A lymphatic route for a hyperbranched heteroglycan from Radix Astragali to trigger immune responses after oral dosing

Carbohydrate Polymers 292 (2022) 119653

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Carbohydrate Polymers
journal homepage: www.elsevier.com/locate/carbpol

A lymphatic route for a hyperbranched heteroglycan from Radix Astragali
to trigger immune responses after oral dosing
Quanwei Zhang a, 1, Lifeng Li a, 1, Shuang Hao b, Man Liu a, Chuying Huo a, Jianjun Wu a,
Hongbing Liu a, Wanrong Bao a, Hongming Zheng a, Zhipeng Li a, Huiyuan Cheng a,
Hauyee Fung a, Tinlong Wong a, Pingchung Leung c, Shunchun Wang d, Ting Li e, Ge Zhang a,
Min Li a, Zhongzhen Zhao a, Wei Jia a, Zhaoxiang Bian a, Timothy Mitchison f, Jingchao Zhang b, *,
Aiping Lyu a, *, Quanbin Han a, *, Handong Sun g
a

School of Chinese Medicine, Hong Kong Baptist University, Hong Kong 999077, China
The First Affiliated Hospital, Zhengzhou University, Zhengzhou 450000, China
c
State Key Laboratory of Research on Bioactivities and Clinical Applications of Medicinal Plants, The Chinese University of Hong Kong, Hong Kong 999077, China
d
Institute of Chinese Materia Medica, Shanghai University of Traditional Chinese Medicine, Shanghai 201203, China
e
State Key Laboratory for Quality Research in Chinese Medicine, Macau University of Science and Technology, Macau 999078, China
f
Laboratory of Systems Pharmacology, Department of Systems Biology, Harvard Medical School, Boston 02115, United States
g
State Key Laboratory of Phytochemistry and Plant Resources in West China, Kunming Institute of Botany, Chinese Academy of Sciences, Kunming, China
b

A R T I C L E I N F O

A B S T R A C T

Keywords:
Radix Astragali
Polysaccharide
Intact
Targeting route
Antitumor immune responses

Gut barrier makes a huge research gap between in vivo and in vitro studies of orally bioactive polysaccharides:
whether/how they contact the related cells in vivo. A hyperbranched heteroglycan RAP from Radix Astragali,
exerting antitumor and immunomodulatory effects in vitro and in vivo, is right an example. Here, we determined
first that RAP's antitumor activity is immune-dependent. Being undegraded and non-absorbing, RAP quickly
entered Peyer's patches (PPs) in 1 h where it directly targeted follicle dendritic cells and initiated antitumor
immune responses. RAP was further delivered to mesenteric lymph node, bone marrow, and tumor. By contrast,
the control Dendrobium officinale polysaccharide did not enter PPs. These findings revealed a blood/microbiotaindependent and selective lymphatic route for orally administrated RAP to directly contact immune cells and
trigger antitumor immune responses. This route bridges the research gap between the in vitro and in vivo studies
and might apply to many other bioactive polysaccharides.

1. Introduction
The gut wall barrier to macromolecules remains an unsolved chal­
lenge for developing orally-delivered macromolecular therapeutics
(Scaldaferri et al., 2012). Many natural polysaccharides, being safe and
effective, show great potential to be medicines (Mohammed et al., 2021;
Yu et al., 2018). However, few of them are developed to medications
because their efficacy on the human body is often doubted due to their
poor bioavailability. Inspiration is found from some polysaccharides
that could quickly affect the immune system after oral dosing (Jiang
et al., 2010; Schepetkin & Quinn, 2006; Yu et al., 2018). It is hypothe­

sized that there might be a blood-independent route for these

polysaccharides to work in body. Understanding how polysaccharides
access the immune system may open new doors in developing oral de­
livery of polysaccharide-based vaccines or drugs.
Here, we take Radix Astragali polysaccharide RAP as a case study to
test the hypothesis. Radix Astragali is the most frequently used ‘Qi’ tonic
herb medicine in tumor therapy with Traditional Chinese Medicine. In
our previous study, a hyperbranched heteroglycan RAP (1334 kDa) was
purified from this herb medicine (Yin et al., 2012). Its chemical structure
was characterized by monosaccharide composition, partial acid hydro­
lysis, methylation analysis, GC-MS, NMR spectra, SEM and AFM mi­
croscopy. The backbone of RAP consists of 1,2,4-linked Rhap, α-1,4linked Glcp, α-1,4-linked GalAp6Me, β-1,3,6-linked Galp, with branched

* Corresponding authors.
E-mail addresses: (J. Zhang), (A. Lyu), (Q. Han).
1
These authors contributed equally.
/>Received 24 January 2022; Received in revised form 5 May 2022; Accepted 22 May 2022
Available online 27 May 2022
0144-8617/© 2022 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY license ( />

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Carbohydrate Polymers 292 (2022) 119653

at O-4 of the 1,2,4-linked Rhap and O-3 or O-4 of β-1,3,6-linked Galp.
The side chains are mainly α-T-Araf and α-1,5-linked Araf with O-3 as
branching points, having trace Glc and Gal. The terminal residues are Tlinked Araf, T-linked Glcp, and T-linked Galp. As a major component, it
looks similar to the reported heteroglycans from the same herb medicine

(Zheng et al., 2020).
RAP is a typical example of the poor bioavailability concern. Orally
administrated RAP exhibited promising antitumor activities in vivo,
particularly the synergism in combination with Taxol in increasing
survival rate of tumor bearing mice, and protection of the bone marrow
of cyclophosphamide-treated mice (Bao et al., 2019; Bao et al., 2021).
The mechanism investigations in vitro have revealed its inducing effects
on macrophages via TLR4 signaling pathway, inducing phenotype po­
larization to antitumor M1 via Notch signaling pathway, and protection
on mice/human hematopoietic stem cells (HSCs) via regulation of FOS
expression (Bao et al., 2021; Wei et al., 2016; Wei et al., 2019). How­
ever, there is a huge gap between these in vivo and in vitro activities:
whether/how the poorly bioavailable polysaccharide contact immune
cells after oral administration (Barclay et al., 2019).
Interestingly, we found that orally administrated RAP might be able
to enter the bone marrow to protect chemotherapy-induced myelosup­
pression (Bao et al., 2021). Our preliminary data further indicated that
RAP could quickly induce immune responses in the Peyer's Patches (PPs)
of the small intestine in 1 h, consistent with what we observed on
macrophages in vitro (Wei et al., 2016). Therefore, a reasonable hy­
pothesis is that intact RAP may enter PPs and directly contact immune
cells to trigger immune responses.
As the immune sensors in the small intestine, PPs, playing an
essential role in linking intestinal antigens and the host mucosal immune
responses (Jung et al., 2010), may provide a possible route. Actually,
PPs have been mentioned as a gateway for glucans, but the detection
solely relied on the fluorescence or radioactivity signals and was poorly
validated (De Jesus et al., 2014; Sakai et al., 2019). There are still a few
doubts, including 1) whether the observed signal was only the fluores­
cent/radioactivity flag rather than the polymer; 2) whether poly­

saccharides stayed intact carbohydrate polymer in the PPs; 3) what is
the direct target of intact polysaccharide in PPs; 4) whether PPs is
accessible to all polysaccharides (Hashimoto et al., 1991; Pedro et al.,
2021; Rice et al., 2005; Smet et al., 2013; Vetvicka et al., 2007; Xie et al.,
2016; Zheng et al., 2022). The gateway via PPs for intact poly­
saccharides needs to be verified.
In this study, we compared RAP's effects on tumor growth between
normal and nude mice first and confirmed that its antitumor effects are
immune system-dependent. We further tracked orally dosed RAP in the
gastrointestinal tract using a series of methods including carbohydrate
testing, fluorescence imaging, HPLC-FLD, and flow cytometry. The re­
sults indicated that RAP, as an undegraded carbohydrate polymer,
quickly entered the PPs to specifically activate DCs and trigger immune
responses. Confocal micrographs further revealed that DCs might sub­
sequently transport RAP from PPs to mesenteric lymph nodes (MLN),
bone marrow, and even tumor tissue. This interesting delivery route for
intact RAP entering PPs must be selective as another control poly­
saccharide DOP from Dendrobium officinale could not take the same
route. These findings provide convincing evidence of an efficient
lymphatic route for intact RAP to directly trigger immune responses
after oral administration. This interesting route might be also applicable
to many other immunomodulatory carbohydrate polymers.

Diego, CA, USA). Phenol‑sulfuric acid, 4′ ,6-diamidino-2-phenylindole
DAPI, fluorescein isothiocyanate isomer I FITC, methyl sulphoxide, and
other related chemical reagents were all purchased from Sigma-Aldrich
Corp. (St. Louis, MO, USA). High glucose Dulbecco Modified Eagle
Medium (DMEM), heated-inactivated fetal bovine serum (FBS), GMCSF, and IL-4 were bought from the Thermo Fisher Scientific (Cleve­
land, OH, USA).
2.2. Mice and cell

BALB/C and nude mice were purchased from the Chinese University
of Hong Kong. Five- to eight-week-old mice were used in this study. The
animals were provided with a standard pellet diet and purified water
and maintained under controlled temperature and humidity conditions,
with 12 h light/dark cycles. All animal experiments followed the Ani­
mals Ordinance guidelines, Department of Health, Hong Kong SAR ((1665) in DH/HA&P/8/2/6, (19-151) in DH/HT&A/8/2/6).
Macrophage RAW264.7 cells were bought from American Type
Culture Collection (ATCC) and cultured in high glucose DMEM with
10% FBS in a humidified incubator at 37 ◦ C under an atmosphere of 5%
CO2.
2.3. Preparation of polysaccharides
RAP was prepared from the water extract of the dried roots of
Astragalus membranaceus and stored at 4 ◦ C (Yin et al., 2012). In our
previous work, its structure was elucidated by monosaccharide
composition, partial acid hydrolysis, and methylation analysis, and
further confirmed by FT-IR, GC–MS, and 1H and 13C NMR spectra, SEM
and AFM microscopy (Yin et al., 2012). Before using the polysaccharide
RAP, the average molecular weight, purity, the 1H NMR spectrum, 13C
NMR spectrum, and monosaccharides composition of RAP was detected.
RAP was labeled with fluorescein isothiocyanate isomer I (FITC) as re­
ported (Li et al., 2019). Briefly, RAP (1.0 g) was dissolved in methyl
sulphoxide (8 mL) containing a few drops of pyridine. FITC (80 mg) was
added to the RAP solution, followed by dibutyltin dilaurate (16 μL). The
mixture was heated for 2 h at 95 ◦ C. After precipitation in ethanol (90%
v/v) to collect the precipitate and remove the free dye, the FITC-RAP
was re-dissolved in water and purified by molecular sieve (3 kDa cutoff). Using a procedure reported previously, control polysaccharide
DOP was prepared from Dendrobium officinale and similarly labeled with
FITC (Li et al., 2019).
2.4. 4T1 breast tumor mouse model and treatment
First of all, we optimized the dosages (50, 100, and 200 mg/kg) by

testing RAP-induced immune responses in PPs, and further evaluated
the selected dosage for RAP's suppression against 4T1 breast tumor in
BALB/c female mice, with cisplatin (4 mg/kg) as the positive control.
Five-week-old BALB/c female and nude female mice were used for
the tumor model (10 mice/group). Before establishing the tumor model,
mice were pre-treated with 100 mg/kg RAP for seven days. In accor­
dance with a previous study (Pulaski & Ostrand-Rosenberg, 2000), the
implantation of 4 T1 cells was performed on the seventh day. Briefly,
4T1 cells (2 × 104/mouse) were implanted with a 27-G needle. RAP
treatment continued until the last day of sacrificing mice. When tumors
begin to develop, a vernier caliper was used to measure tumor diameters
and calculate tumor volume. After three weeks of tumor growth, animals
were sacrificed, and organ samples were harvested for investigation,
including tumor weight were weighted, and immune cells and cytokines
were detected in the tumor and immune system.

2. Materials and methods
2.1. Materials
Flow cytometry antibodies were bought from BioLegend (San Diego,
CA, USA). CD11c primary antibody and goat anti-rat IgG antibody
conjugated with Alexa Fluor 568 were purchased from Abcam (Cam­
bridge, UK). All mouse ELISA kits were purchased from eBioscience (San

2.5. Carbohydrate detection of intestinal contents by phenol‑sulfuric acid
method
The intestinal contents (i.e., stomach, the small intestine, and the
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Carbohydrate Polymers 292 (2022) 119653

large intestine) of BALB/C mice that had received oral doses of RAP (10
mg/mouse) were collected at 1, 2, 3, 4, 5 h after oral administration (n
= 4 each group). Similarly, the intestinal contents of mice that had not
received RAP were collected as control. Carbohydrates in the samples
were detected by the phenol‑sulfuric acid method (Masuko et al., 2005).
In detail, samples were homogenized and centrifuged at 15,000 rpm for
10 min. 50 μL of each supernatant was injected in a well of 96-well
microplate to which 150 μL of concentrated sulfuric acid was added
and mixed rapidly. 30 μL of 5% phenol in water was then added into the
mixed solution, and it was incubated for 5 min at 90 ◦ C in a static water
bath. The plate was then cooled to room temperature and wiped dry for
detection at A490 nm by a microplate reader. Glucose was used as a
reference standard to establish standard curves. The experiments were
repeated three times.

then homogenized and centrifuged at 15,000 rpm for 10 min to obtain
the supernatant for cytokine production assessment using ELISA kits.
According to the manufacturer's instructions, cytokines interleukin 6
(IL-6), IL-12, tumor necrosis factor-alpha (TNF-α), transforming growth
factor-beta (TGF-β), monocyte chemotactic protein-1 (MCP-1), macro­
phage colony-stimulating factor (M-CSF), and interferon-gamma (IFN-γ)
were determined using ELISA kits.
2.10. Immunofluorescence staining and confocal microscopy
Frozen sections of PPs from the ligated loop assay were washed three
times with PBS and blocked with 5% normal goat serum in PBS for 1 h.
Sections were incubated with anti-mouse CD11c antibody overnight at
4 ◦ C. PPs sections were washed three times with PBS and then treated

with Alexa Fluor 568 secondary goat anti-rat antibody for 1 h at RT in
dark, followed by three PBS washes. The cell nuclei were stained with 1
μg/mL 4′ ,6-diamidino-2-phenylindole (DAPI) for 15 min. Sections were
washed three times with PBS and mounted with an anti-fade mounting
medium. Images were captured with a Leica TCS SP8 confocal laser
scanning microscope.

2.6. Dynamic distribution of FITC-RAP after oral administration
FTIC-RAP (10 mg/mouse) was orally administrated to each mouse (n
= 6). The mice were sacrificed at 0, 1, 2, 3, 4, and 5 h later, and blood
and major tissues, including liver, spleen, kidney, stomach, small in­
testine, mesenteric lymph nodes (MLN), Peyer's patches (PPs), caecum,
and colon, were collected and imaged using an IVIS Lumina XR in vivo
imaging system (PerkinElmer) immediately. The molecular size of FITCRAP was monitored using high-performance gel-permeation chroma­
tography. Similarly, the dynamic distribution of FITC-DOP control in
PPs was investigated. For assay by confocal microscopy, cell suspensions
from tissues or bone marrow were collected and detected for RAP dis­
tribution in vivo.

2.11. Cell culture and treatment
Macrophage RAW264.7 cells were used to confirm effects of RAP on
macrophage differentiation as reported in previous study (Wei et al.,
2019). In brief, RAW264.7 cells were treated with RAP at different
concentrations (0.001, 0.01, 0.1, 1, 10, 100 μg/mL). After 24 h incu­
bation, cell suspensions were collected for ELISA assay. According to the
results, we chose an optimal concentration for the following study. To
figure out the effects of RAP on macrophage differentiation, we treated
RAW264.7 cells with RAP at the optimal concentration and detected the
surficial markers of macrophages by flow cytometry, including F4/80,
CD80, CD86, and CD206. Antibodies for flow cytometry are shown in

Table 1.
To figure out the potential receptors of RAP on DCs, bone marrowderived dendritic cells (BMDCs) were isolated as reported in previous
study (Granucci et al., 2012). In brief, femur bones were collected and
transferred into dishes with 70% ethanol on ice for 2 min/time, repeated
three times. BM cells were collected, washed, and resuspended with 1×
RBS water lysis to remove the red blood cells. 2× 106 cells/mL were
prepared with culture medium (RPMI-1640 + 10% FBS + 20 mM
penicillin/streptomycin +20 ng/mL rmGM-CSF). 10 mL cell suspension
was added into each petridish and was incubated at 37 ◦ C, 5% CO2 for 3
days. After removing old medium, 10 mL of fresh culture medium with
20 ng/mL GM-CSF was added to each petridish, and the cells were
incubated for another 3 days. BMDCs were treated with RAP for 24 h at
the optimal concentration and then screened the expression of surficial
toll-like receptors by flow cytometry, including TLR1, TLR2, TLR4,
TLR5, and TLR6. Antibodies are shown in Table 1.

2.7. High-performance gel permeation chromatography coupled with
fluorescence detector (HPGPC-FLD) analysis
The tissues collected as described above were homogenized using a
3-fold volume of 0.1 mol/L phosphate buffer (pH 7.4) and centrifuged at
15,000 rpm for 10 min. PP collected from the ligated loop assay model
was homogenized using 200 μL PBS and centrifuged at 15,000 rpm for
10 min. The supernatant was collected and stored at − 20 ◦ C for chro­
matographic analysis. The separation was achieved on a TSK GMPWXL
column (300 × 7.8 mm i.d., 10 μm) system operated at 40 ◦ C using an
Agilent-1100 HPLC system equipped with FLD. Ammonium acetate
aqueous solution (20 mM) was used as a mobile phase at a 0.6 mL/min
flow rate. The excitation wavelength and emission wavelength of FLD
were 495 and 515 nm, respectively.
2.8. Western blotting (WB)

PPs from different groups were treated with RAP (100 mg/kg),
collected at time points (0, 1, 2, 3, and 4 h) after oral administration, and
prepared for WB. In brief, PPs collected at different time points were
lysed with RIPA protein extraction reagent containing protease and
phosphatase inhibitors for 30 min. Protein samples were separated by
10% SDS–PAGE and transferred to a PVDF membrane. Membranes were
blocked in 5% blocker milk (BioRad) at room temperature (RT) for 1 h
then incubated with primary antibodies at 4 ◦ C overnight with shaking.
According to our previous study (Wei et al., 2016), the primary anti­
bodies were GAPDH, P38, p-P38, P65, p-P65, ERK, and p-ERK. The
membranes were washed three times with PBST (0.1% Tween 20) and
incubated with horseradish peroxidase (HRP)-conjugated secondary
antibodies for 1 h. Protein bands were visualized using enhanced
chemiluminescence (ECL) detection reagent and medical X-ray film.
Gray value of each band was evaluated with ImageJ software.

Table 1
Antibodies used for flow cytometry analysis.

2.9. ELISA for quantitative analysis of cytokines
For ELISA assay, PPs were collected 24 h after 100 mg/kg RAP or
RAP at different concentrations (50, 100, and 200 mg/kg) treatment,
3

Fluorophore

Antibody

Clone

Vendor

APC
FITC
PE
PE/Cyanine7
PE
PE
PE/Cyanine7
PE
PerCP
PE
PE
PE/Cyanine 7
Alexa Fluor647
PE

Anti-CD11c
Anti-MHCII
Anti-CD4
Anti-CD8a
Anti-CD19
Anti-CD80
Anti-F4/80
Anti-CD86
Anti-CD206
Anti-TLR1Anti-TLR2
Anti-TLR4
Anti-TLR5
Anti-TLR6

N418
M5/114.15.2
GK1.5
53–6.7
1D3/CD19
16-10A1
BM8
A17199A
C068C2
TR23
CB225
MTS510
ACT5
418,601

BioLegend
BioLegend
BioLegend
BioLegend
BioLegend
BioLegend
BioLegend
BioLegend
BioLegend
TheremoFisher
BioLengend
BioLegend
BioLegend
R&D systems

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Carbohydrate Polymers 292 (2022) 119653

2.12. Flow cytometry analysis

3. Results

For the detection of RAP-induced immune responses, tumor tissues
and PPs were minced then ground with a syringe plug. PPs collected
from mice were ground with a syringe plug for the isolation and
detection of immune cells. Tissues and cells were collected and rinsed
with ice-cold PBS at 400 g for 5 min, then filtered through a 70 μm filter.
Single-cell suspensions were prepared for antibody staining. Cells were
incubated with antibodies or the matching isotypes for 25 min at room
temperature. The stained cells were rinsed twice, resuspended in PBS,
and analyzed by FACSAria III (BD Biosciences). Data were analyzed with
FlowJo V10 software. Antibodies for flow cytometry are shown in
Table 1.

3.1. RAP's antitumor activity is immune system-dependent
As shown in Supplementary Fig. 1A–D, the average molecular weight
and purity of RAP remain unchanged, and the 1H NMR spectrum, 13C
NMR spectrum, and monosaccharides composition further confirmed its
stable chemistry. The average molecular weight of RAP was 1334 kDa.
And RAP was composed of Rha, Ara, Glc, Gal and GalA in a molar ratio
of 0.03:1.00:0.26:0.37:(0.28).
The antitumor beneficial effects of polysaccharides isolated from

Radix Astragali have been closely associated with its immunomodula­
tory effects (Li et al., 2020; Yang et al., 2013; Lijing Zhou et al., 2017).
Here we first optimized RAP's dosages by testing RAP-induced immune
responses in PPs, and the results showed that 100 mg/kg is the optimal
to induce significant increase the IL-6 and TNF-α production in 24 h after
oral administration (Supplementary Fig. 2). The further evaluation, with
cisplatin as the positive control, confirmed that this dosage significantly
suppressed the growth of 4T1 breast tumor (Supplementary Fig. 3). And
it was much safer than the positive control cisplatin which caused half
death. Therefore, 100 mg/kg was selected for the subsequent tests.
To determine if RAP's antitumor effects are dependent on its
immunomodulatory effects, we compared RAP's effects on the 4T1

2.13. Statistical analysis
Each experiment was independently repeated three times. Statistical
analysis was performed by IBM SPSS Statistics 25 software. As noted in
figure legends, all data are shown as mean ± SD. Statistical differences
between each experimental group were analyzed by Student's t-test or
one-way ANOVA. Differences with P < 0.05 were considered significant.

Fig. 1. RAP's antitumor activity is dependent on the
immune system. (A) Timeline of 4T1 tumor-bearing
mouse model. RAP was pre-treated for 7 d before
4T1 cells were implanted into the mammary fat pads
of BALB/c mice or nude mice (n = 8– 10 for each
group) and then the treatment continued. Mice were
sacrificed after 21 d treatment with RAP. (B and C)
Tumor (B) and tumor weight (C) in BALB/c mice. (D
and E) Tumor (D) and tumor weight (E) in nude mice.
(F and G) Percentage of CD8+ T cells (CTL, F) and

CD4+CD25+ T cells (Treg, G) in the tumor of BALB/c
mice. (H–K) Cytokine production in tumors isolated
from BALB/c mice, including TGF-β (H), IL-10 (I),
IFN-γ (J), and MCP-1 (K), Data are shown as mean ±
SD. Significant difference *P < 0.05, **P < 0.01,
***P < 0.001, ns = no significance.

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Carbohydrate Polymers 292 (2022) 119653

breast tumor growth between BALB/c mice and the immune-deficient
nude mice (Fig. 1A). The results showed that 100 mg/kg RAP signifi­
cantly suppressed tumor growth (P < 0.001; Fig. 1B and C) in BALB/c
mice but became inactive in nude mice (Fig. 1D and E). Body weight of
mice did not show a big difference during the whole experiment (Sup­
plementary Fig. 4). Thus, these findings suggest that RAP has a signifi­
cant antitumor effect, which is related to its immunoregulation
functions.
We further screened immune cells and cytokines to reveal RAPinduced immune responses in tumor tissues by Flow cytometry and
ELISA assay. Our data indicated that RAP could induce antitumor im­
mune responses in tumor tissues, including the increase of CD3+CD8+
cytotoxic T lymphocyte (CTL) (P < 0.05; Fig. 1F), the decrease of
CD4+CD25+ regulatory T cells (Treg, P < 0.05; Fig. 1G) and changes in
antitumor-related cytokines TGF-β (P < 0.05; Fig. 1H), IL-10 (P < 0.05;
Fig. 1I), IFN-γ (P < 0.05; Fig. 1J), and MCP-1 (P < 0.05; Fig. 1K). These
findings suggest that the immune system plays a critical role in RAP's

anti-tumor effects.
Furthermore, a big difference between the tumor tissues of BALB/c
and nude mice was found in the population and differentiation of
macrophages. Flow cytometry results showed that total macrophages of

tumor tissues were significantly decreased in RAP-treated BALB/c mice
(P < 0.01, Fig. 2B) but not in nude mice (Fig. 2D). Further analysis of
macrophage differentiation in BALB/C mice showed that RAP caused a
noticeable increase of the F4/80+CD11bhigh M1-type macrophage
(MTM, P < 0.001) and a significant decrease of F4/80+CD11blow tumorassociated macrophage (TAM, P < 0.01) in BALB/c mice (Fig. 2A and B),
both of which contributes to the inhibition of tumor growth (Franklin
et al., 2014). While in the tumor tissue collected from nude mice, neither
MTM nor TAM was affected by RAP treatment (Fig. 2C and D). To
confirm the effects of RAP on differentiation of macrophages, we also
test the effects of RAP on RAW264.7 cells in vitro. Interestingly, RAP
promoted the differentiation of RAW264.7 cells into F4/
80+CD80+CD86+ M1 type macrophages (Fig. 2E-G) but had no effects
on CD206 expression (Fig. 2H). Besides, RAP also induced RAW264.7
cells to produce IL-6 and TNF-α production (Fig. 2I and J). These in vitro
findings suggest that RAP could induce macrophage differentiating into
MTM. Taken together, the above findings prove that RAP-induced
antitumor effects are dependent on the immune system.

Fig. 2. Differentiation of macrophages induced by
orally administrated RAP. (A) Dot plots of flow
cytometry showing F4/80+CD11b+ macrophages in
4T1 breast tumor-bearing BALB/C mice. (B) Per­
centages of F4/80+ macrophages, M1 type macro­
phages (MTM, F4/80+CD11bhigh macrophages), and
tumor-associated

macrophages
(TAM,
F4/
80+CD11blow macrophages) in the tumor of BALB/c
mice. (C) Dot plots of flow cytometry showing F4/
80+CD11b+ macrophages in 4T1 breast tumorbearing nude mice. (D) Percentage of F4/80+ mac­
rophages, MTM, and TAM in 4T1 breast tumorbearing nude mice. (E-H) Flow cytometry histogram
of F4/80 (E), CD80 (F), CD86 (G), and CD206 (H)
expression on RAW264.7 cells. RAW264.7 cells were
treated with RAP (blue) or without treatment (con­
trol, red). (I and J) Cytokine IL-6 (I) and TNF-α (J)
production from RAW264.7 cells with treatment of
RAP at different concentrations (0, 0.001, 0.01, 0.1,
1, 10, 100 μg/mL). Data are shown as mean ± SD.
Significant difference **P < 0.01, ***P < 0.001,
****P < 0.0001, ns = no significance.

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Carbohydrate Polymers 292 (2022) 119653

3.2. Intact RAP is selectively transported into lymphatic system

tracts after treating with the unlabeled RAP by oral administration. The
results showed that the carbohydrate quickly left the stomach in 1–2 h
(Fig. 3A), mainly stayed in the small intestine for 3–4 h (Fig. 3B), but
shortly occurred in the large intestine only at the third hour (Fig. 3C).

And the carbohydrate content detected in the large intestine was only
around 1/6 of that in the small intestine. These results are consistent
with DOP's destiny where the carbohydrate polymers were quickly
degraded to short-chain fatty acids in the large intestine (Li et al., 2019).
Then we labeled RAP with FITC to improve the detection sensitivity
for subsequent tracking its distribution in organs (Supplementary
Fig. 5). Analysis of fluorescence intensity demonstrated a similar

We previously found the positive signal of FITC-RAP in the bone
marrow (Bao et al., 2021) but did not find it in the serum. So, we hy­
pothesized that there might be a blood-independent way for RAP to
work in the body. To test this possibility, we systematically tracked RAP
in its oral route. We already found that Dendrobium officinale poly­
saccharide (DOP), being indigestible and unabsorbed, ended in regu­
lating gut microbiota as a prebiotic (Li et al., 2019). So, DOP was used as
a control polysaccharide.
We first monitored the carbohydrate contents in the gastrointestinal

Fig. 3. Dynamic distribution of RAP in the gastrointestinal tract. (A-C) Dynamic carbohydrate contents in the stomach (A), the small intestine (B), and the large
intestine (C) collected from normal mice at 1– 5 h after gavage with unlabeled RAP (10 mg/mouse); RAP was detected by the phenol‑sulfuric acid method. Mice
without RAP treatment is the control group. (D) Fluorescence images of major organs, including stomach, small intestine, caecum, colon, liver, kidneys, and spleen
collected from normal mice (n = 6) at 1 to 5 h after gavage with FITC-RAP (10 mg/mouse). Mice without RAP treatment is the control group (0 h). (E) HPGPC-FLD
chromatograms of FITC-RAP in serum. (F-K) Fluorescence intensity of small intestine (F), caecum (G), colon (H), liver (I), kidneys (J), and spleen (K). (L-N) HPGPCFLD chromatograms of FITC-RAP in the small intestine (L), caecum (M), and colon (N). Red dotted lines show the chromotographic retention time of original FITCRAP. Significant difference *P < 0.05, **P < 0.01, ***P < 0.001, ns = no significance.
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Carbohydrate Polymers 292 (2022) 119653

dynamic distribution of FITC-RAP signals in the gastrointestinal tract
(Fig. 3D and F-H), suggesting positive signals of FITC-RAP were detected
in the small intestine (Fig. 3F), caecum (Fig. 3G), and colon (Fig. 3H).
And no positive signals were detected in the serum (Fig. 3E), liver
(Fig. 3I), kidneys (Fig. 3J), or spleen (Fig. 3K), showing that FITC-RAP
could not be absorbed into the blood system, liver, kidney, and spleen.
The HPGPC-FLD chromatograms further indicated that FITC-RAP
remained undegraded in the small intestine (Fig. 3L) but quickly
degraded in the caecum (Fig. 3M) and colon (Fig. 3N). Therefore, the
fluorescence signals in the caecum and colon were proved to be unre­
liable to detect carbohydrate polymers. These data suggest that the small
intestine is the main location where RAP remained an undegraded car­
bohydrate polymer for a long time.
Further examination of the gut-associated lymphoid tissues PPs,
which are the immune sensors of the small intestine, offered interesting
data. The fluorescence images showed that FITC-RAP, but not FITCDOP, transiently accumulated in the PPs about 3–4 h (P < 0.001;
Fig. 4A and B). The procedure was highly consistent with RAP's stay in
the small intestine. As the fluorescence intensity reached maximum at 2
h, HPGPC analysis was used to further confirm the integrity of FITCRAP. The result indicated that FITC-RAP remained intact in PPs
(Fig. 4C), while FITC-DOP failed to enter PPs (Fig. 4D). The confocal

micrographs also clearly displayed the occurrence of FITC-RAP in PPs
(Fig. 4E). These results demonstrate that the PPs gateway for RAP is
efficient and selective.
3.3. RAP directly targets follicle dendritic cells and initiates immune
responses in Peyer's patches
After entering PPs, the cells directly targeted by RAP play a critical
role in the beginning and initiation of its antitumor immune responses.
By screening immune cells in PPs using flow cytometry, we observed
that clearly at 1 h after FITC-RAP treatment, the positive signals of FITCRAP were only detected in the monocytes (Fig. 5C) rather than CD8+ T

cells (Fig. 5A), CD4+ T cells (Fig. 5B), or CD19+ B cells (Fig. 5D).
Furthermore, without a significant increase of CD11c+ DCs (Fig. 5E and
F), a noticeable portion (22.4%) of CD11c+ DCs among those monocytes
could bind with FITC-RAP (Fig. 5G and H), suggesting that CD11c+ DCs
might be RAP's direct target cells. Observations of frozen sections
confirmed that FITC-RAP was directly captured by CD11c+ DCs in PPs
(Fig. 5I). These data suggest that RAP could directly target FDCs after
being transported into PPs.
Further analysis of RAP-induced immune responses in PPs showed
that the proportion of CD11c + DCs was significantly increased by RAP

Fig. 4. Selective transportation and intact detection of RAP in PPs. (A and B) Fluorescence images (A) and fluorescence intensity (B) of PPs separated from small
intestines collected from normal mice at 0 to 5 h after gavage with FITC-RAP and FITC-DOP (10 mg/mouse). Mice without RAP treatment is the control group (0 h).
(C and D) HPGPC-FLD chromatograms of PPs at 2 h after gavage with FITC-RAP (C) and FITC-DOP (D). (E) Confocal microscopic images of PP sections isolated from
mice treated with FITC-RAP (green) for 2 h. DAPI (blue) was used as a DNA-specific stain. Scale bar, 40 μm. Significant difference ****P < 0.0001, ns = no
significance.
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Carbohydrate Polymers 292 (2022) 119653

Fig. 5. Direct contact of RAP with DCs in vivo. (A-D)
Flow cytometry histogram of RAP-bound immune
cells, including CD8+ T cells (A), CD4+ T cells (B),
monocytes (C), and CD19+ B cells (D) of PPs isolated
from mice treated with FITC-RAP (10 mg/kg) for 1 h.
(E and F) Flow cytometry analysis (E) and percentage
(F) of CD11c+ DCs population in the monocytes of

PPs isolated from mice of the control and FITC-RAPtreated groups. (G and H) Flow cytometry histogram
(G) and percentage (H) of RAP-binded CD11c+ DCs
in the CD11c+ DCs shown in Fig. 5E. (I) Confocal
images of PP frozen sections (dome zone) collected
from mice treated with FITC-RAP for 1 h. FITC-RAP
(green), CD11c (yellow), DAPI (blue), and scale bar,
40 μm. Significant difference ****P < 0.0001, ns =
no significance.

treatment at 24 h (Fig. 6A), suggesting that RAP could increase the
population of DCs in the PPs. Flow cytometry analysis also indicated that
CD80 and MHCII expressions of CD11c+ DCs were induced in 24 h after
oral administration of RAP (Fig. 6B), suggesting that RAP would quickly
induce the maturation and differentiation of DCs in PPs. As we previ­
ously found that RAP could quickly induce the phosphorylation of
MAPKs and NF-κB signaling pathways (Wei et al., 2016), we next
examined how orally administrated RAP affected these signaling path­
ways in PPs at different time points (1–4 h). Western blotting results
showed that RAP could quickly induce the phosphorylation of ERK, p38,
and p65, implying the activation of MAPKs and NF-κB signaling in the
PPs in 1 h (Fig. 6C). In addition, cytokines production was also detected,
and the results indicated that RAP was quickly transported into PPs to
trigger immune responses. The results of the ELISA assay indicated that
100 mg/kg RAP could up-regulate the production of IL-6 (P < 0.001;
Fig. 6D), TGF-β (P < 0.05; Fig. 6F), and INF-γ (P < 0.05; Fig. 6G), but
down-regulate M-CSF (P < 0.01; Fig. 6E) in PPs isolated from mice

treated with RAP for 24 h. Taken together, these results demonstrated
that RAP quickly triggered immune responses in PPs by targeting and
activating FDCs, which should be the initiation of RAP-induced immu­

noregulation in vivo.
3.4. Follicle dendritic cells might further transport RAP in the lymphatic
system
To figure out the destiny of RAP after being transported into PPs, we
determined the signal of FITC-RAP in the distant lymphatic organs using
immunofluorescence staining assay. Further tracking the binding of
CD11c+ DCs and FITC-RAP using confocal microscopy revealed that,
after the treatment of FITC-RAP for 24 h, positive signals of FITC-RAP
were detected in mesenteric lymph node (MLN, Fig. 7A), bone marrow
(Fig. 7B), and even tumor tissues (Fig. 7C). Furthermore, the binding of
CD11c+ DCs and FITC-RAP was demonstrated in the merged images of
these organs, as found in PPs. Thus, these observations collectively
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Carbohydrate Polymers 292 (2022) 119653

Fig. 6. Immune responses induced by RAP in PPs.
(A) Flow cytometry histogram of CD11c+ DCs from
PPs of mice treated with or without RAP for 24 h.
Red, control group; blue, RAP-treated group. (B) Dot
plots of cell surface expression of MHCII and CD80 in
CD11c+ DCs shown in Fig. 6A. (C) Signaling path­
ways of NF-κB and MAPKs (p38 and ERK) of PPs at
1– 4 h after oral administration with RAP (100 mg/
kg), as determined by Western blotting assay. GAPDH
was the control. Gray value of protein was analyzed
by ImageJ. (D-G) IL-6 (D), M-CSF (E), TGF-β (F), and

IFN-γ (G) production in PPs of the small intestine
collected from mice 24 h after RAP treatment. PP
homogenate was collected and detected by ELISA
kits. Data are shown as mean ± SD. Significant dif­
ference *P < 0.05, **P < 0.01, ***P < 0.001.

suggest that RAP might be further transported to distant organs via the
lymphatic system by FDCs, providing a chance for RAP to directly
contact diverse cells and to exert beneficial effects. In this regard, this
lymphatic route successfully bridges the gap between the in vivo and in
vitro investigations.

4. Discussion
The current investigations of polysaccharide pharmacokinetics
solely rely on interpreting fluorescence signals; however, this method
needs to be validated before it can be considered reliable. The fluores­
cence signal might arise from the fluorescence reagent itself if the
sample is not pure enough or if the fluorescence flag is released some­
how. So, the first step to ensuring valid fluorescent results is to confirm
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Carbohydrate Polymers 292 (2022) 119653

Fig. 7. Distribution of FITC-RAP in other tissues. (A–C), Confocal microscopic images of cell suspension isolated from mesenteric lymph nodes (MLN, A), bone
marrow (BM, B), and tumor (C). The tumor-bearing mice were treated with FITC-RAP (green) for 24 h. Cell suspensions were stained with CD11c-APC antibody (red).
Scale bar, 20 μm.

the purity and stability of the labeled polymer. In this study, we used
HPGPC-FLD to check the purity (Supplementary Fig. 5A and C) and
HPGPC-CAD to determine whether the FITC-RAP polymer has been
degraded after labeling. The second concern is that the polymer might
be degraded in the gut. This concern is proved by the difference of the
signals in the caecum and colon between Fig. 3G/H and M/N. It was
clearly shown that the positive fluorescence signals observed in the
caecum and colon came from the degraded chemicals instead of the
intact polymer. The fluorescence signal needs to be validated. So, we not
only tested the carbohydrate property using the phenol‑sulfuric acid
method, but also checked the molecular size using HPGPC-FLD to see if
RAP remains an intact carbohydrate polymer in the intestinal organs.
With these validations, the fluorescence signal observed in the confocal
micrograph can be assigned to the polymer with confidence. Here we
present the first evidence to show that RAP remains intact in the
lymphatic system after oral administration.
Most in vitro studies of polysaccharides were focused on macro­
phages (Guan et al., 2020; Kallon et al., 2013; L. Zhou et al., 2017),
while our findings highlight the importance of FDCs in the mechanism of
polysaccharide's bioactivities. The cells that polysaccharides directly
target after passing through the gut cell wall have not been directly
identified before (Wang et al., 2021; Yin et al., 2019; Yue Yu et al., 2018;
Zhao et al., 2020). Our findings clearly reveal that FDCs are RAP's direct
targets in PPs and further deliver RAP to other lymphatics and even
tumor tissues. These results may offer a further explanation about the
initiation of polysaccharide-induced immune responses, which high­
lighted the important role of FDCs.

The multiple immunomodulatory effects of RAP in vivo might result
from a combination of the lymphatic system-dependent effects in the

small intestine and gut microbiota-related effects in the large intestine.
The interaction between prebiotics and gut microbiota needs time and
cannot explain the fast immune responses in the small intestine induced
by orally administrated polysaccharides (Kim et al., 2019; Sakai et al.,
2019). This lymphatic route found in this study works 2 h before RAP
arrives the caecum where gut microbiota mainly works, therefore it is
independent to gut microbiota. But we cannot exclude the possibility of
microbiota-related immune-regulation because a large amount of RAP
entered the caecum and was quickly digested (Fig. 3M and N). Consis­
tent with this finding, many studies have shown that gut microbiota
could digest polysaccharides to produce short-chain fatty acids (SCFAs)
which are the mediators of microbiota and the immune system (Hong
et al., 2020; Li et al., 2019; Liu et al., 2019; Zhou et al., 2021). We
speculate that polysaccharides' beneficial effects are a combination of
activity in both the small and large intestines.
Although we here proved a blood/microbiota independent and se­
lective lymphatic route for RAP to work in vivo, one limitation of this
study is that we are not sure how RAP passes through the gut cell wall.
As shown in the Fig. 4A and B, the control polysaccharide DOP failed to
enter PPs, suggesting that this lymphatic route is selective to poly­
saccharides. There are several possible cell receptors known to poly­
saccharides, such as TLRs, which might also mediate this delivery route
(Batbayar et al., 2012; Taylor et al., 2007; Wei et al., 2016). By screening
surficial TLRs using flow cytometry (Supplementary Fig. 6), we found
TLR4 might be the targeting receptor of RAP on DCs, which is consistent
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Carbohydrate Polymers 292 (2022) 119653

with results shown in previous studies. Interestingly, we also found
TLR1 expression on DCs was induced by RAP, suggesting TLR1 is a
possible related receptor of RAP as well. We speculate the way of RAP
passing through the gut cell wall might be receptor-dependent, which
deserves further investigation.

Sciences.
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.carbpol.2022.119653.

5. Conclusions

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which plays a crucial role in the tumor suppression of RAP. These
findings indicate a blood/microbiota independent and selective
lymphatic route for intact RAP to modulate the immune system. This
route helps bridge the gap between the in vitro and in vivo studies of
natural polysaccharides and spires novel insights into how orally
administered polysaccharides work in vivo.

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CRediT authorship contribution statement
Quanwei Zhang: Conceptualization, Data curation, Formal analysis,
Investigation, Methodology, Software, Visualization, Writing – original
draft. Lifeng Li: Conceptualization, Data curation, Formal analysis,
Investigation, Methodology, Software, Visualization, Writing – original
draft. Shuang Hao: Methodology, Formal analysis, Visualization. Man
Liu: Methodology, Formal analysis, Visualization. Chuying Huo:
Methodology, Formal analysis, Visualization. Jianjun Wu: Methodol­
ogy, Formal analysis, Visualization. Hongbing Liu: Methodology,
Formal analysis, Visualization. Wanrong Bao: Methodology, Formal
analysis, Visualization. Hongming Zheng: Methodology, Formal anal­
ysis, Visualization. Zhipeng Li: Methodology, Formal analysis, Visual­
ization. Huiyuan Cheng: Methodology, Formal analysis, Visualization.
Hauyee Fung: Methodology, Formal analysis, Visualization. Tinlong
Wong: Methodology, Formal analysis, Visualization. Pingchung
Leung: Writing – review & editing. Shunchun Wang: Writing – review
& editing. Ting Li: Writing – review & editing. Ge Zhang: Writing –

review & editing. Min Li: Writing – review & editing. Zhongzhen Zhao:
Writing – review & editing. Wei Jia: Writing – review & editing.
Zhaoxiang Bian: Writing – review & editing. Timothy Mitchison:
Writing – review & editing. Jingchao Zhang: Writing – review & edit­
ing. Aiping Lyu: Writing – review & editing. Quanbin Han: Concep­
tualization, Funding acquisition, Project administration, Resources,
Writing – original draft, Writing – review & editing. Handong Sun:
Writing – review & editing.
Declaration of competing interest
The authors declare that they have no known competing financial
interests or personal relationships that could have appeared to influence
the work reported in this paper.
Acknowledgments
This work was funded and supported by HKSAR Innovation and
Technology Fund (ITF), Tier 3, ITS/311/09, General Research Fund
(12100615, 22100014, 12100818), UGC Research Matching Grant
Scheme (2019-1-10, 2019-1-14, 2019-2-06), Health Medical Research
Fund (11122531, 14150521, 17182681), National Natural Sciences
Foundation in China (81473341, 82173948), the Science and Technol­
ogy Project of Shenzhen (JCYJ20160531193812867), the Key-Area
Research and Development Program of Guangdong Province
(2020B1111110007), Hong Kong Baptist University (RC-Start-up
Grants, MPCF-001-2016/2017, MPCF-002-2021-22, RC-IRMS/14-15/
06, IRMS-20-21-02, FRG2/17-18/060 and FRG2/16-17/002), and Vin­
cent & Lily Woo Foundation. We appreciate the advice from Prof. Ding
Kan at Shanghai Institute of Materia Medica, Chinese Academy of
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