficient K562 cell expansion, which reaches a
significant difference on day 5 after differentiation. Addition of
leucine to K562/SCR (Figure 8C) and WT (data not shown) control
cells did not increase cell numbers.
Genes that regulate ROS levels are differentially expressed in
SDS marrow mononuclear cells
To determine whether the high ROS levels in SBDS-deficient
erythroid cells are secondary to the defect in global translation, we
treated SBDS-deficient K562 cells with leucine and evaluated ROS
Because p53 is activated in some disorders with defective ribosomes and translation,41-43 we asked whether the proapoptotic p53
family proteins were aberrantly expressed in SBDS-deficient cells.
As previously reported,44 our K562 cells were deficient in p53
protein by Western blotting and its sequence showed biallelic
nonsense mutations (data not shown). In addition, p63 protein was
not detectable in K562 cells by Western blot analysis (data not
shown). The proapoptotic p73 isoform TAp73 was expressed in
K562 cells. TAp73 expression decreased during differentiation
without significant difference between the SBDS knockdown and
control cells (supplemental Figure 7A). We also analyzed the
expression of p53, p63, and TAp73 in erythrocytes from BFU-E
colony cells derived from CD34⫹ cells plated in the presence of
erythropoietin and SCFs. After 14 days, only BFU-E colonies were
observed in the clonogenic assays. The expression of p53 and
TAp73 proteins in SDS patients’ BFU-E colony cells was similar to
that of the controls. p63 protein expression was slightly higher (by
15%-50%) in the SDS patients than the normal control cells
(supplemental Figure 7B).
Discussion
Our findings suggest that SBDS is critical for normal erythropoiesis, and its deficiency impaired erythroid cell expansion. These
data support that the anemia seen in most SDS patients is largely
the result of primary bone marrow failure. In contrast to the
impaired cell expansion of SBDS-deficient cells, erythroid differentiation program was maintained because hemoglobinization, TFR2
and GATA1 expression, and the proportion of normoblasts in
patient marrows were not reduced. The mild elevation of proerythroblasts in patient bone marrows and the increase in hemoglobinization and GATA1 and TFR2 expression in K562 cells might
reflect a positive feedback mechanism and stress erythropoiesis.
Interestingly, previous studies using murine SBDS-deficient 32Dcl3
cells reported normal neutrophil differentiation.5 These data along
with ours suggest that the hematopoietic defects in SDS are mainly
the result of reduced expansion capacity of differentiating cells
rather than a defect in the differentiation program itself. It is
intriguing that a dramatic effect on global protein synthesis in
SBDS-deficient cells had no negative effect on differentiation. This
suggests that the balance between pro-differentiation and antidifferentiation factors allows differentiation in SBDS-deficient cells.
ROS play a critical role in inducing exit of HSC/Ps from a
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6414
SEN et al
BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
Figure 6. Ribosome profiles during differentiation of SBDS-knockdown cells. (A) Undifferentiated control and SBDS-knockdown K562 cells
were lyzed using TMK100 buffer with cycloheximide. Equal amounts of
RNA were layered on 5% to 45% sucrose gradients. Absorbance of each
fraction at A254 was done by the Brandel Density Gradient Fractionator.
The 40S, 60S, 80S, and polysomes peaks are seen on the graphs. A
representative diagram of 3 independent experiments is shown.
(B) SBDS-knockdown and control K562 cells were induced to undergo
erythroid differentiation with hemin. After 72 hours, cells were lyzed and
analyzed for ribosome profile as described in panel A. A representative
diagram of 2 independent experiments is shown. Arrows are pointed to the
markedly reduced polysomes in SBDS-knockdown cells. (C) The ribosomal profiles of undifferentiated control and SBDS-knockdown K562 cells
were analyzed by sucrose gradient density centrifugation under conditions
that allowed dissociation of the 40S and 60S subunits. Cells were
harvested using TMK100 lysis buffer with cycloheximide and EDTA. Equal
amounts of RNA were layered on 5% to 35% sucrose gradients. Absorbance of each fraction at A254 was done by the Brandel Density Gradient
Fractionator. The ratios between 60S and 40S subunits were calculated by
measuring area under the curve, using Adobe Illustrator, and are depicted
under the curves. A representative diagram of 2 independent experiments
is shown. (D) Ribosome profiles of differentiated K562 cells after 48 hours
of hemin induction under EDTA dissociating conditions as described in
panel C. A representative diagram of 2 independent experiments is shown.
quiescent state into differentiation.45 This raises the hypothesis that
elevated ROS levels in SBDS-deficient cells play a role in this
process.
For differentiation of K562 cells, we used hemin, which is
known to increase hemoglobinization in normal erythroid precursors. Hemin was extensively used to study the erythroid differentiation program. It is a ferric chloride heme salt, which induces
erythroid differentiation of HSC/Ps46 and erythroleukemia cell
lines.47 The precise mechanism by which hemin induces differentiation is unclear, but it may involve modulating the activity of
transcription factors, such as Oct-1 and GATA-1, or generating
ROS, such as superoxide. When applying identical hemin concentration to the SBDS knockdown and control cells, we identified
major differences between the cell types, suggesting a different
erythrocyte response to differentiation inducers in the presence or
absence of SBDS. Further, the aberrant expansion, survival, and
oxidative stress of SBDS knockdown CD34⫹ HSC/Ps and the
positive effect of antioxidants on SDS marrow cells suggest that the
observed phenotype is not restricted to the usage of hemin and
K562 cells.
We found that SBDS is up-regulated early during erythroid
differentiation. SBDS expression in murine myeloid 32Dcl3 cells
undergoing neutrophil differentiation also showed an early upregulation. Interestingly, the expression pattern of the murine
homologs of a few other inherited bone marrow failure genes, such
as FANCC48 and RPS1949 followed a similar pattern. These data
suggest a higher need for certain inherited bone marrow failure
syndrome genes during early stages of hematopoietic cell development, when most of the expansion occurs. This is in contrast to the
severe congenital neutropenia gene, ELA2, which is expressed
mainly at the promyelocyte stage of neutrophils development.50
We showed that during erythroid differentiation SBDS deficiency leads to accelerated apoptosis. Because oxidative stress can
activate apoptosis and because we have previously found high
levels of ROS in SBDS-deficient cells, we asked whether high ROS
levels impair the expansion of SBDS-deficient erythoid cells. The
high ROS levels during differentiation and the ability of antioxidants to substantially improve cell expansion and colony formation
of SBDS-deficient cells suggest that reduced expansion of differentiating erythroid cells is at least in part the result of high ROS
levels. Furthermore, these ROS abnormalities may be the result of
up-regulation of NOX4 and down-regulation of EPX, CYGB, and
FOXM1 in these cells. Nox4 protein acts as an oxygen sensor and
generates superoxide by transferring electrons from NADPH to
molecular oxygen. Superoxide is converted to H2O2 by superoxide
dismutase. Nox4 is expressed in various cells, including myeloid
cells.51 In addition, 3 genes that are involved in ROS metabolism
were down-regulated. EPX encodes eosinophil peroxidase, which
metabolizes H2O2.38 It is expressed in various cells, including
platelets.52 CYGB encodes cytoglobin, a ubiquitously expressed
hexacoordinate hemoglobin that scavenges various ROS and
provides a protective function during oxidative stress.39 FoxM1
encodes a transcription factor that up-regulates the expression of
ROS scavenger genes, such as catalase.40 In future studies, the
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BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
SBDS ROLE IN ERYTHROPOIESIS
6415
enhanced during differentiation, but to lesser degree than S3, which
is probably the result of a higher level of SBDS (⬃ 10%). Having a
more severe phenotype during erythrocyte maturation suggests that
SBDS regulation of ribosome biogenesis is particularly critical
during differentiation and might be related to the fact that the
Figure 7. Global translation in SBDS-deficient cells. (A) Global translation in
K562/shSBDS-3 and K562/SCR was studied by measuring incorporation of 35Smethionine/cysteine into equal amounts of newly synthesized trichloroacetic acidprecipitable peptides. Data are the mean ⫾ SE of 3 experiments. Statistically
significant results using t test: *P ⬍ .05. The ratios of the knockdown cells to control
are presented in percentages. (B) Global translation in stable SBDS-knockdown cells
after reintroduction of SBDS. Data are the mean ⫾ SE of 3 experiments. *P ⬍ .05,
statistically significant results using 1-way ANOVA and Tukey test for multiple means
against both S-3 ⫹ YPET and S-3 cells.
activity of these enzymes and factors in SBDS-deficient myeloid
cells and the potential rescue by manipulating their levels have to
be evaluated. It is interesting that apoptosis was accelerated in
SBDS-knockdown K562 cells despite high GATA1 expression.
Because apoptosis of SBDS-deficient marrow cells is mainly
through the Fas pathway,10,11,13 it is possible that the positive effect
of GATA1 on BCL-XL and cell survival was not apparent.
Loss of SBDS protein in differentiating erythroid cells resulted
in a more severe reduction in all ribosome subunits, polysomes, and
global translation than in nondifferentiating cells. SBDS deficiency
has been suggested to interfere with TIF6 release from the 60S
ribosomal subunit, thereby impairing its binding to 40S.18 Our
results showing a reduced level of the mature ribosome subunits
(80S) are consistent with these data. During differentiation, depletion of SBDS to very low levels by the shSBDS-3 cassette (⬃ 5%
of normal) resulted in reduced levels of 40S, 60S, 80S, and
polysomes. This is consistent with the complex ribosome maturation defects observed in yeasts depleted of the SBDS homolog
SDO1 by Moore et al.46 They showed a delay in 35S and 23S
pre-rRNA processing, which suggests defects in cleavage of the 5⬘
end of 18S rRNA and in maturation of the 40S ribosome subunit.
They also showed a delay in 27S rRNA processing, which suggests
defects in cleavage between 5.8S and 25S and in maturation of the
60S ribosome subunit. The defect in K562/shSBDS-1 was also
Figure 8. Leucine treatment of SBDS-deficient cells. (A) Translation in S-3 cells
and SCR cells was studied by measuring incorporation of 35S-methionine/cysteine
into equal amounts of newly synthesized trichloroacetic acid-precipitable peptides.
Data are the mean ⫾ SE of 3 experiments. *P ⬍ .05, statistically significant results
(t test). (B) SBDS-knockdown and control K562 cells were induced to undergo
erythroid differentiation by plating 2 ⫻ 104 cells/35-mm dishes with hemin. Replicate
cultures were either treated or not treated with 600 ␮g/mL of leucine and assessed
daily for cell numbers by trypan blue exclusion. The cell growth of the SBDSknockdown K562 cells after 5 days was significantly improved with leucine treatment
(P ⬍ .05). (C) SDS patients and healthy control bone marrow CD34⫹ cells were
plated in duplicates at a density of 1 ⫻ 103 cells/1 mL dish with serum-free containing
methylcellulose and a cytokine cocktail (SCF, GM-CSF, IL-3, IL-6, G-CSF, and
erythropoietin). Additional duplicate cultures were plated in the same conditions, but
with leucine. BFU-E colonies containing 50 cells or more were scored after 14 days
under an inverted microscope.
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6416
BLOOD, 8 DECEMBER 2011 䡠 VOLUME 118, NUMBER 24
SEN et al
hematopoietic system is specifically targeted in SDS. Erythrocytes
have a particularly high demand for rRNA and protein synthesis
during early stages of erythropoiesis. Therefore, deficiency of
SBDS might limit the ability of the cells to meet this demand,
thereby affecting their survival and expansion capacity. Because
ribosome profile remained abnormal after removing dead cells, it is
unlikely that the reduced ribosome biogenesis and translation are
secondary to accelerated apoptosis. However, we cannot exclude
the possibility that aberrant growth signal leads to the reduced
ribosome biogenesis and translation.
An important unanswered question is whether abnormalities in
ribosome biogenesis in SDS are the primary cause of the cell
expansion defect. The protein synthesis stimulator leucine, improved colony production from SDS marrow CD34⫹ cells in 2 of 3
patients. Leucine significantly improved the expansion capacity of
SBDS-knockdown K562 cells, but not of the controls. It is
noteworthy that leucine did not completely abrogate the growth
defects of SDS CD34⫹ cells and SBDS-knockdown K562 cells.
Thus, is it probable that translation insufficiency contributes to, but
is not the sole cause of, the hematopoietic failure in SDS.
Alternatively, it is possible that more potent translation stimulators
are necessary as leucine did not completely correct that global
translation defect in SBDS-deficient cells. Because leucine did not
improve ROS levels, it is probable that the mechanism of increased
ROS levels is independent of the defect in translation insufficiency.
Our gene expression data showing dysregulation of enzymes
involved in ROS regulation are in agreement with this hypothesis.
Our analysis of human myeloid cells showed that SBDS
deficiency leads to a similar reduction of all ribosome subunits.
These data are consistent with results from human SBDS-deficient
fibroblasts, where no reduction in the relative ratio between the 60S
and 40S subunits was shown.19 Together with the results of our
previous study showing reduced expression of multiple ribosomal
protein genes of both the small and large ribosome subunits in SDS
marrow cells,17 we conclude that SBDS is important for the
integrity of all ribosome subunits by maintaining normal levels of
its protein and rRNA components.
p53 has been suggested to mediate apoptosis in disorders of
ribosome biogenesis.41-43,53 The role of p53 in causing expansion
defects of maturing hematopoietic cells in SDS is unknown. The
findings that K562 cells do not express p53 and that the expression
of the protein in SDS erythroid colony cells was not higher than the
controls suggest that the accelerated apoptosis in SBDS-deficient
cells is independent of p53. The expression patterns of p73 and p63
in K562 cells and in patient erythroid cells suggest that also these
genes are not related to accelerated apoptosis in SDS. Because p73
and p63 exist in multiple isoforms, our results do not exclude the
possibility of pathogenetic significance of their particular isoforms.
To further provide evidence of a lack of a role of the p53 family
members in the reduced SDS hematopoietic cell expansion, it
would be useful to inhibit the genes before induction of differentiation. However, the limited numbers of SDS HSC/Ps12 and their
substantial sensitivity to in vitro manipulation render these experiments difficult to perform.
This study suggests that SBDS is critical for normal erythropoiesis. Translation is decreased, apoptosis and ROS levels are
increased, and cell expansion is decreased during erythroid differentiation. We also showed that antioxidants and stimulators of
protein synthesis improve the expansion capability of SBDSdeficient cells, providing the rationale for devising future preclinical and clinical studies to test the safety and efficacy of these agents
as novel therapeutic strategies in SDS.
Acknowledgments
The work was supported by the Canadian Institute for Health
Research (grant HPA161281) and Shwachman-Diamond Syndrome Canada. S.S. is a recipient of an Ontario Graduate Scholarship Award and the Hospital for Sick Children Research Competition Award.
Authorship
Contribution: S.S. performed research, analyzed data, and wrote
the paper; H.W., C.L.N., K.Z., and J.Y. performed research and
analyzed data; C.S.T. and M.S.I. performed research, contributed to
research design, and wrote the paper; and Y.D. conceived and
designed research, analyzed data, and wrote the paper.
Conflict-of-interest disclosure: The authors declare no competing financial interests.
Correspondence: Yigal Dror, Program in Cell Biology, Research Institute, and Marrow Failure and Myelodysplasia Program,
Division of Haematology/Oncology, Department of Paediatrics,
Hospital for Sick Children and the University of Toronto,
555 University Avenue, Toronto, ON M5G 1X8, Canada; e-mail:
[email protected].
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From www.bloodjournal.org by guest on June 17, 2017. For personal use only.
2011 118: 6407-6417
doi:10.1182/blood-2011-02-335190 originally published
online September 30, 2011
The ribosome-related protein, SBDS, is critical for normal erythropoiesis
Saswati Sen, Hanming Wang, Chi Lan Nghiem, Kim Zhou, Janice Yau, Chetankumar S. Tailor,
Meredith S. Irwin and Yigal Dror
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