Infantile Myelofibrosis and Myeloproliferation with CDC42
Dysfunction
verfasst von:
Jeffrey M. Verboon, Dilnar Mahmut, Ah Ram Kim, Mitsutoshi Nakamura, Nour J. Abdulhay, Satish K. Nandakumar, Namrata Gupta, Thomas E. Akie, Amy E. Geddis, Becky Manes, Meghan E. Kapp, Inga Hofmann, Stacey B. Gabriel, Daryl E. Klein, David A. Williams, Haydar A. Frangoul, Susan M. Parkhurst, Genevieve M. Crane, Alan B. Cantor, Vijay G. Sankaran
Studies of genetic blood disorders have advanced our understanding of
the intrinsic regulation of hematopoiesis. However, such genetic studies have only
yielded limited insights into how interactions between hematopoietic cells and their
microenvironment are regulated. Here, we describe two affected siblings with
infantile myelofibrosis and myeloproliferation that share a common de novo mutation
in the Rho GTPase CDC42 (Chr1:22417990:C>T, p.R186C) due to paternal germline
mosaicism. Functional studies using human cells and flies demonstrate that this
CDC42 mutant has altered activity and thereby disrupts interactions between
hematopoietic progenitors and key tissue microenvironmental factors. These findings
suggest that further investigation of this and other related disorders may provide
insights into how hematopoietic cell-microenvironment interactions play a role in
human health and can be disrupted in disease. In addition, we suggest that
deregulation of CDC42 may underlie more common blood disorders, such as primary
myelofibrosis.
Hinweise
Jeffrey M. Verboon and Dilnar Mahmut contributed equally to this
work.
Springer Nature remains neutral with regard to jurisdictional claims in
published maps and institutional affiliations.
Introduction
Recent studies have provided considerable insights into how interactions
between hematopoietic stem/progenitor cells and their microenvironment are regulated
[5, 10, 34, 44]. Despite the substantial progress in this
field, it remains unclear whether perturbation of such interactions can lead to
human disease. Rare experiments of nature provide a potentially effective way to
gain insights into how these interactions function normally in health and can be
perturbed in disease states. For instance, a recent study demonstrated that
chromothriptic disruption of a gain-of-function mutation in CXCR4 in hematopoietic stem cells (HSCs) was curative in a patient
with warts, hypogammaglobulinemia, immunodeficiency, and myelokathexis (WHIM)
syndrome as a result of preferential expansion of the CXCR4-haploinsufficient HSCs
in the bone marrow niche [28]. How
other genetic blood disorders may alter hematopoietic cell-microenvironment
interactions remains largely unknown.
In the context of acquired blood disorders, there is more evidence for
altered interactions between hematopoietic cells and the bone marrow
microenvironment. Mouse models have demonstrated that perturbations of the HSC niche
can impact the function and differentiation of hematopoietic cells, as exemplified
by mutations in the bone marrow microenvironment causing myeloproliferative
disorders [34, 47, 48]. Moreover, studies of human primary myelofibrosis (PMF) have
provided evidence for microenvironment dysfunction contributing to disease
pathogenesis [23], and recent studies
have elucidated pathways that may contribute to this process in mouse models
[3, 12, 38, 40].
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The Rho family of small GTPases composed of the RHO, RAC, and CDC42
proteins, which are best known as signaling molecules that control actin dynamics,
have emerged as potent factors in hematopoiesis [31]. Indeed, this family of proteins is known to regulate
hematopoietic cell shape, polarity, adhesion, and migration. Germline mutations of
the hematopoietic-specific Rho GTPases RAC2 andRHOH have previously been implicated in
inherited immunological conditions, impacting myeloid and lymphoid cell functions
[11, 50]. Recently, human mutations in CDC42 have presented with diverse developmental phenotypes, and a
specific missense mutation, R186C, has been reported in a few patients with a
syndrome involving immune dysregulation [14, 22, 27].
Here, we have identified two children born to unaffected parents
presenting with infantile myelofibrosis and myeloproliferation. Both affected
patients harbor the identical CDC42 R186C mutation due to low-level paternal
mosaicism. We demonstrate dysfunctional activity of this mutant form of CDC42.
Introduction of this mutation into primary human hematopoietic stem/progenitor cells
disrupts the ability of the cells to migrate towards the key hematopoietic
chemokine, CXCL12/SDF-1 [10]. We
additionally demonstrate that this mutant acts in a dominant manner to disrupt the
ability of Drosophila hematopoietic cells to
migrate appropriately. Our studies extend the phenotypic spectrum of disorders
resulting from mutations of Rho GTPases and specifically implicate the CDC42 R186C
mutation in altering interactions of hematopoietic cells with the microenvironment.
Moreover, studies of acquired primary myelofibrosis suggest that disruption of CDC42
may occur in hematopoietic progenitor cells in this disease, connecting the
observations made in these rare cases to the pathogenesis of other, more common,
acquired malignant hematopoietic disorders.
Methods
Study Approval
All family members provided written informed consent to participate
in this study. The institutional review boards of Boston Children’s
Hospital and Massachusetts Institute of Technology approved the study
protocols.
Whole-Exome Sequencing and Related Genetic Analyses
The patients described in this paper are part of a rare blood
disorder cohort that has been studied through the use of whole-exome sequencing
(WES), as previously described [1,
19, 20, 45]. WES in these cases was performed using genomic DNA
obtained from peripheral blood samples of the patients. The resultant variant
call file (in hg19 coordinates) was annotated with VEP v89 [30], and rare variants (based on ExAC v0.3.1
and GnomAD r2.0.2) [24] (http://gnomad.broadinstitute.org/) were identified using a combination of the Genome Analysis
Toolkit, Bcftools, and Gemini [25,
29, 33]. No rare (< 0.01% allele
frequency in ExAC and GnomAD) loss-of-function or missense variants were
identified in any known blood cell disorder genes. All mutations were confirmed
from genomic DNA samples of the patients or family members by Sanger
sequencing.
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Targeted Amplicon Sequencing
A 205-bp region containing the mutation was amplified from all
family members and unrelated controls. The PCR amplicons were processed, and
paired-end sequencing was performed using a MiSeq instrument (Illumina).
Lentiviral Constructs
CDC42 wild type (WT) and R186C mutant cDNA were cloned into the HMD
lentiviral vector with EcoRI and XhoI digestion and ligation. FLAG-tagged CDC42
WT and FLAG-tagged CDC42 R186C cDNA were also cloned into HMD lentivirus vector
with EcoRI and XhoI digestion and ligation. Transduced cells were identified
based on GFP expression driven by the IRES-GFP from the HMD vector.
Cell Culture and Lentiviral Transduction
HEK293T and NIH-3T3 cell lines were obtained and maintained in
Dulbelcco’s Modified Eagle Medium (DMEM; Gibco) supplemented with 10% of
fetal bovine serum (FBS; Atlanta Biologicals) and 1% of penicillin/streptomycin
(GIBCO). Human primary adult bone marrow-derived
CD34+ hematopoietic stem and progenitor cells
(HSPCs) were obtained from Fred Hutchinson Cancer Research Center and maintained
in StemSpan II and 100X CC100 (Stem Cell Technologies) with recombinant
thrombopoietin at 50 ng/mL (TPO; Peprotech).
CDC42 WT and R186C mutant lentiviral constructs were transfected
into HEK293T with transfection reagent FuGene (Promega) and helper plasmids, as
previously described [19,
26]. The primary HSPCs were
infected with virus during day 1 of culture. Primary HSPCs were then sorted for
GFP on day 3 and processed for assays. Multiple donors were used for downstream
assays and produced similar results. NIH-3T3 cells were infected with lentivirus
and 48 h later, used for further downstream applications.
Fly Strains and Genetics
Flies were cultured and crossed on yeast-cornmeal-molasses-malt
extract media and maintained at 25 °C. The following
hemocyte-specific drivers were used: P{Pxn-GAL4}II, P{UAS-GFP}II and
P{Pxn-GAL4}III, P{UAS-GFP}III [43].
Generation of Cdc42(R186C) Fusion Constructs and Transgenics
A Cdc42(R186C) mutation was made by introducing the R186C point
mutation into the wild type Drosophila Cdc42
cDNA by PCR. UAS-Cdc42(R186C) was made by cloning the resulting Cdc42(R186C)
cDNA into the KpnI-XbaI sites of the pUASp vector [37]. These constructs were used to make germline
transformants, as previously described [18]. The resulting transgenic lines (P{w+; UAS-Cdc42(R186C)}) were mapped to a single chromosome and shown
to have non-lethal insertions. mCh-Cdc42 flies have been previously described
[2].
Transwell Migration Assay
A transwell migration assay (Costar; Corning) was performed with
the lentiviral-infected and GFP-positive sorted human
CD34+ HSPCs. Briefly,
3X105 CD34+ cells were
suspended in StemSpan II media in the upper chamber of the transwell plate. The
cells were allowed to migrate to the lower chamber in which there was
CXCL12/SDF-1 at a concentration of 100 ng/mL in StemSpan II medium.
Migration was performed over the course of 4 h at 37 °C.
After incubation, number of cells that migrated to the lower chamber were
quantified.
Cell Cycle Assay
Infected and GFP-positive sorted CD34+
cells were also evaluated for cell cycle phase progression differences. These
cells were suspended in StemSpan II media (Stem Cell Technologies) and
stimulated with CXCL12/SDF1-a at 300 ng/mL (Peprotech) for 1 h at
37 °C. During this 1 h incubation, 10 μM of
EdU-Alexa647 (ThermoFisher) was also added to the cell suspension. The cells
were fixed, permeabilized, and stained with Click-iT™ EdU Alexa Flour
647™ Flow Cytometry kit (ThermoFisher) following the
manufacturer’s protocols. Cells were also stained with propidium iodide
(Biolegend) at 200 ng/mL during the final step of the protocol. Cell
cycle distribution was acquired by using a BD LSR II flow cytometer (BD
Biosciences), and the FlowJo software was used to determine the percentage of
cells in different phases of cell cycle.
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Flow Cytometry
Surface staining of CXCR4/CD184 was measured in infected and
GFP-positive sorted CD34+ cells by flow cytometry.
Cells were recovered in CD34+ culture media after
sort and washed with cold PBS twice. Cells were re-suspended in 1% BSA in PBS
and stained with 5-μL CD184-APC from a stock of 200 μg/mL
(306509; BD Biosciences) for 30 mins on ice. Cells were washed once with
PBS after staining and re-suspended in 1% BSA in PBS with propidium iodide
(Biolegend) at 200 ng/mL. CXCR4 surface staining FACS was acquired using
an Accuri C6 flow cytometer (BD Biosciences), and mean fluorescence
intensity of CXCR4 was measured by using the FlowJo software.
Mammalian Cell Microscopy
3T3 cells were transfected with either FLAG-tagged WT CDC42 or
CDC42 R186C and were plated on fibronectin-coated slides and stained with
Rhodamine-Phalloidin following manufacturer protocols (BK005; Cytoskeleton Inc).
Mouse, anti-Flag antibody was used for CDC42 visualization (166,355; Santa
Cruz). Cells were mounted with DAPI-containing medium (0100–20; Southern
Biochem). Confocal micrographs were imaged on a Zeiss LSM 700 using a Zeiss
40x/1.4 Plan-Apochromat objective.
Cell Morphology Analyses
Infected 3T3 cells were washed with PBS, treated with 0.25% trypsin
(GIBCO) for 4 min at 37 C, deactivated with 10% FBS supplemented medium,
and re-plated onto fibronectin
(2 μg/cm2)-coated plates (Sigma
Aldrich) for 45 mins. Morphology differences were noted and captured by
using light microscopy at different magnifications.
Western Blotting
After 48 h of infection, approximately 1 million 3T3 cells
were plated onto fibronectin-coated plates and incubated at 37 °C
for 1 h. Post-incubation, cells were washed with PBS twice and then lysed
with cold radioimmunoprecipitation assay buffer supplemented with protease and
phosphatase inhibitor cocktails (Santa Cruz). Proteins were quantified with DC
protein assay (BioRad), run using the Mini-Protean TGX gel system (BioRad), and
transferred to polyvinylidene fluoride membrane. Signal was detected by ECL
Amersham Hyperfilm (GE Healthcare). Western blotting was performed with primary
antibodies anti-CDC42 at 1:500 dilution (610928; BD Biosciences), anti-PAK1/2/3
at 1:1000 dilution (2604; CST), anti-pPAK at 1:1000 dilution (2601s; CST), and
anti-GAPDH at 1:10,000 dilution (clone 6C5; Santa Cruz). HRP-conjugated goat
anti-mouse and anti-rabbit (BioRad) were used at 1:20,000 dilution as secondary
antibodies.
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Fly Microscopy
For migration studies, fly embryos were collected on grape agar
plates at either 25 °C for 1 h and then aged at
18 °C for 16–20 h or at 25 °C for
1 h and then aged 12–15 h at 25 °C. The two
regimens yielded indistinguishable results. Embryos were then dechorionated,
dried for 5 min, transferred onto strips of glue dried onto No. 1.5
coverslips, and covered with series 700 halocarbon oil (Halocarbon Products,
River Edge, NJ). For migration studies, 30-μm stacks were taken once
every 5 min with a 0.5-μm step size over the course of
1.5 h. Individual hemocytes were imaged from top to bottom with a
0.5–1-μm step size. All imaging was performed at room temperature
(22 °C). For live imaging, the following microscope setup was
used: Revolution WD systems (Andor Technology Ltd., Concord, MA, USA) mounted on
a Leica DMi8 (Leica Microsystems Inc., Buffalo Grove, IL, USA) with a 20x/0.7 NA
objective lens and controlled by using the MetaMorph software. Images and videos
were acquired with 488 nm, using an Andor iXon Ultra 897 EMCCD cameras
(Andor Technology Ltd., Concord, MA, USA). For individual hemocyte images, the
following microscope setup was used: Zeiss LSM 780 spectral confocal microscope
(Carl Zeiss Microscopy GmbH, Jena, Germany) fitted with Zeiss 40x/1.4 oil
Plan-Apochromat objective at 4× zoom. FITC (Alexa 488) fluorescence was
excited with the 488-nm line of an argon laser, and detection was between 498
and 560 nm. Pinhole was typically set to 1.0 Airy units. Confocal
sections were acquired at 0.5–1.0-μm spacing. All images were
analyzed with Fiji [39].
Measurements of percent cell body protrusions were done manually. Measurements
of average hemocyte velocity over 30 min were done using the manual
tracking plugin in Fiji. Five random hemocytes per embryo were measured.
Microarray Analysis
Normalized expression sets for PMF and control samples were
accessed using the R package GEOquery from the GEO dataset GSE53482.
Differential expression was performed as previously described using the
Bioconductor package Limma [36].
Immunohistochemical Analysis of Primary Myelofibrosis
PMF (n = 10
independent samples) or non-diagnostic findings/normocellular (n = 7 independent samples) bone
marrow specimens were identified in the pathology archives at Weill Cornell/New
York Presbyterian Hospital. Bone marrow specimens underwent standard histologic
processing and were stained with an antibody against CDC42 (PA1-092,
ThermoFisher Scientific) following optimization of staining conditions. All
measured samples were stained together to ensure comparable staining. Level of
CDC42 staining in megakaryocytes was performed using the Halo Image analysis
platform (Indica Labs). Between 3 and 14 megakaryocytes were quantified per
individual sample to ensure consistency.
Statistical Analyses
All pairwise comparisons were performed using the 2-tailed
Student’s t test, unless otherwise
indicated. Differences were considered significant if the p value was less than 0.05. Statistical testing for
deep sequencing was done using a one-sided binomial test, Bin(q,n,p), whereq is the number of alternate alleles,n is the depth at the variant position,
and p is the site-specific error rate
determined from the unrelated samples.
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Data Availability
The whole-exome sequencing data are available in the dbGaP database (http://www.ncbi.nlm.nih.gov/gap) under the accession number phs000474.v2.p1.
Results
Case History
A description of the clinical course of the proband has previously
been reported [41]. Briefly, he is
a Caucasian male born at 37 weeks of estimated gestational age following
labor induction due to maternal hypertension. No dysmorphic features were noted.
On day of life 1, he developed a patchy and scaly erythematous rash that
intermittently disappeared and recurred, and was urticarial at times. At
1 week of age during evaluation for his rash, he was found to be
thrombocytopenic with a platelet count of 36 K/μL. He then
developed fever and lethargy and was admitted to the hospital where he underwent
a sepsis rule out, skin biopsy, and bone marrow biopsy. The skin biopsy revealed
perivascular and interstitial inflammation with eosinophils. The bone marrow
biopsy showed a hypercellular marrow with predominantly maturing myeloid
elements, a high myeloid:erythroid ratio, and presence of megakaryocytes. Blasts
were < 2% and the karyotype was normal 46 (X,Y). No hemophagocytosis
was noted. He was discharged home, but about 1 week later, he was
re-admitted after developing hematochezia associated with a platelet count of
10 K/μL. Hepatomegaly was noted for the first time, which
progressively worsened. He also developed persistent tachypnea and intermittent
hypoxia requiring supplemental oxygen. A chest X-ray was unremarkable. It was
thought that his tachypnea was due to mechanical causes related to his marked
hepatomegaly. Liver biopsy revealed extensive extramedullary hematopoiesis. He
subsequently developed splenomegaly. He continued to have a recurrent
polymorphic rash as well as intermittent angioedema and nodules on his hands. A
ferritin level at 18 days of life was 594 ng/mL, and fibrinogen
was 211 mg/dL. Repeat skin biopsy showed inflammatory infiltrates with
lymphocytes, histiocytes, and rare neutrophils and eosinophils. He developed
anemia and neutropenia in addition to his thrombocytopenia and became both red
blood cell and platelet transfusion dependent. He also developed mild
eosinophilia with absolute eosinophil counts ranging from 0.52 to
3.2 K/μL. Immunoglobulin levels were all elevated (IgM
202 mg/dL, IgG 1830 mg/dL, IgE 353 IU/mL, and IgA
81 mg/dL) except for IgD. Peripheral lymphocyte subset studies were
normal. Repeated bone marrow biopsies revealed a hypercellular bone marrow with
megakaryocyte dysplasia and moderate to marked fibrosis that was positive by
reticulin and trichrome stains (Fig. 1). The karyotype continued to be normal, and no excess of
blasts was found. In vitro GM-CSF hypersensitivity testing to assess for
possible juvenile myelomonocytic leukemia was negative. He was diagnosed with
idiopathic infantile myelofibrosis. He was treated with a 4-day course of
cytosine arabinoside (Ara-C) at 20 mg/m2/day
to try to reduce his hepatosplenomegaly, but he did not have a clinical
response. He continued to have baseline hypoxia with massive hepatomegaly
causing restrictive lung disease. Oxygen saturations were 88% breathing room
air, but increased to > 94% with supplemental oxygen. Echocardiography
did not show evidence of pulmonary hypertension, and chest CT did not show
evidence of parenchymal disease. At 2 months of age, he underwent bone
marrow transplantation (BMT) with his 4-year-old female HLA-matched sibling
serving as the donor. He was conditioned for BMT with busulfan
(1.2 mg/kg/dose every 6 h for 16 doses, with busulfan
pharmacokinetics to achieve a range of 600 to 900 nm/mL) followed by
cyclophosphamide, 50 mg/kg for 4 days. There was no radiation as
part of the formal BMT conditioning regimen. However, he did receive
300 cGy to his spleen prior to BMT conditioning to try to reduce the
spleen size because of mechanical interference with his respiration. The
patient’s respiratory compromise improved in the first few days following
BMT, and he no longer required supplemental oxygen. However, beginning on day 21
following BMT, he developed worsening tachypnea and hypoxia. There were no
infiltrates on his chest X-ray. Cardiac catheterization demonstrated severe
pulmonary hypertension with baseline right pulmonary arterial pressure of
70 mmHg compared with a systemic pressure of 65 mmHg. He was
placed on diuretics and calcium channel blockers, but his respiratory disease
worsened and he died on day 32 following BMT. Autopsy revealed severe intimal
hyperplasia of the pulmonary arterioles.
×
The proband’s younger brother (Fig. 2a, II-3) was born at 35 2/7 weeks of
gestation by cesarean section due to antenatally detected hepatomegaly and
reduced fetal movements. He was blue and floppy at birth (SaO2 in the 50% range
with poor chest movement) and was intubated. Apgar scores were 3 at
1 min, 6 at 5 min, and 9 at 10 min. He was also noted to
have a purpuric rash, hepatosplenomegaly, anemia (initial hematocrit 34%),
reticulocytosis (initial reticulocyte count 9.8%), thrombocytopenia (initial
platelet count 11 K/μL), leukoerythroblastosis (Fig. 1), and an elevated C-reactive protein level
(73.5 mg/dL). A bone marrow aspirate performed on day of life 1
demonstrated moderate erythrocytic dysplasia. The karyotype was normal showing
46(X,Y). No hemophagocytosis was observed. Flow cytometric analysis revealed
< 3% myeloblasts. A bone marrow core biopsy was not successful. He was
weaned to continuous positive airway pressure (CPAP) by day of life 4, but
continued to require supplemental oxygen. There was no evidence of pulmonary
hypertension by echocardiography. It was thought that his severe hepatomegaly
was causing restrictive lung disease. He was given supportive care while
awaiting bone marrow transplantation. He remained platelet and red blood cell
transfusion dependent, and continued to have elevated C-reactive protein levels
(38.7–96.7 mg/dL). At about 4 weeks of life, he developed
labored breathing and tachycardia, followed by bradycardia and decreased
respiratory rate. He was re-intubated and diagnosed with presumed septic shock.
Chest X-rays showed total atelectasis of the left lower lobe with bronchiectasis
with the lobe. There was marked hyperexpansion of the left upper lobe and heavy
markings of the right lung. Chest CT was initially concerning for possible
congenital lobar emphysema, but this was subsequently ruled out. Direct
laryngoscopy and bronchoscopy showed bronchomalacia below the right mainstem
bronchus. He subsequently developed multi-organ failure and died at about
2.5 months of age. Autopsy revealed a hypercellular vertebral bone marrow
comprised exclusively of early myeloid cells (Fig. 1g), which are positive for myeloperoxidase (MPO) via
immunohistochemistry (Fig. 1h) and
negative for CD3, CD43, CD34, and CD117. Mild fibrosis could be highlighted by
reticulin stain (Fig. 1i), and patchy
involvement was noted of the spleen, bilateral kidneys (Fig. 1j), and lymph nodes (Fig. 1k). There was diffuse trichome-positive fibrosis
of the liver. The lungs were bilaterally consolidated with left upper lobe
hemorrhage and necrotizing pneumonia (Fig. 1l) present in the left upper, left lower, and right lower
lobes. There was no evidence of pulmonary hypertension. No fibrosis was noted.
Lung cultures showed moderate growth of Pseudomonas
aeruginosa.
×
The proband’s 3 sisters (II-1, II-4, II-5) and both parents
(I-1, I-2) are all healthy. Sanger sequencing of peripheral blood mononuclear
DNA from the proband and his affected younger brother failed to identify
mutations in the coding sequences or intron/exon boundaries of GATA1, JAK2, orMPL.
Genetic Analyses
Given the observed inheritance pattern, we initially searched for
X-linked or autosomal recessive causes of this phenotype using WES data
generated from all family members (Fig. 2a). This analysis was unrevealing for any putative causal
mutations showing appropriate segregation. However, analysis of potential de
novo variants revealed a single recurrent event in both affected children,
Chr1:22417990:C>T (hg19 coordinates), resulting in the R186C mutation inCDC42 (Fig. 2b). Given that such recurrent de novo mutations can be
associated with low-level parental germline mosaicism [35] and that the father had shown a single
read with this variant in WES, we performed deep sequencing to
> 100,000-fold of the mutated region of CDC42 in blood-derived genomic DNA from all family members and
unrelated controls and observed an increased level of mosaicism from 0.07 to
0.15% in controls to 0.59% in the unaffected father (I-1; Fig. 2c). This finding suggests paternal germline
mosaicism as the source for the recurrent mutations noted in the two affected
children. This mutation was located in an evolutionarily conserved C-terminal
polybasic region of CDC42 (Fig. 2d) and
was recently described in patients with a distinct disorder characterized by
immune dysregulation [14,
22].
Functional Studies
CDC42 is a member of the well-studied Rho family of small GTPases
and, through interacting proteins known as effectors, regulates signaling
pathways that control diverse cellular functions including cell cycle
progression, cell migration, and cytoskeletal dynamics [13]. Rho GTPases have been shown to play
critical roles in hematopoietic cells, including hematopoietic stem/progenitor
cells, as well as cells of the myeloid and lymphoid lineages [31]. In addition, Rho GTPases, particularly
RHOA, RHOH, and the GTPase most closely related to CDC42, RAC, have been
implicated in malignant transformation. CDC42 is best studied for its role in
regulating and enabling signaling pathways to intersect with cytoskeletal
activity. Importantly, CDC42 has been shown to have a critical role in HSCs and
other hematopoietic progenitors through the study of mutant mice [52, 53]. Conditional deletion of Cdc42 in the mouse hematopoietic system led to a rapidly fatal
myeloproliferative disorder [53].
Finally, a different variant in CDC42 (Y64C)
in a human patient has been associated with myelofibrosis observed in adulthood
[7]. Therefore, the observed
mutation appeared to be potentially relevant to hematopoiesis, although the
human phenotype described in this kindred is distinct from what has been
reported in animal models and other human patients previously.
We initially examined the location of the mutation in CDC42 in a
structure involving the complex of CDC42 with its critical guanine nucleotide
exchange factor, RhoGDI (encoded by the ARHGDIA gene in humans; Fig. 3a) [17]. The
R186 residue is located near the geranylgeranyl moiety and is predicted to alter
several key interactions with RhoGDI (Fig. 3a), as has recently been demonstrated through functional
analyses [22]. Consistent with
this, introduction of either wild type or the R186C mutant CDC42 into 3T3 cells
using lentiviral transduction led to abnormal cytoskeletal structure and cell
morphology. When transduced cells were plated on fibronectin, we found that the
mutant, but not control or WT, CDC42-transduced cells inhibited the formation of
filipodia, which depend upon CDC42 activity (Fig. 3b) [16]. After
short-term plating of 3T3 cells onto fibronectin for 45 min, we noted
that significantly more control or WT CDC42-transduced cells showed a flattened
morphology compared with the CDC42 mutant-transduced cells that demonstrated a
predominantly rounded morphology (Fig. 3
c, d). These results suggested that the mutant was acting in a dominant-negative
or potentially neomorphic manner. Additionally, the phosphorylation of PAK1, a
known CDC42 effector protein, was reduced in the mutant-transduced 3T3 cells
plated on fibronectin compared with WT CDC42-transduced cells (Fig. 3e).
×
While these studies suggested dominant-negative or neomorphic
effects and deregulated activity of the mutant CDC42, we wanted to gain more
precise insights into the function of this mutant protein in hematopoietic
cells. Since no hematopoietic cells were available from the patients, we
utilized CD34+ HSPCs from healthy individuals. These
cells were transduced using lentiviral vectors expressing either the WT or the
R186C mutant form of CDC42. Two days after infection and immediately after flow
cytometric sorting of the transduced cells, we assessed cell migration in a
CXCL12/SDF-1 gradient using a transwell migration assay (Fig. 3f). While we observed no change in the
expression of CXCR4 (CD184), the receptor for CXCL12/SDF-1 (Fig. 3 g, h), we noted a > 2-fold reduction
in chemokine-directed migration of cells (Fig. 3f). We next assessed cell cycle progression in vitro using
BrdU staining in the HSPCs transduced with either the control, WT, or mutant
CDC42 in the presence of multiple cytokines including thrombopoietin, stem cell
factor, interleukin 3 (IL-3), and the chemokine, CXCL12. Under these conditions,
we noted no difference in cell cycle progression (Fig. 3i), suggesting that the observed phenotypes in the patients
rely upon disruption of key interactions in the context of an intact niche, as
has been observed in conditional knockout mice [53].
As this CDC42 residue is highly conserved (Fig. 2d), and hematopoiesis in Drosophila melanogaster has many shared features with mammalian
hematopoiesis, including critical interactions between hematopoietic cells and
their microenvironment; we further explored this mutation in vivo in Drosophila [4]. Rho GTPases have previously described important functions
in the fly hematopoietic system, where Rho1 and its downstream effector Wash
have key roles in the migration of embryonic hemocytes, while Cdc42 has been
shown to be critical for maintaining appropriate cell polarity during hemocyte
migration towards wounds [43,
46].
In order to explore this specific mutation in Drosophila, we made transgenic flies which express
the CDC42 R186C mutation using the inducible UAS-Gal4 system in Pxn-Gal4/UAS-GFP
transgenic flies to express this mutant CDC42 specifically in Drosophila hemocytes that also expressed GFP
[6, 37]. We explored hemocyte function in wild
type flies containing the Gal4 driver only (control), flies overexpressing wild
type Cdc42 (Cdc42 WT), and flies overexpressing the Cdc42 R186C mutation (Cdc42
mutant) and found that in contrast to control and Cdc42 WT flies, Cdc42 mutant
flies had reduced protrusions in migrating hemocytes (Fig. 4a–c). In order to quantify this defect,
we measured the percentage of the hemocyte cell body with protrusions and found
a profound loss of protrusions in the Cdc42 mutant compared with control and
Cdc42 WT hemocytes (p < 0.0001) (Fig. 4d). Given the defective migration observed with the CDC42
mutant in human hematopoietic cells, we wondered whether Drosophila hemocytes may also fail to migrate appropriately from
head to tail in response to cues supplied by their microenvironment. During fly
embryogenesis, hemocytes are formed from the head mesoderm, while the head and
tail are adjacent. A subset of hemocytes migrate to the tail from the head,
dependent on the Rho GTPase, RhoL, before the tail retracts during morphogenesis
to the posterior [42]. Once tail
retraction is complete and in a chemotactic response to the PVR ligands, Pvf2
and Pvf3, head hemocytes migrate towards the posterior and tail hemocytes
migrate anteriorly along the ventral midline towards each other and subsequently
distribute throughout the entire embryo [9, 51]. This
stereotypic process starts in stage 10 embryos and completes in
~ 2 h in normal embryos. We found that this migration process was
severely disrupted in embryos expressing the Cdc42 mutant, but not in control or
Cdc42 WT embryos (Fig. 4e–g,n = 20 replicates per
condition). Despite this defect, hemocytes in all conditions were able to
receive cues to migrate from head to tail and vice versa as hemocytes migrate
from both the head and tail and can be seen along the ventral midline (Fig.
4g, 90 min). However, their
capacity for migration was greatly diminished as Cdc42 mutant-expressing
hemocytes showed ~ 2-fold reduction of migration speed, as compared with
the other conditions (p < 0.0001, n = 5 hemocytes per embryo, 20 embryos per condition).
Thus, the CDC42 R186C mutation in flies also functions in a dominant-negative or
neomorphic manner, impacts cell protrusions, and impairs the ability of
hematopoietic cells to migrate in response to tissue microenvironmental cues,
leading to an overall defective hematopoietic system.
×
Relationship to Primary Myelofibrosis
Primary myelofibrosis is a condition associated with somatic
mutations in JAK2 or other factors that activate the thrombopoietin signaling
pathway [15, 49]. Given the previously characterized role
of Rho GTPases in hematopoietic cell localization within the marrow
microenvironment [8, 31] and the disruption of this process
observed in PMF [23], we
hypothesized that the severe infantile myelofibrosis phenotype that results from
the CDC42 R186C mutation may provide insights into the more common acquired PMF
cases. CDC42 was broadly expressed in normal human hematopoiesis across the full
hematopoietic hierarchy, including in HSCs and other early hematopoietic
progenitors (Fig. 5a). To examine
whether CDC42 mRNA expression may be altered
in PMF patients, we compared gene expression in peripheral blood-derived
CD34+ HSPCs from 42 PMF cases with 16 healthy
donors [32]. CDC42 was among the most downregulated genes in PMF
compared with normal donors with expression being ~ 5-fold less across
all the samples (Fig. 5b, c). To
validate these observations, we assessed bone marrow sections from 10
individuals with PMF and 7 individuals with normocellular bone marrows. We opted
to examine megakaryocytes, given that they are easily distinguished
morphologically and highly express CDC42 (Fig. 5a). We observed reduced CDC42 protein expression using
immunohistochemistry in the megakaryocytes across the PMF patients, bolstering
and extending our results on CDC42 dysfunction occurring more commonly in PMF
(Fig. 5d, e, f).
×
Discussion
Our findings complement recent studies of patients with a distinct
disorder involving immune deregulation [14, 22] and suggest
that CDC42 dysfunction can result in congenital myelofibrosis and myeloproliferation
presenting during infancy. The data reported here extend the phenotypic spectrum
resulting from CDC42 R186C mutations and suggest that patients presenting with
congenital myeloproliferation or myelofibrosis be screened for this germline
mutation. The differences between the phenotypes reported here and those reported
recently with similar CDC42 mutations are of
interest and warrant further investigation [14, 27]. We would
note that there is clear phenotypic overlap between the patients reported here and
the other patients previously reported to have a syndrome of immune dysregulation
due to the R186C mutation [22]. Some of
the phenotypes reported in the other patients, such as the observed alterations in
hematopoiesis, may be attributable to altered interactions between hematopoietic
progenitors and the microenvironment, as we demonstrate here. In addition, immune
dysregulation may also be contributing to the observed phenotypes and perturbation
of hematopoiesis. Further studies to dissect these varying contributions will be
valuable.
Additional study of these disorders might provide insights into the
role of CDC42 in the intrinsic regulation of hematopoietic cells in their
microenvironment, as we suggest through our functional studies of human and fly
hematopoietic cells. It is notable that both the upstream receptor, CXCR4, and the
downstream effector, WASP, which act in concert with CDC42, have key roles in
enabling HSCs to home to and occupy their appropriate niche in the bone marrow
[21, 28]. Through rare experiments of nature, we demonstrate how
disruption of these key hematopoietic cell-microenvironment interactions may result
in human disease and may also have implications for more common acquired blood
disorders, such as PMF.
Acknowledgments
We are grateful to the family described in this paper for their interest and
willingness to participate in this study. We thank members of the Sankaran
laboratory, as well as J. Orange, S. Morrison, Y. Zheng, S. Orkin, M. Fleming, and
D. Nathan for valuable comments.
Compliance with Ethical Standards
Conflict of Interest
All family members provided written informed consent to participate
in this study. The institutional review boards of Boston Children’s
Hospital and Massachusetts Institute of Technology approved the study
protocols.
The authors declare that they have no competing interests.
Open Access This article is licensed
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provide a link to the Creative Commons licence, and indicate if changes were
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Infantile Myelofibrosis and Myeloproliferation with CDC42 Dysfunction
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