M cells of mouse and human Peyer''s patches mediate the lymphatic absorption of an Astragalus hyperbranched heteroglycan

Carbohydrate Polymers 296 (2022) 119952

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

M cells of mouse and human Peyer's patches mediate the lymphatic
absorption of an Astragalus hyperbranched heteroglycan
Quanwei Zhang a, Shuang Hao b, Lifeng Li a, Man Liu a, Chuying Huo a, Wanrong Bao a,
Huiyuan Cheng a, Hauyee Fung a, Tinlong Wong a, Wenjie Wu 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, *
a

School of Chinese Medicine, Hong Kong Baptist University, Hong Kong 999077, Hong Kong, 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, Hong Kong,
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, 999078, Macau
f
Laboratory of Systems Pharmacology and Department of Systems Biology, Harvard Medical School, Boston 02115, United States
b

A R T I C L E I N F O

A B S T R A C T

Keywords:
Radix Astragali polysaccharide
Microfold cell
Lymphatic absorption
Transcytosis
Immune regulation

The gut cell wall is considered an impenetrable barrier to orally administrated polysaccharides. We recently
reported a selective lymphatic route for Radix Astragali polysaccharide RAP to enter Peyer's patches (PPs) to
trigger immune responses. However, how RAP enters PPs is unclear. Herein, we screened the intestinal epithelial
cells of mice and found that the follicle-associated epithelium cells were specifically bound with FITC-RAP.
Further studies in vitro and in vivo revealed that RAP was efficiently transported by microfold (M) cells. We
also confirmed that M cell-transported RAP directly contacted dendritic cells. More importantly, for the first
time, we verified this interesting M cell-mediated transcytosis of RAP in the human distal ileum. Mechanistically,
we identified M cells to be the transporter cells that independently deliver RAP into the lymphatic system to
trigger immune responses. This interesting transcytosis mechanism might apply to many other immunomodu­
latory polysaccharides orally dosed to human body.

1. Introduction
The gut wall barrier to macromolecules remains an unsolved chal­
lenge for developing orally-delivered macromolecular therapeutics
(Moroz et al., 2016). Recently proposed solutions have focused on
highly engineered systems such as self-orienting microinjectors
(Abramson et al., 2019), but such complexity creates challenges for
manufacture and regulation (Hussain, 2016). In puzzling over this, we
noticed that polysaccharides from traditional herbal medicines quickly
affect the immune system after oral dosing (Jiang et al., 2010; Sche­
petkin & Quinn, 2006; Yu et al., 2018). Our previous study revealed that
the Radix Astragali polysaccharide RAP enters Peyer's patches (PPs) in

the small intestine intact, and subsequently initiates mucosal immune
responses by targeting follicle dendritic cells (FDCs) (Zhang et al.,
2022). In other words, this appears to be a lymphatic route for

polysaccharides crossing the gut wall barrier.
Radix Astragali (RAP, Huang qi in Chinese) is one of the most widely
used traditional Chinese medicine (TCM). It could promote Yang and
replenish Qi, which is considered the primitive application of the herb's
immune regulation effects. Being first recorded and classified as the top
grade in Shennong's Materia Medica Classic, RA exists various therapeutic
activities, such as antioxidant, anti-cancer, anti-diabetic, and immuno­
modulatory effects (Chen et al., 2020; Fu et al., 2014; Liu et al., 2011).
Statistics showed that RA was found in about 200 Chinese medicine
prescriptions, and water decoctions were the major ways to use RA in
clinics (Cheng et al., 2019). Polysaccharides of RA, as one of the most
important compounds in its water decoction, play a critical role in im­
mune functions of RA (Li et al., 2020; Zheng et al., 2020). A better un­
derstanding of how polysaccharides of RA work in vivo would be helpful
for the development and application of RA-based prescriptions.

* Corresponding authors.
E-mail addresses: (J. Zhang), (A. Lyu), (Q. Han).
/>Received 6 June 2022; Received in revised form 16 July 2022; Accepted 1 August 2022
Available online 4 August 2022
0144-8617/© 2022 The Authors. Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license ( />

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Carbohydrate Polymers 296 (2022) 119952

The hyperbranched heteroglycan RAP (1334 kDa) was purified from
Radix Astragali (Yin et al., 2012). Its chemical structure was charac­
terized by monosaccharide composition, partial acid hydrolysis,
methylation analysis, GC–MS, NMR spectra, SEM, and AFM microscopy.
It was composed of Rha, Ara, Glc, Gal, and GalA in a molar ratio of
0.03:1.00:0.26:0.37:(0.28). As shown in Supplementary Fig. 1 in our
newly reported study (Zhang et al., 2022), its average molecular weight
and purity remain unchanged, and the 1H NMR spectrum, 13C NMR
spectrum, and monosaccharides composition further confirmed its sta­
ble chemistry. As a major component, it looks similar to the reported
heteroglycans from the same herb medicine (Zheng et al., 2020).
Although our recent study revealed a selective route for RAP entering
PPs to trigger immune responses quickly (Zhang et al., 2022), it did not
explain how RAP actually crosses the cell wall. Transcytosis by micro­
fold (M) cells (Dillon & Lo, 2019) is one possible mechanism. These
specialized epithelial cells, located on immune sensors of the small in­
testine PPs, deliver pathogens and food molecules from the gut lumen to
the lymphatic system (Neutra et al., 1996; Okumura & Takeda, 2017).
The M cell-targeted transcytosis has been a promising route for oral
vaccination, especially for bioactive macromolecules (Kim & Jang,
2014; Saraf et al., 2020). Considering that RAP quickly triggers immune
responses in PPs 1 h after oral administration, and M cell-mediated
transport can be completed in 15 min (Sakhon et al., 2015), we hy­
pothesize that M cells are the transporter for RAP to pass through the gut
cell wall to initiate immunoregulation in vivo.
In this study, we collected comprehensive evidence in vitro and in
vivo, on both animal and human subjects, that supports this hypothesis.
We first screened the intestinal epithelial cells using flow cytometry and
found that RAP was specifically bound to the follicle-associated
epithelial (FAE) cells. We further tested RAP's transport on an in vitro

FAE model. A control nanoparticle and the specific M-cell marker GP2
were used to validate the specific transport function of M-like cells in
this model. Immunofluorescence staining assay, flow cytometry, and
high-performance permeation chromatography were performed to
detect the transport of intact FITC-RAP in the lower layer of transwell
inserts. In order to exclude the influence of FITC, we also tested unla­
beled RAP. ELISA assay was used to assess the effects of transported RAP
on macrophages. The results suggested that M-like cells transported RAP
could reach a significant concentration of 1 μg/mL. In the in vivo ex­
periments, the FAE M cells were characterized by two specific cell
markers, GP2 and NKM 16-2-4, in immunofluorescence staining. The
RAP transport by M cells was clearly demonstrated in 3D confocal mi­
crographs. This interesting transport of RAP by M cells was also verified
in human subjects. Taken together, evidence showed that M cells were
the specific transporter cells.

primary antibody, goat anti-rabbit or rat IgG antibody conjugated with
Alexa Fluor 647, and goat anti-rat IgG antibody conjugated with
Alexa488 was purchased from Abcam. Matrigel basement membrane
matrix phenol red-free, phenol-sulfuric acid, 4′ ,6-diamidino-2-phenyl­
indole DAPI, anti-microfold cell antibody (NKM 16-2-4), and methyl
sulfoxide were obtained from Merck. Fluoresbrite® Multifluorescent
Microspheres (0.20 μm) were purchased from Polysciences. DMEM, fetal
calf serum (FBS), 4 % paraformaldehyde (PFA), and all ELISA kits were
purchased from Thermo Fisher Scientific.
2.3. Preparation of polysaccharides and FITC-labeled polysaccharides
RAP was previously prepared from the water extract of the dried
roots of Astragalus membranaceus (Yin et al., 2012) and stored at 4 ◦ C.
RAP is a hyperbranched heteroglycan with a molecular weight of 1334
kDa. The quality was stable compared to multiple newly prepared RAP

samples. This sample displayed high consistency in the HPGPC molec­
ular distribution patterns and the HPLC oligo-saccharide profile pro­
duced by partial hydrolysis (Zhang et al., 2022). As reported, RAP was
labeled with fluorescein isothiocyanate isomer I (FITC) (Li et al., 2019).
2.4. Isolation of intestinal epithelial cells
The small intestine and Peyer's patches (PPs) were isolated from five
mice and were washed five times with cold PBS. All the samples were
stirred in 20 mL of 0.5 mM EDTA/PBS at 37 ◦ C with 300 g for 20 min,
followed by vigorous shaking for 15 s. Samples were washed three times
and filtered with 70 μm nylon filter mesh, and then we would get cell
suspensions. Cell suspensions were prepared as shown in the method of
flow cytometry.
2.5. In vitro M-like cell model
An in vitro M-like cell model was used to evaluate the transport of
RAP (Beloqui et al., 2017). In brief, Caco-2 cells were cultured in flasks
in 10 % (v/v) fetal calf serum DMEM at 37 ◦ C in a 10 % CO2 watersaturated atmosphere. Caco-2 cells (6 × 106) grew on Transwell poly­
ester inserts (0.3 μm pore size, 12 mm diameter) which were coated with
100 μg/mL Matrigel basement membrane matrix prepared in pure cold
DMEM without phenol red. Raji B cells (5 × 105) were resuspended in
DMEM and added to the basolateral chamber of 7-day-old Caco-2 cell
monolayers. The co-cultures were maintained for 5 days. Monoculture of
Caco-2 cells, cultivated as above but without the Raji B cells, was used as
control. Inserts were used for all the following experiments, including
assessment of M cell functionality by nanoparticle transport measure­
ment using flow cytometry, M cell demonstration in co-cultured
monolayers using an immunofluorescence staining assay, and determi­
nation of transported RAP using chromatographic analysis and ELISA
assay.

2. Materials and methods

2.1. Human subjects, animals, and cells
For the study of human subjects, this study was performed following
the established ethical guidelines. It was approved (2019-KY-373) by the
research ethics committee of the School of Medicine, Zhengzhou Uni­
versity, Zhengzhou, China.
C57 BL/6 and BALB/c mice were purchased from the Chinese Uni­
versity of Hong Kong. Five- to eight-week-old mice were used. All animal
experiments followed the Animals Ordinance guidelines, Department of
Health, Hong Kong SAR.
Caco-2 cells, Raji B cells, and RAW264.7 cells were obtained from
American Type Culture Collection (ATCC).

2.6. Assessment of M cell transport functionality
The transport function of this in vitro M cell model was validated
using nanoparticles (Beloqui et al., 2017). Fluoresbrite multifluorescent
microspheres (green, 200 nm diameter) were used in transport experi­
ments. The nanoparticles (400 μL, 5 × 109/mL) were added to the upper
layer of mono-culture and co-culture inserts and then incubated for 2 h
at 37 ◦ C under 5 % CO2. Solutions of the lower inserts were collected and
measured using a cytometer FACSAria (BD Biosciences).
2.7. Identification of M-like cells using immunofluorescence staining

2.2. Antibodies and reagents

Three co-culture and monoculture inserts were used to detect the
presence of M-like cells by immunofluorescence staining. All inserts
were washed three times with PBS and fixed with 4 % paraformaldehyde
(PFA) for 15 min and then blocked for 1 h in 5 % of normal goat serum in

PE-conjugated rabbit anti-mouse glycoprotein 2 (GP2) antibody

(2F11-C3), rat anti-mouse GP2 antibody (2F11-C3), and mouse antihuman GP2 antibody (3G7-H9) was from MBL International. CD11c
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Carbohydrate Polymers 296 (2022) 119952

PBS. Rat anti-human GP2 antibody was used as the primary antibody
and incubated overnight at 4 ◦ C. After washing three times with PBS, all
inserts were incubated with Alexa Fluor 647-conjugated goat anti-rat
secondary antibody for 1 h in the dark. DAPI was used to stain cell
nuclei for 15 min in the dark. All inserts were washed three times with
PBS, and the polyester membranes were cut and mounted using an antifade mounting medium. Slides were examined with a Leica TCS SP8
confocal laser scanning microscope.

2.13. Whole-mount staining
For whole-mount staining, PPs were isolated from the small intestine
of mice treated with/without FITC-RAP. After being washed three times,
the PPs were fixed in 4 % paraformaldehyde overnight at 4 ◦ C and then
permeabilized with 0.1 % Triton X-100 for 30 min. PPs were blocked
with 5 % normal goat serum in PBS for 1 h. Primary antibodies,
including rat anti-mouse glycoprotein 2 (GP2) and rabbit anti-mouse
NKM 16-2-4, and 1 μg/mL DAPI were used to incubate PPs for 1 h at
RT, followed by rinsing 3 × 5 min with PBS. The second goat anti-rat IgG
antibody conjugated with Alexa Fluor 647 and goat anti-rabbit IgG
antibody conjugated with Alexa Fluor 488 were further stained for 1 h at
RT. PPs were washed 3 × 5 min with PBS and mounted with the antifade reagent. Images were captured by a confocal laser microscope
(SP8, Leica Microsystems).

2.8. Treatment of FITC-RAP in the M-like cell model
FITC-RAP (2 mg/mL) in 400 μl DMEM was added to the upper layers
of monoculture and co-culture inserts and then incubated for 2 h at 37 ◦ C
under 5 % CO2. All inserts were removed, and lower solutions were
collected for chromatographic analysis and flow cytometry analysis. The
lower solution collected from the co-culture group treated with unla­
beled RAP was further used to treat RAW264.7 cells in a 96-well plate
for 24 h, and the supernatants were sampled for IL-6 and IL-12 detection
using ELISA kits.

2.14. H&E staining
Frozen sections of PPs from mice were washed three times with PBS.
In H&E staining, the sections were stained with Harris' hematoxylin,
then treated with 1 % acid alcohol, and finally stained with 1 % eosin.
Excess eosin was removed by washing in running water. Sections were
mounted on slides with permount. Images of H&E staining were taken
with light microscopy.

2.9. High-performance gel permeation chromatography coupled with
fluorescence detector (HPGPC-FLD) analysis
500 μL lower layer solution/insert were collected and detected using
HPGPC-FLD assay according to our reported method (Li et al., 2019). In
brief, all solutions were centrifuged at 15,000 rpm for 10 min. The
separation was performed 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.

2.15. Study of human subjects
Twenty volunteers who needed surgery were enrolled in this study.
The age of the participants ranged from 20 to 50 years. All distal ilea of
the small intestine were collected after their surgeries. Some of them
were not qualified because they had no PPs. Most of them only have one
PP. 32 PPs isolated from twenty samples of distal ilea were used in this
study. In brief, the small intestines with PPs were stored in ice-cold PBS
with 5 % FBS. Each small intestine segment was ligated with a surgical
suture for the ligated loop assay. It was then injected with 10 mL FITCRAP solution (5 mg/mL) followed by incubation at 37 ◦ C in an atmo­
sphere of 5 % CO2 for 2 h. 20 PPs were collected for whole-mount
staining, and 12 PPs were for immunofluorescence staining.

2.10. Flow cytometry analysis
200 μL lower layer solution was collected and measured using
FACSAria (BD Biosciences) cytometer. To identify the transporter of
FITC-RAP in the intestinal epithelial cells (IECs) in vivo, cell suspension
of IECs was stained with PE-conjugated GP2 for 25 min at RT and
washed two times. The stained cells were rinsed twice, resuspended in
PBS, and analyzed by a cytometer FACSAria. All data were analyzed
with FlowJo V10 software.

2.16. Statistical analysis
Each experiment was independently repeated at least three times
with consistent results. As noted in figure legends, all data are shown as
mean ± SD. Statistical differences between each experimental group
were analyzed by a t-test or one-way ANOVA. Significant differences
with P < 0.05 were considered significant.

2.11. ELISA for quantitative analysis of cytokines
The supernatants of RAW264.7 cells collected from the above

method were centrifuged at 3000g for 10 min. According to the manu­
facturer's instructions, cytokines Interleukin 6 (IL-6) and IL-12 were
determined using ELISA kits.

3. Results
3.1. RAP specifically appears in FAE cells isolated from Peyer's patches
Our newly published data showed that intact Radix Astragali poly­
saccharide RAP entered Peyer's patches (PPs) within 1 h to initiate im­
mune responses (Zhang et al., 2022). We further screened the intestinal
epithelial cells that may have been involved in the lymphatic transport.
Firstly, we confirmed the fast transport of intact RAP into PPs by an
intestinal loop assay. Images taken by the living system showed that
within 1 h after sample loading to the loop, PPs were found to contain
FITC-RAP (P < 0.001, Fig. 1A and B). High-performance gel permeation
chromatography confirmed that the fluorescence signal came from
intact FITC-RAP (Fig. 1C). After confirming the transport, we isolated
epithelial cells from mice orally treated with FITC-RAP. Before analysis
by flow cytometry, we firstly confirmed the gate of intestinal epithelial
cells (IECs) and excluded the CD45+ cells as shown in Supplementary
Fig. 1. With the similar gate strategy of IECs, flow cytometry analysis of
these isolated cells indicated that only the cells from the FAE of PPs-no

2.12. Immunofluorescence staining
Frozen sections of PPs from mice treated with/without FITC-RAP
were washed three times with PBS and blocked with 5 % normal goat
serum in PBS for 1 h. Sections were incubated with anti-mouse GP2 and
anti-mouse NKM 16-2-4 antibodies overnight at 4 ◦ C. PPs sections were
washed three times with PBS and then treated with Alexa Fluor 647
secondary goat anti-rabbit antibody, Alexa Fluor 488-conjugated antirat antibody, and Alexa Fluor 594-coujugated anti-rat antibody for 1 h
at RT, followed by three consecutive 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 antifade mounting medium. Images were captured with a Leica TCS SP8
confocal laser scanning microscope.
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Carbohydrate Polymers 296 (2022) 119952

other IECs-specifically accumulated FITC-RAP (P < 0.001, Fig. 1D–G).
Therefore, we concluded that the transport of intact RAP into PPs is
associated with the FAE cells.

M cell-specific marker GP2 (Fig. 2B). The flow cytometry analysis
showed that the model group containing Caco-2 cells co-cultured with
Raji B cells was permeable to nanoparticles, while the Caco-2 monolayer
was not (Fig. 2C and D). Thus, the induction of M-like cells, the integrity
of Caco-2 monolayers, and the transport function of M-like cells were
confirmed.
In the tests with FITC-RAP, specifically in the immunofluorescence
staining assay, we found that FITC-RAP was perfectly co-localized with
GP2+ M-like cells (Fig. 3A). To figure out how much FITC-RAP was
transported by the M-like cell model, we used flow cytometry (FCM) to
detect the amount of FITC-RAP in the lower layer supernatant. Flow

3.2. Transport of intact RAP by M-like cells in vitro
We further tested the possibility that M cells may be the transporter
using an in vitro FAE cell model. As shown in Fig. 2A, we first evaluated
the expression of the M cell marker and validated its transporter func­

tion using control nanoparticles (Beloqui et al., 2017). Confocal images
of inserts showed that M-like cells were successfully characterized with

Fig. 1. Selective transport of RAP in PPs. (A) Images of PPs (n = 8) collected from mice in a ligated intestinal loop mouse model after FITC-RAP treatment using IVIS
Lumina XR Small Animal Imaging System. (B) Fluorescence intensity of the region of interest (ROI) of the ex vivo images taken above. (C) HPGPC-FLD chromatograms
of PPs with and without the FITC-RAP treatment for 1 h. (D and E) Flow cytometry dots (D) and percentage (E) of intestinal epithelial cells (IECs) in cell suspension
isolated from PPs and small intestine segments without PPs (SI). These tissues were collected from mice treated with FITC-RAP (10 mg/kg) by oral administration. (F
and G) Flow cytometry histograms (F) and percentage (G) of FITC-RAP-binding IECs gated from D. Error bar indicates SD (n = 10). Data are shown as mean ± SD and
are representative of at least three independent experiments. Significant difference ***P < 0.001, ****P < 0.0001.
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Fig. 2. Establishment and validation of the in vitro M cell-like model. (A) Protocol used to establish the in vitro M cell model and its application. (B) Confocal
microscopic images of co-culture transwell inserts stained with anti-GP2 mAb and subsequently with Alexa 647-conjugated secondary antibody (red). DAPI (blue)
was used as a DNA-specific stain. Scale bars, 50 μm. (C) Flow cytometry detection of nanoparticles in the supernatant collected from the lower layer of monoculture
and co-culture groups (n = 3). (D) Percentage of nanoparticles transported by the M cell model. Data are shown as mean ± SD. Significant difference ****P < 0.0001.
All in vitro experiments repeated at three times, respectively.

cytometry analysis of the lower layer of transwell inserts indicated much
more FITC-RAP was detected in the co-culture group, suggesting the
polysaccharide RAP was transported by M-like cells in the in vitro model
(Fig. 3B and C). Furthermore, we also detected whether FITC-RAP was
degraded in the transport process using HPGPC assay. The HPGPC
chromatograms clearly showed that the FITC-RAP in the lower layer
supernatants had the same molecular weight with primary FITC-RAP,
suggesting the transported FITC-RAP remained intact (Fig. 3D). By

contrast, the monolayer of Caco-2 cells was not permeable for FITC-RAP.
We also tested unlabeled RAP to exclude the effect of FITC using the
M-like cell model. Our previous studies have demonstrated that RAP
strongly induced cytokine production by macrophages, such as
RAW264.7 cell line, in a dose-dependent manner (Li et al., 2017; Wei
et al., 2016; Wei et al., 2019). Therefore, cytokine production of mac­
rophages here was used to detect the transported RAP in the lower layer
of the co-culture by comparing with RAP treatment at different con­
centrations. RAP standard solutions of varying concentrations (0.1, 1,
10, and 100 μg/mL) were used as control. The results indicated that in
terms of both IL-6 and IL-12 production, the transported RAP in the coculture group exhibited significant inducing effects (Fig. 3E and F),

which were comparable with 1 μg/mL of RAP standard solution. It is
suggested that M-cell's transport could be so efficient that the trans­
ported RAP could reach a concentration as high as 1 μg/mL.
3.3. Transport of RAP by M cells in vivo
The role of the M cell as the key transporter cell was further verified
in vivo. Firstly, we need to identify M cells in PPs tissues. Using wholemount staining of PPs, we found the dome zone of FAE in PPs
(Fig. 4A), which is the location of M cells. We compared two reported
specific M-cell markers, GP2 and NKM 16-2-4, using confocal micro­
scopy and found that M cells were successfully labeled with both
markers (Fig. 4B–D) (Hase et al., 2009; Nochi et al., 2007). We also
prepared frozen sections of PPs and identified GP2+NKM16-2-4+ M
cells in the FAE of PPs (Fig. 4E). The H&E staining image showed the
structure of the FAE of PPs (Fig. 4F), where M cells are located in. Thus,
these two specific markers, GP2 and NKM 16-2-4, were well validated
for definitely identifying M cells in vivo.
After identifying M cells in vivo, we tracked orally administrated
FITC-RAP. With the same method in Fig. 1D, we gated IECs for verifying
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Fig. 3. Transport of RAP in the M cell-like model. (A)
Confocal microscopic images of co-culture transwell
inserts treated with FITC-RAP (green) and then
stained with GP2 mAb and subsequently with Alexa
647-conjugated secondary antibody (red). DAPI
(blue) was used as a DNA-specific stain. Scale bars,
25 μm. (B and C) Flow cytometry histogram of FITCRAP (B) and transport percentage (C) in the super­
natant collected from lower layers of monoculture
and co-culture groups (n = 9 per group). (D) HPGPCFLD chromatograms of FITC-RAP in the supernatants
collected from lower layers of monocultures and cocultures (n = 9 per group). (E and F) IL-6 (E) and
IL-12 (F) secretion of RAW264.7 cells treated with
supernatants collected from the lower layer of
monocultures and co-cultures (n = 9 per group) 2 h
after loading 200 μL RAP (2 mg/mL), compared with
cytokines produced from RAW 264.7 cells treated
with RAP at different concentration (0.1, 1, 10, and
100 μg/mL). Data are shown as mean ± SD and are
representative of three independent experiments.
Significant difference *P < 0.05, ****P < 0.0001, ns
= no significance.

the transport role of M cells using the flow cytometry analysis. As shown
in Fig. 5A and B, results of flow cytometry showed that GP2+ cells in
IECs of PPs were much higher than that in IECs of SI, suggesting that

GP2+ M cells are located in the FAE of PPs. Further analysis of these
GP2+ M cells indicated that GP2+ M cells instead of other IECs accu­
mulated FITC-RAP (Fig. 5C and D). The specific transport of FITC-RAP
by GP2+ M cells was further demonstrated by confocal microscopic
images of fixed and stained whole-mount preparation (Fig. 5E) of PPs
isolated from mice 1 h after FITC-RAP treatment. Consistently, threedimensional confocal micrographs demonstrated the accumulation of
GP2 around FITC-RAP, which was internalized into M cells from the
apical side to the basal side (Fig. 6A–C and Supplementary movie 1).
Taken together, these data showed that M cells of PPs in the small in­
testine could independently transport RAP.
In our previous study, we reported that follicle dendritic cells (FDCs)
were RAP's direct target immune cells in the PPs. Here, we further
confirmed this finding using an immunofluorescence staining assay to
observe the co-localization of RAP, M-cell and FDCs. As shown in Fig. 6D
and Supplementary movie 2, GP2+ M cells in FAE are localized with

FITC-RAP in the FAE of PPs, and FITC-RAP is simultaneously localized
with CD11c+ DCs in the sub-epithelial dome, which is consistent with
the results of our previous flow cytometry analysis (Zhang et al., 2022).
These results precisely show that RAP will directly contact FDCs after
being transported into PPs.
3.4. Transport of RAP by M cells in the human distal ileum
As M cells exist in both mice and humans, we hypothesize that this
interesting M-cell mediated transcytosis may also occur in the human
body. So, we collected live distal ileum explants from surgery patients
and screened them for PPs (Fig. 7A and B) and incubated the ileum
containing PPs with FITC-RAP. M cells are located in the follicle asso­
ciated epithelium (FAE) of PPs (Sakhon et al., 2015). In photos of stained
whole-mount of sections, the FAE zone of PPs is denoted by a white
circle (Fig. 7C). To confirm the location of M cells and M cell-associated

RAP transport, we first collected PPs and stained with GP2 cell marker.
As shown in Fig. 7D, FITC-RAP was recruited of puncta, most of which
were positive for the M-cell marker GP2, suggesting GP2+ M cells
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Fig. 4. M cells in the small intestine recognized by
the specific markers GP2 and NKM 16-2-4. (A)
Confocal microscopic images of whole-mount PPs
stained with DAPI (blue). DAPI (blue) was used as a
DNA-specific stain. The location of FAE in PPs is
marked by a dotted white circle. Scale bars, 250 μm.
(B) Confocal microscopic images of the FAE area of
whole-mount PP stained with anti-NKM 16-2-4 mAb
and anti-GP2 mAb and subsequently with Alexa 488labeled (green) and Alexa 647-labeled (red) second­
ary antibody. Scale bars, 250 μm. (C) Confocal
microscopic image of the FAE area of whole-mount
PP stained with anti-NKM 16-2-4 mAb and anti-GP2
mAb. Scale bars, 75 μm. (D) The enlarged area was
marked in C by a dotted white frame. Scale bars, 25
μm. (E) Confocal microscopic images of frozen PP
tissue isolated from wild type BALB/c mice stained
with anti-NKM 16-2-4 mAb and anti-GP2 mAb and
subsequently with Alexa 488-conjugated (green) and
Alexa 647-conjugated (red) secondary antibody. Scale
bars, 25 μm. (F) Cryosections of ileal PPs stained with

H&E and visualized by light microscopy. The location
of FAE M cells in PPs is marked by a dotted white line.
Scale bars, 100 μm. Abbreviations: FAE, follicleassociated epithelium; SED, sub-epithelial dome; LF,
lymphoid follicle.

located in the FAE of PPs could transport FITC-RAP like what happed in
mice. The two signals were not coincidental, which might be due to the
dynamic transcytosis procedure. Three-dimensional images further
confirmed and demonstrated the dynamic transport of FITC-RAP in M
cells from the apical surface to the subepithelial dome of PPs (Fig. 7E
and Supplementary movie 3). Confocal microscopic images of frozen
sections of PPs confirmed that FITC-RAP has been transported into PPs
(Fig. 7F). These findings demonstrate that RAP is transported by human
M cells into PPs.

be determined if the transported polysaccharides were already
degraded. Few reports addressed this because the detection solely relied
on the fluorescence signal. As we found previously when we tracked the
orally dosed FITC-RAP in the caecum and colon using HPGPC-FLD, the
fluorescence signal might come from degraded chemicals. Fourth, the
unlabeled polysaccharide should be tested by using other biochemical
assays. In the current report, we also observed a weak HPLC signal in the
lower layer of the monoculture insert loaded with FITC-RAP. But that
loaded with unlabeled RAP was completely inactive in the ELISA assay.
Therefore, the weak signal observed FITC-RAP monoculture control
might be attributable to a false positive result. It has been reported that
this kind of weak signal always exists in the in vitro M-like cell model and
is hard to remove (Ahmad et al., 2017; Beloqui et al., 2017). It is also
possible that FITC labeling changed the transcytosis property of RAP in
the in vitro cell model; this possibility deserves further investigation. It is

suggested that the fluorescence signal is unreliable and needs to be
validated with other analyses. An isotype control of FITC might be
helpful to address this concern.
The popularly used in vitro M-like cell model has its own limitations.
This model does not simulate all factors in the transport of intestinal
antigens. The small intestinal epithelial cells are heterogenous and act as
a passive barrier to prevent the transit of antigens and pathogens
(Nagler-Anderson, 2001; Romero et al., 2015; Vancamelbeke & Ver­
meire, 2017). Some of the epithelial cells are specialized for producing
mucus as a vital physicochemical barrier (Grondin et al., 2020; Pela­
seyed et al., 2014). This model only addresses the similar transcytosis
function. Herein, using an in vivo model, we screened the small in­
testine's epithelial cells and proved for the first time that GP2+ M cell
was the specific transporter cell for polysaccharide RAP. The in vivo
study showed several advantages. First, the in vitro cell model includes
multiple time-consuming operations over 3 weeks, while the in vivo

4. Discussion
An in vitro M-like cell model has been used to evaluate whether M
cells transport polysaccharides (Feng et al., 2020; Li et al., 2021; Xiang
et al., 2020; Zheng et al., 2022), but the model needs validation. First, it
does not include a nanoparticle control to test the transport function
(Rieux et al., 2005; Rieux et al., 2007). The results would be doubtable
without control. Using the well-studied nanoparticle as a control will be
helpful to judge whether the model is successfully established. Second,
the presentation of M-like cells should be verified using a specific cell
marker. Actually, this cell model is often directly used to test for
transcytosis without determining if M-like cells existed in this model
(Gibb et al., 2021). Even when the M-like cells were labeled in a few of
these tests, the cell marker in labeling was not specific. For example,

Ulex europaeus agglutinin-1 (UEA-1) was used as the marker of M-like
cells (Jiang et al., 2019), but it had been found to be non-specific for M
cells (Terahara et al., 2008; Zheng et al., 2022). The current study is the
first in which GP2, which has been identified as a specific M-cell marker,
was used to test for polysaccharide transcytosis. In in vivo studies, we
even used two cell markers, GP2 and NKM 16-2-4 (Hase et al., 2009;
Nochi et al., 2007), to validate the existence of M cells. Third, it should
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Carbohydrate Polymers 296 (2022) 119952

Fig. 5. Transport of RAP by M cells in vivo. (A and B) Flow cytometry density plot (A) and percentage (B) of GP2+ M cells in intestinal epithelial cells (IECs), which
are gated in FSC-SSC flow cytometry dots by analyzing cells isolated from PPs and small intestine without PPs (SI). The gate strategy of IECs is same with the IECs
gate in Fig. 1D. These tissues (n = 10) are collected from mice treated with FITC-RAP (10 mg/kg) by oral gavage. (C and D) Flow cytometry histogram (C) and
percentage (D) of FITC-RAP-binding GP2+ M cells gated from A. (E) After the ligated intestinal loop assay with FITC-RAP, PPs were stained with anti-GP2 mAb and
DAPI (blue). Uptake of FITC-RAP (green) by GP2 + M cells (red). Scale bars, 25 μm. Data are shown as mean ± SD and are representative of three independent
experiments. Significant difference **P < 0.01, ****P < 0.0001.

assay could be quickly completed in 2–3 h. Second, the in vivo study
involved the real, multi-cellular organ/tissue environment rather than a
single type of cells. Third, the in vivo study enabled further observation
of which cell the polysaccharide directly targeted in PPs. Thus, the in
vivo approach we established here might be a better option to investigate
polysaccharides' intestinal lymphatic absorption through M-cell
transcytosis.
The M cell cell-mediated transcytosis of RAP might be receptormediated. There are three possible pathways for the M cell-mediated
antigen transcytosis: non-specific transcytosis, receptor-mediated

transcytosis, and extension of DCs through M cell transcellular pores
(Dillon & Lo, 2019; Kim & Jang, 2014; Lelouard et al., 2012; Mabbott
et al., 2013; Neutra et al., 2001). In our previous study, Dendrobium
officinale polysaccharide DOP showed an identical destiny after oral
administration (Li et al., 2019). Being indigestible and nonabsorbable, it
ended in regulating gut microbiota as it degraded to short-chain fatty
acids. DOP was used as a control in the tests of the lymphatic route for
RAP to enter PPs (Zhang et al., 2022). The results showed that only RAP
rather than DOP entered PPs. It is suggested that this lymphatic route is
selective. Here M cells have been identified as the specific transporter
cell in this route, so we hypothesize that RAP's transcytosis might be
mediated by some specific receptors of M cells. TLR4 is one possible
receptor because it has been reported to mediate the RAP-induced

signaling pathway of macrophages (Wei et al., 2016). GP2 is another
possibility because it is not only the specific M-cell marker but also the
receptor mediating the antigen sampling of PPs (Hase et al., 2009).
Furthermore, GP2 is common in both mice and humans, and we
demonstrated that human M cells have the same function in transporting
RAP. Therefore, we hypothesize that TLR4/GP2 is the receptor that
mediates the M-cell transcytosis of RAP. This hypothesis deserves
testing.
The M cell-mediated lymphatic route demonstrated here with human
subjects is scientific evidence for the potential medicinal value of many
immunomodulatory polysaccharides. Whether these polysaccharides
are found to be effective for mice is not necessarily relevant, although
mouse studies are common. There are big differences between the mice
model and the human clinical trial. Many promising drug candidates
that fail are called “mouse drugs” because they are eventually found to
be effective in mice but not humans. Despite this risk, mice models are

still the best choice for most medicine R&D projects due to the concerns
of cost and regulations. Therefore, there is a huge research gap between
the mice study and the clinical trial of polysaccharide-based medicines.
Efficacy in mice is not a guarantee of efficacy in humans. Another
problem is studies of mechanisms. It is hard to investigate the underlying
mechanisms in clinical trials. Here we demonstrated that the efficient M
cell-mediated lymphatic route is common between mice and humans,
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Carbohydrate Polymers 296 (2022) 119952

Fig. 6. Follicle dendritic cells (FDCs) are the direct target immune cell of M cell-transported RAP in PPs. (A) The X-Y angle of three-dimensional confocal image of
whole-mount PP stained for anti-GP2 mAb (red) and DAPI (blue). PPs were isolated from the ligated intestinal loop assay with FITC-RAP (green). The threedimensional movie was shown in Movie S1. (B) Clockwise 45 degree-rotation image of A. (C) Clockwise 45 degree-rotation and amplified image of B. In this
image, FITC-RAP was transcytosed by an M cell from the apical side to the M cell pocket which marked by a dotted line. (D) Confocal images of PP frozen sections
(dome zone), collected from the ligated intestinal loop assay treated with FITC-RAP (green), and stained with GP2 (red), CD11c (yellow), and DAPI (blue). Threedimensional image of PP indicating that FITC-RAP (green) was transported into a sub-FAE layer and contacted CD11c+ dendritic cells (yellow). Arrows indicate
FITC-RAP. The three-dimensional movie was shown in Supplementary movie 2.

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Carbohydrate Polymers 296 (2022) 119952

Fig. 7. Transport of RAP by M cells of human ileum PPs. (A) Section of human distal ileum with PPs indicated by a black arrow. (B) Enlarged view of the distal ileum
tissue section in A, with a black arrow pointing to a PP (2.5×). (C) Confocal microscopic images of the whole-mount PP domes stained with DAPI (blue), as a DNAspecific stain. The dome zone, which is the location of FAE in PPs, is marked by a dotted white circle. Scale bars, 100 μm. (D) Confocal microscopic images of the
whole-mount PP domes stained with anti-human GP2 antibody and DAPI, showing that FITC-RAP (green) was bound to GP2+ M cells (red). Scale bars, 100 μm. (E)

Three-dimensional confocal microscopic images of FAE. FITC-RAP (green) was internalized by human GP2+ M cells (red). The three-dimensional movie was shown
in Supplementary movie 3. Scale bar, 8 μm. (F) Confocal microscopic image of frozen sections of human ileal PPs stained with DAPI (blue). FITC-RAP (green) was
found in the SED of PPs. Scale bars, 25 μm. Abbreviations: FAE, follicle-associated epithelium; SED, sub-epithelial dome; LF, lymphoid follicle.

successfully bridging this research gap. RAP has been verified to have
diverse beneficial effects, such as antitumor, but it has never been tested
on humans. Considering that most of RAP's bioactivities are associated
with the immune system and that RAP has been known to quickly enter
PPs and directly target FDCs to trigger immune responses, we

hypothesize that RAP follows the same lymphatic route to exhibit
similar immune-related beneficial effects in human bodies. In other
words, RAP's bioactivities and mechanisms observed in the mice model
should also apply to human bodies. And this route may be the way many
other active polysaccharides work. This finding may attract great
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Carbohydrate Polymers 296 (2022) 119952

interest in developing polysaccharide-based medications.

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In summary, our studies provide compelling evidence that M cells in
the FAE of PPs mediate the transcytosis of the Radix Astragali poly­
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Supplementary data to this article can be found online at https://doi.
org/10.1016/j.carbpol.2022.119952.
CRediT authorship contribution statement
Quanwei Zhang: Conceptualization, Data curation, Formal analysis,
Investigation, Methodology, Software, Visualization, Writing – original
draft. Shuang Hao: Methodology, Formal analysis, Visualization.
Lifeng Li: Methodology, Formal analysis, Visualization. Man Liu:
Methodology, Formal analysis, Visualization. Chuying Huo: Method­
ology, Formal analysis, Visualization. Wanrong Bao: Methodology,
Formal analysis, Visualization. Huiyuan Cheng: Methodology, Formal
analysis, Visualization. Hauyee Fung: Methodology, Formal analysis,
Visualization. Tinlong Wong: Methodology, Formal analysis, Visuali­
zation. Wenjie Wu: 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 & editing. Aiping Lyu: Writing – review & editing. Quanbin
Han: Conceptualization, Funding acquisition, Project administration,
Resources, Writing – original draft, 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.
Data availability
No data was used for the research described in the article.
Acknowledgments
This work was funded and supported by HKSAR Innovation and

Technology Fund (ITF, ITS/311/09), General Research Fund
(12100615, 22100014, 12100818, 12101322), UGC Research Matching
Grant Scheme (2019-1-10, 2019-1-14, 2019-2-06), Health Medical
Research Fund (11122531, 14150521, 17182681), National Natural
Science Foundation of China (81473341, 82173948), the Science and
Technology Project of Shenzhen (JCYJ20160531193812867), the KeyArea 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-20-21-2, FRG2/17-18/060 and FRG2/16-17/002), and Vin­
cent & Lily Woo Foundation.

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