From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
Regular Article
RED CELLS, IRON, AND ERYTHROPOIESIS
Cytokinesis failure in RhoA-deficient mouse erythroblasts involves
actomyosin and midbody dysregulation and triggers p53 activation
Diamantis G. Konstantinidis,1,* Katie M. Giger,1,* Mary Risinger,2 Suvarnamala Pushkaran,1 Ping Zhou,1 Phillip Dexheimer,3
Satwica Yerneni,3 Paul Andreassen,1 Ursula Klingmüller,4 James Palis,5 Yi Zheng,1 and Theodosia A. Kalfa1
1
Cancer and Blood Diseases Institute, Cincinnati Children’s Hospital Medical Center, University of Cincinnati College of Medicine, Cincinnati, OH; 2College
of Nursing, University of Cincinnati, Cincinnati, OH; 3Biomedical Informatics, Cincinnati Children’s Hospital Medical Center, University of Cincinnati College
of Medicine, Cincinnati, OH; 4Systems Biology of Signal Transduction, German Cancer Research Center, Center for Molecular Biology Alliance, Heidelberg,
Germany; and 5Department of Pediatrics, Center for Pediatric Biomedical Research, University of Rochester Medical Center, Rochester, NY
RhoA GTPase has been shown in vitro in cell lines and in vivo in nonmammalian
organisms to regulate cell division, particularly during cytokinesis and abscission,
when 2 daughter cells partition through coordinated actomyosin and microtubule
• RhoA GTPase activates
machineries. To investigate the role of this GTPase in the rapidly proliferating
pMRLC and localizes to the
mammalian erythroid lineage, we developed a mouse model with erythroid-specific
site of midbody formation to
deletion of RhoA. This model was proved embryonic lethal as a result of severe
regulate erythroblast
anemia by embryonic day 16.5 (E16.5). The primitive red blood cells were enlarged,
cytokinesis.
poikilocytic, and frequently multinucleated, but were able to sustain life despite
• Cytokinesis failure in
experiencing cytokinesis failure. In contrast, definitive erythropoiesis failed and the
erythroblasts caused by RhoA mice died by E16.5, with profound reduction of maturing erythroblast populations
deficiency triggers p53within the fetal liver. RhoA was required to activate myosin-regulatory light chain
mediated DNA-damage
and localized at the site of the midbody formation in dividing wild-type erythroresponse, cell-cycle arrest,
blasts. Cytokinesis failure caused by RhoA deficiency resulted in p53 activation and
and apoptosis.
p21-transcriptional upregulation with associated cell-cycle arrest, increased DNA
damage, and cell death. Our findings demonstrate the role of RhoA as a critical
regulator for efficient erythroblast proliferation and the p53 pathway as a powerful quality control mechanism in erythropoiesis.
(Blood. 2015;126(12):1473-1482)
Key Points
Introduction
The first circulating “primitive” erythroid cells in the mouse embryo
emerge in blood islands of the yolk sac at around embryonic day 7.5
(E7.5) and remain the only circulating erythroid cells until E12.5,
transporting oxygen to all tissues of the rapidly growing embryo.1
They are characterized by their large size, the presence of a nucleus,
and the expression of embryonic hemoglobins.2 Primitive erythroblasts continue to mature and divide in circulation and enucleate
between E12.5 and E16.5 after interactions with the macrophages of
the fetal liver.1,3 As the embryo increases in size, growth and life
cannot be sustained by the limited potential of primitive erythropoiesis; the vastly more numerous definitive red blood cells (RBCs) begin
to be released from the fetal liver at ;E12.5, enucleated and containing
adult hemoglobin.4 When primitive erythropoiesis fails, embryos
do not survive beyond E9.5 to 10.5, whereas disruption of genes
necessary for definitive erythropoiesis causes fetal demise after
;E15.5.5 No other normal mammalian tissue proliferates as fast as
the erythroid lineage, which produces in the adult human at steadystate 2 million new RBCs per second. The erythroid proliferation rate
is even faster during embryonic development in which a 70-fold
increase in the red cell mass has been estimated to occur in fetal
mice in the period E12.5 to E16.5 of gestation.6 It is clear that any
disruption of the cell division mechanism would have a detrimental
effect on the efficiency of erythropoiesis.
RhoA, a member of the Rho GTPase family of proteins, is a major
regulator of actomyosin contractility and vesicular trafficking,7,8
processes that play a significant role in cytokinesis, the final stage
of cell division.9 Studies in urchin and frog cells have shown that
microtubules creating the mitotic spindle determine the position of
the cleavage furrow via localization of active RhoA to this zone.10
After actomyosin ring contraction and cleavage furrow ingression,
the 2 daughter cells remain connected via the midbody, a minute
cytoplasmic bridge that contains microtubules.11 Abscission, the
separation of the 2 daughter cells, requires new membrane formation,
likely through vesicular trafficking.11,12 Our understanding of the
role of RhoA in cytokinesis in mammalian cells has come mainly
from work in cell lines using dominant-negative and constitutively
active mutants of RhoA and its effectors to inhibit or overstimulate RhoA-related signaling. Evaluation of these pathways in vivo
Submitted December 9, 2014; accepted July 20, 2015. Prepublished online as
Blood First Edition paper, July 30, 2015; DOI 10.1182/blood-2014-12-616169.
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.
*D.G.K. and K.M.G. contributed equally to this study.
The online version of this article contains a data supplement.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
© 2015 by The American Society of Hematology
1473
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
1474
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
KONSTANTINIDIS et al
Figure 1. EpoR-CreTg/1; RhoAflox/flox (RhoAD/D) mice
with erythroid-specific RhoA deficiency die in utero
by E16.5 because of failure of definitive erythropoiesis. (A) Representative results of embryo genotyping
by PCR (somatic DNA) show bands for EpoR-Cre (top,
WT:431bp, Cre1:679bp) and RhoA (bottom, WT:482bp,
flox:633bp) for all possible genotypes of the offspring of
an EpoR-CreTg/1;RhoAWT/flox 3 EpoR-Cre2/2;RhoAflox/flox
timed pregnancy. (B) Immunodetection of RhoA protein in peripheral WT and RhoAD/D blood cells from
E11.5 embryos (top panel) and in CD711 cells from
WT and RhoAD/D E13.5 fetal livers (bottom panel),
using a size-based capillary electrophoresis instrument,
shows significant reduction of RhoA in the RhoAD/D
embryos. (C-D) Stereoscopic images of E13.5 embryos showing severe anemia in RhoAD/D yolk sac (C),
as well as pallor and small fetal liver in RhoAD/D
embryos (D). The scale bar represents 1 mm. (E-F)
Light microscopy images of peripheral blood cytospins
showing poikilocytosis with large and frequently multinucleated primitive RhoAD/D erythroid cells in contrast to the
homogeneous population of WT primitive red cells (E);
hematoxylin and eosin–stained yolk sacs show the paucity
of primitive erythroid cells in the RhoAD/D yolk sac vessels
(F). The scale bar represents 10 mm. (G) The offspring of
EpoR-CreTg/1;RhoAWT/flox 3 EpoR-Cre2/2;RhoAflox/flox follow Mendelian ratios up to E14.5. By E15.5, only 6% of
live embryos are RhoAD/D, with none being alive by
E16.5. Data based on genotyping of .60 embryos per
each time point. (H) RhoAD/D embryos exhibit anemia
already by E11.5, worsening significantly by E14.5. Data
are represented as mean 6 SEM of blood counts for at
least 3 embryos per genotype per each time point. *P , .05
of RhoAD/D vs WT.
has been hampered by the fact that mice with constitutional deletion
of RhoA could not be created because of very early embryonic
lethality.13,14
In this study, we investigate the role of RhoA in vivo in the
erythroid lineage using a Cre-lox recombination system in which
Cre-recombinase expression is controlled by the erythropoietin
receptor (EpoR) promoter, thereby resulting in erythroid-specific
deletion of the floxed RhoA gene.15 We found that RhoA is essential for cytokinesis in both primitive and definitive erythroid
lineages. Defective cytokinesis in RhoA-deficient erythroids manifested as polyploidy and maturation delay and was accompanied by
increased phosphorylation of p53 and transcriptional upregulation
of p21, leading to cell-cycle arrest and increased cell death. Although
frequently multinucleated and dysplastic, RhoA-deficient primitive erythroid cells were able to support survival of the embryo,
whereas failure of definitive erythropoiesis led to in utero demise
by E16.5. These data reveal the important role of RhoA during
maturation and expansion of the rapidly proliferating erythroid
lineages, the associated quality control mechanisms that manifest in
RhoA-deficient cells, and their differential effects in definitive vs
primitive erythropoiesis.
Methods
Mice
All mouse protocols were approved by the Institutional Animal Care and Use
Committee of Cincinnati Children’s Hospital Medical Center. Our experimental
mouse colony was established by crossing mice with conditional RhoA alleles
(RhoAflox/flox),16 where exon 3 of the RhoA gene is flanked by loxP sites
(supplemental Figure 1, available on the Blood Web site), with EpoR-CreTg/1
mice,15 where Cre recombinase expression is controlled by the promoter
of the erythropoietin receptor. The resulting EpoRCreTg/1;RhoAWT/flox and
EpoRCre2/2; RhoAWT/flox siblings were crossed together and their offspring
were backcrossed for at least 8 generations on a C57/BL6 background. To
have an easily detectable Cre-reporter, the EpoR-CreTg/1 mice were bred with
B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J, where a loxP-flanked STOP cassette prevents transcription of the downstream red fluorescent protein variant
tdTomato, in the cells not expressing Cre.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
CYTOKINESIS FAILURE IN RhoA-DEFICIENT ERYTHROIDS
1475
methanol to a final concentration of 90% methanol. Samples were immediately vortexed and kept on ice for 30 minutes. Cells were then stained with
anti-gH2AX antibody (Millipore; 1:300) 30 minutes at RT, followed by
staining with anti-mouse-AF488 (1:200), anti-Ter119-APC (BD Biosciences), and DAPI (5 mg/mL final concentration) and visualized on ImageStreamX (603/0.9NA objective). Erythroblast populations were gated as
previously described,18 and multinucleated cells were identified by direct visual
observation of all events within the Ter1191DAPI1 population with large
nuclear size (increased area_DAPI) and elongated nuclear shape (low aspect
ratio_DAPI). Total and multinucleated Ter1191DAPI1 cells were analyzed for
staining intensity for gH2AX.
Statistics
Statistical analysis was performed using KaleidaGraph software, v.4.1 (Synergy
Software). Comparison between 2 groups with values assuming normal distribution was performed using Student t test for unpaired data with equal
variance (results presented as mean 6 standard error of the mean [SEM]).
The ImageStreamX data of gH2AX intensity in the experiments shown in
Figure 6B-C were exported for statistical evaluation. Because the distribution
of the values for gH2AX intensity appears asymmetric indicating nonnormality, we used the Wilcoxon rank-sum test, which makes no assumptions
about the underlying probability distributions and is thus appropriate for
Figure 2. RhoAD/D primitive erythroid cells are often multinuclear. (A) PI staining of
E12.5 primitive circulating erythroid cells reveals a significant number of 4N, .4N, as well
as ,2N cells in the RhoAD/D population compared with the almost exclusively 2N cells of
the WT population. The graphs are representative of 3 biological repeats. Insets:
representative imaging flow cytometry pictures of brightfield and Syto-16–stained WT
and RhoAD/D primitive erythroid cells within the 2N (top) and 4N gate (bottom). (B)
Graphic representation as mean 6 SEM from the data of 3 experiments (3 different
biological repeats) of PI staining of E12.5 primitive erythroid cells. *P , .005 of RhoAD/D
vs WT. (C) Imaging flow cytometry reveals that tetraploid (4N) and .4N RhoAD/D
primitive erythroid cells do not appear to be undergoing mitosis. Cells were stained with
the nuclear stain Syto16. Micronuclei or nuclear fragments (arrows) can be observed in
the .4N population.
Timed pregnancies and embryo harvest
After pairing EpoRCreTg/1;RhoAWT/f with EpoRCre2/2;RhoAf/f mice for
24 hours, females were pulled and housed in separate cages. On embryonic days
E11.5 to 16.5 (based on the date of pairing), pregnant females were euthanized
and embryos were collected, as detailed in supplemental Methods.
Fixation, permeabilization, and staining for imaging
flow cytometry
Fixation, permeabilization, and staining of dividing erythroblasts for
imaging flow cytometry was performed as previously described.17 In brief,
wild-type (WT) mice subjected to phlebotomy (removal of 300 mL blood
for 3 consecutive days to induce stress erythropoiesis) were euthanized
3 days after last phlebotomy and the spleens collected. Splenocytes were
pelleted at 1600 rpm/3 min in a bench-top centrifuge, fixed in phosphatebuffered saline containing 4% formaldehyde for 15 minutes at room
temperature (RT), and permeabilized by consecutive suspensions in icecold 50% acetone, 100% acetone, and again 50% acetone solution. Cells
were then incubated with anti-RhoA (1:50; Santa Cruz Biotechnology),
washed in fluorescence-activated cell sorting buffer, and incubated with
anti–b-tubulin-AF488 (1:200; Cell Signaling), anti-mouse-AF594 (1:100;
Life Technologies), and the nuclear stain 49,6 diamidino-2-phenylindole
(DAPI) (Life Technologies). Samples were counted on the ImageStreamX
(Amnis) using a 603/0.9NA objective. At least 10 000 events per
experimental sample were collected and analyzed with the associated
Image Data Exploration and Analysis Software (IDEAS; Amnis).
DNA damage detection through gH2AX
Fetal liver cells were fixed with 3.7% formaldehyde in PBS for 10 minutes at
37°C and then put on ice for 1 minute. Permeabilization was attained by
adding (without removing the formaldehyde) 900 mL of ice-cold 100%
Figure 3. Fetal liver cellularity is severely reduced and erythroblast maturation
delayed in RhoAD/D embryos. (A) Beginning at E12.5, RhoAD/D fetal livers have
reduced cellularity compared with the WTs. Cellularity is unaffected in the heterozygous
(RhoAWT/D) fetal livers. Data are represented as mean 6 SEM of the cell count of at least
5 fetal livers for each embryonic date. *P , .05 of RhoAD/D vs WT. (B) The number of cells
per fetal liver with BFU-E (left panel) and CFU-E (right panel) activity is unaffected in E12.5
RhoAD/D embryos compared with the WT. Data are represented as mean 6 SEM of 3
fetal liver samples per each genotype, each sample evaluated by colony assay in triplicate
plates. (C-D) Flow cytometry analysis of fetal liver cells from E12.5 embryos shows that
RhoAD/D erythroblasts diminish in number as they move through the stages of erythroid
maturation. Flow cytometry dot plots are representative of 3 different biological repeats
(C). Bar graph of mean 6 SEM of the cell count per each erythroid population as defined
by CD71-Ter119 analysis of 3 fetal livers per each genotype (D). *P , .05 of RhoAD/D vs
WT. (E) Touch preps prepared from E13.0 fetal livers show an abundance of larger, more
immature cells in RhoAD/D fetal livers compared with the WT. Binucleated erythroblasts
(arrows) are evident in RhoAD/D fetal livers, though they are less prominent than among
the primitive erythroid cells in circulation. The scale bar represents 10 mm.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
1476
KONSTANTINIDIS et al
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
Figure 4. RhoAD/D fetal liver cells exhibit increased cell-cycle arrest and cell death. (A) Fetal liver sections stained for the cell-proliferation marker Ki67 show that the RhoAD/D
binucleated cells in the fetal liver (arrows) are not actively proliferating, implying an abnormal arrest in the binucleated state and failure to complete cytokinesis. The scale bar represents
10 mm. (B) Graphic representation of the gating used to separate fetal liver cell populations in Figure 4C-E, defining the erythroid progenitors S0 (CD71–;Ter119– cells) and
S1 (CD711;Ter119– cells) based on CD71 level, and the erythroid precursors E1-E4 based on CD71 and Ter119 levels and size (forward scatter), as detailed in supplemental
Figure 2. (C-D) Cell-cycle analysis of WT and RhoAD/D fetal liver cells from E14.5 embryos using in vivo BrdU assay shows a significant increase of the polyploid cells (.4N)
and the G2/M population in the RhoAD/D late erythroblasts (E2 and E3 populations) along with a decrease in the S phase. Flow cytometry dot plots are representative of
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
CYTOKINESIS FAILURE IN RhoA-DEFICIENT ERYTHROIDS
1477
non-normal data. For each set of measurements, we report the mean and
median for each group and the 2-tailed P value according to the Wilcoxon
rank-sum test. Significance was set at P , .05.
The remaining methods for this article are detailed in the supplemental
Materials.
Results
Erythroid-specific RhoA deficiency results in embryonic
lethality caused by failure of definitive erythropoiesis
To investigate the role of RhoA in erythropoiesis, we generated mice
with conditional RhoA alleles16 and Cre-recombinase expression
controlled by the EpoR promoter,15 resulting in erythroid-specific
RhoA deficiency (Figure 1A-B). Although primitive erythropoiesis
is not completely dependent upon EpoR,19 EpoR mRNA expression
is detected in yolk sac blood islands in E8.5 embryos,20 and EpoRnull embryos generate only 10% normal numbers of late-stage primitive erythroblasts.21 The associated Cre recombinase appears to
be active at variable levels, as is frequently the case with the Cre
system,22 and causes deletion of the floxed RhoA gene in both
primitive and definitive erythroblasts, as demonstrated by significant
decrease of the RhoA protein in E11.5 peripheral blood cells and
E13.5 fetal liver erythroblasts (Figure 1B). Mice not expressing Cre
(EpoR-Creneg;RhoAflox/flox or EpoR-Creneg;RhoAWT/flox, hereafter called
WTs) or heterozygotes for RhoA deletion (EpoR-CreTg/1;RhoAWT/flox or
RhoAWT/Δ) were born healthy and fertile, whereas mice with erythroidspecific RhoA deficiency (EpoR-Cre Tg/1 ;RhoAflox/flox , hereafter
called RhoAD/D) died in utero by E16.5. Timed pregnancies between
EpoR-Cre2/2;RhoAflox/flox and EpoR-CreTg/1;RhoAWT/flox parents were
used to investigate the causes of the lethality of erythroid-specific RhoA
deficiency. Anemia was pronounced in the RhoAD/D embryos with
the yolk sac blood vessels being barely visible as a result of the paucity
of RBCs and the embryos being pale with small livers (Figure 1C-D).
The primitive erythroid cells in their circulation were generally larger
than the WT, heterogeneous in size and shape, and frequently
binucleated or multinucleated (Figure 1E-F). Genotypes of live
embryos roughly followed the expected Mendelian ratio with 25%
EpoR-CreTg/1;RhoAflox/flox embryos up to E14.5, but that ratio
dropped to zero by E16.5 (Figure 1G). Peripheral blood counts of
RhoAD/D embryos demonstrated anemia already at E11.5, indicating
effects on primitive erythropoiesis, which worsened precipitously by
E14.5 (Figure 1H), leading to fetal death. To pinpoint the stage of
erythroid differentiation in which Cre recombinase starts being expressed and hence RhoAflox/flox becomes deleted, we bred EpoR-CreTg/1
mice with Gt(ROSA)26Sortm9(CAG-tdTomato)Hze mice to use tdTomato as
a Cre-reporter.23 Analysis of the erythroid progenitors and precursors
in fetal livers from EpoR-CreTg/1;Gt(ROSA)26Sortm9(CAG-tdTomato)Hze
embryos using the CD71-Ter119 flow cytometric assay24 showed
that EpoR expression drives tdTomato expression already in a
fraction of the S0 subpopulation and in 100% of the S1 subpopulation containing erythroid colony-forming cells (CFU-E),25 as well
as in the erythroblasts throughout the further stages of erythroid
differentiation in population E1 and all Ter1191 cells (supplemental
Figure 2).
Figure 5. RhoA regulates myosin regulatory light-chain phosphorylation and
microtubule organization during erythroblast cytokinesis. (A) Myosin regulatory
light-chain phosphorylation is decreased in E14.5 RhoAD/D fetal liver cells as shown
by western blotting. (B) RhoA colocalizes with b-tubulin in WT erythroblasts undergoing mitosis, possibly participating in the formation of the microtubule-derived
midbody structure observed in cytokinesis. The nuclear stain DAPI was used to
indicate the nucleus. Four representative cells are shown of at least 30 dividing cells
with similar morphology. Images were obtained with a 603 objective lens by
ImageStreamX. (C) Citron kinase was significantly decreased in E13 RhoAD/D
Ter1191 fetal liver cells, as shown by capillary electrophoresis and immunodetection.
Circulating primitive RhoA-deficient erythroid cells
exhibit polyploidy
A striking characteristic of the RhoAD/D embryos was that the primitive
erythroids frequently exhibit polyploidy with $2 nuclei, indicating
a failure to complete cytokinesis. At E12.5, the majority of circulating
WT primitive erythroid cells are nucleated and predominantly diploid
(2N), with a few tetraploid (4N) primitive erythroblasts and the rest
being either enucleated primitive or definitive RBCs (,2N). In
contrast, the majority of the RhoAD/D circulating primitive erythroblasts
were tetraploid and hypertetraploid; cells with ,2N DNA content were
also increased, likely because of the presence of cells with nuclear
fragmentation as well as cytoplasmic fragments (Figure 2A-B).
Analysis by imaging flow cytometry confirmed that WT cells within
the 4N gate are undergoing mitosis, whereas the tetraploid RhoAD/D
cells did not appear to undergo cytokinesis (bottom panels in Figure 2A,
insets). RhoAD/D multinucleated cells frequently contained additional
micronuclei (Figure 2C), which may result from nonsegregation
of chromosomes in mitosis associated with cytokinesis failure or
formation of a multipolar spindle and unbalanced chromosome
segregation in tetraploid cells that fail to arrest in G1.26,27
RhoA is necessary for efficient definitive erythropoiesis in the
fetal liver
Despite the abnormal phenotype of the circulating primitive erythroid
cells, RhoAD/D embryos survived the period supported exclusively by
primitive erythropoiesis, indicating that primitive RBCs are able to
perform their function even with abnormal nuclear material, but died
by E16.5 when survival depends on fetal liver–derived definitive
erythropoiesis. Fetal livers were small and pale in RhoAD/D embryos
Figure 4 (continued) 5 different biological repeats (C). Bar graphs of mean 6 SEM of the percentage of each cell-cycle stage per erythroid population of 5 fetal livers for each
genotype (D). *P , .05 of RhoAD/D vs WT. (E) No significant difference was found in early apoptosis of the RhoAD/D erythroblasts (left panel); however, late apoptosis was
increased in E2, E3, and E4 RhoAD/D fetal liver cells, as evidenced by an increased number of cells in those populations positive for both annexin V and 7-AAD (middle panel).
Clearly necrotic cells (negative for annexin V but positive for 7AAD) were also found to be increased in RhoAD/D fetal liver erythroblasts (right panel). Data are shown as
mean 6 SEM from 6 WT and 4 RhoAD/D fetal livers. *P , .05 of RhoAD/D vs WT.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
1478
KONSTANTINIDIS et al
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
Figure 6. Cytokinesis failure results in DNA damage
in RhoAD/D fetal liver erythroblasts, as evidenced
by staining for the DNA damage marker gH2AX. (A)
Fetal liver cells from E14.0 WT and RhoAD/D embryos
were analyzed by imaging flow cytometry as shown
before,18 up to the gating of erythroblasts (Ter1191
DAPI1 cells). Further analysis based on the size of the
nucleus (area_DAPI) and the shape of the nucleus
(aspect ratio_DAPI5ratio of the minor axis/major axis)
gave a population (gated by the pink line) enriched in
binucleated cells. Truly binucleated or multinucleated
cells were then identified by visual observation and
marked blue in the dot plot. (B-C) Concurrent staining
for the DNA damage marker gH2AX revealed that
gH2AX, as measured in arbitrary units of fluorescence,
was significantly higher in the RhoA D/D erythroblasts
compared with WT. Considering a threshold for H2AX
positivity at 1.5 3 105, the percentage of gH2AXpositive cells was 18.5 6 4.8% for RhoAΔ/Δ Ter1191
nucleated fetal liver cells vs 3.1 6 1.3% for the WT
counterparts (B). The difference was more significant in
the multinucleated erythroblasts: 39 6 13.7% in RhoAD/D
vs 2.3 6 1.4% in the WT (C). Mean and median values of
the fluorescence intensity of gH2AX in the WT and
RhoAD/D erythroid precursors are shown (P , .0001 of
RhoA D/D vs WT). At least 5000 Ter1191;DAPI1 cells
were analyzed in each experiment and results are
representative of 3 biological repeats for each genotype.
(D) Binucleated RhoAD/D cells were strongly positive for
gH2AX, indicative of DNA damage. In contrast, binucleated WT cells were negative for gH2AX and likely normal
mitotic cells. Three representative cells are shown for
each genotype of at least 250 RhoAD/D and 40 WT
hyperdiploid cells with similar morphology. Images were
obtained with a 603 objective lens by ImageStreamX.
DAPI was used as a nuclear stain. (E) Fetal liver cells
from E14 embryos were stained for gH2AX foci and
imaged by confocal microscopy. Cells with .4 foci per
nucleus were counted as positive, and at least 50 cells
were evaluated per sample. The percentage of gH2AXpositive cells was 38.5% for RhoAΔ/Δ vs 2% for the WT.
Bottom panels show the nuclear stain merged with the
corresponding brightfield image to demonstrate the binucleated and dysplastic RhoAΔ/Δ cells. The scale bar
represents 10 mm.
compared with those of their WT littermates. From E12.5 on, fetal liver
cellularity of RhoAD/D embryos was significantly reduced (Figure 3A).
To investigate the stage at which definitive erythropoiesis fails, we
first performed colony assays using fetal liver cells (Figure 3B).
Erythroid burst-forming unit (BFU-E) colony numbers were normal
as expected, because EpoR-Cre expression is not established until
the CFU-E stage (supplemental Figure 2). CFU-E numbers were also
similar between the RhoAD/D and WT embryos despite the fact that
excision of the floxed RhoA gene is attained at this stage. CD71-Ter119
analysis of fetal liver cells24,28 demonstrated that RhoAD/D erythroblasts diminish in number as they mature (Figure 3C-D), resulting in
a severely decreased number of reticulocytes (population E4).29 Cell
sorting of cells after similar analysis revealed an increased incidence
of dysplastic, binucleated RhoAD/D fetal liver cells at stages E1-E3,
indicating pathologic cytokinesis of definitive erythroblasts (supplemental Figure 3). RhoAD/D fetal liver erythroblasts were larger
and had fewer condensed nuclei than their WT counterparts, as
demonstrated in touch-preps of the tissue. Binucleated erythroblasts were also more frequent in RhoAD/D fetal livers, though not as
prominent as within the primitive erythroid lineage in circulation
(Figure 3E). The increased ratio of early erythroblasts and the
paucity of mature erythroblasts suggested a differentiation defect
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
CYTOKINESIS FAILURE IN RhoA-DEFICIENT ERYTHROIDS
1479
Figure 7. DNA damage in RhoAD/D fetal liver erythroblasts leads to p53 and caspase activation. (A)
Gene Set Enrichment Analysis (GSEA) from RNA-Seq
data of CD711 cells isolated from E13.0 WT and RhoAD/D
fetal livers (n 5 3 for each genotype) demonstrating
upregulation of p53-related DNA damage response
genes39 in the RhoA-deficient erythroblasts. (B) The
upregulation of PHLDA3 and p21 (CDKN1A) mRNA
found by RNA-Seq was confirmed using RT-PCR on
RNA isolated from CD711 cells from E13.0 WT and
RhoAD/D fetal livers (n 5 3 for each genotype). Data
are represented as mean 6 SEM of fold-expression
relative to WT; expression normalized vs Actin-B and
vs glyceraldehyde-3-phosphate dehydrogenase (GAPDH)
is shown. (C) Increased p53 phosphorylation at Ser-15
is demonstrated by western blotting in CD711 cells
isolated from E13.0 RhoAD/D fetal livers compared with
WT. (D) Immunostaining on fetal liver touch-preps further
confirmed increased phosphorylation of p53 phosphorylated at Ser-15 in E13.0 RhoAD/D fetal liver cells. The
scale bar represents 10 mm.
(ie, reduced cell-cycle and proliferation capacity) and/or increased
loss of the RhoA-deficient erythroid precursors.
RhoA-deficient fetal liver erythroblasts exhibit disrupted cell
cycle and increased cell death
Based on the increased incidence of binucleated cells in RhoAD/D fetal
livers, we evaluated the proliferation status of these cells. Immunohistochemistry for the cell-proliferation marker Ki67 revealed that in
contrast to binucleated WT fetal liver cells that were positive for Ki67,
binucleated RhoAD/D fetal liver cells did not stain for Ki67, indicating
that they are not actively proliferating and thus implying cell-cycle
arrest (Figure 4A). Quantitative evaluation in vivo with BrdU assay
demonstrated that the cell cycle was similar in RhoAD/D and WT
erythroid progenitor S0-S1 populations and early erythroblasts
(E1).25,29 However, the RhoA-deficient terminal erythroblasts in the
E2 and E3 populations (basophilic to orthochromatic stages) exhibited decreased S and increased G2/M phase compared with WT
(Figure 4B-D). Moreover, there was a marked increase in the
percentage of hypertetraploid cells in the RhoAD/D cells, accumulating to ;15% in the E2 population. Of note, even more mature
CD71high;Ter119high erythroblasts, if hypertetraploid, may still
appear to belong in the E2 population because of their large size
and corresponding increase in forward scatter (Figure 4C-D and
supplemental Figure 3). Although no difference was found in
annexin V–only positive cells between RhoA D/D and WT erythroblasts, an increased rate of late apoptosis (annexin V1;7AAD1)
and necrosis (annexin V–;7AAD1) was observed in RhoA-deficient
Ter119high cells (Figure 4E).
RhoA organizes myosin and microtubules during
erythroblast cytokinesis
To further investigate the binucleated phenotype observed in
RhoAD/D fetal liver erythroblasts, we examined the effect of RhoA
on the phosphorylation of the myosin regulatory light chain (MRLC),
which is necessary for effective assembly of the actomyosin contractile
ring at the cleavage furrow during cytokinesis.30 RhoAD/D fetal liver
cells had decreased phosphorylation of MRLC (Figure 5A) in agreement with the action of RhoA in smooth muscle cell contraction,31 as
well as in cytokinesis of cultured cells in vitro32 and of hematopoietic
progenitors in vivo.33
RhoA has also been shown to regulate microtubules during cytokinesis, taking part in the formation of the midbody, a transient structure
composed of microtubules connecting the 2 daughter cells just before
abscission.34 In WT fetal liver cultures, inhibition of microtubules with
taxol resulted in a binucleated phenotype similar to that observed in
the RhoAD/D fetal liver cells (supplemental Figure 4). Therefore, we
explored the localization of RhoA in dividing WT splenic erythroblasts
by imaging flow cytometry. RhoA stained strongly at the site where
b-tubulin marked the midbody formation before abscission (Figure 5B).
Citron kinase (Cit-K) has been shown before as the major downstream
RhoA effector that localizes to the midbody of Drosophila and HeLa
cells during cytokinesis, mediating the regulation of microtubule
organization by RhoA during these final steps of cytokinesis.35-37
Cit-K protein levels were significantly decreased in the RhoAΔ/Δ
erythroid (Ter1191) fetal liver cells (Figure 5C), indicating a likely
association of this RhoA effector with the abscission failure in
RhoA-deficient erythroblasts.
Cytokinesis failure caused by RhoA deficiency results in DNA
damage and p53 phosphorylation
Because cytokinesis failure is known to be associated with DNA
damage,38 we next investigated the RhoAD/D fetal liver cells for
evidence of DNA damage. DNA damage was significantly increased
in RhoAD/D fetal liver erythroid precursors (Ter1191DAPI1 cells)
compared with WT (Figure 6A-B), as demonstrated by staining for
phosphorylated histone H2AX (gH2AX) in imaging flow cytometry.
Moreover, we explored the multinucleated erythroblasts for gH2AX
positivity. Multinucleated erythroblasts were enriched within the
Ter1191 DAPI1 population with large nuclear size (“increased
area_DAPI”) and elongated nuclear shape (“aspect ratio_DAPI,”
calculated by ratio of the minor axis/major axis of nuclear stain; a low
value indicates an elongated area of nuclear stain, whereas a high
value of approximately 1 indicates a round nucleus). Truly binucleated
or multinucleated cells were identified by direct visual observation
within this population (blue dots in Figure 6A). Bi- and multinucleated
RhoA-deficient Ter1191 cells had significantly increased staining
for gH2AX (Figure 6C). Although the binucleated WT cells appeared
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
1480
KONSTANTINIDIS et al
to be normal mitotic cells and were negative for gH2AX, the binucleated
RhoAD/D cells were strongly positive, indicative of DNA damage
(Figure 6D-E).
RNA-Seq analysis of magnetically sorted CD711 fetal liver cells
confirmed a DNA-damaging stress response in the RhoAD/D fetal liver
erythroblasts with upregulation of p53-related DNA damage-response
genes39 (Figure 7A). Follow-up real-time polymerase chain reaction
(RT-PCR) verified the increased expression of PHLDA3 and CDKN1A
that encodes p21, a cyclin-dependent kinase inhibitor (Figure 7B). Both
PHLDA3 and p21 are downstream transcriptional targets of p53; the
first contains a PH domain that competes with the PH domain of AKT
and promotes apoptosis40 and the second mediates cell-cycle arrest.41
These data pointed to a p53-mediated pathway leading to cell-cycle
arrest and/or programmed cell death in RhoAD/D fetal liver erythroblasts. Stabilization of p53 by increased phosphorylation at Ser-15
was indeed found by western blotting and immunofluorescence
(Figure 7C-D) in RhoAD/D fetal liver erythroblasts compared with
WT. Increased early and late apoptosis was also found in the primitive
circulating RhoAD/D erythroids (supplemental Figure 5). The fact that
early apoptosis (annexin V–only positivity) was easily observed in
primitive but not definitive erythroid cells may indicate faster
progression to cell death for the definitive erythroblasts because of
their proximity to macrophages in the fetal liver.
Discussion
In this study, we identify a critical role for RhoA during cytokinesis
of mouse embryonic erythroid cells both in primitive and in definitive
erythropoiesis. We show that erythroid cells deficient in RhoA often
exhibit a multinuclear phenotype indicating failure of cytokinesis.
Cytokinesis failure in erythroblasts was also reported recently in
a mouse model with germline deletion of mDia2,42 a formin, which
catalyzes the nucleation and polymerization of unbranched actin
filaments downstream of Rho GTPases.43 mDia2-deficient mice die
in utero by E12.5, showing anemia at E11.5 with about 15% multinucleated cells in peripheral blood and a decreased number of BFU-E
and CFU-E colonies in colony assays from fetal liver cells,42 therefore
exhibiting a more severe phenotype than the phenotype of our mice
with EpoR-Cre–driven RhoA deficiency. This may be caused by
germline deletion of mDia2 likely contributing to cytokinesis failure
very early in hematopoietic and other fast-growing embryonal tissues.
Moreover, a deletion of mDia2 may display effects downstream of
multiple Rho GTPases (RhoA and B, Rac1 and 2, Cdc42, and Rif) with
which this formin has been shown to have direct and/or indirect
interactions.43-48 Cytokinesis failure in mDia22/2 cells was attributed
to either impaired F-actin accumulation at the cleavage furrow or to
F-actin localization at aberrant sites.42 We found decreased phosphorylation of MRLC in RhoAD/D fetal liver erythroblasts compared with
WT in agreement with previous in vitro work with cell-free systems and
pharmacologic and genetic studies in Drosophila and C. elegans cells,
which have shown RhoA-downstream kinases (ROCK1 or ROCK2,
MLCK, and Cit-K) to phosphorylate MRLC on Ser19.9,49 In addition,
we demonstrated in WT-dividing erythroblasts that RhoA marked the
presumptive abscission site along with the microtubule bundle that
forms the midbody, the transient structure that connects 2 daughter cells
at the end of cytokinesis, as it has been observed before in HeLa
cells.12,34 These 2 molecular mechanisms used by RhoA in cytokinesis
may contribute to the impaired contractility or aberrant localization of
the actomyosin contractile ring observed in mDia22/2 erythroblasts.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
Germline deletion of ROCK1 was found to have no effect on
baseline erythropoiesis and provided a survival advantage with
increased response to stress erythropoiesis, associated with downregulation of p53 phosphorylation, decreased reactive oxygen species
production, and reduced caspase-3 activation in erythroid cells.50 In
contrast, in our model, RhoA-deficient fetal liver erythroblasts exhibited
higher levels of cell death compared with WT, with an increased rate of
late apoptosis (annexin V1;7AAD1) and necrosis (annexin V–;7AAD1).
This difference may be caused by Cit-K and not ROCK mediating the
effect of RhoA to the midbody during the final steps of cytokinesis.
Failure of cytokinesis and development of binuclear and multinuclear
cells has been previously shown in Drosophila and HeLa cells upon
Cit-K ablation or mutation,35-37 similar to what we observe with RhoA
deficiency. Indeed, we found that Cit-K protein levels were significantly
decreased in RhoAΔ/Δ fetal liver erythroblasts.
The RhoA-deficient fetal liver cells had increased p53 phosphorylation at S15, and thus increased p53 activity within the cell,51 and
increased p21 expression, which has been shown as a transcriptional
target of p53 to regulate cell-cycle arrest, apoptosis, or DNA repair.52 It is
unclear how cytokinesis failure leads to p53 activation. p53 has been
named a “guardian of ploidy” and has been shown, as a tumor suppressor,
to either induce cell-cycle arrest or apoptosis of tetraploid cells, which,
without p53, proceed to develop DNA damage including chromosomal
instability or aneuploidy.25,53,54 An initial hypothesis of a “tetraploidy
checkpoint” causing p53 stabilization in response to tetraploidy and
centrosome amplification with subsequent arrest of binucleated cells
was found not to be universally true when it was demonstrated that
mammalian binucleated cells after inhibition of cytokinesis by lowdose cytochalasin were able to start again DNA synthesis (ie, re-enter
mitosis).55 It has been suggested that the connection of cytokinesis
failure to cell-cycle arrest noted in higher doses of cytochalasin may be
caused by a more persistent disorganization of the actin cytoskeleton,
even after washing out the drug.53,55 Genetic RhoA deficiency would
cause a similar disorganization of the cellular actin cytoskeleton. Recently, the Hippo pathway was proposed to mediate cell-cycle arrest after cytokinesis failure by activation of LATS2 kinase, which
stabilizes p53 and inhibits the transcriptional regulators YAP and
TAZ.56 Further investigation will be required to evaluate whether this
pathway is active and exercises quality control during erythropoiesis.
It is of interest that despite defective cytokinesis and pronounced
multinuclearity, primitive erythropoiesis is able to sustain the embryo
through the time definitive erythropoiesis begins to take over at E11.5
to E12.5.57 In contrast, the cytokinesis defect during definitive erythropoiesis proves to be lethal with severe progressive anemia leading to
death by E16.5. This difference might be caused by the increasing
erythropoietic demands of the mouse embryo in the period E12.5 to
E16.5 of gestation,6 whereas an additional factor contributing to the
earlier demise and engulfment/consumption of multinucleated definitive erythroblasts might be the presence of the macrophages in
close proximity to the abnormal cells in the fetal liver posing an
external quality control mechanism.
Based on data from in vivo and in vitro models of DiamondBlackfan anemia and 5q- syndrome, it has been suggested that the
p53 pathway plays an important role in regulating erythropoiesis and
preventing malignant transformations in the rapidly proliferating
erythroid lineage.58 Loss or silencing of specific genes coding for
ribosomal proteins triggers increased activation of p53, which in turn
results in cell-cycle arrest via p21 and in apoptosis.59,60 Cell-cycle
arrest mediated by the p53-p21 pathway was also observed in the
RhoA-deficient erythroblasts, offering another example of p53 applying quality control in erythropoiesis. Binucleated and multinucleated erythroid cells are often observed in certain types of congenital
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
dyserythropoietic anemias (CDAs), a group of human genetic disorders
with heterogeneous and, for many of them, not yet fully clarified
causality.61 CDA type III, characterized by large, multinucleated erythroblasts in the patients’ bone marrow, is caused by mutations in the
KIF23 gene, which codes for a mitotic protein essential for cytokinesis.62 It is intriguing to speculate whether any of the as yet nonelucidated CDAs may be related to RhoA-associated pathways.
In conclusion, by using an erythroid-specific model of RhoAdeficiency, we identify an essential role of the RhoA GTPase in
erythroblast cytokinesis both in primitive and definitive erythropoiesis.
Understanding the unique and overlapping roles of RhoA and associated
signaling molecules in primitive vs definitive erythropoiesis will provide
valuable insights into erythroid lineage development and may reveal
potential targets for improving RBC production in vivo in patients with
hypoproductive anemias and in vitro as a transfusion resource.
Acknowledgments
The authors thank the Research Flow Cytometry Core and the Sequencing Core at Cincinnati Children’s Hospital for excellent technical support.
CYTOKINESIS FAILURE IN RhoA-DEFICIENT ERYTHROIDS
1481
This work was supported by the National Institutes of Health
grants R01HL116352 (National Heart, Lung, and Blood Institute; to
T.A.K.) and P30 DK090971 (National Institute of Diabetes and
Digestive and Kidney Diseases; to Y.Z.).
Authorship
Contribution: D.G.K., K.M.G., and T.A.K. designed and performed
research, analyzed data, and wrote the manuscript; M.R., S.P., P.Z.,
P.D., and S.Y. performed research and analyzed data; and U.K.,
P.A., J.P., and Y.Z. contributed valuable reagents and instrumental
suggestions on research design, data analysis, and writing of the
manuscript.
Conflict-of-interest disclosure: The authors declare no competing
financial interests.
Correspondence: Theodosia A. Kalfa, Division of Hematology/
Oncology, Cincinnati Children’s Hospital Medical Center, 3333 Burnet
Ave, MLC 7015, Cincinnati, OH 45229-3039; e-mail: theodosia.kalfa@
cchmc.org.
References
1. Kingsley PD, Malik J, Fantauzzo KA, Palis J. Yolk
sac-derived primitive erythroblasts enucleate
during mammalian embryogenesis. Blood. 2004;
104(1):19-25.
2. Kingsley PD, Malik J, Emerson RL, et al.
“Maturational” globin switching in primary primitive
erythroid cells. Blood. 2006;107(4):1665-1672.
3. Fraser ST, Isern J, Baron MH. Maturation and
enucleation of primitive erythroblasts during
mouse embryogenesis is accompanied by
changes in cell-surface antigen expression.
Blood. 2007;109(1):343-352.
4. Baron MH, Isern J, Fraser ST. The embryonic
origins of erythropoiesis in mammals. Blood.
2012;119(21):4828-4837.
5. McGrath K, Palis J. Ontogeny of erythropoiesis in
the mammalian embryo. Curr Top Dev Biol. 2008;
82:1-22.
6. Russel ES, Thompson MW, McFarland E.
Analysis of effects of W and f genic substitutions
on fetal mouse hematology. Genetics. 1968;58(2):
259-270.
7. Wheeler AP, Ridley AJ. Why three Rho proteins?
RhoA, RhoB, RhoC, and cell motility. Exp Cell
Res. 2004;301(1):43-49.
8. Ridley AJ. Rho GTPases and actin dynamics in
membrane protrusions and vesicle trafficking.
Trends Cell Biol. 2006;16(10):522-529.
9. Piekny A, Werner M, Glotzer M. Cytokinesis:
welcome to the Rho zone. Trends Cell Biol. 2005;
15(12):651-658.
10. Bement WM, Benink HA, von Dassow G. A
microtubule-dependent zone of active RhoA
during cleavage plane specification. J Cell Biol.
2005;170(1):91-101.
11. Barr FA, Gruneberg U. Cytokinesis: placing and
making the final cut. Cell. 2007;131(5):847-860.
gene targeting. J Biol Chem. 2013;288(51):
36179-36188.
15. Heinrich AC, Pelanda R, Klingmüller U. A mouse
model for visualization and conditional mutations
in the erythroid lineage. Blood. 2004;104(3):
659-666.
16. Melendez J, Stengel K, Zhou X, et al. RhoA
GTPase is dispensable for actomyosin regulation
but is essential for mitosis in primary mouse
embryonic fibroblasts. J Biol Chem. 2011;286(17):
15132-15137.
17. Konstantinidis DG, Pushkaran S, Giger K,
Manganaris S, Zheng Y, Kalfa TA. Identification of
a murine erythroblast subpopulation enriched in
enucleating events by multi-spectral imaging flow
cytometry. J Vis Exp. 2014;(88).
18. McGrath KE, Bushnell TP, Palis J. Multispectral
imaging of hematopoietic cells: where flow meets
morphology. J Immunol Methods. 2008;336(2):
91-97.
19. Wu H, Liu X, Jaenisch R, Lodish HF. Generation
of committed erythroid BFU-E and CFU-E
progenitors does not require erythropoietin or the
erythropoietin receptor. Cell. 1995;83(1):59-67.
20. McGann JK, Silver L, Liesveld J, Palis J.
Erythropoietin-receptor expression and function
during the initiation of murine yolk sac
erythropoiesis. Exp Hematol. 1997;25(11):
1149-1157.
21. Malik J, Kim AR, Tyre KA, Cherukuri AR, Palis J.
Erythropoietin critically regulates the terminal
maturation of murine and human primitive
erythroblasts. Haematologica. 2013;98(11):
1778-1787.
22. Nagy A. Cre recombinase: the universal reagent
for genome tailoring. Genesis. 2000;26(2):99-109.
synchronized with the cell cycle clock through
mutual inhibition between PU.1 and S-phase
progression. PLoS Biol. 2010;8(9):e1000484.
26. Andreassen PR, Lohez OD, Lacroix FB, Margolis
RL. Tetraploid state induces p53-dependent
arrest of nontransformed mammalian cells in G1.
Mol Biol Cell. 2001;12(5):1315-1328.
27. Borel F, Lohez OD, Lacroix FB, Margolis RL.
Multiple centrosomes arise from tetraploidy
checkpoint failure and mitotic centrosome clusters
in p53 and RB pocket protein-compromised cells.
Proc Natl Acad Sci USA. 2002;99(15):9819-9824.
28. Zhang J, Socolovsky M, Gross AW, Lodish HF.
Role of Ras signaling in erythroid differentiation of
mouse fetal liver cells: functional analysis by
a flow cytometry-based novel culture system.
Blood. 2003;102(12):3938-3946.
29. Kalfa TA, Pushkaran S, Zhang X, et al. Rac1
and Rac2 GTPases are necessary for early
erythropoietic expansion in the bone marrow but
not in the spleen. Haematologica. 2010;95(1):
27-35.
30. Iwasaki T, Murata-Hori M, Ishitobi S, Hosoya H.
Diphosphorylated MRLC is required for
organization of stress fibers in interphase cells
and the contractile ring in dividing cells. Cell Struct
Funct. 2001;26(6):677-683.
31. Kimura K, Ito M, Amano M, et al. Regulation of
myosin phosphatase by Rho and Rho-associated
kinase (Rho-kinase). Science. 1996;273(5272):
245-248.
32. Yamashiro S, Totsukawa G, Yamakita Y, et al.
Citron kinase, a Rho-dependent kinase, induces
di-phosphorylation of regulatory light chain of
myosin II. Mol Biol Cell. 2003;14(5):1745-1756.
23. Madisen L, Zwingman TA, Sunkin SM, et al.
A robust and high-throughput Cre reporting and
characterization system for the whole mouse
brain. Nat Neurosci. 2010;13(1):133-140.
33. Zhou X, Florian MC, Arumugam P, et al. RhoA
GTPase controls cytokinesis and programmed
necrosis of hematopoietic progenitors. J Exp Med.
2013;210(11):2371-2385.
13. Heasman SJ, Ridley AJ. Mammalian Rho
GTPases: new insights into their functions from in
vivo studies. Nat Rev Mol Cell Biol. 2008;9(9):
690-701.
24. Koulnis M, Pop R, Porpiglia E, Shearstone JR,
Hidalgo D, Socolovsky M. Identification and
analysis of mouse erythroid progenitors using the
CD71/TER119 flow-cytometric assay. J Vis Exp.
2011;(54).
34. Hu CK, Coughlin M, Mitchison TJ. Midbody
assembly and its regulation during cytokinesis.
Mol Biol Cell. 2012;23(6):1024-1034.
14. Zhou X, Zheng Y. Cell type-specific signaling
function of RhoA GTPase: lessons from mouse
25. Pop R, Shearstone JR, Shen Q, et al.
A key commitment step in erythropoiesis is
12. Gai M, Camera P, Dema A, et al. Citron kinase
controls abscission through RhoA and anillin. Mol
Biol Cell. 2011;22(20):3768-3778.
35. Naim V, Imarisio S, Di Cunto F, Gatti M,
Bonaccorsi S. Drosophila citron kinase is required
for the final steps of cytokinesis. Mol Biol Cell.
2004;15(11):5053-5063.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
1482
KONSTANTINIDIS et al
36. Paramasivam M, Chang YJ, LoTurco JJ. ASPM
and citron kinase co-localize to the midbody ring
during cytokinesis. Cell Cycle. 2007;6(13):
1605-1612.
37. Madaule P, Eda M, Watanabe N, et al. Role of
citron kinase as a target of the small GTPase Rho
in cytokinesis. Nature. 1998;394(6692):491-494.
BLOOD, 17 SEPTEMBER 2015 x VOLUME 126, NUMBER 12
and function in cytokinesis. Mol Biol Cell. 2010;
21(18):3193-3204.
46. Staus DP, Taylor JM, Mack CP. Enhancement
of mDia2 activity by Rho-kinase-dependent
phosphorylation of the diaphanous autoregulatory
domain. Biochem J. 2011;439(1):57-65.
38. Hayashi MT, Karlseder J. DNA damage
associated with mitosis and cytokinesis failure.
Oncogene. 2013;32(39):4593-4601.
47. Ji P, Jayapal SR, Lodish HF. Enucleation of
cultured mouse fetal erythroblasts requires Rac
GTPases and mDia2. Nat Cell Biol. 2008;10(3):
314-321.
39. Amundson SA, Do KT, Vinikoor L, et al. Stressspecific signatures: expression profiling of p53
wild-type and -null human cells. Oncogene. 2005;
24(28):4572-4579.
48. Pellegrin S, Mellor H. The Rho family GTPase Rif
induces filopodia through mDia2. Curr Biol. 2005;
15(2):129-133.
40. Kawase T, Ohki R, Shibata T, et al. PH domainonly protein PHLDA3 is a p53-regulated repressor
of Akt. Cell. 2009;136(3):535-550.
49. Amano M, Ito M, Kimura K, et al. Phosphorylation
and activation of myosin by Rho-associated
kinase (Rho-kinase). J Biol Chem. 1996;271(34):
20246-20249.
41. Waldman T, Kinzler KW, Vogelstein B. p21 is
necessary for the p53-mediated G1 arrest in
human cancer cells. Cancer Res. 1995;55(22):
5187-5190.
42. Watanabe S, De Zan T, Ishizaki T, et al. Loss of
a Rho-regulated actin nucleator, mDia2, impairs
cytokinesis during mouse fetal erythropoiesis. Cell
Reports. 2013;5(4):926-932.
43. Lammers M, Meyer S, Kühlmann D, Wittinghofer
A. Specificity of interactions between mDia
isoforms and Rho proteins. J Biol Chem. 2008;
283(50):35236-35246.
50. Vemula S, Shi J, Mali RS, et al. ROCK1 functions
as a critical regulator of stress erythropoiesis and
survival by regulating p53. Blood. 2012;120(14):
2868-2878.
51. Shieh SY, Ikeda M, Taya Y, Prives C. DNA
damage-induced phosphorylation of p53
alleviates inhibition by MDM2. Cell. 1997;91(3):
325-334.
52. Vousden KH, Prives C. Blinded by the Light: The
Growing Complexity of p53. Cell. 2009;137(3):
413-431.
44. Kühn S, Geyer M. Formins as effector proteins of
Rho GTPases. Small GTPases. 2014;5:e29513.
53. Stukenberg PT. Triggering p53 after cytokinesis
failure. J Cell Biol. 2004;165(5):607-608.
45. Watanabe S, Okawa K, Miki T, et al. Rho and
anillin-dependent control of mDia2 localization
54. Aylon Y, Oren M. p53: guardian of ploidy. Mol
Oncol. 2011;5(4):315-323.
55. Uetake Y, Sluder G. Cell cycle progression after
cleavage failure: mammalian somatic cells do not
possess a “tetraploidy checkpoint”. J Cell Biol.
2004;165(5):609-615.
56. Ganem NJ, Cornils H, Chiu SY, et al.
Cytokinesis failure triggers hippo tumor
suppressor pathway activation. Cell. 2014;
158(4):833-848.
57. Palis J. Primitive and definitive erythropoiesis in
mammals. Front Physiol. 2014;5:3.
58. Raiser DM, Narla A, Ebert BL. The emerging
importance of ribosomal dysfunction in the
pathogenesis of hematologic disorders. Leuk
Lymphoma. 2014;55(3):491-500.
59. Danilova N, Sakamoto KM, Lin S. Ribosomal
protein S19 deficiency in zebrafish leads to
developmental abnormalities and defective
erythropoiesis through activation of p53
protein family. Blood. 2008;112(13):
5228-5237.
60. Dutt S, Narla A, Lin K, et al. Haploinsufficiency
for ribosomal protein genes causes selective
activation of p53 in human erythroid progenitor
cells. Blood. 2011;117(9):2567-2576.
61. Iolascon A, Heimpel H, Wahlin A, Tamary H.
Congenital dyserythropoietic anemias: molecular
insights and diagnostic approach. Blood. 2013;
122(13):2162-2166.
62. Liljeholm M, Irvine AF, Vikberg AL, et al. Congenital
dyserythropoietic anemia type III (CDA III) is
caused by a mutation in kinesin family member,
KIF23. Blood. 2013;121(23):4791-4799.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
2015 126: 1473-1482
doi:10.1182/blood-2014-12-616169 originally published
online July 30, 2015
Cytokinesis failure in RhoA-deficient mouse erythroblasts involves
actomyosin and midbody dysregulation and triggers p53 activation
Diamantis G. Konstantinidis, Katie M. Giger, Mary Risinger, Suvarnamala Pushkaran, Ping Zhou,
Phillip Dexheimer, Satwica Yerneni, Paul Andreassen, Ursula Klingmüller, James Palis, Yi Zheng and
Theodosia A. Kalfa
Updated information and services can be found at:
http://www.bloodjournal.org/content/126/12/1473.full.html
Articles on similar topics can be found in the following Blood collections
Red Cells, Iron, and Erythropoiesis (796 articles)
Information about reproducing this article in parts or in its entirety may be found online at:
http://www.bloodjournal.org/site/misc/rights.xhtml#repub_requests
Information about ordering reprints may be found online at:
http://www.bloodjournal.org/site/misc/rights.xhtml#reprints
Information about subscriptions and ASH membership may be found online at:
http://www.bloodjournal.org/site/subscriptions/index.xhtml
Blood (print ISSN 0006-4971, online ISSN 1528-0020), is published weekly by the American Society
of Hematology, 2021 L St, NW, Suite 900, Washington DC 20036.
Copyright 2011 by The American Society of Hematology; all rights reserved.