Control of Homeostasis and Dendritic Cell
Survival by the GTPase RhoA
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Shuai Li, Bastian Dislich, Cord H. Brakebusch, Stefan F.
Lichtenthaler and Thomas Brocker
Published September 25, 2015, doi:10.4049/jimmunol.1500676
The Journal of Immunology
Control of Homeostasis and Dendritic Cell Survival by the
GTPase RhoA
Shuai Li,* Bastian Dislich,† Cord H. Brakebusch,‡ Stefan F. Lichtenthaler,†,x,{ and
Thomas Brocker*
C
onventional dendritic cells (cDCs) are professional APCs,
which, depending on the stimulus, initiate either adaptive
immunity or immune tolerance (1). Although cDCs represent only a very low percentage of all immune cells, the presence of appropriate numbers of cDCs with defined life spans in
lymphoid organs is crucial for a balance between immunity and
tolerance. Accordingly, elevated numbers of cDCs and experimentally prolonged dendritic cell (DC) life spans caused exaggerated T and B cell responses, defective tolerance induction,
and autoimmunity (2–5). Conversely, experimentally reduced cDC
frequencies or life spans led to defective priming of T cell responses (6–10), as well as defective DC-mediated negative selection, T cell–mediated autoimmunity (9), inflammation, and
myeloproliferation (11). Although these examples show the importance of maintaining precise physiological numbers of cDCs, it
remains unclear which signals and molecular mechanisms control
cDC homeostasis and turnover in vivo.
*Institute for Immunology, Ludwig-Maximilians-University, 80336 Munich, Germany;
†
German Center for Neurodegenerative Diseases, 81377 Munich, Germany;
‡
Biotech Research and Innovation Center, Molecular Pathology Section, 2200
Copenhagen, Denmark; xNeuroproteomics, Technical University Munich, 81675 Munich,
Germany; and {Munich Cluster for Systems Neurology, 81377 Munich, Germany
Received for publication March 20, 2015. Accepted for publication September 1,
2015.
This work was supported by the Deutsche Forschungsgemeinschaft (SFB914
TPA06 and SFB1054 TPB03 to T.B.) and by Joint Programming Neurodegenerative
Diseases “Risikofaktoren und modifizierende Mechanismen bei Fronto-Temporaler
Demenz” (to S.F.L.). S.L. was supported by a Ph.D. grant from the China
Scholarship Council.
The mass spectrometry proteomics data presented in this article have been submitted
to the ProteomeXchange Consortium via the PRoteomics IDEntifications (PRIDE)
database repository with the dataset identifier PXD001636.
Address correspondence and reprint requests to Prof. Thomas Brocker, Institute for
Immunology, Ludwig-Maximilians-University, Goethestrasse 31, D-80336 Munich,
Germany. E-mail address: [email protected]
Abbreviations used in this article: 7-AAD, 7-aminoactinomycin D; BAD, Bcl-2–
associated death promoter; BM, bone marrow; BMDC, BM-derived DC; cDC, conventional dendritic cell; DC, dendritic cell; DCRhoA-ko, DC-specific RhoA deficient; Flt3L, Flt3 ligand; GO, gene ontology; HSC, hematopoietic stem cell; ko,
knockout; preDC, precursor DC; proDC, progenitor of preDC; qPCR, quantitative
PCR; RIP, receptor-interacting protein; wt, wild-type.
Copyright Ó 2015 by The American Association of Immunologists, Inc. 0022-1767/15/$25.00
www.jimmunol.org/cgi/doi/10.4049/jimmunol.1500676
Initially cDCs in lymphoid organs were thought to be nondividing and short-lived (2, 12–14). However, more refined analyses
of cDCs in parabiont experiments indicated that $5% of cDCs in
spleen are in the cell cycle (15, 16). The constant replenishment of
cDC pools by DC precursors from the circulation maintains the
homeostasis of a defined number of cDCs in lymphoid organs
(16). Therefore, cell cycle control and in situ proliferation of cDCs
play important roles in the maintenance of correct cDC numbers
and immune homeostasis. Control of cytokinesis might also regulate precursor–product relationships between circulating precursor
DCs (preDCs) and cDCs, which occupy distinct niches and have
different proliferative capacities. For cDCs, these mechanisms are
not described; knowledge of cytokinesis regulation in DCs may
increase our knowledge of the regulation of key checkpoints of
immune homeostasis.
The two best-studied cytokines controlling the DC lineage are
GM-CSF and Flt3 ligand (Flt3L). Although lack of GM-CSF does
not abrogate generation of cDCs in mice (17), it caused a reduction in CD103+ and CD11b+ cDC subsets in various nonlymphoid
organs by apoptosis, suggesting its role as a DC survival factor
(18). The receptor for Flt3L, Flt3 (CD135), is expressed on various
DC progenitors and drives differentiation of cDCs from hematopoietic stem cells (HSCs) (18). It also contributes to maintenance
of homeostatic DC numbers by regulating proliferation (19). GMCSF signals involve RhoA GTPase pathways in various cell types
(20, 21). Also, signaling via mutated Flt3 in acute myeloid leukemia exploits a pathway involving RhoA for survival and proliferation of tumor cells (22). Therefore, RhoA might be involved
in the integration of signals through GM-CSFR and Flt3, and it
could play a pivotal role in the control of DC proliferation, longevity, and homeostasis.
RhoA is a member of the Rho GTPase family, which cycles
between active GTP-bound and inactive GDP-bound states in response to various external stimuli. RhoA plays a central role in
regulating cytoskeleton organization, adhesion, proliferation, migration, cytokinesis, and cell survival (23). However, most of our
knowledge about RhoA function is based on overexpression of
dominant-negative or constitutively active mutants in cell lines
with little physiological relevance and nonspecific effects on other
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Tissues accommodate defined numbers of dendritic cells (DCs) in highly specific niches where different intrinsic and environmental
stimuli control DC life span and numbers. DC homeostasis in tissues is important, because experimental changes in DC numbers
influence immunity and tolerance toward various immune catastrophes and inflammation. However, the precise molecular mechanisms
regulating DC life span and homeostasis are unclear. We report that the GTPase RhoA controls homeostatic proliferation, cytokinesis,
survival, and turnover of cDCs. Deletion of RhoA strongly decreased the numbers of CD11b2CD8+ and CD11b+Esamhi DC subsets,
whereas CD11b+Esamlo DCs were not affected in conditional RhoA-deficient mice. Proteome analyses revealed a defective prosurvival
pathway via PI3K/protein kinase B (Akt1)/Bcl-2–associated death promoter in the absence of RhoA. Taken together, our findings
identify RhoA as a central regulator of DC homeostasis, and its deletion decreases DC numbers below critical thresholds for immune
protection and homeostasis, causing aberrant compensatory DC proliferation. The Journal of Immunology, 2015, 195: 4244–4256.
The Journal of Immunology
GTPases (23, 24). Cell-specific RhoA deletion demonstrated that
its effects vary greatly in different cell types. For example, RhoA
deficiency results in cytokinesis failure in keratinocytes (25),
migration defects in mouse embryonic fibroblasts (26), and necrotic loss of HSCs (27).
In this study, we used CD11c-Cre mice crossed with RhoAfl/fl
mice to study the role of RhoA in cDCs in vivo. We demonstrate
that RhoA controls DC homeostasis, and loss of RhoA results in
a subset-specific decrease in cDC numbers in lymphoid organs.
RhoA deficiency causes cytokinesis failure that is accompanied
by increased rates of apoptosis via inhibition of the prosurvival
pathway PI3K/Akt1/Bcl-2–associated death promoter (BAD),
which likely causes a reduction in DC numbers. Our results indicate that RhoA controls DC homeostasis by regulating cytokinesis and cell survival.
Materials and Methods
Mice
Flow cytometry and sorting
Histology
For preparation of cryosections, spleen was embedded in O.C.T. compound (no. 4583; Sakura Finetek) and snap frozen; 5-mm sections were
cut with a cryostat (Jung Frigocut 2800 E; Leica), air-dried overnight,
fixed in acetone (220˚C for 10 min), and air-dried for a minimum of
12 h. Sections were rehydrated for 15 min in PBS containing 0.25% BSA
and blocked for 15 min in PBS containing 0.25% BSA and 10% mouse
serum. Rat anti-mouse CD8-FITC, CD11c-PE, MOMA-allophycocyanin
(all from BD Pharmingen), and DAPI (1 mg/ml; Molecular Probes) diluted
in blocking buffer were added to the sections and incubated for 30 min. After
washing, sections were mounted in Fluoromount-G (Southern Biotech).
Sections were analyzed on an Olympus BX41TF-5 microscope (Olympus
CELL-BND-F software) equipped with an F-View II Digital Micro
Camera (Olympus).
Image stream
For cell mitosis analysis, 3 3 106 BMDCs or splenic DCs isolated by
MACS were stained with CD11c and MHC class II Abs. Cells were fixed
in BD Cytofix/Cytoperm buffer for 30 min at 4˚C, washed with Perm/Wash
buffer, and stained with DAPI in Perm/Wash buffer at 4˚C for 10 min.
After washing, cells were resuspended with 50 ml PBS. Mitosis was
analyzed using ImageStream X Mark II (Amnis, Seattle, WA). Flow
cytometry and image data were acquired and analyzed by INSPIRE and
IDEAS software, respectively. Different populations were discriminated
based on combined features of size, shape, and fluorescent intensity.
BrdU labeling in vivo
Mice were injected i.p. with 1 mg BrdU in PBS and then continuously
received drinking water containing 0.8 mg/ml BrdU for 3 d. After various
times, splenocytes were harvested and processed using a BrdU flow kit
(BD Pharmingen).
Spleens or lymph nodes were digested in a solution of Liberase and
DNase I (Roche) in serum-free medium for 30 min at 37˚C. Unless stated
otherwise, we used pooled axillary, brachial, and inguinal lymph nodes
for analyses. RBCs were removed with ACK buffer (0.15 M NH4Cl,
1 mM KHCO3, and 0.1 mM Na2EDTA [pH 7.4]) for 5 min at room temperature, and cells were filtered through a 70-mM nylon mesh strainer
(BD Biosciences). The following Abs were used for flow cytometry:
CD3 (145-2C11), CD4 (GK1.5), CD11b (M1/70), CD11c (N418),
MHCII (M5/114), Esam (1G8), F4/80 (BM8), and Flt3 (A2F10) (eBioscience); CD43 (S7), B220 (RA3-682) CD45RA (14.8), CD103 (M290),
NK1.1 (PK136), Sirpa (P84), and Ki67 (BD Pharmingen); CD8a
(MCD0826) (Invitrogen); and CD45.1 (A20), CD45.2 (104), and PDCA-1
(129c1) (BioLegend). Dead cells were identified by Aqua LIVE/DEAD
dye (Invitrogen). Progenitors of preDCs (proDCs) and preDCs were sorted
from Flt3L culture on day 3.5 using a FACSAria. ProDCs were identified as Lin2 CD11c2 MHCII2 CD43+Ly6C2 , whereas preDCs were
identified as Lin2CD11c+MHCII2CD43+Sirpaint. To sort preDCs from
spleen, total splenocytes were coated with biotin-conjugated CD3, B220,
NK1.1, and CD19 Abs, and Lin+ cells were depleted by Streptavidin
MicroBeads (Miltenyi Biotec)–mediated column sorting. The flow-through
Lin2 cells were used for preDC sorting using a FACSAria.
TUNEL+ DCs in spleen were detected using an ApopTag Fluorescein
In Situ Apoptosis Detection Kit (Millipore, Billerica, MA), per the
manufacturer’s instructions. Briefly, splenocytes were labeled with mAbs
specific for surface markers, and cells were fixed in 1% paraformaldehyde in PBS for 30 min at room temperature. Cells were permeabilized
with 0.1% Triton X-100 on ice for 2 min. Then, cells were incubated
with working- strength TdT enzyme at 37˚C for 30 min. Adding stop/
wash buffer stopped the reaction. After three washes in PBS, cells were
incubated with anti-digoxigenin fluorescein conjugate for 30 min at room
temperature, and DNA of cells was stained with DAPI. TUNEL+ DCs
were determined by flow cytometry.
OT-I and OT-II T cell adoptive transfer
Proteome analysis
OT-I (CD8+) and OT-II (CD4+) T cells were isolated from lymph nodes
and spleens of OT-I or OT-II donor mice using magnetic CD8 or CD4
negative selection (.96% purity, MACS; Miltenyi Biotec). Either 1 3
106 OT-I or 2 3 106 OT-II CFSE-labeled T cells were injected i.v. into
the lateral tail veins of age- and sex-matched recipient mice. One day
later, recipient mice were immunized by injecting 100 mg OVA protein in
PBS (10 mg LPS) into the tail vein. OT-I and OT-II T cell proliferation
was analyzed by CFSE dilution 3 d after immunization.
Cell pellets were mixed with 400 ml lysis buffer (4% SDS, 100 mM DTT,
100 mM Tris [pH 7.6]), incubated for 5 min at 95˚C, sonicated for 5 min in
a sonication bath, and incubated again for 5 min at 95˚C. The samples were
cleared by centrifugation at 13,000 rpm for 5 min. Proteins were digested
as described previously (29). In brief, seven volumes of 8 M urea were
mixed with the lysate, and the sample was loaded onto a spin column
(30 kDa nominal m.w. cut off; Sartorius Stedim). Alkylation was performed with iodoacetamide, and samples were digested using trypsin
(Promega) for 16 h. Peptides were eluted in double-distilled water and
subsequently fractionated into three fractions using strong anion exchange
in a pipet tip–based format, as described (30). Fractions were generated
using elution buffers with a pH of 11, 6, and 3. Sample clean-up and
desalting were achieved on homemade C18 STAGE tips (31). Samples
were analyzed in liquid chromatography–mass spectrometry–mass spectrometry mode on the LTQ Orbitrap Velos Pro Mass Spectrometer coupled to an
Easy-nLC II Liquid Chromatograph (both from Thermo Fisher Scientific)
using homemade 15-cm columns (2.4 mm resin) and 3-h gradients. A total of
3 mg peptides was injected per run. Five biological replicates with two
technical replicates each were analyzed, alternating control and knockout (ko) samples. Label-free quantification and data analysis were performed using the freely available MaxQuant and Perseus software
Generation of Flt3L DCs and bone marrow–derived DCs
DCs were generated from bone marrow (BM) in the presence of Flt3L.
BM was extracted from femur and tibiae, and erythrocytes were lysed
with ACK buffer. BM cells were cultured at 1 3 106 cells/ml in RPMI
1640 medium (with 10% FCS, Pen-Streptomycin, and 50 mM 2-ME) in
the presence of 20 ng/ml homemade recombinant murine Flt3L. At day
3, cells were collected and used for proDC and preDC sorting, as described (28). For BM-derived DC (BMDC) culture, BM cells were
cultured at 1 3 106 cells/ml in IMDM (Sigma) (with 10% FCS, PenStreptomycin, and 50 mM 2-ME) in the presence of 20 ng/ml homemade
rGM-CSF. The medium was refreshed on days 3, 7, and 10. BMDCs can
be used from days 10 to 13.
Generation of BM chimeras
BM was harvested from femurs and tibiae of 6–8-wk-old wild-type (wt;
CD45.1) and DC-specific RhoA-deficient (DCRhoA-ko) donor (CD45.2)
mice. RBCs were lysed with ACK buffer, and 5 3 106 total BM cells (1:1
ratio of BM cells from both donors) was injected i.v. into lethally irradiated
(600 rad; split dose at days 21 and 0) recipient mice (age 10–12 wk).
Chimeras were analyzed 8–10 wk after BM reconstitution.
TUNEL assay
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RhoAfl/fl mice were described previously (25). Six- to eight-week-old
CD11c-Cre and CD11c-Cre RhoAfl/fl mice were used for this study.
Thy1.1+ OT-I and OT-II mice with a transgenic TCR recognizing
pOVA257–264 or pOVA323–339, respectively, were originally obtained
from The Jackson Laboratory. All mice were bred and maintained at the
animal facilities of the Institute for Immunology, Ludwig-MaximiliansUniversity, in accordance with established guidelines of the Regional
Ethics Committee of Bavaria.
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(http://www.maxquant.org). Only proteins that were quantifiable with two
or more unique peptides in at least three of five biological were included in
the dataset. The p value was corrected after false discovery rate–based
multiple hypothesis testing. The mass spectrometry proteomics data were
submitted to the ProteomeXchange Consortium (32) (http://proteomecentral.
proteomexchange.org) via the PRIDE partner repository with the dataset
identifier PXD001636.
Quantitative PCR
Western blots
The following Abs were used for Western blot: actin, RhoA (67B9), PI3K-g
(D55D5), Akt, phospho-Akt (Ser473), Bad (D24A9), and phospho-Bad
(Ser136) (Cell Signaling). Proteins from BMDCs and splenic DCs were
separated by SDS-PAGE (12%). After transferring to a nitrocellulose membrane, proteins were incubated with primary Ab, washed, and incubated with
secondary HRP-labeled Ab. Then, membranes were visualized using luminescent substrate ECL (Amersham, Piscataway, NJ). Western blot bands
were quantified by ImageJ software.
Detection of Flt3L in serum
Serum concentrations of Flt3L were tested by mouse Flt3L ELISA (R&D
Systems), according to the manufacturer’s instructions.
Statistical analysis
Statistical differences between the experimental groups were determined
by the Student two-tailed t test. The p values ,0.05 were considered
significant.
Results
RhoA is essential for the homeostasis of DCs in spleen
To explore the function of RhoA in DCs, we crossed mice carrying loxP-flanked RhoA alleles (RhoAfl/fl) (25) with CD11c-Cre
mice (33) to generate DCRhoA-ko mice. We confirmed RhoA
deficiency by qPCR and Western blot analyses of MHCII+CD11c+
cDCs (Fig. 1A). Flow cytometry analyses revealed that frequencies of cDC populations were significantly decreased in spleens of
DCRhoA-ko mice (Fig. 1B). The percentages of CD11c+MHCII+
cDCs were reduced by .50% compared with controls (Fig. 1B).
Further analyses of CD11b+ and CD8+ DC subsets of the spleen
showed that both subsets were substantially reduced (Fig. 1B).
CD11b+ DCs of the spleen can be subdivided into Esamlo and
Esamhi populations, with the latter subpopulation being dependent
on signals through Notch2 and lymphotoxin b-receptor and having a higher turnover rate (10, 34, 35). Deletion of RhoA results in
a dramatic, but highly selective, reduction in Esamhi DCs, whereas
the total number of Esamlo DCs is unaffected and, therefore, seems
independent of RhoA (Fig. 1B, 1C). Similar results were obtained
when CD4 was used as a marker for DCs, with CD4+ DCs being
more affected than CD42 DCs (Fig. 1B). In lymph nodes, the total
numbers of migratory and resident RhoA-ko DCs were substantially reduced (Fig. 1D). Although migratory DC subpopulations
(CD103+, CD11b+, CD1032CD11b2) were equally affected by
lack of RhoA, in the resident DC subpopulation primarily CD8+
DCs were reduced in frequency and total number; reductions in
RhoA-deficient CD11b+ DCs were not significant (Fig. 1D). A
similar picture was found in mesenteric lymph nodes draining the
intestine, with migratory CD103+CD11b2 and CD103+CD11b2
DC subsets and CD8+ resident DCs being the most affected by
RhoA deficiency, whereas the numbers of CD1032CD11b+ migratory and CD11b+ resident DCs were normal (Fig. 1E). However, the numbers of other cell types, such as CD4+ T cells, B220+
B cells, Ly6C+CD11b+ monocytes, F4/80+ macrophages (Fig. 1F),
and NK cells (data not shown), were unaltered. CD8+ T cells,
which partially express CD11c-driven transgenes, also were reduced in number to some extent (Fig. 1F). Histological analyses
revealed that DCRhoA-ko mice have an intact overall architecture
of the spleen with normal white/red pulp organization (Fig. 1G).
As previously described, CD8+ DCs are primarily located in T cell
zones (36), as well as in marginal zones, and fewer numbers are
found in the red pulp of the spleen (37), whereas CD11b+ DCs are
preferentially located in the bridging channels (10) and red pulp
of the spleen. Although both DC subsets could be identified in
spleens of DCRhoA-ko mice in their proper locations, their total
numbers were markedly decreased in the absence of RhoA
(Fig. 1G), confirming the flow cytometry data (Fig. 1B, 1C). As
a consequence, priming of adoptively transferred CD8+OT-I or
CD4+OT-II T cells with OVA protein was strongly impaired in
DCRhoA-ko mice (Fig. 1H). Neither OT-I nor OT-II T cells
proliferated to the same extent as observed in control animals
(Fig. 1H). Taken together, our results indicate that RhoA controls homeostasis of the majority of DC subsets; upon deletion
of RhoA, DC numbers decrease beyond a critical level for
T cell priming.
Deletion of RhoA does not affect generation of cDCs from
progenitors
cDCs develop from preDCs, which originate from hematopoietic
progenitor cells in the BM (38–40). It was reported recently that
RhoA deficiency in hematopoietic progenitor cells causes multilineage hematopoietic failure (27). To control whether the observed
decrease in cDC numbers in DCRhoA-ko mice was due to defects
in preDCs or occurred later in cDC development, we next analyzed
preDCs from BM and spleen. Because preDCs constantly leave the
BM and enter lymphoid organs (40), we also analyzed preDCs after
their migration to spleens of DCRhoA-ko mice. The frequency or
total cell number of preDCs (Lin2CD11c+MHCII2CD43+Sirpaint)
in BM and spleen was not affected by RhoA deficiency (Fig. 2A).
PreDCs were identified according to their expression of CD11c, the
regulatory elements of which control expression of Cre recombinase
and, therefore, deletion of the floxed RhoA gene. Despite CD11c
expression, preDCs from BM did not show reduced levels of RhoA
mRNA (Fig. 2B). Only those preDCs that had emigrated from the
BM and reached the spleens of DCRhoA-ko mice showed significantly lower expression levels of RhoA mRNA (Fig. 2B). However,
this loss of RhoA mRNA apparently did not affect the size of the
preDC pool in spleens in vivo (Fig. 2A). To control the abilities of
preDCs to generate cDCs in vitro, we analyzed Flt3L culture–
enriched proDCs (Lin2CD11c2CD43+Ly6C2) and preDCs (Lin2
CD11c+MHCII2CD43+Sirpaint), as described previously (28), and
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Total RNA was isolated from 1 3 106 BMDCs or splenic DCs with TRIzol
reagent (Invitrogen) and reverse transcribed into cDNA using a QuantiTect
Reverse Transcription Kit (QIAGEN). The TaqMan assay was performed
with the LightCycler TaqMan Master Kit and the Universal Probe Library
Set mouse (Roche), according to the manufacturer’s instructions, on a
CFX96 Real-Time System (Bio-Rad, Hercules, CA). The housekeeping
gene HPRT was used for normalization. The mRNA level of genes was
measured by quantitative PCR (qPCR), according to the Universal Probe
Library system (Roche) with the following primers and Universal Probe
Library Probes (Roche): HPRT: forward, 59-TCCTCCTCAGACCGCTTTT-39, reverse, 59-CCTGGTTCATCATCGCTAATC-39, and probe #95;
RhoA: forward, 59-CCTGTGTGTTTTCAGCACCTT-39, reverse, 59-ACCTCTGGGAACTGGTCCTT-39, and probe #46; PI3K-g: forward, 59-GGAGACTGAATCTCTGGACCTG-39, reverse, 59-GTGGCATCCTTTACAATCTCG-39, and probe #29; Ripk1: forward, 59-TACCTCCGAGCAGGTCAAAT-39, reverse, 59-AAACCAGGACTCCTCCACAG-39, and
probe #50; Ripk3: forward, 59-CCGACGATGTCTTCTGTCAA-39, reverse,
59-GACTCCGAACCCTCCTTTG-39, and probe #6; Glud1: forward, 59GGTCATCGAAGGCTACCG-39, reverse, 59-TCAGTGCTGTAACGGATACCTC-39, and probe #76; TNF-a: forward, 59-CCCCAAAGGGATGAGGTAGT-39, reverse, 59-CTAACCCGTCTTGCTTGTGAG-39, and probe
#104; Fth1: forward, 59-GCCAGAACTACCACCAGGAC-39, reverse,
59-CCACATCATCTCGGTCAAAAT-39, and probe #75; TNFR1: forward,
59-TGTCAATTGCTGCCCTGTC-39, reverse, 59-GATGTATCCCCATCAGCAGAG-39, and probe #40. The relative expression of target genes was
calculated using the 22DDCt method.
RhoA INDUCES SURVIVAL OF DCs
The Journal of Immunology
4247
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FIGURE 1. RhoA is essential for the homeostasis of DCs. (A) RhoA deficiency is confirmed in mRNA and protein levels by qPCR and Western blot in
spleen cDCs isolated via MACS. (B) The frequency of cDCs (CD11c+MHCII+) is significantly decreased in the spleen of DCRhoA-ko mice. cDCs can be
separated into CD8+ and CD11b+ DC subsets. Expression of Esam and CD4 was further analyzed in CD11b+ DCs. The percentages represent mean 6 SEM
(n = 3). (C) Bar graph shows total cell number of spleen cDCs and different spleen DC subsets (n = 3). (D) Surface lymph node DCs from pooled inguinal,
brachial, and axillary lymph nodes were analyzed based on the expression of CD11c and MHC class II. Resident DCs (MHCIIint) were further analyzed for
expression of CD8 and CD11b. Migratory DCs (MHCIIhi) were analyzed for expression of CD103 and CD11b. Percentages represent mean 6 SEM.
Bar graphs show the total cell number of DCs and DC subsets in lymph node (n = 3). (E) Same analysis as shown in (D) for resident and migratory DCs
from mesenteric lymph nodes. (F) Bar graph shows total number of CD19 + B cells and CD4+ and CD8+ T cells in spleen (Figure legend continues)
4248
RhoA INDUCES SURVIVAL OF DCs
monitored their abilities to generate cDCs in vitro (Fig. 2C).
ProDCs first develop into CD11c+ preDCs and differentiate
further into CD11c+MHCII+ cDCs (Fig. 2C). These analyses
showed that proDCs, as well as preDCs, from BM and spleen of
DCRhoA-ko mice could differentiate normally into preDCs and
cDCs or cDCs, respectively (Fig. 2C). ProDCs isolated from BM
and preDCs from BM or spleens of DCRhoA-ko mice did not
show any deficiency with regard to the generation of their expected progeny in vitro. This suggests that, in DCRhoA-ko mice,
neither the seeding of preDCs from BM to spleen nor the development of cDCs from their precursors is affected (Fig. 2B, 2C).
of CD11b+ and CD8+ DCs (35), accumulated in DCRhoA-ko–
derived DCs proportionally compared with the other DC populations (Fig. 3A) but were strongly reduced with regard to total
cell number (Fig. 3B). B and T cells served as internal controls
because, as a result of the specificity of the CD11c promoter
RhoA, they should not be deleted. Accordingly, both lymphocyte
subsets were not reduced when derived from DCRhoA-ko BM;
rather, they were present in slightly increased numbers (Fig. 3B).
Taken together, these results suggest that the reduction in cDCs
is cell intrinsic in DCRhoA-ko mice and directly due to the loss
of RhoA.
DCRhoA-ko phenotype is DC intrinsic
RhoA-deficient cDCs show decreased long-term survival
We next assessed whether the reduction in cDCs found in
DCRhoA-ko mice is cell intrinsic or caused by external or secondary factors. To this end, we reconstituted lethally irradiated B6
mice with a 1:1 BM mixture derived from CD11c-Cre (CD45.1)
and DCRhoA-ko (CD45.2) mice. In this competitive situation,
DCRhoA-ko–derived DCs reconstituted the cDC pool in spleens
very inefficiently (13%, Fig. 3A). Reductions in different cDC
subsets were also detected (Fig. 3A); similar to nonchimeric
DCRhoA-ko mice (Fig. 1C), the total cell numbers of RhoAdeficient CD8+ and CD11b+Esamhi DCs were markedly decreased (Fig. 3B). In contrast, CD11b+Esamlo DCs showed only
a slight decrease in cell number (Fig. 3B). In addition, a distinct
population of CD82 CD11b2 cDCs, the natural immature stages
The above experiments showed that lack of RhoA caused lower
numbers of DCs. To determine whether this is the result of altered
survival and turnover of cDCs, we next performed in vivo BrdUlabeling studies (Fig. 4). Three days of continuous BrdU labeling
revealed a statistically significantly higher BrdU incorporation
rate into cDCs and CD8+ and CD11b+ DCs in DCRhoA-ko mice
(Fig. 4A). When we analyzed the Esamlo and Esamhi DC subsets
at day 9 of the experiment, we detected significantly reduced
frequencies of BrdU+ DCs in CD8+, Esamhi, and Esamlo subsets,
indicating faster turnover of all subsets analyzed (Fig. 4B). A
more refined analysis was then performed after a 2-h short-term
BrdU pulse in vivo (Fig. 4C); this revealed strongly elevated
incorporation of BrdU into RhoA-ko DC subsets, suggesting
(left panel) and numbers of CD11b+Ly6C+ monocytes or F4/80+ macrophages (right panel) (n = 3). (G) Histology staining shows an intact spleen structure
but markedly decreased DC numbers in DCRhoA-ko mice. (H) A total of 1 3 106 CFSE-labeled CD8+ OT-I T cells or 2 3 106 CFSE-labeled CD4+ OT-II
T cells was injected i.v. into the indicated recipient mice on day 21. Then mice were immunized with 100 mg OVA protein/10 mg LPS i.v. On day 4, mice
were sacrificed to measure CFSE dilution of OT-I or OT-II cells by gating on live Thy1.1+CD8+ or Thy1.1+CD4+ cells, respectively. Bar graphs show the
percentage (upper panel) or total numbers (lower panel) of divided T cells in spleen. Figures show representative data of one of at least six (A–C and F) or
three (E, G, H) experiments with similar outcome. Asterisks indicate statistical significance. *p , 0.05, **p , 0.01, ***p , 0.001.
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FIGURE 2. Deletion of RhoA does not affect generation of cDCs from progenitors. (A) Identification of preDCs in BM and spleen. preDCs are recognized as Lin2CD11c+MHCII2CD43+Sirpaint. Bar graphs show the percentage and total number of preDCs in BM and spleen (n = 3). Lin: CD3/CD19/
NK1.1/B220. (B) RhoA mRNA in preDCs. Bar graph shows RhoA mRNA transcript in preDCs sorted from BM culture supplemented with Flt3L or directly
from spleen (n = 3). (C) Development of preDC progenitors and preDCs in vitro after reculture. The expression of CD11c and MHC class II was monitored
over 5 days by flow cytometry. Bar graphs show the generation of preDCs or cDCs from preDC progenitors or preDCs (n = 3). Representative data of one of
three experiments with similar outcome are shown. Asterisks indicate statistical significance. *p , 0.05, **p , 0.01, ***p , 0.001.
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4249
FIGURE 3. DC reduction caused by RhoA deficiency is cell intrinsic. (A) Irradiated B6 mice (CD45.2) were reconstituted with an equal mixture of
CD45.1 wt and CD45.2 DCRhoA-ko BM cells. Spleen cDC and DC subsets were characterized by flow cytometry. (B) Bar graph shows the ratio of total
cell numbers of CD45.2+ DCRhoA-ko–derived cells/CD45.1+-derived wt cells. Identical cell numbers result in a ratio of 1 (dashed line). Bars represent
mean 6 SEM for three mice. Figure shows representative data of one of four experiments with similar outcome.
RhoA deficiency indirectly enhances DC proliferation but
controls DC cytokinesis intrinsically
DCs were thought to be nondividing cells, until recent studies
showed that ∼5% of cDCs were dividing at any time in the
spleen (15, 16). Therefore, cDC proliferation is considered an
important mechanism to maintain homeostatic cDC numbers in
lymphoid organs. RhoA inhibition by dominant-negative mutants was previously reported to enhance proliferation of hematopoietic stem and progenitor cells (41). Furthermore, RhoA
controls cytokinesis by regulation of cortical contractility and
cleavage furrow formation during proliferation (42, 43). To test
whether RhoA controls DC proliferation, we analyzed spleen
cDCs for DNA content and expression of the proliferation
marker Ki67 (Fig. 5). As described previously, ∼5% of CD11c+
MHCII+ spleen cDCs were in the cell cycle (DAPI+Ki67+) in
control mice (Fig. 5A). However, two-fold more cDCs were
dividing in DCRhoA-ko spleens compared with DCs in control
mice (Fig. 5A). As a consequence of such compensatory proliferation, we found similar total numbers of Ki67+ DCs in
controls and DCRhoA-ko mice (data not shown). To test
whether such increased proliferation was caused by deletion of
RhoA itself or a secondary effect, we generated mixed BM
chimeras, as described above (Fig. 3). Cre control–derived
cDCs should develop in normal numbers in the same host as
RhoA-deficient cDCs and fill the cDC pool in a competitive
situation. Analyses of cDCs in mixed chimeras showed that Cre
control and RhoA-deficient cDCs had the same normal proliferation rate (∼5%) (Fig. 5B). As a consequence of this normal
proliferation rate of RhoA-ko DCs in the competitive condition
in mixed chimeras, the total numbers of Ki67+ RhoA-ko DCs
were strongly decreased (data not shown). These results indicate that the compensatory proliferation of cDCs in DCRhoAko mice (Fig. 5A) was most likely due to secondary effects
caused by disturbed cDC homeostasis and reduced cDC numbers in the absence of RhoA but was not due to RhoA deficiency itself. It was reported that the reduction in cDC numbers
as a result of diphtheria toxin–mediated ablation causes myeloproliferation because of increased levels of Flt3L (11). Because cDCs are major “consumers” of Flt3L among peripheral
hematopoietic cells (44), loss of cDCs might cause elevated
levels of Flt3L, triggering increased cDC proliferation. Therefore, we tested for levels of Flt3L and found significantly elevated
concentrations in the serum of DCRhoA-ko mice compared with
control animals (Fig. 5C). In contrast, when wt DCs could fill up
the DC compartment in mixed BM chimeras, Flt3L levels returned
to normal and were comparable to those in wt mice (Fig. 5C). This
finding suggests that reductions in cDC numbers as a result of the
absence of RhoA leads to elevated Flt3L concentrations in serum,
which, in turn, induces enhanced cDC proliferation in DCRhoAko mice but not in mixed chimeras. These results suggest that
deficiency of RhoA does not intrinsically lead to enhanced DC
proliferation and that RhoA is even dispensable for proliferation
of cDCs.
We next analyzed the distribution of RhoA-ko cDCs in the
different stages of the cell division cycle (G1, S, G2/M) (25, 26).
CD11c +MHCII + cDCs from spleens of DCRhoA-ko mice
(Fig. 5D) or mixed BM chimeras (Fig. 5E) showed a similar cell
cycle distribution, with significantly lower frequencies of DCs in
the G1 phase and more DCs in the S/G2-M phases in the absence
of RhoA (Fig. 5D, 5E). These data suggest that RhoA regulates
cell cycle progression in cDCs. We next analyzed mitosis of
spleen DCs using ImageStream analyses (Fig. 5F). Ex vivo–isolated spleen cDCs from DCRhoA-ko mice showed a statistically
significant accumulation of both telophase-like and anaphase-like
cells, which normally coincide with cytokinesis (45). Also, the
fact that frequencies of DCs in metaphase were not significantly
altered and the nuclei were properly separated (Fig. 5F) suggests
that RhoA controls cytokinesis in cDCs.
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increased proliferation (Fig. 4C). CD8+ and CD11b+Esamhi DCs
were most affected and showed the highest BrdU-incorporation rates (Fig. 4C). However, these DCRhoA-ko–derived cDC
subsets showed normal BrdU incorporation rates during the 3-d
pulse phase in a competitive transplantation model in mixed
BM chimeras, indicating normal proliferative behavior, despite
the absence of RhoA, when the cDC pool was filled up with
competitor control cDCs (Fig. 4D). BrdU analyses during the
12-d chase period revealed that survival and turnover of RhoAdeficient cDCs were much shorter compared with control DCs
(Fig. 4A, 4D, days 6–12). In mixed BM chimeras, a strongly
reduced survival was apparent, and CD11b+ cDCs were most
dramatically affected, showing a .50% reduced half-life
compared with the internal competitor control CD11b+ cDCs
(Fig. 4D). When we analyzed DC subsets at day 9 of this experiment, we found that CD8+ and Esamlo DCs, but not the
Esamhi subset, were affected (Fig. 4E, left panel). However, 3 d
later, after 9 d of BrdU-free chase, differences also became
apparent in Esamhi DCs, which showed significantly higher
turnover without RhoA (Fig. 4E, right panel). This suggests
that the reduced half-life of all DC subsets was a direct consequence of RhoA deficiency and demonstrates that RhoA
controls survival of cDCs.
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RhoA INDUCES SURVIVAL OF DCs
RhoA controls GM-CSF–induced cDC survival
To test whether loss of cDCs in the absence of RhoA was due
to apoptosis, we next performed TUNEL assays of spleen cDCs.
DCRhoA-ko cDCs had significantly higher frequencies of TUNEL+
DCs in spleen, where deletion of RhoA resulted in 2-fold increased
rates of apoptosis (Fig. 6A). Primarily, the CD8+ and Esamhi subpopulations were affected (Fig. 6A, bar graph). In mixed BM chimeras, the competitive condition between Cre control cDCs and
RhoA-ko cDCs led to a 4-fold increase in TUNEL+ cDCs from
DCRhoA-ko mice compared with control cDCs within the same
chimeras (Fig. 6A, lower panels), and the frequencies of TUNEL+
Esamhi DCs were even more pronounced (Fig. 6A). Also, when
we quantified apoptotic cDCs using Annexin V and 7-aminoactinomycin D (7-AAD), we detected a statistically significant
increase in apoptotic (AnnexinV+7-AAD2) CD8+ and CD11b+
Esamhi cDCs, whereas CD11b+Esamlo cDCs did not show significant changes (Fig. 6B). As controls, we analyzed MHCII+CD11c2
cells, including B cells, and CD8+CD11c2 cells, including CD8+
T cells, which did not contain increased frequencies of AnnexinV+
7-AAD2 apoptotic cells (Fig. 6B). To further confirm that RhoA
promotes DC survival, we generated DCs in vitro by culturing
BM cells from wt and DCRhoA-ko mice in the presence of GMCSF. Upon GM-CSF removal, DC survival was monitored by
flow cytometry. Consistent with the in vivo findings (Fig. 6A, 6B),
RhoA-deficient BMDCs showed significantly impaired survival
kinetics in the absence of the growth factor compared with
BMDCs from Cre control mice (Fig. 6C). Together, these findings
indicate that RhoA is critical for the homeostasis of DCs because it
regulates cDC survival, and lack of RhoA leads to cell death by
apoptosis.
Proteome analyses of RhoA-deficient cDCs
To explore the survival pathway controlled by RhoA in cDCs, we
performed proteome analyses to compare BMDCs from Cre control
and DCRhoA-ko mice (Fig. 7A). A total of 2699 proteins was
quantifiable, with inclusion criteria being quantification of at least
two unique peptides/protein in at least three biological replicates.
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FIGURE 4. RhoA-deficient cDCs show decreased long-term survival. (A) Control and DCRhoA-ko mice received in vivo BrdU labeling for 3 d (pulse
phase), followed by a BrdU-free chase. The frequency of BrdU+ splenic DCs was examined by flow cytometry at days 3, 6, 9, and 12. Spleen DCs were
first gated on MHCII+CD11c+ cells (data not shown). The percentage of BrdU+ DCs at each time point within total cDCs, CD8+ DCs, or CD11b+ DCs is
displayed (n = 3). (B) At day 9 of this experiment (3 d of BrdU pulse and 6 d of BrdU-free chase), Esamlo and Esamhi DC subsets, as well as CD8+ DCs
of the spleen, were analyzed. (C) BrdU incorporation rate was measured after 2 h of in vivo labeling with BrdU for the indicated spleen DC subsets. Bar
graph shows the percentages of BrdU+ DCs in different DC subsets (n = 3). (D) Mixed BM chimeras were generated by reconstituting irradiated wt mice
(CD45.1+) with a 1:1 mixture of BM from control (CD45.1+) and DCRhoA-ko (CD45.2+) mice. Chimeras were used for experiments 8 wk postreconstitution. Reconstitution always showed a disadvantage for BM of DCRhoA-ko origin, similar to the experiments shown in Fig. 3B. A 12-d BrdU
pulse-chase experiment was performed in mixed BM chimeras, as described in (A). Spleen DC subsets were identified first, as shown in Fig. 3A, by
gating on CD45.2+ or CD45.1+ MHCII+CD11c+ cells (data not shown) and were analyzed for CD8 and BrdU, as shown in (A). (E) Esamlo, Esamhi, and
CD8+ DC subsets from spleens of mixed chimeras were analyzed at day 9 (left panel) or day 12 (right panel) of the pulse-chase experiment shown in
(D). Figure shows representative data from one of three (C) or two (A, B, D, and E) experiments with similar outcome. Asterisks indicate statistical
significance. *p , 0.05, **p , 0.01, ***p , 0.001.
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4251
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FIGURE 5. RhoA deficiency causes enhanced proliferation of cDCs due to increased serum Flt3L and cytokinesis failure. (A) The proliferating spleen
DCs were detected as Ki67+ in S-G2-M phase (DAPIhi) (n = 3). (B) Spleen cDC proliferation was measured in mixed BM chimeras (n = 3). (C) Serum Flt3L
of DCRhoA-ko, wt, and mixed BM chimeric mice was detected by ELISA (n = 6). (D) Cell cycle of spleen cDCs of the indicated mice was analyzed by
DAPI staining of nuclei. Representative line graphs show the cells in different phases (G1, S, G2-M). Bar graph indicates the percentages of spleen DCs in
different phases (n = 3). (E) Same analysis as shown in (D), but spleen cDCs were isolated from mixed BM chimeras. (F) Representative images of spleen
DCs in different phases of mitosis were selected at random from more than 1000 images. Bar graph shows the percentages of spleen DCs in different phases
during mitosis (n = 3). Figure shows representative data from one of three (A), five (B and D), two (C), or three (E and F) experiments with similar outcome.
Asterisks indicate statistical significance. *p , 0.05, **p , 0.01, ***p , 0.001.
Fifty-four proteins were upregulated and 39 proteins were downregulated .3-fold in RhoA-deficient BMDCs (Fig. 7A). Among those,
several proteins related to gene ontology (GO) terms “cell cycle
regulation” (GO:0022402) and “regulation of programmed cell
death” (GO:0043067), such as E-cadherin, a- and b-catenin,
NEDD4, and FLNA, were deregulated. The second messenger
PI3K-g was reduced .6-fold in RhoA-ko BMDCs (Fig. 7A).
This was of potential interest because it plays an important role
in promoting cell survival and growth through activation of protein kinase B (or AKT) (46, 47). Active AKT phosphorylates
4252
RhoA INDUCES SURVIVAL OF DCs
proapoptotic protein BAD, thereby blocking BAD-induced apoptosis (48). To investigate whether RhoA deletion deregulates the
PI3K . AKT . BAD signaling pathway in DCs, we verified the
levels of PI3K-g, AKT, and BAD by Western blot of lysates from
BMDCs and purified spleen cDCs. As shown in Fig. 7B, RhoA
was effectively deleted from both sources of RhoA-ko DCs,
with some residual RhoA protein remaining in the BMDCs of
DCRhoA-ko mice (Fig. 7B). PI3K levels decreased in BMDCs
and spleen cDCs of DCRhoA-ko origin compared with wt controls
(Fig. 7B), confirming proteomics data (Fig. 7A). This decrease
was also confirmed on the mRNA level for DCRhoA-ko DCs,
indicative of an influence of RhoA on PI3K gene transcription
(Fig. 7C). Although total amounts of Akt protein were not altered
in DCRhoA-ko DCs (Fig. 7B), phospho-AKT was strongly decreased (Fig. 7B), possibly due to the reduction in the upstream
activator PI3K. Consequently, RhoA-ko DCs contained decreased
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FIGURE 6. RhoA deficiency leads to
increased apoptosis. (A) Apoptotic spleen
cDCs were detected by TUNEL assay in
the steady-state and in mixed BM chimeras. Cells not treated with TdT enzyme
were used as negative controls. Bar graphs
show the percentage of TUNEL+ splenic
cDCs, as well as distinct DC subpopulations (n = 3). (B) Annexin V and 7-AAD
were used to detect apoptotic cells in CD8+,
Esamhi, and Esamlo spleen DC subsets.
Bar graphs shows the percentage of all
Annexin V+ cells in DC subsets, as well as
B cells and CD8+ T cells as internal control (n = 3). (C) Apoptosis of BMDCs was
induced by removing GM-CSF from the
medium. BMDC survival was monitored
over 4 d (n = 3). Data are representative of
two (A) or three (B and C) experiments
with similar outcome. Asterisks indicate
statistical significance. *p , 0.05, **p ,
0.01; ***p , 0.001.
levels of phospho-BAD, whereas total protein levels of BAD were
not altered (Fig. 7B). These results suggest that deletion of RhoA
leads to reduced mRNA and protein levels of PI3K, causing inefficient engagement of the PI3K/AKT/BAD survival pathway,
which could be responsible for loss of cDCs in the absence of
RhoA. It was reported that RhoA is implicated in necrotic cell
death rather than apoptosis (27). In RhoA-deficient HSCs, the
TNF receptor-interacting protein (RIP) kinase pathway was shown
to regulate programmed necrosis upon upregulation of TNF-RIP–
associated genes (49). Therefore, we also analyzed necrosisrelevant genes, including Tnfa, Ripk3, Fth1, and Glud1 (27, 50),
but could not detect significant alterations in gene expression
(Fig. 7D). Therefore, our findings suggest that, in cDCs, RhoA
induces survival by inhibiting apoptosis. Lack of RhoA permits
the induction of apoptosis via the PI3K . AKT . BAD signaling
pathway.
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4253
Discussion
The Rho GTPase family has several members, with RhoA, Cdc42,
and Rac1 being the best characterized (51). They are known to
integrate signals for regulation of diverse cellular processes, such
as cell cycle progression, cytoskeleton rearrangement, and gene
expression (52). Furthermore, they control the Ag-presenting functions of DCs, as well as their maturation and migration capacities
(53–58). However, little is known about the DC-specific functions of
RhoA. Studies on SWAP70, a regulator of RhoA, indirectly showed
its involvement in motility, endocytosis (59), and MHC class II
surface localization in DCs (60). Our findings in DCRhoA-ko mice
suggest that RhoA regulates DC survival by inhibiting apoptosis.
Constant replenishment of DC precursors from blood, DC division, and cell death maintain DC homeostasis in lymphoid organs
in a dynamic balance (16). Deletion of RhoA in DCs led to
a significant decrease in DC numbers in lymphoid organs. Both
CD8+ DCs and CD11b+Esamhi DCs were dramatically decreased,
whereas the total number of CD11b+Esamlo DCs did not change.
Even in the competitive chimera model, in which the decrease in
RhoA-deficient CD8+ DCs and CD11b+Esamhi DCs became more
severe, the Esamlo population was affected much less than the
other DC subsets. The marker Esam was recently used to subdivide CD11b+ DCs into Esamhi and Esamlo subpopulations, which
are either more specialized for CD4+ T cell priming or secrete
proinflammatory cytokines, respectively (35). In contrast to the
other DC subsets, CD11b+Esamlo DCs are most likely derived
from monocytes (35, 61). This subset of DCs also was shown to
proliferate less than other DCs (35). Eventually, the low intrinsic
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FIGURE 7. RhoA promotes DC survival through PI3K/Akt/Bad pathway. (A) BMDCs of wt controls and DCRhoA-ko mice were used for proteome
analysis. A total of 2699 proteins was quantified by label-free quantification. Each black circle represents a protein that was quantifiable in at least three
biological replicates. The log2-fold change in the mean DCRhoA-ko/CD11c-CRE ratio is plotted against the 2log10 of the p value. The vertical dashed
lines mark a fold change $ 3. The dashed horizontal line shows the cutoff for statistical significance (p value # 0.05). Proteins that are up- or downregulated .3-fold in DCRhoA-ko mice compared with wt controls are considered significantly altered. Among those proteins, cell survival–related proteins
(red color) and cell cycle–related proteins (green color) are indicated. (B) The downregulation of PI3K-g was confirmed in BMDCs and spleen DCs by
Western blot. Spleen DCs were isolated by CD11c mAb-coated magnetic beads. Phosphorylation of Akt and BAD was confirmed by Western blot. Bar
graphs show quantification of Western blot bands done using ImageJ software. (C) Pi3kg mRNA was determined in spleen cDCs by qPCR. (D) qPCR to
examine expression of necrosis-induced genes from spleen cDCs. (E) RhoA signaling cascade regulating apoptosis of DCs. RhoA deletion caused
downregulation of PI3K-g, and the phosphorylation of its downstream effector Akt and BAD was inhibited. Inefficient phosphorylation of proapoptosis
protein BAD results in active BAD, leading to apoptosis. Asterisks indicate statistical significance. *p , 0.05, **p , 0.01, ***p , 0.001.
4254
DCs. In contrast, inhibition of PI3K alone in human cancer cell
lines was insufficient to induce apoptosis via BAD inactivation,
because other signaling pathways compensate for the lack of PI3K
signals (48). However, recent data from PI3K-g–deficient mice
also showed increased rates of apoptosis in normal leukocytes (68)
and strongly reduced DC numbers (69). We show that expression
of PI3K was significantly reduced in RhoA-deficient DCs, which
led to decreased phosphorylation of its downstream effector, Akt.
Thus, the proapoptosis protein BAD cannot be inhibited effectively by phosphorylation due to low levels of active Akt, resulting
in apoptosis of RhoA-ko DCs. It is unclear how RhoA regulates
protein levels of PI3K-g, but we also found that PI3K-g mRNA
transcripts were significantly reduced in RhoA-deficient DCs.
Previous reports showed transcriptional regulation by RhoA via
transcription factor GATA-4 as a nuclear mediator of RhoA signaling (70). Also, the c-Fos serum response element is used by
RhoA to control transcription in some cells (71). However, it is
unclear whether these or other RhoA-responsive transcription
factors transcriptionally regulate PI3K. Therefore, further work is
necessary to differentiate whether apoptosis of RhoA-ko DCs is
caused by defective mitosis, direct regulation of PI3K by RhoA, or
both. However, there is indirect evidence from the literature that
receptors for DC growth factors GM-CSF (20, 21) and Flt3L (22)
can signal via RhoA. In cells bearing the oncogenic form of Flt3,
a signaling pathway via PI3K is activated (22). Therefore, RhoA may
be a sensor for DC growth factors, and its absence could lead to cell
death by apoptosis, either directly via PI3K pathways or indirectly
via the block in mitotic cell division.
In summary, our findings indicate that RhoA is crucial for the
homeostasis of cDCs in spleen. Deletion of RhoA caused a significant decrease in cDCs due to decreased DC survival. Further
unraveling of the signaling pathways used by DC-specific growth
factors might help us to understand the feedback mechanisms that
control DC homeostasis in the steady-state and inflammation.
Acknowledgments
We thank Christine Ried for histological analyses and Dr. Jan Kranich
for critical discussions.
Disclosures
The authors have no financial conflicts of interest.
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proliferation rate and the constant replenishment of this subset
from monocytes or some earlier monocytic precursors might be
the reason why deletion of RhoA did not affect Esamlo DCs in the
steady-state.
Proliferation of cDCs was enhanced .2-fold upon deletion of
RhoA, and it compensated, to some degree, for the overall loss of
DC numbers. However, as shown in competitive transplantation
experiments, this effect was not caused directly by lack of RhoA
but, rather, by environmental factors. Flt3L was elevated in the
sera of DCRhoA-ko mice and may have become available in
higher amounts as a result of the loss of competitor DCs, as described in other models (11). However, although the experimental
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numbers, which was accompanied by a 1.5-fold increase in Flt3L.
Yet, such a modest increase in Flt3L was sufficient to cause DC
proliferation, as well as mild myeloid proliferative syndrome
(data not shown), as observed upon complete deletion of DCs
(11). These data illustrate that levels of DCs must be maintained at
optimal numbers, and RhoA seems to play a central role in this
balance. Even a subtle change in homeostasis leads to activation of
the feedback loop and increased compensatory DC proliferation.
Similarly, increased proliferation was observed previously upon
RhoA suppression by a dominant-negative mutant approach in
hematopoietic stem and progenitor cells (41), as well as in HSCs of
genetically RhoA-deficient mice (27). The investigators speculated
that secondary effects caused by acute hematopoietic stress were the
reason for such increased proliferation (27). It is also possible that,
in these models, increased availability of unidentified HSC factors/
niches caused RhoA-deficient cells to proliferate more strongly.
However, in another study, RhoA deletion in B cells using CD19Cre did not affect B cell proliferation, but it abrogated B cell differentiation (62). Together, these data indicate that proliferation of
hematopoietic cells seems to be independent of RhoA, but lack of
RhoA can cause increased compensatory proliferative responses in
some, but not all, cell types.
We identified reduced survival of RhoA-ko cDCs as a RhoAautonomous effect. In DCRhoA-ko mice, more cDCs were
TUNEL+ and AnnexinV+, indicating increased apoptosis rates. In
contrast, cell cycle control was deregulated in the absence of
RhoA, because a high frequency of DCs accumulated in mitotic
anaphase. It was proposed earlier in megakaryocytes that RhoA is
needed for cleavage furrows to complete separation of daughter
cells (63, 64). Defects in RhoA or its effector/regulatory proteins
caused various phenotypes of failure of cell separation (65, 66),
often accompanied by the presence of multinucleated cells (27,
63, 64). We found multinucleated cells in in vitro BMDC cultures
(data not shown) but not in RhoA-ko DCs in vivo. This might be
another example of RhoA’s effects being influenced by environmental differences, as we described recently for in vitro versus
in vivo conditions (25). Eventually, compensatory effects by RhoB
and RhoC might also cause such differences, as discussed previously (25). Because mitosis and apoptosis are connected, it is
possible that extended mitotic arrest due to the absence of RhoA
leads to mitotic cell death of cDCs. Recently, a genome-wide
screen identified BAD and other components of the mitochondrial apoptosis pathway as contributors to mitotic cell death (67).
We also identified lower amounts of phospho-BAD in RhoA-ko
cDCs; therefore, the link among mitochondrial apoptosis, mitotic
arrest, and RhoA deficiency likely contributes to the death of
RhoA-ko cDCs.
In addition, protein levels of PI3K-g, which has well-described
prosurvival functions, were significantly reduced in RhoA-ko
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