Abstract
Metronomic photodynamic therapy is a long-term, low-dose treatment
strategy that employs optical devices with continuous photosensitizer
administration and requires stable device attachment with a consistent
power source. These factors significantly limit patient mobility.
Currently, no metronomic photodynamic therapy modality can operate
independent of external devices, underscoring the critical need for in
vivo light sources that function without external energy inputs. In
this study, we integrate self-luminous bacteria with a photosensitizer
in alginate microcapsules to create a self-driven metronomic
photodynamic therapy that can be securely implanted within a tumour,
thereby enabling continuous light emission without requiring an
external energy source or ongoing replenishment of photosensitive
reactants. By harnessing nutrients from the tumour microenvironment,
this system sustains the generation of reactive oxygen species. A
single injection effectively eliminates larger tumours (>300 mm^3) in
an opaque melanoma mouse model and transplanted hepatocarcinoma rabbit
model. Self-driven metronomic photodynamic therapy demonstrates
advantages over traditional photodynamic therapy, indicating its
potential as a versatile therapeutic approach for cancer treatment with
deeply situated lesions.
Subject terms: Cancer therapy, Biomaterials, Nanotechnology in cancer
__________________________________________________________________
Metronomic photodynamic therapy is a long-term, low-dose treatment
strategy limited by the need of continuous photosensitizer
administration. Here this group reports combining the
self-bioluminescent bacteria with a neutral red photosensitizer in
alginate microcapsules as a self-driven metronomic photodynamic
therapeutic system with preclinical anti-cancer effects.
Introduction
Recently, significant progress has been made in the development of
light-based technologies for diagnosis, therapy, and surgical
interventions^[44]1. Photodynamic therapy (PDT) is an efficacious and
clinically validated treatment for cancers within the superficial
layers of various organs, including the oesophagus, stomach, lung, and
cervix^[45]2–[46]4. In standard clinical practice, optical fibres
channel light towards internal lesions, delivering short-term,
high-intensity photoirradiation (lasting several tens of minutes at
intensities >100 mW/cm^2)^[47]5,[48]6. However, excessive light
exposure can induce thermal damage to tissues, causing necrosis and
inflammation that potentially compromise organ functionality^[49]7.
Conversely, inadequate light exposure fails to produce the desired
anti-tumour effects^[50]8. Thus, maintaining an optimal balance between
minimising radiation-induced harm and effectively eliminating cancer
cells remains a major challenge in PDT.
To improve both the safety and effectiveness of PDT, an method called
metronomic photodynamic therapy (mPDT) has been developed^[51]9. This
approach involves prolonged, low-intensity photoirradiation sessions
(ranging from a few hours to several tens of hours at intensities
<10 mW/cm^2), which markedly diminishes the likelihood of thermal
damage while effectively eliminating cancer cells. Nevertheless, mPDT
requires continuous administration of a photosensitizer (PS) and relies
on an implantable PDT device powered by batteries^[52]10 and
wireless^[53]11 technology, potentially complicating treatment. To
date, no mPDT modalities that can function without external supporting
devices exist, highlighting the urgent need for in vivo light sources
that can operate independently of external energy sources.
Currently, bacteria-based bioactive drugs are receiving increasing
attention for their tumour-targeting capabilities and roles in
enhancing immune responses^[54]12,[55]13. Drawing inspiration from the
bioluminescent organs of deep-sea anglerfish, scientists discovered
bioluminescent bacteria capable of sustaining light emission by
utilising host nutrients regulated by quorum sensing^[56]14. Although
these bacteria exhibit the potential for self-driven photodynamic
treatment, maintaining the safety, stability, and functionality of
bacteria-based optical devices in vivo poses a substantial challenge
for the practical deployment of fully implantable mPDT systems^[57]15.
Therefore, employing this strategy for internal tumour treatment
requires stable fixation of bioluminescent bacteria and PS to enable
continuous, localised light delivery and the generation of reactive
oxygen species (ROS).
In this work, we combine bioluminescent bacteria and Neutral red (NR)
photosensitizer^[58]16 within alginate microcapsules (MCs)^[59]17 to
develop the self-driven metronomic photodynamic system (Sd-PDT) for
cancer therapy. This system diverges from traditional mechanical- and
chemical-driven PDT methods by enabling continuous, uniform light
emission without external energy support or the continuous
replenishment of photosensitive reactants, and produces prolonged,
low-dose light emissions, sustaining continuous emission for
approximately 50 h. Grafting NR onto the surface of bacteria@alginate
microcapsules (PB@MCs) significantly prolongs self-supported ROS
generation. The effectiveness of PB@MCs is evaluated in four cancer
cell lines and two animal tumour models^[60]18. A single injection of
PB@MCs is sufficient to effectively eradicate large tumours within 2
days. Additionally, our results demonstrate that Sd-PDT through PB@MCs
triggers a strong antitumour immune response, effectively inhibiting
tumour metastasis and recurrence. This study highlights the potential
of Sd-PDT as a therapeutic strategy applicable to certain tumours,
offering promising clinical prospects.
Results
Engineering and characterisation of PB@MCs
Initially, our experimental approach involved constructing the
anticipated PB@MCs by subjecting bioluminescent bacteria to
physiological conditions, encapsulating them within MCs, and chemically
grafting PS molecules onto the surfaces of the MCs (Fig. [61]1A). We
selected the bioluminescent bacteria Vibrio harveyi BB170 (V.H.BB170),
Aliivibrio fischeri-bio115653 (F.A.115653) and Aliivibrio fischeri-7744
(F.A.7744) (Supplementary Fig. [62]1), which are typically cultured in
2216E medium at 25 °C. These three bacterial strains were evaluated for
viability and bioluminescent activity under physiological conditions.
Acclimatisation was gauged by measuring the bacterial density using the
optical density (OD) at 600 nm (Supplementary Fig. [63]2). Despite the
pronounced suppression of growth and bioluminescence in F.A.115653 and
F.A.7744 at elevated temperatures from 25 °C to 37 °C, V.H.BB170
demonstrated adaptability, maintaining standard growth and luminescent
output. Consequently, V.H.BB170 bacteria were selected for the further
development of PB@MCs (Fig. [64]1B and Supplementary Fig. [65]2). For
photosensitisation and initiation of ROS generation,
N-hydroxysuccinimide neutral red (NHS-NR) and N-hydroxysuccinimide
chlorin e6 (NHS-Ce6) photosensitizer are both commonly used PS
molecules. Since the V.H.BB170 strain emits robust bioluminescence
within the 400–600 nm range, we compared the UV–Vis absorption spectra
of these two PS molecules to identify the one best matching the
bacterial emission range. As shown in Supplementary Fig. [66]3, the
emission of V.H.BB170 ideally aligns with the absorption range of
NHS-NR. Additionally, NR functions as a Type I photosensitizer,
primarily generating hydroxyl radicals (•OH)^[67]19,[68]20, while Ce6
acts as a Type II photosensitizer, mainly generating singlet oxygen
(¹O[2]). The generation of •OH is less dependent of oxygen
availability, suggesting that NR, as a Type I photosensitizer, may
offer greater advantages in hypoxic tumour regions.
Fig. 1. Fabrication, demonstration, viability, stability, and
self-luminescence duration of PB@MC living composites.
[69]Fig. 1
[70]Open in a new tab
A Diagram outlining PB@MC construction. B Bioluminescence (BL)
intensity of three types of bioluminescent bacteria at 37 °C (n = 3
independent experiments). The data are presented as mean ± SD.
Differences were considered statistically significant at ***p < 0.001,
compared to the data for the F.A. 115653 and F.A. 7744 groups according
to one-way ANOVA test combination with Tukey’s multiple comparison test
performed using GraphPad Prism 9 XML project software. C Representative
images of B@MCs both with and without PLL coating. Coated and uncoated
PLL-B@MCs were cultured in 2216E medium for 8 h before collection for
microscopy. D Growth curves for B@MCs and PLL-B@MCs cultured in 2216E
medium at 37 °C for 14 h. Microcapsules (50 µL) were gathered within
96-well transparent plates, and their optical density at 600 nm was
measured using a microplate reader (n = 6 independent experiments). The
data are presented as mean ± SD. E Confocal imaging of PB@MCs. Empty
MCs, MCs containing bacterial cells (B@MCs), and fully constructed
PB@MCs were imaged for BL using confocal microscopy with a 400–500 nm
channel. F BL emission spectrum of B@MCs and PB@MCs. 500 µL samples
were cultured in equal volumes of DMEM to analyse BL emission spectra
from 300 to 700 nm using an Edinburgh FS5 spectrophotometer. The
spectra were analysed using Gaussian function fitting on OriginLab 8.0
software. G BL images of PB@MCs in various media. PB@MCs were cultured
in PBS, DMEM, and B16 tumour homogenate (BTM) in transparent penicillin
bottles at 37 °C for 32 h in vitro. Photos were taken with a 3.2 s
exposure. Medium was refreshed at 12 and 26 h (blue arrow). H In vitro
experiments to investigate persistent irradiation energy (mW/cm²)
capabilities of PB@MCs. PB@MCs (100 µL, 3.6 × 10⁴/mL) were cultured in
DMEM medium in 96-well black plates at 37 °C. BL intensity was measured
at 450 nm for 50 h, medium was refreshed at 12 and 26 h (orange arrow).
Source data are provided as a Source data file.
To develop a durable and persistent Sd-PDT system in vivo, an ionically
crosslinked alginate hydrogel was selected as the matrix owing to its
well-known biocompatibility with cell culture^[71]17,[72]21. PB@MCs
were engineered by encapsulating bioluminescent bacteria within MCs via
electrostatic attraction using an electrostatic droplet generation
system. PS molecules were chemically grafted onto the surface of the
MCs as follows (Fig. [73]1A): First, the sodium alginate hydrogel
precursor was blended with V.H.BB170 bacteria to achieve a uniform
mixture. Second, the solution was dispersed into consistent droplets
under an electrostatic attraction, and the droplets were immersed in a
CaCl[2] solution. Ca^2+ ions permeated the droplets, effectuating ionic
crosslinking of alginate chains and encapsulation of the bacterial
cells within the hydrogel matrix (denoted as the B@MC composite).
Third, the alginate-calcium microspheres were coated with poly-L-lysine
(PLL), a water-soluble polycation that is resistant to enzymatic
breakdown and prevents microbial escape^[74]17 (Fig. [75]1C and
Supplementary Figs. [76]4 and [77]5). The morphology and size of the
B@MCs prepared by an electrostatic attraction were controlled to be
within 150 µm, making this small size particularly suitable for
engineering biocompatible Sd-PDT for internal tumours. These PLL-B@MCs
were further cultured in 2216E medium at 37 °C, allowing the bacteria
to proliferate to the platform stage (Fig. [78]1D). Fourth, PLL, which
comprises abundant amino functional groups (−NH[3]) enabled subsequent
chemical conjugation. The reaction between PLL and NHS-NR formed
covalent amide linkages via the active NHS ester, which interacted
specifically with the −NH[3] group. Subsequently, the NR was
efficiently conjugated to the MCs, resulting in the formation of
PB@MCs. The bacterial density and PS (NHS-NR) binding affinity of each
PB@MC were thoroughly assessed by plotting the linear relationship
between bacterial OD and CFU/mL (Supplementary Fig. [79]6) and
measuring the absorbance of unreacted NHS-NR at 452 nm after collecting
the resultant solution (Supplementary Fig. [80]7). Upon calculation,
the formula for the average of PB@MC (1 × 10^3 bacteria cells and
0.028 μg of PS per MC) was established.
Confocal laser scanning microscopy (CLSM) was used to examine PB@MCs.
Notably, while empty MCs displayed no bioluminescence, microspheres
containing bioluminescent bacteria, specifically B@MCs and PB@MCs,
emitted vivid blue light. This demonstrated that the PS coating on the
surfaces of the MCs did not affect the light emission ability of the
bacteria and that the bacteria were capable of independent light
emission without external excitation while efficiently converting
nutrients in the culture medium into light (Fig. [81]1E). These results
confirmed that the encapsulated bioluminescent bacteria maintained
their light emission characteristics. Confocal microscopy analysis
demonstrated that bacterial cells within the MCs self-assembled into
structurally stable aggregates while maintaining sustained
bioluminescent activity, with no detectable bacterial bioluminescence
signals outside the MCs (Fig. [82]1E). Furthermore, we evaluated the
emission spectra of B@MCs before and after PS modification.
Predictably, both groups of encapsulated bioluminescent bacteria
exhibited a Gaussian emission profile centred at ~480 nm with a full
width at half-maximum of 100 nm (Fig. [83]1F).
In practical applications, PB@MCs can be deployed as long-term Sd-PDT
devices in tumour microenvironments. However, maintaining long-lasting
light emission poses a challenge in vitro because persistent
luminescence requires continuous nutrient availability. Once nutrients
in the culture medium are depleted, bacterial luminescence
decreases^[84]22. To mimic the nutritional environment within actual
tumours, in vitro experiments were conducted with frequent
replenishment of the culture medium to simulate a continuous nutrient
supply. PB@MCs were evaluated for their long-term bioluminescence
capabilities in physiological solutions and various medium
formulations, including Dulbecco’s modified Eagle medium (DMEM) and B16
tumour homogenate (BTM). As shown in Fig. [85]1G, when bioluminescence
diminished, immediate supplementation of fresh culture medium
revitalised the light emission of PB@MCs, which was maintained for 32 h
in transparent bottles. A laser power meter was used to plot the linear
relationship between the bioluminescence intensity (10⁴ cps) and the
power density (mW/cm²) of PB@MCs (Supplementary Fig. [86]8), and
further calculations revealed that PB@MCs can emit bioluminescence at a
power density of 1–5 mW/cm² in DMEM medium for up to 50 h
(Fig. [87]1H), which is suitable for the mPDT photoirradiation
dose^[88]23 and significantly surpasses the duration of all currently
available chemical-driven PDTs^[89]14.
Anti-tumour cells and ROS generation test of PB@MCs
We explored the efficacy of Sd-PDT using PB@MCs on cancer cells. This
study assessed the effect on four distinct cancer cell lines, two liver
cell lines (Hep3B and VX2) and two melanoma cell lines (A375 and B16).
The 24-Transwell and 6-Transwell setups prevented direct physical
contact between the PB@MCs and cancer cells, facilitating the
assessment of the Sd-PDT effects. PB@MCs and cancer cells were placed
in the upper chamber and the lower chamber, respectively (Fig. [90]2A).
Cell viability was assessed using the LIVE/DEAD staining assay
(Calcein/PI Assay Kit) and visualised using CLSM. After 8 h, a
significant red signal increase in cell mortality was observed in all
cell lines in the PB@MC group, suggesting a potent cytotoxic effect
(Fig. [91]2B and Supplementary Fig. [92]9). In contrast, the cells
treated with B@MCs and P@MCs, which contained only bacteria or PS,
maintained over 90% cell viability, indicating their lower toxicity.
Cellular metabolic functions were further analysed using a colorimetric
CCK-8 assay (Fig. [93]2C and Supplementary Fig. [94]10) and aligned
with the results from LIVE/DEAD staining (Calcein/PI Assay Kit),
demonstrating significant cytotoxic effects in the PB@MC groups across
all tested cancer cell lines. Apoptosis induced by PB@MCs was
investigated using flow cytometry (Fig. [95]2D, E and Supplementary
Fig. [96]11), Annexin V-FITC(+) and DAPI(−) cell populations are
defined as early apoptotic cells, while Annexin-FITC(+) and DAPI(+)
cell populations are defined as late apoptotic cells. Toposide
VP16^[97]24, a typical anti-tumour drug, was used as a positive
control. The proportions of cells exhibiting late apoptosis (DAPI
fluorescence signal) were 2.67%, 9.19%, 19.5% and 43.3% in the Control,
P@MCs, B@MCs and PB@MCs groups, respectively, demonstrating that
cytotoxic effects were predominantly observed in cells treated with
PB@MCs.
Fig. 2. Anti-tumour efficacy of PB@MC living composites.
[98]Fig. 2
[99]Open in a new tab
A Diagram depicting the anti-tumour action of PB@MCs using a Transwell
model. Images are created by figdraw.com. B Fluorescence microscopy
images following LIVE/DEAD staining (Calcein/PI Assay Kit) in vitro.
Hep3B and VX2 cells were treated with 200 µL of either PBS, Empty MCs
linked to photosensitizer (P@MCs, contain 0.22 μmol NR), MCs
encapsulating V.H.BB170 bacteria (B@MCs), or PB@MCs for 8 h, followed
by staining with a LIVE/DEAD kit for microscopy visualisation. Red
fluorescence represents dead cells and green fluorescence represents
live cells. Scale bars: 100 µm. C In vitro cell viability assessment.
Hep3B and VX2 cells were exposed to 200 µL of P@MCs, B@MCs, or PB@MCs
in dark conditions for 8 h. Viabilities were determined using an CCK-8
assay (n = 3 independent experiments). The data are presented as
mean ± SD. Statistical significance is noted with ***p < 0.001 and
**p < 0.01, compared to the data for P@MC, B@MC and control group
according to one-way ANOVA with Tukey’s multiple comparisons performed
using GraphPad Prism 9 XML project software. D In vitro test of
apoptotic cell detection via flow cytometry. Representative images and
E quantitative analysis. B16, A375, VX2 and Hep3B cells (5 × 10⁵
cells/well) in 6-Transwell plates were treated with bare B@MCs, P@MCs
(contain 0.44 μmol NR), and PB@MCs at concentrations of 400 µL or
400 µg/mL VP-16. Following 8 h of incubation, Annexin V-FITC(+) and
DAPI(−) cells are defined as early apoptotic cells, whereas
Annexin-FITC(+) and DAPI(+) cells are defined as late apoptotic cells
for analysis by flow cytometry. Source data are provided as a Source
data file.
To verify that these anti-tumour effects were predominantly
attributable to ROS production, we analysed the ROS generation
capabilities of PB@MCs both in vitro (physiological solutions) and in
vivo (tumour environments). For in vitro testing, PB@MCs were incubated
in 24-well Transwell at 37 °C for 2 h, with no treatment (vehicle
control), and B@MCs and P@MCs as control groups. The ROS levels in
Hep3B and A375 cells were assessed using
2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA) staining^[100]25,
which measures ROS activity by detecting the oxidation of
nonfluorescent DCFH to fluorescent 2′,7′-dichlorodihydrofluorescein
(DCF) (Fig. [101]3A and Supplementary Fig. [102]12). In contrast to the
limited signals from the cells treated with B@MCs, intense fluorescence
was observed in the cells treated with PB@MCs, indicating more ROS
generation. We also evaluated the bioluminescence performance and
duration of PB@MCs in mice. As shown in Supplementary Fig. [103]13,
PB@MCs exhibited bioluminescent signals in tumours for at least 10 h,
confirming their bioluminescent capability within hypoxic tumour
microenvironments. However, the bioluminescent signals of PB@MCs
gradually attenuated over time, with signal weakening initiating from
the tumour core and progressively diminishing toward the periphery.
This phenomenon may be attributed to the hypoxic gradient within
tumours, where oxygen deprivation intensifies toward the core while
relatively oxygenated conditions prevail in the peripheral regions.
Consequently, stronger self-luminescent signals were observed in areas
distal to the tumour centre. The in vivo bioluminescence duration of
PB@MCs was shorter than the 50-h duration observed in vitro, possibly
because the signal intensity fell below the IVIS system’s detection
limit (potentially influenced by melanin absorption in melanoma and
limited tissue penetration depth), or more likely due to ongoing oxygen
consumption during sustained bacterial bioluminescence within tumours.
Following the experiment, tumours were aseptically harvested, weighed,
and processed according to the Tissue •OH assay kit (BBoxiProbe O28
probes)^[104]26 to evaluate the in vivo •OH levels. As shown in
Fig. [105]3B, mice treated with PB@MCs exhibited significantly higher
fluorescence intensity compared to the other groups, demonstrating
enhanced •OH generation and confirming the in vitro findings. To verify
that the cumulative ROS production depends on the bioluminescence
intensity and exposure duration of PB@MCs, ROS generation was
continuously monitored for approximately 50 h. The duration and
intensity of ROS generated by Sd-PDT were compared with those of
conventional PDT (external LED at 300 mW/cm^2 irradiating NR for 1 h
per session). For the Sd-PDT group, PB@MC microspheres loaded with an
equivalent NR dose were used. ROS fluorescence intensity was quantified
using the DCFH probe at each time point. As shown in Fig. [106]3C, the
LED group exhibited a significant increase in ROS fluorescence signals
during 1-h irradiation, indicating rapid and strong initial ROS
generation. However, ROS production ceased immediately after
irradiation ended. Conversely, the PB@MC group displayed persistent ROS
generation throughout the treatment session (~50 h), despite lower
initial fluorescence intensity compared to the LED group. These results
confirm the sustained ROS production of the Sd-PDT system in a single
treatment session, reflecting their persistent effectiveness in the
tumour environment. The cumulative ROS quantity in the PB@MC group
significantly exceeded that of the LED group (Fig. [107]3D). These
findings provide substantial evidence of the durable ROS-generating
capacity of PB@MCs.
Fig. 3. Characterisation of ROS generation efficiency in vitro and in vivo by
PB@MCs.
[108]Fig. 3
[109]Open in a new tab
A Confocal microscopy of Hep3B cells using DCFH-DA probe in vitro.
Hep3B cells, seeded in 24-Transwell plates at 5 × 10^4 cells/well, were
treated with 100 μL P@MCs, B@MCs and PB@MCs and subsequently stained
with the ROS-specific probe DCFH-DA after 2 h. B In vivo detection of
ROS content. ROS production was assessed using the Tissue Hydroxyl
Radicals Assay Kit (BBoxiProbe O28 probe). For comparison, B16
tumour-bearing mice were treated with either 50 μL PBS (control group),
empty MCs linked to photosensitizer (P@MCs, contain 0.055 μmol NR), MCs
encapsulating V.H.BB170 bacteria (B@MCs), or PB@MCs for 10 h; 190 μL of
B16 tumour homogenate (50 mg/mL) and 10 μL of BBoxiProbe O28 working
solution were added to a 96 black well plate, the plate was incubated
at 37 °C in darkness for 20 min, and fluorescence intensity was
detected using a microplate reader (excitation at 488 nm, emission at
520 nm) (n = 3 independent experiments). The data are presented as
mean ± SD. Statistical significance is noted with ***p < 0.001,
compared to the data for the P@MC, B@MC and control group according to
one-way ANOVA with Tukey’s multiple comparisons performed using
GraphPad Prism 9 XML project software. C ROS generation duration test
in vitro by PB@MCs (30 μL, 3.6 × 10^4/mL). Fluorescence intensity of
DCF was monitored ~50 h using a microplate reader (n = 3 independent
experiments). The LED radiation group (300 mW/cm² for 60 min LED
radiation of with an equivalent dose of NHS-NR, 0.32 mM in 100 μL
medium) was used for comparison. D Quantitative analysis of ROS
generation after a single treatment of PB@MCs and LED group (n = 3
independent experiments). The data are presented as mean ± SD.
Statistical significance is noted with **p < 0.001, compared to the LED
radiation group according to two tailed t-test using GraphPad Prism 9
XML project software. Source data are provided as a Source data file.
Metabolism analysis of tumours after PB@MCs treatment
In addition to the effects of ROS, metabolomic analyses were performed
to elucidate the anti-tumour mechanisms of PB@MCs. Melanoma
tumour-bearing mice treated with PB@MCs, B@MCs, PS and PS with LED
radiation were further investigated to decipher the intrinsic
mechanisms through which PB@MCs impede tumour progression utilising
metabolomic detection and analysis, focusing on specific pathways of
metabolic reorganisation within the tumour microenvironment
post-treatment^[110]27,[111]28. Using non-targeted metabolomics, the
levels of various metabolites in the tumour microenvironment after
PB@MC treatment were compared with those in the control groups.
Principal component analysis highlighted the metabolite distribution
profiles across different treatments (Fig. [112]4A). PB@MCs induced
significant alterations in the metabolic profile, markedly
distinguishing these profiles from those of the other groups,
indicating the potential metabolic reorganisation induced by PB@MCs. As
shown in Fig. [113]4B, a Venn diagram revealed that the metabolite
composition among the groups was similar, suggesting that PB@MCs may
inhibit tumour progression by suppressing the expression of components
within the same metabolic pathways. Supporting this hypothesis, volcano
plot analysis (Fig. [114]4C) identified 90 significantly downregulated
and 50 upregulated components in the PB@MC group compared to those in
the control group (fold change (FC) > 2 or <0.5 and p < 0.05). We
further performed cluster analysis of the metabolites derived from
metabolic profiling. As shown in Fig. [115]4D, metabolite expression
levels from tumour tissues under different treatments were standardized
using Z-score normalization. The top 30 metabolites ranked by relative
abundance were selected for the clustered heatmap. The results
demonstrate significant differences in metabolite expression between
the PB@MC-treated group and the other groups. PB@MCs significantly
reduced the metabolites involved in critical pathways such as glucose
and lipid metabolism, including glucose 6-phosphate,
phosphatidylethanolamine (Pe), glyceraldehyde and phosphatidylcholine
(Pc). Glucose-6-phosphate plays a central role in carbon metabolism by
linking glycolysis to the pentose–phosphate pathway. Pc and Pe are the
key components of eukaryotic cell membranes. The decrease in these
metabolites suggests that PB@MC treatment reduced the utilisation of
carbon sources by tumour cells and the synthesis of NADPH and nucleic
acid-5-phosphate. This disrupts the redox balance in tumour cells,
leading to high levels of ROS and impeding the synthesis of
macromolecules such as nucleotides and glycerolipids that interfere
with tumour cell proliferation. Pathway enrichment analysis of
differential metabolites using the Kyoto Encyclopedia of Genes and
Genomes (KEGG) database revealed a strong concentration of lipid and
amino acid metabolism pathways (Fig. [116]4E). Further analysis
revealed that metabolic pathways closely associated with glutathione
metabolism, including cysteine and methionine metabolism, biosynthesis
of cofactors, ABC transporters, and thyroid hormone synthesis, were
significantly suppressed following PB@MC treatment, contributing to
increased ROS stress and antitumour effects (Fig. [117]4F and
Supplementary Fig. [118]14). Additionally, phospholipid biosynthesis
pathways, including glycerolipid and sphingolipid metabolism pathways,
were notably downregulated. Because glycerolipids and sphingolipids are
crucial for maintaining cell membrane integrity, their downregulation
limits the supply of the essential membrane components required for
tumour cell expansion and repair, thereby enhancing ROS-mediated
cytotoxicity. Moreover, pathways related to energy and nucleotide
metabolism, such as oxidative phosphorylation, citric acid cycle,
glutamate metabolism, and purine–pyrimidine metabolism, were markedly
downregulated, further inhibiting the energy supply and macromolecule
production necessary for tumour cell proliferation. The significant
metabolic changes induced by PB@MCs suggest that their anti-tumour
effects are mediated by the induction of oxidative stress, inhibition
of energy metabolism, and suppression of synthesis and repair
mechanisms in tumour cells.
Fig. 4. Metabolism analysis of tumours after PB@MCs treatment.
[119]Fig. 4
[120]Open in a new tab
A Principal component analysis. Orange ellipses indicate PB@MCs, and
other colours indicate control groups. The metabolic profiles of the
PB@MC and control groups were highly distinct. B Venn diagram of
metabolites of PB@MCs, B@MCs, PS, PS-LED and Control groups (n = 5
independent experiments). C Volcano plot of metabolites. Down and
upregulated metabolites are marked by blue and red dots, respectively,
and the grey dots represent metabolites with no significant difference.
The vertical dotted line denotes |log2(fold change)| = 1, and the
horizontal dotted line shows -lg(p value) = −lg(0.05). p values are
determined by two-sided student’s t-test. D Cluster analysis of the top
30 differential metabolites. Each column represents a sample, and each
row represents a metabolite. The heat map value shows the relative
expression level (n = 5 independent experiments). E KEGG compound
analysis. Bar colour distinguishes the category of the pathways. F KEGG
enrichment analysis of metabolites. −lg(p value) > −lg(0.05) is
considered significant. Each animal was administered a single
intratumoural injection. After 24 h, tumour samples were then stored in
liquid nitrogen for subsequent metabolomics analysis. p-values are
determined by two-sided Student’s t-test and made for Benjamini and
Hochberg FDR (BH) adjustments. The metabolism analysis data reported in
this paper have been deposited on Zenodo Dataverse (DOI
10.5281/zenodo.16792981).
Immunology analysis of PB@MCs in tumour
PB@MC-induced Sd-PDT can cause effective immunogenic cell death (ICD)
in cancer cells, promoting the expression of damage-associated
molecular patterns (DAMPs)^[121]29. This process enhances the
maturation and recruitment of DCs^[122]30 to the tumour
microenvironment. Subsequently, mature DCs activate cytotoxic T
lymphocytes, thereby intensifying the anti-tumour immune responses.
Among the critical DAMPs for ICD, calreticulin (CRT) is particularly
significant for DC activation. As illustrated in Fig. [123]5A, B16
cells treated with PB@MCs, B@MCs, PS and PS-LED displayed CRT
translocation from the endoplasmic reticulum to the cell membrane, with
expression levels 6-, 1.35-, 1.85- and 1.8-fold higher than those
observed in the control group. A similar pattern was observed in the
VX2 cells (Fig. [124]5B), where the CRT levels were 1.85-, 1.33-, 1.4-
and 1.25-fold higher than those in the control groups. The
concentration of interferon-γ (IFN-γ), a cytokine secreted by T cells,
was measured in mouse serum using an enzyme-linked immunosorbent assay.
Treatment with PB@MCs led to a pronounced increase in IFN-γ levels,
indicating significant immune activation (Fig. [125]5C). Additionally,
the maturation of DCs, marked by increased expression of CD80 and CD86,
was quantified using flow cytometry. Compared to the control group, the
proportion of mature DCs in the PB@MC-treated group significantly
increased from 25% to 60% in vitro (Fig. [126]5D and Supplementary
Fig. [127]15). These results indicate that PB@MCs effectively modified
the immune microenvironment to suppress tumour growth. Encouraged by
the promising responses observed in vitro, we further explored the in
vivo effects of this immunotherapy. Intratumoural infiltration of T
lymphocytes was assessed using flow cytometry. The PB@MC group showed a
substantial increase in the presence of T cells within the tumour,
particularly CD8^+ T cells, which reached ~33% of total CD3^+ cells in
tumours treated with PB@MCs, which was significantly higher than that
in other groups (p < 0.001) (Fig. [128]5E and Supplementary
Fig. [129]16). Consequently, the CD8^+/CD4^+ ratio increased
(Fig. [130]5F).
Fig. 5. In vitro and in vivo demonstration of immune response by PB@MCs.
[131]Fig. 5
[132]Open in a new tab
A CRT expression levels in B16 and B VX2 cells after various treatments
as analysed by flow cytometry in vitro. After 6 h post-treatment, flow
cytometry was used to quantify the mean fluorescence intensities for
the PB@MC, a PS (NHS-NR, 0.033 μmol, 30 μL), PS with 450 nm LED light
radiation (300 mW/cm²) (PS-LED, 0.033 μmol, 30 μL), B@MCs, and control
groups (n = 3 independent experiments). C IFN-γ secretion levels in
C57B6 mouse blood serum following various treatments for 3d (n = 3
independent experiments). D Percentage of DC maturation (CD80⁺CD86⁺ DCs
out of total CD11c+ DCs) in B16 cells after exposure to different
treatments in vitro (n = 3 independent experiments). E Quantitative
analysis of CD8⁺ T cells (% of total CD3 + T cells) in B16 tumour
tissues (n = 3 independent experiments). F CD8⁺/CD4⁺ ratio within
tumour tissues (n = 3 independent experiments). Error bars, mean ± SD.
Statistical significance is noted with ***p < 0.001, compared to the
data for PS-LED, PS, B@MCs and control group according to one-way ANOVA
with Tukey’s multiple comparisons test performed using GraphPad Prism 9
XML project software. Source data are provided as a Source data file.
Photodynamic therapeutic effects of PB@MC treatment in vivo
To validate the effect of PB@MC treatment in vivo, a melanoma syngeneic
tumour model was created in mice via the subcutaneous introduction of
B16 cells. When the tumours had grown to approximately 300 mm^3
(approximately 18 days), the mice were randomly allocated to one of
five distinct therapeutic groups: no treatment (vehicle control),
PB@MCs, B@MCs, PS treatment, or PS with LED irradiation (60 min,
300 mW/cm^2). Figure [133]6A shows the treatment regimen. Following a
singular injection on day 0, the tumour size and body mass were
monitored daily. Observations revealed rapid tumour enlargement in the
untreated group, whereas PS and PS-LED treatments had negligible
effects on melanoma progression. Conversely, the B@MC group experienced
a recovery in tumour growth from day 4–6 onwards. Significantly, the
volume metrics substantiated that PB@MCs provided the most pronounced
antitumour response, leading to complete tumour eradication (Fig.
[134]6B–D). In terms of tolerance, mice treated with PB@MCs showed no
notable weight fluctuations (Fig. [135]6E). Survival analysis indicated
that all mice except those treated with PB@MCs succumbed or died within
14 days. In stark contrast, PB@MCs not only completely treated the
condition within 4 days but also notably extended survival to two
months, with nine out of ten mice living beyond 60 days (Fig. [136]6F,
G and Supplementary Fig. [137]17). These results suggest that PB@MCs
are an effective Sd-PDT for melanoma in murine models. For biosafety
evaluation, PB@MCs were administered to healthy mice intraperitoneally
or subcutaneously. The blood test analysis performed on days 1, 3 and 7
revealed no significant abnormalities in WBC levels across all groups
within the 7 day period. Serum procalcitonin (PCT) levels measured on
days 1, 3 and 7 showed that compared to the control group, the PB@MC
group exhibited elevated PCT on day 1. PCT levels in all groups
decreased by days 3 and 7. These findings suggest that PB@MCs may
induce a degree of inflammation, likely attributable to their
generation of ROS, but this response gradually subsides after 3 days
(Supplementary Fig. [138]18). Tissue samples were also collected for
histological examination. Haematoxylin and eosin staining confirmed the
absence of organ-specific pathological changes (Supplementary
Figs. [139]19–[140]21). Recognising the critical role of bacterial
immune adjuvants in initiating immune activation^[141]31, we postulated
that PB@MCs could also amplify immunotherapy by stimulating adaptive
immune activity within the tumour microenvironment. This was
substantiated by immunostaining for TUNEL, CD4, CD8, CD3, CD206, and
CD86 in a melanoma syngeneic tumour model. As shown in Fig. [142]6H–J,
PB@MC treatment significantly ameliorated CD4^+ and CD8^+ production,
enhanced CD3 expression, and reduced CD206 expression in tumour
sections, indicating the activation of T cells and reduced M2-like
macrophage infiltration within the tumour
microenvironment^[143]31,[144]32.
Fig. 6. In vivo therapeutic effects of PB@MCs on tumour growth in melanoma
mice.
[145]Fig. 6
[146]Open in a new tab
A Schematic representation of PB@MC treatment in melanoma mice. Images
are created with biogdp.com. On day 18 post-B16 cell (1 × 10^6/mL)
implantation (~300 mm³ in size), 30 µL of PB@MCs (3.6 × 10⁴/mL) were
intratumourally injected into tumour-bearing mice. B Tumour growth
curves for treatment groups in vivo. Following treatment, tumour
volumes were measured every two days and growth curves were obtained
(n = 6 mice). C Average tumour volumes on day 8 post-treatment (n = 6
mice). The data are presented as mean ± SD. Statistical significance is
noted with ***p < 0.001, compared to the data for PS-LED, PS, B@MCs and
control group according to one-way ANOVA with Tukey’s multiple
comparisons test performed using GraphPad Prism 9 XML project software.
ns not significant. D Optical photographs of the dissected tumours
according to the treatment type. Mice were sacrificed on day 8
post-treatment for tumour visualisation. Ex vivo images were captured
using a Canon ZHS2402 camera (Japan). E Changes in the body weight of
mice during in vivo treatment for 11 days (n = 6 mice). The data are
presented as mean ± SD. F Kaplan–Meier curves showing the survival
rates of the indicated mice at 60 days (n = 10 mice). The data are
presented as ***p < 0.001, compared to the data for other treatment
groups of tumour-bearing mice according to the log rank performed using
GraphPad Prism 9 XML project software. G Optical photographs of
tumour-bearing mice subjected to different treatments for 30 days.
Scale bar: 10 mm. H TUNEL staining of tumour tissues across treatment
groups in which green fluorescence represents FITC-labelled
deoxyuridine triphosphate (dUTP) detected in apoptotic cells. Scale
bar: 200 µm. I Representative immunohistochemical staining images
(TUNEL, CD3, CD206, and CD86) of various treatment groups after 1 day
of treatment. Scale bar: 100 µm. J CD4^+ and CD8^+ immunofluorescence
staining of tumour tissues in various treatment groups.Scale bar:
200 µm. Source data are provided as a Source data file.
Effects of PB@MC implants on hepatocarcinoma
We further investigated the in vivo effects of Sd-PDT on
hepatocarcinoma via direct implantation of PB@MCs into rabbit livers.
Based on the outlined treatment protocol (Fig. [147]7A), rabbits
afflicted with hepatocarcinoma were administered a single intratumoural
injection of PB@MCs and monitored for a duration of 70 days. The
progression of tumour growth was monitored using CT imaging during this
period (Fig. [148]7B). Given the propensity of hepatocarcinoma to
metastasise to the lungs, we assessed the lungs via CT imaging.
Significant metastatic activity was evident in the lungs of untreated
rabbits by day 14; extensive formation of tumour foci was observed by
day 28. In contrast, the PB@MC treatment group exhibited no detectable
lung metastases on day 70 (Fig. [149]7C, D). Oxaliplatin, a typical
anti-tumour drug, was used as a clinical drug control. While tumour
volumes in the oxaliplatin-treated rabbits increased to 3.8 cm^3 by day
14 and to 11.4 cm^3 by day 28 (Fig. [150]7E), the administration of
PB@MCs arrested tumour expansion, and tumour dimensions reduced to
indiscernible small cystic formations by the 14th day. Subsequent
haematoxylin and eosin (H&E) staining confirmed the presence of swollen
liver cells, necrotic liver cells, and fibrotic tissue at the
periphery, whereas normal liver cells were present externally
(Fig. [151]7F). Survival analysis using Kaplan–Meier plots demonstrated
a significant survival benefit for the PB@MC-treated rabbits compared
to controls (Fig. [152]7G). CD3⁺ immunofluorescence and TUNEL staining
of tumour tissues from various groups showed that PB@MC treatment
induced strong antitumour immunity (Fig. [153]7H). H&E staining was
performed to evaluate the histopathological features of hepatocarcinoma
tumours and their pulmonary metastases. (Fig. [154]7I). These results
demonstrate that PB@MC treatment markedly curtailed tumour growth,
effectively inhibiting tumour metastasis and recurrence, and boosted
survival rates in rabbits with hepatocarcinoma.
Fig. 7. In vivo therapeutic effects of PB@MCs on tumour growth and metastasis
in rabbit hepatocarcinoma.
[155]Fig. 7
[156]Open in a new tab
A Schematic of PB@MC treatment in hepatocarcinoma-bearing rabbits.
Images are created with biogdp.com. B CT images depicting tumour growth
in vivo. Tumours in rabbits treated with and without PB@MCs were
compared with oxaliplatin-treated groups using CT imaging at predefined
intervals (0–70 days). Tumour margins are highlighted with a yellow
outline. Scale bars: 1 cm. C CT images of pulmonary tumour metastases.
Lungs were imaged at the same intervals post-treatment to identify
pulmonary metastases, indicated by yellow arrows. Scale bars: 1 cm. D
Ex vivo photographs of livers and lungs from healthy, tumour-bearing
control, PB@MCs, and oxaliplatin-treated rabbits. Animals were
sacrificed on day 28 post-treatment. E Tumour dimensions were
quantified over a 28-day post-treatment (n = 3 rabbits). The data are
presented as mean ± SD. Statistical significance is noted with
**p < 0.01, *p < 0.05 compared to the data for oxaliplatin-treated and
control group according to one-way ANOVA test. F H&E staining of liver
tumour sites treated with PB@MCs. The annotations include purple
squares (b) indicating normal liver cells, blue squares (c) indicating
fibrotic tissue, cyan squares (d) indicating necrotic liver cells, and
red squares (e) indicating swollen liver cells. Scale bar: 100 µm. G
Kaplan–Meier curves showing the survival rates of the indicated mice at
70 days (n = 6 rabbits). The data are presented as ***p = 0.0005
compared to the data for control groups and **p = 0.0036 compared to
the data for oxaliplatin-treated groups according to the log rank test
performed using GraphPad Prism 9 XML project software. H CD3⁺
immunofluorescence (red fluorescence represents Cy5.5-labeled
anti-CD3-PerCP and blue fluorescence DAPI represents the nucleus) and
TUNEL staining red fluorescence represents Cy5.5-labeled dUTP and blue
fluorescence DAPI represents the nucleus. Scale bar: 50 μm. I H&E
staining of hepatocarcinoma tumours and pulmonary metastases. Purple
arrow: pulmonary metastasis, Scale bar: 250 µm. Source data are
provided as a Source data file.
Recently, various PDT methodologies have been developed to overcome
penetration limitations associated with existing methods. For instance,
employing long-wavelength light sources (e.g., near-infrared II) can
modestly extend the penetration depth of PDT (approximately
2 cm)^[157]33,[158]34. Alternatively, the use of high-energy light
(e.g., X-rays) to excite scintillator photosensitive materials
facilitates deeper penetration^[159]35, although prolonged exposure to
high-energy radiation entails potential safety concerns. Direct
excitation strategies, such as those using long-afterglow luminescent
materials^[160]6, micromagnetic induction LED lights^[161]36, inserted
optical fibres^[162]5, chemiluminescence^[163]37 and
bioluminescence^[164]18, can effectively mitigate the risks associated
with prolonged exposure at a high penetration depth. However, these
approaches typically require external energy sources to support or
continuously replenish photosensitive reactants. The development of a
persistent, long-term light source is critical for maintaining ROS
generation at tumour sites. In our study, we utilised the
bioluminescence from bacteria as an implantable light source within
tumours to continuously activate the photosensitizer NR, thereby
enhancing cancer mPDT. Owing to the persistent light emission from
PB@MCs after intratumoural implantation, harnessing the chemical energy
of host organisms can provide adequate ROS for ongoing PDT activation.
This method offers a significant advantage over those requiring
external light sources, which typically have limited penetration
capability in physiological environments. Although the bioluminescence
intensity was considerably lower than that of standard clinical PDT
lights or lasers, PB@MC-enhanced Sd-PDT showed superior efficacy,
particularly in treating large tumours exhibiting high light
absorption, for which traditional PDT light penetration is inadequate
and may cause photodamage. Additionally, our findings indicate that
treating tumours with a high-dose pulsed light (0.33 W/cm^2, the
maximum permissible exposure of skin according to the American National
Standard) curbs tumour growth less effectively than treating tumours
with PB@MCs, thus supporting the utility of PB@MCs for Sd-PDT.
PB@MCs, a generation microbial ball complex, holds significant promise
for clinical applications in cancer treatment. Using ROS detection
assays, metabolomics, and immunohistochemistry, we elucidated the
mechanisms underlying their potent antitumour effects (Fig. [165]8).
Initially, tumour cells exploit metabolic reprogramming to achieve
uncontrolled proliferation. To maintain the redox balance within the
tumour microenvironment, the production of antioxidants, such as
glutathione (GSH), is crucial^[166]38. The existing literature has
explored the feasibility of enhancing antitumour effects by
exacerbating redox imbalances in tumour cells^[167]39,[168]40. Our
combined results from metabolomics and ROS detection experiments
demonstrated that PB@MCs significantly activated ROS and inhibited GSH,
ultimately inducing oxidative stress in tumour cells. The mechanisms by
which PB@MCs lead to GSH depletion were further investigated.
Metabolomic analyses revealed that the GSH metabolic pathway was
suppressed after treatment with PB@MCs, along with the downregulation
of amino acid metabolism and the pentose–phosphate pathway, which
reduced the amount of substrate necessary for GSH synthesis and
regeneration. Additionally, pathways related to glutamine metabolism,
TCA cycle, and oxidative phosphorylation were inhibited, indicating
that the primary energy supply routes for tumour cells were
disrupted^[169]41. Furthermore, the suppression of purine, pyrimidine,
and lipid metabolism suggests that tumour cells lose the ability to
synthesise the essential membrane structures and genetic material
necessary for proliferation. Moreover, an increase in IFN-γ expression;
the increase in the ratio of CD8^+ T cells^[170]42,[171]43 and mature
DCs^[172]44; and the reduced M2-like macrophage infiltration in tumour
sections^[173]45 collectively indicate that PB@MCs activate the
antitumour immune microenvironment within tumour tissues. While
V.H.BB170 treatment alone (B@MCs) showed only partial efficacy in
tumour suppression, its combination with the photosensitizer NR-boosted
mPDT was markedly effective. This strategy of integrating living
bioluminescent bacteria with photosensitizer-enhanced mPDT
simultaneously inhibited tumour growth and prevented tumour recurrence
by activating both innate and adaptive immune responses. Our findings
highlight the utility of strategically integrating bioluminescent
bacteria with mPDT with immunotherapy, leveraging bioluminescent
bacteria-activated PS as a durable source of ROS for sustained mPDT
activation and an influential immunostimulatory agent that amplifies
PDT-induced ICD. We are optimistic that this approach, combining
bioluminescent bacteria and enhanced mPDT with the immunostimulatory
capabilities of bacteria, offers significant potential for clinical
applications owing to its exceptional anti-tumour efficacy and
acceptable safety profile.
Fig. 8. Mechanisms of anti-tumour effects of PB@MCs.
[174]Fig. 8
[175]Open in a new tab
PB@MCs inhibit the malignant biological behaviour of tumours through
several mechanisms. (1) PB@MCs disrupt the redox balance in tumour
cells. This process is triggered synchronously by the production of
reactive oxygen species (ROS) output and glutathione depletion. (2)
PB@MCs lead to metabolic reprogramming of tumour cells. The synthesis
pathways of biomolecules essential for tumour cell proliferation, such
as phospholipids and nucleic acids, were significantly suppressed.
Furthermore, energy metabolism pathways related to tumour cells,
including glycolysis, oxidative phosphorylation, and glutamine
metabolism, were markedly inhibited. This results in a significant
reduction in the synthesis of glutathione, a key metabolite that
alleviates oxidative stress, making it difficult for tumour cells to
maintain their malignant phenotypes associated with unlimited
proliferation. (3) PB@MCs facilitate metabolic-immune co-evolution of
the tumour microenvironment (TME), transforming it from an
immunosuppressive to an immunoactive state. The content of lactic acid,
a marker of immune-suppressive TME, significantly decreased, while
levels of INFγ and ROS, markers of immune activation, rose
significantly. This process relieved the exhaustion of CD4^+ T cells,
CD8^+ T cells, and mature dendritic cells (DCs) in the tumour immune
microenvironment (TIME), reactivating their anti-tumour functions and
promoting the conversion of tumour-associated macrophages (TAM) from
the M2 to the M1 type. Ultimately, this cascade of events induced
tumour cell necrosis, apoptosis, or immunogenic cell death. Images are
created by figdraw.com.
In summary, we developed an Sd-PDT system which enables continuous and
uniform light emission throughout a tumour without requiring an
external energy source or continuous replenishment of photosensitive
reactants. By utilising nutrients within the tumour microenvironment,
this system produces prolonged, low-dose light emission that can be
sustained for up to 50 h, which significantly surpasses the duration of
all currently available chemically driven PDT methods. Moreover, PB@MCs
significantly prolonged the self-supported generation of ROS. A single
injection of PB@MC is required to effectively eradicate large tumours,
including melanoma and hepatocarcinoma tumours in mice and rabbits.
Furthermore, our findings indicate that PB@MC-enhanced Sd-PDT not only
suppresses tumour growth but also evokes strong antitumour immunity.
Consequently, the successful development of Sd-PDT offers significant
potential for advancing tumour biology studies and cancer treatments.
This method may emerge as a promising generalised therapeutic strategy
for treating various cancer types and tumour sizes.
Methods
Ethics statement
All animal experiments were performed in accordance with the guidelines
approved by the Laboratory Animal Welfare and Ethics Committee of The
Second Affiliated Hospital of Zhejiang University School of Medicine
(ARIB-2023-1520). The maximum tumor size permitted by the ethics
committee/institutional review board is 2 cm^3 for mice or 60 cm^3 for
rabbit. The tumor sizes in all experiments have never exceeded this
threshold.
Reagents and materials
Sodium alginate (a molecular weight of 460 kDa; molar ratio of
mannuronic acid to guluronic acid ratio, 2:1) was procured from Qingdao
Bright Moon Seaweed Group Co., Ltd. in Qingdao, China. Chemical
reagents such as dimethyl sulfoxide (DMSO), methanol, and ether were
sourced from Sinopharm Chemical Reagent Co., Ltd. Anhydrous calcium
chloride (CaCl[2]; Thermo Fisher Scientific, C614-500); dibasic sodium
phosphate (Na[2]HPO[4]; ≥98.5%, Cat. 7558-79-4) were obtained from
Calbiochem®. The Annexin V-FITC/PI apoptotic cell death assay kit
(abs50001) was acquired from Absin Bioscience Inc., Shanghai, China.
Corning, New York, USA, supplied the PC membrane Transwell-24 (Cat.
3422) and Transwell-6 (Cat. 3452). The 2216E medium (catalog No.
bio-54545) was purchased from the Institute of Hydrobiology, Wuhan,
China, and CCK-8 assay kits came from Target Molecule Corp., USA.
LIVE/DEAD staining kits, penicillin, streptomycin, trypsin-EDTA, and
DMEM were sourced from ThermoFisher Scientific, Grand Island, NY, and
Hyclone Laboratory, South Logan, Utah, USA, respectively. RPMI 1640
medium and fetal bovine serum (FBS) were procured from Gibco and
Gemini, Woodland, USA, respectively. Beyotime Biotechnology, Shanghai,
China, provided the ROS Assay Kit and Hoechst 33258.
Dibenzocyclooctyne-PEG4-N-hydroxysuccinimidyl (DBCO-PEG4-NHS) ester was
acquired from Xi’an Ruixi Biological Technology Co., Ltd. The cellular
reactive oxygen species detection assay kit (DCFH-DA) was ordered from
Abcam. Propidium iodide (PI) and 4’,6-diamidino-2-phenylindole (DAPI)
were obtained from Shanghai Yuanye Beyotime Co., Ltd, China. Pure water
(resistivity: 18.2 mΩ·cm) was produced using an ELGA Purelab classic
UVF system for preparing working solutions and buffers, while sodium
hydroxide (NaOH, ≥ 98.0%) was purchased from Sigma-Aldrich. Abcam
supplied the Alexa Fluor® 488-conjugated CRT primary antibody
(ab196158, British). Poly-L-Lysine (MW: 15,000-30,000, P4832),
BMS-345541 (Cat. 401480), were purchased from Sigma-Aldrich (St. Louis,
MO, USA). Chemstan (CS-13629, China) provided Etoposide VP-16. Tissue
•OH Assay Kit (BBoxProbei O28, BB-46072, BestBio, Shanghai, China);
tissue freezing medium (14020108926, Leica, Germany); the ELISA kit for
IFN-γ measurement (Solarbio, SEKM-0145, China); anti-CD4-PE (Biolegend,
Cat. 100511), anti-CD45-APC-Cy7 (Cat.557659), anti-CD3-PerCP-Cy5.5
(Cat. 551163), anti-CD11c-FITC (Cat. 117305), anti-CD80-PE (Cat.
560016), and anti-CD8-FITC (Cat. 553030) were purchased from BD, USA.
Details of all antibodies used are provided in Supplementary
Table [176]1
Cultural conditions of bacteria
Vibrio harveyi BB170 (catalog No. bio-108879), Aliivibrio
fischeri-bio115653 (catalog No. bio-105653), and Aliivibrio
fischeri−7744 (catalog No. bio-72794) were obtained from Biobw,
Beijing, China, and acclimatized under controlled physiological
conditions by varying growth factors, including temperature (25–37 °C)
and medium composition (2216E to DMEM). These modifications ensured the
survival of the bioluminescent bacteria under physiological conditions.
A sterile pipette was employed to transfer approximately 0.5 mL of
2216E medium into a 2 mL Eppendorf tube (Centrifuge 5810 R, Eppendorf).
This reconstituted bacterial suspension was incubated overnight at
37 °C in a sterile, clear BeyoGold™ Bacteria Culture Tube containing
5 mL of 2216E medium, agitated at 10 × g. Simultaneously, 100 μL of the
suspension was cultured on a 2216E agar plate at 37 °C for 12–16 h to
cultivate monoclonal colonies. The optical density (OD) at 600 nm was
periodically measured to establish the bacterial growth curve. Once the
bacteria reached the logarithmic phase, the strain was inoculated into
a fresh culture medium. For CFU/mL calculation, 100 μL of the bacterial
suspension was taken when the bacteria reached the logarithmic phase
(OD value of around 0.8) and dilute stepwise to 16-fold. Dilutions (0,
2, 4, 8 and 16) of the bacterial suspension were added to a 96-well
plate to recorded the OD600 values. Subsequently, 20 μL of five
different proportions of bacterial suspension was mixed with 180 μL PBS
buffer and further diluted stepwise from 10^−1 to 10^−8. Then, 100 μL
of bacterial suspension was cultured on a 2216E agar plate. After
overnight incubation, monoclonal bacterial colonies were counted.
Finally, a standard linear graph of bacterial count (CFU/mL) was
plotted against the OD600 reading.
PB@MC construction
Vibrio harveyi BB170 was collected by centrifugation at 100 × g for
10 min using a Centrifuge 5810R (Eppendorf) in a 15 mL Eppendorf tube
and subsequently resuspended in 2216E medium for PB@MC construction.
The bacterial suspension was merged with a 1.5 wt% sodium alginate
solution. After agitating the mixture, it was extruded through a 5 mL
syringe equipped with a 0.5 mm needle into a 0.2 M CaCl[2] solution
(gelling bath) using an electrostatic droplet generator to form
alginate-Ca beads. These beads were then soaked in a 0.05 wt%
poly-L-lysine solution (volume ratio 1:1) for 10 min to form a
poly-L-lysine coating. The NHS-Neutral Red photosensitizer (NHS-NR) was
produced through Cu-free click chemistry, 200 µL of the NR-N[3] stock
solution (25 mM) was added to 2 mL of DBCO-PEG4-NHS ester (50 mM),
which was freshly prepared by dissolving it with dry DMSO. The reaction
was incubated at RT for 4 h and then dialyzed against PBS for
purification. For conjugation, 500 µL of poly-L-lysine-coated
bacteria@MCs (PLL-B@MCs) at 3.6 × 10^4/mL was mixed with 2 mg/mL NHS-NR
(500 μL, 2.2 mM) at 37 °C for 20 min. Subsequently, the medium was
renewed with 2216E medium. The optical density was evaluated at 452 nm
to quantify the unreacted NHS-NR (extinction coefficient
ε[max] = 27,500 M^−1•cm^−1), revealing a binding efficiency of
approximately 49%. The encapsulated microcapsules were reinstated in
15 mL of 2216E medium and analyzed under a microscope. All formulated
PB@MCs were cultured at 35–37 °C for subsequent applications in a
HerryTech KE-200 incubator, Shanghai, China.
Characterization of PB@MCs
The morphology and size of empty MCs, bacteria@MCs (B@MCs),
poly-L-lysine coating bacteria@MCs (PLL-B@MCs), and PB@MCs were
determined by microscopy (FV1200, Japan). For self-bioluminscence
testing, empty MCs, B@MCs and PB@MCs at (500 μL, 3.6 × 10^4/mL) were
added to 24-well chambers to assess their self-bioluminescence
activities using CLSM (LSM900 Carl Zeiss, Germany). Images were
collected with 5× and 10× objectives at the 488–620 nm channel.
For long-term bioluminscence (BL) intensity testing of PB@MCs, the
tested PB@MCs (50 μL, 3.6 × 10^4/mL) were incubated in 96-well black
plates (FCP966, Beyotime Biotechnology, China) with 150 μL 2216E
medium. The BL intensity was continuously monitored at 37 °C every
hour, with light emission detected by a Microplate Reader (Tecan Spark,
Switzerland) until the BL reached its maximum intensity. After removal
of the residual 2216E medium, DMEM medium (150 μL) and B16 tumour
homogenate (150 μL, diluted 1:1 with PBS) were added when the BL
intensity decreased. BL intensity was recorded for another 50 h.
For photo imaging of PB@MCs, 6 mL of freshly prepared PB@MCs were
suspended in 50 mL Eppendorf tubes (Centrifuge 5810 R, Eppendorf) with
20 mL of 2216E medium at 35 °C, in an incubator (HerryTech KE-200,
Shanghai, China) until self-bioluminescence reached its maximum
intensity. After removing the residual 2216E medium, 2 mL of PB@MCs
were suspended in a 14 mL bacteria culture tube (BeyoGold™ Bacteria
Culture Tube, Clear, Sterile) and refreshed with PBS (3 mL), DMEM
medium (3 mL), and B16 tumour homogenate (3 mL, diluted 1:1 with PBS)
when the BL intensity decreased. BL intensity was recorded for another
32 h using a Xiaomi 14 smartphoto with an exposure time of 5 s.
For self-BL spectra of B@MCs and PB@MCs, 500 μL, 3.6 × 10^4/mL B@MCs
and PB@MCs were suspended in 3.5 mL quartz glass cuvette (s2840-04-2EA,
aladdin, China) with equal volumes of DMEM to analyze their self-BL
emission spectrum from 300 to 700 nm without excitation using an
Edingbour NanoSpectralyzer fluorimetric analyzer (Applied
NanoFluorescence, FLS980). The data was fit to a Gaussian function
using OriginLab 8.0 software.
To obtain the in vitro ROS release profile, freshly made PB@MCs (30 μL,
3.6 × 10^4/mL) were incubated in 96-well black plates with 150 μL of
2216E meidum. The BL intensity was continuously monitored at 37 °C
every hour, with light emission detected by a Microplate Reader
(bioluminescence model) until the BL reached its maximum intensity.
After removal of the residual 2216E medium, DMEM medium (100 μL) was
added to each well of the 96-well black plates. For comparison, the
photosensitizer NR (0.32 mM) in 100 μL of DMEM with LED irradiation
(450 nm laser LED, Xi’an Lei Ze Electronics Tech Co., Ltd, Shanxi,
China) was used as a tradional PDT control group^[177]46 (LED
radiation). The LED radiation group in 96-well plates were exposed to
300 mW/cm^2 LED radiation for four intervals (20 min each, 15 min On
and 5 min Off). DCFH was pre-prepared by dissolving DCFH-DA (1 mM,
1 mL) in NaOH (0.01 M, 4 mL) at room temperature for 30 min, then
balanced with Na[2]HPO[4] (25 mM, 20 mL) to make a stock solution at
40 μM in the dark. ROS generation was determined by adding 40 μL of
DCFH (40 μM) at 0, 0.25, 0.5, 0.75, 1, 2, 5, 12, 16, 18, 24, 28, 30,
36, 40 and 48 h. After 15 min of incubation, the residual medium was
transferred to a 96-well black plate, and fluorescence intensity of DCF
was immediately recorded by a Microplate Reader (Tecan spark,
Switzerland) at Ex/Em: 488/525 nm. Fresh DMEM and 40 μL DCF were also
included as negative and positive control.
For In vitro PB@MCs energy radiation density, PB@MCs (100 μL,
3.6 × 10^4/mL) were incubated in 24-well black plates (3 wells in
parallel) with 500 μL of 2216E meidum. The BL intensity was
continuously monitored at 37 °C every 2 h, with light emission detected
by a Microplate Reader (Tecan Spark, Switzerland) until the BL
intensity reached 100, 200, 500, 700 and 1000 × 10^4 cps. The energy
radiation density of PB@MCs was immediately recorded using a Laser
Power Meter (YanbangTech VLP2000, Beijing, China) with a distance of
~0.2 cm between the detector and the plate surface. The plate was
wrapped in aluminum foil with a hole in the center of each well for
light transmission detected by the Laser Power Meter. All testing was
conducted in the dark.
Cell culture
A375 (Cat. CL-0014), Hep3B (Cat. CL-0102), and B16 (Cat. CL-0319) cell
lines were procured from Procell system (Wuhan, China). VX2 (Cat.
BFN60700420) cell line were procured from BLUEFBIO™ Product (Shanghai,
China). and cultured in RPMI 1640 medium or DMEM, supplemented with 10%
FBS (Gemini, Woodland, USA) and 1% penicillin-streptomycin solution
(Hyclone Laboratory, 10,000 U/mL) in a cell culture incubator. A375,
Hep3B, B16 and VX2 cells were morphologically confirmed according to
the information provided by the cell-source center. STR analysis was
performed to authenticate A375, Hep3B, B16 cells in Supplementary
Tables [178]2–[179]4. STR loci are amplified using fluorescently
labeled PCR primers that flank the hypervariable regions. For VX2
cells, a certificate of analysis were performed in Supplementary files.
The cells were all negative in mycoplasma test. Bone Marrow
(BM)-Derived Dendritic Cells (BMDCs) were produced from the BM of
8-week-old C57B6 mice. The procedure involved cutting the hind legs of
the mouse, removing the attached tissue, and sterilizing the femur and
tibia by soaking them in 75% alcohol for 5–10 s. The bones were then
washed with PBS, and both ends were cut off. A syringe needle was
inserted into the bone to flush the BM into the medium. The BM extract
was carefully agitated, and cells were separated by centrifugation at
400 × g for 5 min. The cell pellet was dissolved, and the cell density
regulated to 1 × 10^6/ml prior to being introduced into RPMI 1640
medium containing recombinant mouse GM-CSF (20 ng/ml) and IL-4
(10 ng/mL). The medium was exchanged every 72 h, and immature DCs were
harvested on day 8.
Cell viability test
Tested cells were plated in 24-well transwell plates at 5 × 10^4 cells
per well. Following an overnight incubation, the culture medium was
substituted with 1 mL of fresh medium, to which 200 μL of PBS, empty
MCs connected to a photosensitizer (P@MCs, contain 200 μg, 0.22 μmol
NR), MCs containing V.H.BB170 bacteria (B@MCs, 1 × 10^3 bacteria cells
per MC), and PB@MCs (1 × 10^3 bacteria cells per MC and contain 200 μg,
0.22 μmol NR) were introduced into the upper chamber. The cells were
then incubated at 37 °C for eight hours. Subsequent to the removal of
the supernatants and any residual P@MCs, B@MCs, and PB@MCs, each well
received 500 μL of a 10% CCK-8 solution (diluted in DMEM). The plates
were further incubated at 37 °C for an additional hour. Absorbance was
recorded at OD 450 nm using a Microplate Reader (Tecan Spark,
Switzerland). Cell viability was calculated (1):
[MATH: Cell viability%=(AN−AB)(AC−AB)×100% :MATH]
1
where A[N], A[C] and A[B] represent the absorbance at 450 nm of the
treated, untreated, and blank samples, respectively.
Confocal microscopy imaging
For LIVE/DEAD staining. B16, VX2, Hep3B, and A375 cells were cultured
in 24-well transwell plates at 5 × 10^4 cells per well and incubated
for 24 h. Subsequently, the supernatants were exchanged with fresh RPMI
1640 or DMEM medium, and the cells were subjected to 200 μL of
P@MCs(P@MCs, (contain 200 μg, 0.22 μmol NR), B@MCs (1 × 10^3 bacteria
cells per MC), and PB@MCs (1 × 10^3 bacteria cells per MC and contain
200 μg, 0.22 μmol NR) in the upper chamber for 8 h. After treatment,
the cells were twice washed with PBS and stained using a Calcein/PI
viability assay kit, diluted in culture media, for 20 min at 37 °C. All
cell samples were examined under a confocal laser scanning microscope
(FV 1200, Olympus, Japan) utilizing 5× or 10× immersion objective
lenses at excitation wavelengths of 517 nm and 617 nm.
In vitro ROS content detection. To assess intracellular ROS generation,
a DCFH-DA staining kit was employed. Hep3B and A375 cells were cultured
in 24-well transwell plates at 37 °C with 5% CO[2] at 5 × 10^4 cells
per well after an overnight pre-incubation. Fresh RPMI 1640 or DMEM
medium containing 10% v/v FBS replaced the supernatants. The cells
underwent treatment with a vehicle control (no treatment), PB@MCs
(100 μL, 3.6 × 10^4/mL, contain 100 μg, 0.11 μmol NR), B@MCs (100 μL,
3.6 × 10^4/mL), and P@MCs (100 μL, 3.6 × 10^4/mL, contain 100 μg,
0.11 μmol NR) in the upper chamber in darkness at 37 °C for 2 h.
Subsequently, the cells were incubated in fresh medium containing
DCFH-DA (10 μM) for 30 min, washed thrice with PBS, and stained with
10 μg/mL Hoechst 33342 (Beyotime Biotechnology, Shanghai, China) in PBS
for five minutes. DCF fluorescence was promptly visualized using CLSM
(LSM900 Carl Zeiss, Jena, Germany) at 488 nm.
In vivo ROS content detection. To evaluate the •OH levels produced by
PB@MCs in mouse tumour tissues in vivo. Four groups of B16
melanoma-bearing mice were injected intratumourally with 50 μL PBS
(control), B@MCs, P@MCs or PB@MCs. After 10 h, tumours were aseptically
collected, weighed, and processed according to the manufacturer’s
instructions. Tumour tissues were homogenized by cryo-grinding at 60 Hz
for 120 s, then adjusted to 50 mg/mL with sterile PBS. Subsequently,
190 μL of tumour homogenate (50 mg/mL) and 10 μL of BBoxiProbe O28
working solution were added to a 96-well black plate and incubated at
37 °C in the dark for 20 min. Fluorescence intensity was measured using
a microplate reader (excitation: 488 nm; emission: 520 nm). Protein
concentrations in the homogenate were quantified using the Bradford
Protein Assay Kit (P0006, Beyotime Biotechnology, Shanghai, China).
Tissue •OH levels were expressed as fluorescence intensity normalized
by protein concentration.
Flow cytometry analysis
To evaluate if PB@MCs treatment prompted apoptotic cell deaths, B16,
VX2, Hep3B and A375 cells (5 × 10^5 cells/well) were cultured in
6-transwell plates using RPMI 1640 or DMEM for 16 h. Fresh media
containing 400 µg/mL etoposide (serving as the positive control for the
VP-16 group) and 400 μL of P@MCs (contain 400 μg, 0.44 μmol NR), B@MCs
(1 × 10^3 bacteria cells per MC), or PB@MCs (1 × 10^3 bacteria cells
per MC and contain 400 μg, 0.44 μmol NR) were then introduced to the
upper chamber for 8 h. Following incubation, cells were harvested and
stained using an Annexin V-FITC/DAPI apoptosis kit, then analyzed via
flow cytometry.
To measure IFN-γ levels, whole blood was collected from 8-week-old
C57B6 mice by eyeball blood collection on day 3 after treatment,
including an intratumoural injection of 30 μL saline, intratumoural
injection of 1.5 mg/kg PS (NHS-NR, 0.033 μmol, 30 μL) (PS group), PS
coupled with 300 mW/cm^2 blue light LED radiation for 60 min (PS-LED
group), intratumoural injection of 30 μL (3.6 × 10^4/mL) of bacterial
encapsulate of MCs (B@MCs), and PS-modified B@MCs (PB@MCs group). Blood
was centrifuged at 100 × g for 5 min, and the serum was extracted.
Levels of IFN-γ were quantified using an ELISA kit (Solarbio,
SEKM-0145, China) as per the provided instructions.
For CRT expression assay. CRT expression was assessed through flow
cytometry. A375 and B16 cells were plated in 6-well plates at 5 × 10^5
cells per well and incubated for 24 h. Treatments administered included
PS (NHS-NR, 0.033 μmol, 30 μL/well) (PS group), PS in conjunction with
300 mW/cm^2 of blue light LED radiation for 60 min (PS-LED group), and
intratumoural injections of 30 μL of bacterial encapsulate of MCs
(B@MCs), and 30 μL of PS-modified B@MCs (PB@MCs group) for 6 h.
Post-treatment, cells were rinsed with cold PBS (1 mL) at 4 °C and
fixed in 0.25% paraformaldehyde (1 mL) for 10 min, then incubated with
Alexa Fluor®488-conjugated CRT primary antibody (ab196158, Abcam, UK)
for 30 min before undergoing flow cytometry.
To explore in vitro DC maturation, BMDCs were cultured from the BM of
8-week-old C57B6 mice. B16 cells were pre-treated with the PS group
(NHS-NR, 0.033 μmol, 30 μL), PS-LED group, 30 μL of B@MCs, or 30 μL of
PB@MCs at the same concentration for 6 h. Subsequently, 1 × 10^6
immature DC cells were co-cultured with these pre-treated B16 cells for
24 h. The maturation of DC cells was then assessed by staining with
anti-CD11c-FITC, anti-CD80-PE, and anti-CD86-APC antibodies and
analyzed using flow cytometry (BD, FACSCanto II). Flow cytometry gating
strategy for the analysis of DC were shown in Supplementary
Fig. [180]22.
For intratumoural infiltration of T lymphocytes. To prepare a tumour
single-cell suspension, tumours were excised on day 3 post-treatment,
sectioned into small pieces, and enzymatically digested in DMEM
containing 1 mg/mL collagenase IV (C8160, Solarbio, China) and
0.2 mg/mL DNase I (D8071, Solarbio, China) for 45 min at 37 °C.
Isolated cells were stained with anti-CD45-APC-Cy7,
anti-CD3-PerCP-Cy5.5, anti-CD4-PE, and anti-CD8-FITC antibodies for
30 min and subsequently analyzed by flow cytometry. Flow cytometry
gating strategy for the analysis of T cells were shown in Supplementary
Fig. [181]22.
Metabolomics analysis
Mice inoculated with B16 cells were prepared for further interventions
once tumours reached 300 mm^3. The qualified subjects were randomly
allocated into five groups (n = 5 per group) to receive treatments
including an intratumoural injection of 30 μL saline, intratumoural
injection of 1.5 mg/kg PS (NHS-NR, 0.033 μmol, 30 μL) (PS group), PS
coupled with 300 mW/cm^2 blue light LED radiation for 60 min (PS-LED
group), intratumoural injection of 30 μL (3.6 × 10^4/mL) of bacterial
encapsulate of MCs (B@MCs), and PS-modified B@MCs (PB@MCs group). Each
animal was administered a single intratumoural injection, anesthetized
with a lethal dose of sodium pentobarbital (400 mg/kg), and euthanized
24 h later. tumour samples were then stored in liquid nitrogen for
subsequent metabolomics analysis.
Animal treatment
Healthy female C57B6 mice (6–8 weeks old, 20 g) were acquired from the
Experimental Animal Center at Hangzhou Medical College, China. Female
New Zealand White rabbits (6 months old, weighing between 2 and 2.5 kg)
were procured from Qingdao Kangda Rabbit Co., Ltd. (Qingdao, China).
All animals were maintained under controlled environmental conditions
(22–25 °C), with a 12-h light/dark cycle and relative humidity ranging
from 40 to 70%, in plastic cages using sterilized wood shavings as
bedding. The B16 melanoma tumour model was established in mice by
subcutaneously injecting 100 µL of B16 cell suspension (1 × 10^6 cells)
into the hind leg. Tumour volumes were recorded bi-daily using a
specific formula:
[MATH: Tumor
volume(mm3)=wid<
mi>th2×length2 :MATH]
2
Once tumours reached roughly 300 mm³, preparations for further
interventions began. Qualified animals were randomly distributed into
five groups for various treatments, which included a single
intratumoural injection of either 30 µL saline (vehicle control),
1.5 mg/kg PS (NHS-NR, 0.033 μmol, 30 μL) (PS group), PS combined with
300 mW/cm² of blue light LED radiation for 60 min (PS-LED group), 30 µL
(3.6 × 10^4/mL) of bacterial encapsulate in MCs (B@MCs), or PS-modified
B@MCs (PB@MCs group), all administered on day 18.
For inducing hepatic VX2 tumours in rabbits, VX2 cell suspensions
(2 × 10^6 cells, 200 μL) were implanted in the thigh muscles of donor
rabbits. When tumours grew beyond 2 cm, typically within two weeks,
donor rabbits were euthanized via intravenous administration of a
lethal dose (2 mL/kg) of xylazine hydrochloride for tumour harvesting.
Each tumour was meticulously minced into 1 mm³ pieces under sterile
conditions. Recipient rabbits were sedated using intramuscular
injections of xylazine hydrochloride (250 µL/kg), and the minced tissue
was precisely inserted into the subcapsular parenchyma of the left
hepatic lobe guided by a 16-slice CT spiral scan (Brilliance-16,
Philips, USA). tumour development was monitored by CT until they
reached about 1 cm³. Hepatocarcinoma-bearing rabbits of similar tumour
sizes were randomly assigned to three groups: vehicle control,
oxaliplatin treatment, and a PB@MCs group. Each received a singular
intratumoural injection of oxaliplatin (100 µL, 6.62 mg/rabbit) or
PB@MCs suspension (500 µL, 3.6 × 10^4/mL) on day 14. Tumour growth and
lung metastasis were periodically assessed via CT (MHCT Brilliance 16,
Philips, Holland). Harvested organs were documented using a Canon
camera (Japan). Both species were ultimately euthanized using an
overdose of sodium pentobarbital (400 mg/kg) to collect tumours and
organs for cytokine quantification or for preservation in liquid
nitrogen. Rabbit tissues were processed for H&E staining and
immunostaining for TUNEL and CD3 markers, whereas mouse tissues were
subjected to immunostaining for TUENL, CD4, CD8, CD3, and CD206
markers.
In vivo biosafety assessment
Healthy female C57B6 mice (age: 6–8 weeks; weight: 20 × g) were
randomized into four groups. Mice underwent intraperitoneal or
subcutaneous injections of either PB@MCs suspensions (30 μL,
3.6 × 10^4/mL) or an equivalent volume of PBS. Post-injection
observations of behavior and physical condition were conducted
periodically. Mice were euthanized via CO[2] inhalation on days 1, 3
and 7 to procure organs and blood. Collected tissues were subsequently
prepared for H&E staining.
Statistics and reproducibility
All experiments were repeated at least thrice with three to ten
replicates. Data were expressed as mean ± standard deviation (SD) from
at least three replicates. All confocal imaging and immunohistochemical
staining imaging were repeated at least three replicates. Data analysis
was performed by two-tailed Student’s t-test, one-way ANOVA or Log rank
test via by GraphPad Prism 9 XML project software. The difference was
regarded as statistical significance if p < 0.05.
Reporting summary
Further information on research design is available in the [182]Nature
Portfolio Reporting Summary linked to this article.
Supplementary information
[183]Supplementary Information^ (4.1MB, pdf)
[184]Reporting Summary^ (179.6KB, pdf)
[185]Transparent Peer Review file^ (2MB, pdf)
Source data
[186]Source Data^ (232.4KB, xlsx)
Acknowledgements