Synergistic actions of FGF2 and bone marrow transplantation mitigate radiation-induced intestinal injury
Kim et al. Cell Death and Disease
Synergistic actions of FGF2 and bone marrow transplantation mitigate radiation- induced intestinal injury
Byoung Hyuck Kim 0 1 2
Hee-Won Jung 1
Seok Hyun Seo 1
Hyemi Shin 4
Jeanny Kwon 3
Jae Myoung Suh 1 4
0 Division of Biological Warfare Preparedness and Response, Armed Forces Medical Research Institute , Daejeon , Republic of Korea
1 Graduate School of Medical Science and Engineering , KAIST, Daejeon , Republic of Korea
2 Department of Radiation Oncology, Seoul National University College of Medicine , Seoul , Republic of Korea
3 Department of Radiation Oncology, Chungnam National University College of Medicine , Daejeon , Republic of Korea
4 Biomedical Science and Engineering Interdisciplinary Program, KAIST , Daejeon , Republic of Korea
Unwanted radiological or nuclear exposure remains a public health risk for which effective therapeutic countermeasures are lacking. Here, we evaluated the efficacy of fibroblast growth factor-2 (FGF2) in treating radiationinduced gastrointestinal syndrome (RIGS) incurred by lethal whole-body irradiation (WBI) when administered in conjunction with bone marrow transplantation (BMT). In vitro experiments indicated FGF2 treatment increased proliferation, reduced apoptosis, and upregulated AKT-GSK3β/β-catenin signaling in irradiated IEC-6 cells. We next established and analyzed mice cohorts consisting of sham irradiation (Group Sh); 12 Gy WBI (Group A); WBI with BMT (Group B); WBI with FGF2 treatment (Group F); and WBI with BMT and FGF2 treatment (Group BF). At 2 weeks postirradiation, Group BF showed a dramatic increase in survival over all other groups. Intestinal epithelium of Group BF, but not Group B or F, showed augmented proliferation, decreased apoptosis, and preserved crypt numbers and morphology. Furthermore, Group BF maintained intestinal barrier function with minimal inflammatory disturbances in a manner comparable to Group Sh. In accordance, transcriptomic analyses showed significant upregulation of intestinal barrier and stem cell markers in Group BF relative to Groups A and B. Taken together, parenteral FGF2 synergizes with BMT to confer potent mitigation against RIGS.
type of radiation. First, hematopoietic syndrome can
occur at doses of 1 Gy or more but adequate infection
control, supportive care, and, more aggressively, bone
marrow transplantation (BMT) can help subjects to
regain hematopoietic function3,4. However, for subjects
exposed to radiation doses high enough to induce acute
RIGS, there are no effective treatments available
currently, resulting in 100% mortality in about 2 weeks5,6.
While BMT can readily support recovery from
hematopoietic syndrome, its role in resolving
radiationinduced gastrointestinal syndrome (RIGS) upon
highdose exposure remains controversial and of limited
efficacy7,8. Radioprotectors, such as amifostine, are
effective only when administered prior to radiation
exposure9. Therefore, the development of radiation
mitigators that can reduce toxicity after radiation
exposure are essential to the arsenal of medical
countermeasures for unexpected radiation incidents.
Intestinal stem cells located in crypts are required for
maintaining tissue homeostasis and regeneration upon
intestinal injury10–12. Previous studies demonstrate that
crypt stem cell apoptosis is a primary cause for
irreparable intestinal damage following radiation injury.
Several growth factors, such as fibroblast growth
factor2 (FGF2), keratinocyte growth factor, and insulin-like
growth factor, were shown to protect crypt stem cells
from apoptosis and increase their survival after
irradiation, albeit with limited efficacy, through
mechanisms yet to be clarified13–15. Notably, the FGF family of
secreted growth factors has been implicated in a broad
range of biological processes including the maintenance
and proliferation of stem cells, tissue remodeling, and
homeostasis16–20. FGF2 has relatively higher
thermostability in comparison to the closely related paralog
FGF1 and exerts target cell effects through binding and
activation of FGF receptor tyrosine kinases, typically in
a paracrine or autocrine fashion. Endogenous FGF2
expression was shown to be upregulated in the small
intestine after irradiation injury with suggested roles in
preserving intestinal stem cells21. However, detailed
physiological and molecular functions of endogenous
FGF2 in intestinal tissue remodeling or regeneration
after intestinal radiation damage remain unclear.
In clinical settings, the proposed dose range for a
BMT referral is limited to 7–10 Gy as exposures
exceeding 10 Gy negates any benefits of hematopoietic
rescue by BMT due to fulminant RIGS3,22. Herein, we
investigate whether FGF2 treatment alone or FGF2
treatment in conjunction with BMT, would counteract
RIGS resulting from high-dose radiation exposure.
Such approaches utilizing combined biologics may
potentially extend the upper dose range for BMT
referrals and lead to novel and effective treatment
strategies for ameliorating RIGS.
FGF2 improves cell viability and clonogenic potential after irradiation in vitro
We tested multiple members of the FGF family for the
potential to mitigate radiation damage in vitro using IEC-6
cells, which derive from rat small intestine. Among FGF
family members tested, only FGF2 treatment for 24 h
following irradiation significantly increased clonogenic cell
survival as compared to other treatments (Fig. 1a). In
addition to increased clonogenicity, cell viability also
increased by FGF2 treatment as compared to other
treatments (Fig. 1b). Next, we examined apoptotic signaling
pathways and found that FGF2 treatment decreased the
levels of Bax and cleaved Casp3. The decrease in apoptotic
signals was accompanied by an increase in signaling
through the AKT–GSK3β/β-catenin axis (Fig. 1c).
FGF2 synergizes with BMT to enhance survival of mice exposed to lethal WBI
To evaluate the in vivo effects of FGF2, either in the
absence or presence BMT, a total of five groups of mice
were established: sham irradiation (Group Sh); 12 Gy
wholebody irradiation (WBI) without any treatment (Group A);
WBI followed by syngeneic BMT (Group B); WBI followed
by parenteral FGF2 treatment (Group F); and WBI followed
by both syngeneic BMT and parenteral FGF2 treatment
(Group BF) (Fig. 2a). Following a lethal dose of 12 Gy WBI,
most mice from all groups died within 7 days; however,
Group BF showed a significant increase in survival
compared to all other experimental groups that received WBI
(p < 0.001, Fig. 2b). Group BF also showed a significant
increase in mean survival time of mice that died in the first
2 weeks after WBI (Fig. 2c). Body weight was measured daily
for 13 days as a surrogate index for gastrointestinal function
and general health of the mice. Irrespective of treatment
group, body weight continuously decreased in all mice that
died within the first 7 days after irradiation. Nevertheless, the
trend of body weight loss was significantly dampened in
Group BF relative to Group A (Fig. 2d). Among surviving
mice, body weight decreased comparably in Group BF and
Group B during the initial 5 days following irradiation,
however, the rate of recovery to normal body weight from
day 7 to 13 showed an increased trend for Group BF
compared to Group B (Fig. 2e).
To assess the degree of hematological toxicity after
irradiation, we performed complete blood count
examinations. Analyses of subjects at day 5 post-irradiation
demonstrated that severe leukopenia and mild anemia
were present in all groups other than Group Sh
(Supplementary Figure 1). Groups A and B did not
significantly differ in terms of the white blood cell (WBC)
count, hemoglobin level, and platelet count in peripheral
blood (Supplementary Figure 1). These results indicate
that transfused donor marrow had not undergone
complete engraftment at the time of analysis. However,
in Group BF, the administration of FGF2 did result in
significantly higher hemoglobin levels (10.4 ± 0.3 vs. 11.9
± 0.6, p = 0.008) and hematocrit values (36.0 ± 1.1 vs.
40.4 ± 2.8, p = 0.032) compared to Group B
(Supplementary Figure 1). There were no significant differences
in WBC and platelet count among all irradiated groups.
FGF2 synergizes with BMT to restore intestinal epithelium damage from RIGS
Since 12 Gy WBI resulted in the loss of crypts within 3
to 5 days after irradiation (Supplementary Figure 2), we
examined histologic characteristics and functional
permeability of intestines on days 3 and 5. In contrast to
intestines from Group Sh, which showed normal tissue
architecture, those from Group A, F, and B showed
massive infiltration of inflammatory cells, ulceration with
detachment of the mucosal layer, destruction of crypt/villi
structure, and lengthened basal lamina (Fig. 3a). In
striking contrast, Group BF showed sparing of intact
crypt/villi structure accompanied by a greatly attenuated
inflammatory response with levels comparable to that of
Group Sh. Quantitative analyses of histologic parameters
showed that Groups A, F, and B, in comparison to Group
Sh, had decreased crypt counts per circumference (5.6,
8.2, 8.3 vs. 129.7), shorter villi heights (160.4, 176.0, 184.4
vs. 264.2 μm), and longer basal lamina (77.6, 79.2, 71.1 vs.
44.6 μm). In addition, Group BF was the least affected in
terms of crypt counts while villi height and basal lamina
length were comparable to those of Group Sh (Fig. 3b).
Cytokine profiles revealed pro-inflammatory cytokines
IL1α and IL-6 from intestinal tissue lysates were reduced in
FGF2-treated groups at day 3 post-irradiation (Fig. 3c). At
12 Gy WBI, intestinal permeability is known to be
significantly increased due to compromised barrier function;
however, compared to other treatments, the increase in
intestinal permeability was remarkably attenuated in
Group BF and nearly indistinguishable to Group Sh
(Group BF vs. B, p = 0.038; Group BF vs. A, p = 0.027;
Fig. 3d). These changes in Group BF were accompanied
by enhanced expression of Cldn15, a major component of
epithelial tight junction complexes (Fig. 3e). Interestingly,
the mitigative effects against RIGS were not observed
when FGF2 treatment was performed ex vivo to isolated
donor bone marrow prior to transplantation (data not
FGF2 treatment leads to enhanced proliferation of crypt stem cells in damaged intestine
We next examined the functional integrity of
proliferating crypt structures in the small intestine, which is
critical for normal homeostasis of intestinal tissue. The
fraction of proliferating crypts, as determined by
immunohistochemistry of bromodeoxyuridine
(BrdU)incorporated nuclei, increased significantly in Group BF
compared to other groups (Fig. 4a, b). Therefore, the
improved intestinal tissue integrity in Group BF (Fig. 3a)
may be attributed to increased functionality of crypt stem
cells that are required to replenish damaged intestinal
epithelium following radiation injury. We further
evaluated this notion by analyzing the pool of proliferating
precursors with Ki-67 immunofluorescent staining.
Similar to BrdU incorporation, Group BF intestines
showed an increase in Ki-67-positive cells within crypt
structures (Fig. 4c, d). Terminal deoxynucleotidyl
transferase dUTP nick end labeling (TUNEL) staining also
showed that crypt cell apoptosis in Group BF at day 3 was
significantly reduced (Fig. 4c, e). Interestingly, protein
levels of phospho-GSK3β (p-GSK3β), the inactive form of
GSK3β which acts as negative regulator of the
Wnt/βcatenin axis, and the intestinal stem cell marker Lgr5 were
both increased in Group BF compared to other groups
(Fig. 4f). In agreement with these results, Lgr5 mRNA
expression was elevated in Group BF (Fig. 4g). Taken
together, administration of FGF2 in conjunction with
BMT enhances crypt stem cell proliferation after
irradiation injury, possibly through inhibiting GSK3β activity
and increasing Lgr5 levels.
FGF2 protects against radiation-induced alterations of
inflammatory serum cytokines
Irradiation induced a significant increase in serum
cytokines IL-6, TNFα, and IL-10, consistent with expected
radiation-induced systemic inflammation. Serum cytokine
analyses on day 3 revealed an increase of
antiinflammatory cytokine IL-10 in Group BF, whereas
proinflammatory TNFα decreased in FGF2-treated groups
(Fig. 5a, b). IL-4 and IL-6 levels were not significantly
different among treatment groups that received
irradiation on both day 3 and day 5 (Fig. 5c, d). Other cytokines,
such as IL-1α, IL-1β, IL-12p70, IL-2, and IFNγ, remained
below the detection limit of the assay (data not shown).
FGF2 treatment reveals transcriptomic signatures of
reduced inflammation and restoration of intestinal tissue
To characterize molecular changes elicited by FGF2
treatment, we examined gene expression changes in
intestinal tissues from three different treatment groups
through mRNA-sequencing (RNAseq) analyses. Global
gene expression analyses revealed that Groups A and B
clustered together while Group BF showed a distinct
pattern (Fig. 6a). We found 26, 2799, and 2960 genes with
significant changes in expression levels (>twofold) in
Groups A/B, A/BF, and B/BF, respectively (Fig. 6b).
Interestingly, in relationship to Groups A and B, Group
BF showed higher expression levels of genes involved in
intestinal tight junction protein (Cldn15, Epcam, and
Vil1), intestinal stem cell maintenance (Lgr5 and Olfm4),
GSK3β/β-catenin signaling (Wnt3, Gskip, and Ctnnb1),
and lipid transport (Apoa1 and Fabp2) (Fig. 6c).
Conversely, Group BF showed lower expression levels for
genes encoding pro-inflammatory factors (Il1α, Il6, Ccl2,
and Cxcl13) and extracellular matrix degradation-related
transcripts (Mmp2, Mmp3, Ctgf, and Timp2). RT-qPCR
analyses for representative genes showed similar
expression changes obtained from RNAseq analyses (Fig. 6d).
Therefore, transcriptomic analyses indicated a shift in
global gene expression towards an enhanced recovery of
damaged intestinal tissue by FGF2 treatment when
provided in conjunction with BMT.
The underlying mechanisms of radiation-induced
gastrointestinal injury are complex. Although BMT can
rescue hematopoietic failure after acute high-dose radiation
exposure, efficacious treatment strategies to ameliorate
RIGS are yet to be established. A number of different
growth factors have been implicated in the response to
radiation injury; to this end, we sought to further
investigate the biological role and therapeutic utility of FGF2
concerning RIGS. Survival analyses following high-dose
irradiation revealed a role for FGF2 as a radiomitigator in
mice receiving BMT. Analysis of lethally irradiated mice
that received BMT followed by FGF2 treatment, Group
BF, showed enhanced proliferation of intestinal crypt cells
accompanied by histologic and functional recovery from
intestinal radiation damage. On the other hand, intestinal
epithelium of Group B displayed insignificant changes
compared to Group A, indicating that BMT by itself
cannot account for the improved intestinal tissue
dynamics and functional recovery observed in Group BF.
Therefore, FGF2 treatment ameliorates intestinal damage
in RIGS through a newly identified synergism with BMT
and thus suggests a novel treatment strategy to mitigate
the effects of RIGS.
Numerous research efforts have been aimed towards the
development of effective therapies to improve recovery
from RIGS. Previous studies suggested that growth
factors, including members of the FGF family, can have
broad effects influencing the survival, proliferation, and
differentiation of intestinal cells13,21,23. Endogenous FGF2
mRNA and protein in the small intestine increases 12 h
after high-dose irradiation, reaching peak levels at
48–120 h after irradiation21. Radiation exposure induces
expression of FGF2 in several cell lines in vitro as well24,25.
These results suggest that endogenous FGF2 may play a
role in the protective response to radiation exposure.
Some studies show that exogenous FGF2 alone failed to
increase crypt survival when administered after
irradiation21,26, whereas others suggested that FGF2 as an
independent treatment can improve crypt survival and
restore gastrointestinal function, albeit with limited
effect9,27. Others have reported that FGF2 injection a total
of three times before and after radiation exposure (30 min
before, 5 min before, and 30 min after) attenuated
radiation-induced intestinal stem cell apoptosis through
the Akt-p53 signaling pathway28. The aforementioned
studies were performed under different experimental
conditions, which may account for the varying outcomes.
Together with our results, further investigations are
warranted to determine the optimal course of FGF2
treatment for mitigating damage due to RIGS.
FGF2 has also been implicated in the attenuation of
radiation-induced increase in intestinal permeability,
which is considered an important cause of sepsis after
WBI29,30. Results from our transcriptomic studies (Fig. 6c)
indicate FGF2 may enhance barrier function through
increased expression of Cldn15, a tight junction protein
that also has a role in facilitating glucose absorption31. In
addition, relative to Group A and B, Group BF showed
decreased expression of many pro-inflammatory factors
and MMPs in the intestine as well as a reduction in local
IL-1α, IL-6, and serum TNFα cytokine levels. Therefore,
FGF2 may exert effects against RIGS by enhancing
intestinal barrier function and consequent resolution of
both local and systemic inflammation.
Interestingly, FGF2 exerted radiomitigative properties
in vivo, Group BF, as well as in vitro, FGF2-treated IEC-6
intestinal cells, as both showed decreased
radiationinduced apoptosis. Analyses of intracellular signaling
pathways suggested that the protective effects of FGF2
might work through the AKT–GSK3β axis, which in turn
regulates nuclear β-catenin accumulation. GSK3β has
been shown to induce apoptosis in various conditions,
including DNA damage, while specific inhibitors of
GSK3β are able to protect against radiation-induced
damage32,33. On the other hand, GSK3β inhibition
enhances β-catenin signaling by preventing its
degradation, which in turn can activate a transcriptional program
for the regeneration of the damaged intestinal
epithelium34,35. Analyses of intestinal tissue from Group BF
showed GSK3β inhibition and Gskip/Ctnnb1 upregulation
suggesting that the AKT–GSK3β/β-catenin signaling axis
may be involved in the radiomitigative effects of FGF2.
Further studies are required to delineate the molecular
mechanisms underlying the synergistic actions of FGF2
and BMT in mitigating radiation injury.
A number of reports show that mesenchymal stem cells
(MSCs) can facilitate the repair of tissue damage including
intestinal injury and that MSCs have a potential for
replacing injured intestinal stem cells36–38. In similar
fashion, transplanted bone marrow-derived MSCs can
promote engraftment of stem cells at the site of intestinal
damage incurred by WBI, although the engraftment rates
were quite low39,40. Other studies found that transplanted
bone marrow cells could be detected in the recipient’s
intestines as early as 2 days after transplantation41,42. In
the present study, significant attenuation of
radiationinduced intestinal damage occurred only when FGF2 was
administered in combination with BMT. This suggests a
cell type(s) and/or secreted factor(s) in the transfused
bone marrow interacts to synergize with FGF2 to achieve
the full range of effects observed in Group BF.
RIGS causes death at much earlier time points than
death resulting from hematopoietic syndrome43. Our
results suggest that FGF2 treatment, when combined with
BMT, represents a viable intervention strategy for
targeting the critical timeframe after radiation exposure but
before RIGS-induced death occurs. Although BMT would
help recover intestinal function once fully engrafted at
several weeks post-irradiation, we speculate that FGF2
might enhance the incorporation of bone marrow-derived
stem cells at an earlier time point during intestinal
remodeling to offset the acute early stage mortality due to
RIGS8,44. FGF2 has versatile functions in reducing local
and systemic inflammation, supporting stem cell
migration, engraftment, and proliferation in the intestine, all of
which may contribute to FGF2-mediated mechanisms of
In summary, our current studies indicate that FGF2
combined with BMT can mitigate intestinal injury after
acute high-dose irradiation. In treating radiation victims,
the recommended therapeutic window for BMT is rather
narrow (7–10 Gy) due to the limited efficacy of BMT at
radiation doses exceeding 10 Gy. Our studies suggest that
combining FGF2 treatment together with BMT could
widen this window if translatable to humans. Further
experiments are required to clarify the underlying
mechanisms, optimal dose, treatment sequence, and
potential adverse effects of combined therapy of FGF2 and
BMT in RIGS. Nevertheless, given the paucity of available
therapeutic strategies for RIGS, our findings point to a
novel strategy to mitigate intestinal radiation injury and
offer new insights into the pharmacologic actions of
Materials and methods
Cell lines, recombinant proteins, animals, and irradiation
Rat intestinal epithelial cell line IEC-6 was obtained
from the Korean Cell Line Bank (Seoul, South Korea) and
maintained in DMEM with 10% FBS and 1%
penicillin–streptomycin. Human recombinant proteins
were purchased from Prospec (East Brunswick, NJ, USA).
A total of 8–9-week-old male C57BL/6 J mice (Central
Lab Animal Inc., Seoul, South Korea) were housed at
room temperature under 12-h light–dark cycle and
allowed access to water and chow ad libitum. All animal
experiments were approved by the Institutional Animal
Care and Use Committee at the Korea Advanced Institute
of Science and Technology (KA2015-35). Mice and cells
were irradiated at a dose rate of 2.16 Gy/min using a
Cs137 irradiator (Gammacell 3000 Elan, Best Theratronics,
Ottawa, ON, Canada). Irradiation was performed without
anesthesia on mice held in a 50-ml conical tube on a
rotating turntable. Dosimetric quality assurance was
performed using nanoDots (Al2O3:C) optically stimulated
luminescence dosimeters (Landauer, Glenwood, IL, USA),
which were read using a MicroStar OSL reader (Landauer,
Glenwood, IL, USA).
Clonogenic survival and CCK-8 viability assays
Clonogenic survival assays were performed as
previously described47. Briefly, cells were plated immediately
after irradiation at the same density into wells of a 6-well
culture plate for each treatment group and cultured for
10–14 days to allow for colony formation. Colonies were
fixed with methanol and stained with 0.5% crystal violet
solution (Sigma-Aldrich, St. Louis, MO, USA). Colonies
containing 50 or more cells were counted using ImageJ
software (version 1.50, NIH, Bethesda, MD, USA), and the
surviving fraction was calculated. For CCK-8 cell viability
assays, cells were seeded in 96-well plates and treated with
FGFs (10 ng/ml) for 48 h after irradiation. Viable cells
were quantified by 450 nm absorbance measurement
acquired 2 h after treatment with CCK-8 reagent (Dojindo
Molecular Technologies, Gaithersburg, MD, USA)
Mouse experimental groups and bone marrow transplantation
We performed BMT in mice 4–6 h after irradiation to
evaluate the progression of gastrointestinal damage and
subsequent recovery while minimizing the effects of
hematopoietic system damage. Starting from 7 days
before irradiation, the recipient mice were given acidified
(pH 2.7) antibiotic water containing 1.1 mg/ml neomycin
sulfate (N1876, Sigma-Aldrich, St. Louis, MO, USA) and
106 U/l polymyxin B (P-1004, Sigma-Aldrich, St. Louis,
MO, USA). The mice were kept on antibiotic water for
2 weeks after irradiation. The procedure of generating
bone marrow-transplanted mice was based on previously
published studies, with minor modifications44,48.
Recipient mice received at least 5 × 106 bone marrow cells via
tail vein injection. For Group F and BF, mice were injected
intraperitoneally with 1 mg/kg human recombinant FGF2
in three doses delivered 1, 3, and 5 days after irradiation
while other groups were injected with vehicle (PBS, Gibco,
Grand Island, NY, USA) at identical time points.
Peripheral blood count
Whole blood was collected in EDTA-coated tubes
(Greiner Bio-One, KremsmÜnster, Austria) from mice at
the time of sacrifice. Peripheral blood cell counts were
obtained using a veterinary hemocytometer XN-9000
(Sysmex Co., Kobe, Japan), according to the
Tissue harvest and histological analysis
Upon killing of mice, small intestinal tissues were quickly
harvested from the interval spanning 5 cm distal to the
gastroduodenal junction to 5 cm proximal to the ileocecal
valve. Harvested intestinal tissues were trisected and the
resulting segments were immediately processed as follows:
fixation in neutral buffered formalin for histological
analysis, immersion in RNAlater (Thermo Fisher Scientific,
Wilmington, DE, USA) for RNA extraction, and snap
freezing in liquid nitrogen for tissue lysate preparation. For
histology samples, formalin-fixed tissues were embedded in
paraffin and processed for routine histological examination.
Tissue sections were deparaffinized, rehydrated, and used
for immunohistochemistry and hematoxylin and eosin
staining. For quantitative comparison, the proliferating
crypts in the circumference of jejunal transverse
crosssections were counted. Proliferating crypts were defined as
containing five or more adjacent chromophilic non-Paneth
cells and a lumen. At least 10 circumferences were
evaluated per mouse. To analyze morphological changes, villus
height and basal lamina length of 10 enterocytes from the
middle of randomly selected villi were measured. The
lengths of the 10 longest villi in each slide were used for
analysis in each sample.
Immunohistochemistry and immunofluorescence staining
Proliferating cells in the jejunum were labeled with BrdU
(ab142567, Abcam, London, UK), which was injected
intraperitoneally into mice at 10 mg/kg 2 h before
sampling. Briefly, deparaffinized and rehydrated sections were
submerged in pH 6.0 sodium citrate buffer and heated to
95 °C for 15–30 min for antigen retrieval. Next, the sections
were incubated with peroxidase-blocking solution (S2023,
Dako, Carpinteria, CA, USA), followed by incubation with
1% normal goat serum for 30 min at room temperature to
block nonspecific binding. Sections were washed with PBS
three times and incubated with a primary antibody against
BrdU (#5292, Cell Signaling Technology, Danvers, MA,
USA) for 2 h at room temperature, followed by 1 h
incubation with a species-specific Envision+ kit (Dako,
Carpinteria, CA, USA), according to the manufacturer’s
instructions. Liquid DAB + (K3467, Dako, Carpinteria, CA,
USA) was used for visualizing. For immunofluorescence
staining, anti-Ki-67 antibody (#12202, Cell Signaling
Technology, Danvers, MA, USA) and DAPI (D9542,
Sigma-Aldrich, St. Louis, MO, USA) were used. For
TUNEL staining, In situ Cell Death Detection Kit
(#1168481-7910, Roche, Indianapolis, IN, USA) was used
according to the manufacturer’s instructions.
Total RNA was obtained from tissues using TRIzol
(Invitrogen, Carlsbad, CA, USA) according to the
manufacturer’s protocol. RNA concentrations were measured
with a NanoDrop spectrophotometer (Thermo Fisher
Scientific). For RNA sequencing, RNA quality was
assessed on an Agilent 2100 Bioanalyzer (Agilent
Technologies, Palo Alto, CA, USA). RNA samples with RIN > 7.0,
A260/280 > 1.5, A260/230 > 1.0, and rRNA ratio > 1.0
were used for subsequent analyses.
Quantitative reverse transcriptase-PCR (qRT-PCR) analysis
After RNA quantification, 1 μg of total RNA was used to
generate complementary DNA using HelixCript™ Easy
cDNA Synthesis kit (Nanohelix, Daejeon, South Korea).
To assess individual gene expression, real-time PCR was
performed with Nanohelix premier qPCR premix
(Nanohelix, Daejeon, South Korea) and ViiA 7 Real-Time
PCR system (Applied Biosystems, Foster city, CA, USA).
Relative quantification was based on the ΔΔCt method,
and RPLP0 gene was used as a reference control. PCR
primers were designed using NCBI Primer-BLAST.
RNA sequencing libraries were prepared using the
Illumina TruSeq Sample Preparation Kit and sequenced
on a HiSeq 2000 system (Illumina, San Diego, CA, USA),
according to the manufacturer’s instructions. Sequencing
was performed at a multiplexing level sufficient to
generate >80 million reads per sample. The reads were
aligned to the appropriate reference mouse genome
(UCSC mm10) using TopHat. Aligned reads were
converted to fragments per kilobase of transcript per million
(FPKM) mapped reads values, calculated by the Cufflinks.
After data preprocessing and QC processing,
normalization was carried out based on the value of log (FPKM
+ 1) transformation49. When comparing two groups, we
defined differentially expressed genes as those having a
minimum of two-fold change in the expression of pooled
samples. After analysis of RNA sequencing results, genes
involved in intestinal homeostasis and tissue
inflammation were chosen for further validation using qRT-PCR.
Differential transcriptional profiles were presented as a
form of heatmap using PermuteMatrix software50.
Tissue and serum cytokine assays
Cytokines were analyzed using the Luminex
Performance Assay kit (R&D systems, Minneapolis, MN, USA),
according to manufacturer’s instructions. Briefly, duplicate
serum samples or tissue lysates were added to a 96-well
filter plate preconfigured with a panel of anti-cytokine
antibodies covalently linked to unique polystyrene beads.
Individual cytokines were identified and classified by their
bead color using red laser excitation. Classified cytokines
were quantified as a form of mean fluorescent intensity via
green laser excitation. Standard curves for each cytokine
were generated using the reference cytokine
concentrations supplied by the manufacturer.
Intestinal permeability assay
Serum concentration of FITC–dextran (FD4,
SigmaAldrich, St. Louis, MO, USA) was used a measure of
intestinal permeability as previously described51. Briefly,
FITC–dextran was dissolved in PBS at a concentration of
100 mg/ml and was administered through oral gavage of
44 mg/100 g body weight. After 4 h, serum was collected by
cardiac puncture. Fluorescence intensity of diluted sera was
measured on a TriStar2 S LB 942 Modular
Monochromator Multimode Reader (Berthold Technologies, Bad
Wildbad, Germany). Serum from mice not administered
with FITC–dextran was used to determine the background.
Cells and tissues were lysed in modified RIPA buffer (50
mM Tris-HCl, 150 mM NaCl, 1% NP-40, and 0.5% sodium
deoxycholate) and homogenized with FastPrep-24 (MP
biomedicals, Orangeburg, NY, USA). Tissue homogenates were
sonicated and insoluble debris was cleared by centrifugation.
Protein concentrations of cleared lysates were quantified with
Pierce BCA Protein Assay Kit (Thermo Fisher Scientific,
Wilmington, DE, USA). Lysates were loaded on
SDSpolyacrylamide gels and transferred onto polyvinylidene
fluoride membranes. The membranes were blocked for 30
min at room temperature with 3% BSA in TBST and were
then incubated with antibodies against p-AKT (Thr308,
#9275, Cell Signaling Technology, Danvers, MA, USA),
pGSK3β (Ser9, #9331, Cell Signaling Technology, Danvers,
MA, USA), p-β-catenin (PY489, Developmental Studies
Hybridoma Bank, Iowa City, IA, USA), total β-catenin
(ab32572, Abcam, London, UK), p-ERK (Thr202/Thr204,
#9101, Cell Signaling Technology, Danvers, MA, USA),
cleaved Casp3 (#9664, Cell Signaling Technology, Danvers,
MA, USA), LGR5 (UMAB210, Origene, Rockville, MD,
USA), α-Tubulin (sc-23948, Santa Cruz Biotechnology, Santa
Cruz, CA, USA), Bax (sc-20067, Santa Cruz Biotechnology,
Santa Cruz, CA, USA), and LC3B (#2775, Cell Signaling
Technology, Danvers, MA, USA). Following incubation with
HRP-conjugated primary host-specific secondary antibody
(Cell Signaling Technology, Danvers, MA, USA), blots were
visualized on a ChemiDoc™ XRS + System (BioRad,
Hercules, CA, USA). α-Tubulin was used as the loading control.
All values were expressed as the mean ± standard error of
the mean (SEM), unless otherwise specified. The mean
values between experimental groups were analyzed using
Student’s t-test. Mouse survival curves were analyzed using
Wilcoxon test. We considered p values < 0.05 as statistically
significant. Data analysis was performed using GraphPad
Prism version 6 (GraphPad Software, San Diego, CA, USA).
We gratefully acknowledge Ju Eun Kim, Jung Hae Sunwoo, and Hee-Saeng
Jung for expert technical support and Jong Min Park in the Seoul National
University Hospital for dosimetric quality assurance. This work was funded by
Armed Forces Medical Research Institute (AFMRI) Republic of Korea Army
institutional R&D grant 2016-AFMRI-02/2017-AFMRI-02 and National Research
Foundation of Korea (NRF) grants NRF-2016R1A2B1011083,
NRF2016M3A9B6902871. H.W.J was supported by the National Research
Foundation of Korea (NRF) Global Ph.D. Fellowship Program
Conflict of interest
The authors declare that they have no conflict of interest.
Springer Nature remains neutral with regard to jurisdictional claims in
published maps and institutional affiliations.
Supplementary Information accompanies this paper at https://doi.org/
Kim et al. Cell Death and Disease
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