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. Author manuscript; available in PMC: 2023 Jan 1.
Published in final edited form as: Biomaterials. 2021 Dec 9;280:121315. doi: 10.1016/j.biomaterials.2021.121315

Co-delivery of Dihydroartemisinin and Pyropheophorbide-Iron Elicits Ferroptosis to Potentiate Cancer Immunotherapy

Wenbo Han 1,†, Xiaopin Duan 1,3,†, Kaiyuan Ni 1,†, Youyou Li 1, Christina Chan 1, Wenbin Lin 1,2,*
PMCID: PMC8724418  NIHMSID: NIHMS1764241  PMID: 34920370

Abstract

Dihydroartemisinin (DHA) has shown cytotoxicity against various tumor cells in vitro in an iron-dependent manner, but its in vivo antitumor efficacy is compromised by its rapid degradation and clearance. Here we show the induction of ferroptosis by DHA in an immunogenic fashion and the maximization of in vivo antitumor efficacy of DHA by co-delivering a cholesterol derivative of DHA (Chol-DHA) and Pyropheophorbide-iron (Pyro-Fe) in ZnP@DHA/Pyro-Fe core-shell nanoparticles. ZnP@DHA/Pyro-Fe particles stabilize DHA against hydrolysis and prolong blood circulation of Chol-DHA and Pyro-Fe for their enhanced uptake in tumors. Co-delivery of an exogenous iron complex and DHA induces more ROS production and causes significant tumor inhibition in vivo. By increasing tumor immunogenicity, the combination of DHA and Pyro-Fe sensitizes non-immunogenic colorectal tumors to anti-PD-L1 checkpoint blockade immunotherapy. These findings suggest the potential of using nanotechnology to repurpose DHA and other drugs with excellent safety profiles for combination with immune checkpoint blockade to treat cancers.

Introduction

Recent clinical success in cancer immunotherapy has demonstrated its potential to stimulate the host’s immune system for tumor eradication, thereby paving a new avenue to providing durable treatment responses for patients with advanced and aggressive tumors.1, 2 In particular, neutralizing negative immune checkpoints with anti-programmed cell death protein 1 (PD-1) or anti-programmed death ligand 1 (PD-L1) antibodies have produced impressive antitumor responses for several types of cancers, including melanoma, head and neck cancer, lung cancer, kidney cancer, bladder cancer, Merkel cell carcinoma, and Hodgkin disease. Despite these successes, responses to immunotherapy remain fairly restricted to a subset of patients, with the majority of patients refractory to PD-1/PD-L1 blockade.3, 4 The responders are found to more likely have immunogenic tumors that are characterized by greater T-cell infiltration and Th1 cytokine expression, while the non-responders tend to have immunologic “cold” tumors with sparse T-cell infiltration.5–8 In this context, combination therapies designed to increase tumor immunogenicity and attract CD8+ T cells into tumors may broaden the response of cancer patient population to PD-1/PD-L1 blockade.9, 10

Dihydroartemisinin (DHA) is an active metabolite of artemisinin, which has been used extensively as an anti-malarial drug with few side effects.11 In recent years, DHA has been shown to exhibit cytotoxicity against a variety of cancer cells in vitro.12, 13 It is believed that free radicals generated from DHA can oxidize lipids and damage membrane proteins to induce cancer cell death. The radical generation process was proposed to be iron-dependent via reductive scission of the peroxide bridge to generate carbon-centered radicals or the Fenton-reaction-related production of hydroperoxide or hydroxyl radicals.14–18 However, DHA has a very short blood circulation half-life (<15 min) and rapidly hydrolyzes in water, severely limiting the potential use of DHA and related anti-malarial drugs as antitumor drugs.19–21

DHA has also been reported to induce ferroptosis, an iron-dependent type of cell death.22–28 We recently identified DHA as one of only a handful of immunogenic cell death (ICD) inducers that synergistically enhanced tumor immunogenicity of oxaliplatin to potentiate cancer immunotherapy.29 Since the iron center in heme is crucial to DHA toxicity to malaria parasites30, 31 and insufficient endogenous iron supply causes parasite resistance to artemisinin treatment, we hypothesize that heme-iron may also be critical to DHA’s ability to induce ICD and ferroptosis. Unfortunately, solid tumors tend to be iron deficient due to the proliferation of cancer cells, presenting another hurdle for the use of DHA and derivatives as anticancer therapeutics.32–35

Herein, we report the induction of ferroptosis by DHA in an immunogenic fashion and the co-delivery of a cholesterol derivative of DHA (Chol-DHA) and Pyropheophorbide-Iron (Pyro-Fe) in ZnP@DHA/Pyro-Fe core-shell nanoparticles to maximize ferroptosis and in vivo anticancer efficacy of DHA. Due to the low concentration of thiol-based reductants in blood circulation, Pyro-FeIII cannot catalyze the decomposition of Chol-DHA via reductive scission of the peroxide bridge or Fenton-like reactions. Upon endocytosis by cancer cells, glutathione and other thiol-based reductants rapidly reduce Pyro-FeIII to Pyro-FeII, which catalyzes the decomposition of DHA to cytotoxic carbon- and oxygen-based radicals. ZnP@DHA/Pyro-Fe particles prolong circulation half-lives of Chol-DHA and Pyro-Fe to increase their uptake in tumors. Enhanced delivery of Chol-DHA and Pyro-Fe to tumors induces more ROS production to elicit significant tumor inhibition in vivo. By increasing tumor immunogenicity, the combination of DHA and Pyro-Fe enhances intratumoral CD8+ T cell infiltration and potentiates anti-PD-L1 immune checkpoint blockade in non-immunogenic colorectal tumors. Our work shows that DHA and other drugs with excellent safety profiles can be repurposed with nanotechnology for combination therapy with immune checkpoint blockade to treat cancers.36

Results and Discussion

Preparation of ZnP@DHA, ZnP@Pyro-Fe and ZnP@DHA/Pyro-Fe

In the presence of 1,2-dioleoyl-sn-glycero-3-phosphate sodium salt (DOPA), coordination polymerization between Zn2+ and pyrophosphate ions afforded spherical and monodispersed Zn-pyrophosphate (ZnP) core particles as determined by transmission electron microscopy (TEM) (Figure 1b).37 Dynamic light scattering (DLS) measurements of ZnP particles showed a Z-average diameter of 43.9 ± 0.4 nm and a polydispersity index (PDI) of 0.139 ± 0.001 (Figure S3). After coating with a mixture of lipids containing 1,2-dioleyl-sn-glycero-3-phosphocholine (DOPC), cholesterol, and 1,2-diastearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)2000] (DSPE-PEG2k) in the presence of Chol-DHA and/or Pyro-FeIII, core-shell ZnP@DHA, ZnP@Pyro-Fe, or ZnP@DHA/Pyro-Fe nanoparticles were obtained, respectively (Figure 1a). Chol-DHA and Pyro-Fe were incorporated into lipid bilayer at high loadings of 10.6 ± 0.5% and 13.2 ± 0.6 wt%, as determined by liquid chromatography-mass spectrometry (LC-MS) and inductively coupled plasma-mass spectrometry (ICP-MS), respectively. ZnP@DHA, ZnP@Pyro-Fe, and ZnP@DHA/Pryo-Fe were well-dispersed (Figures 1c–d and S4) with similar Z-average diameters of ~90 nm (Figure 1e). In addition, ZnP@DHA/Pyro-Fe exhibited constant size and PDI after incubation with 5 mg/mL bovine serum albumin (BSA) for up to 24 h (Figure S5), suggesting favorable structural stability of ZnP@DHA/Pyro-Fe under physiological conditions. The small size and high stability should help the nanoparticles to avoid clearance by the reticuloendothelial system, prolong blood circulation, and improve tumor accumulation, making them an optimal delivery system for in vivo therapeutic applications.38–40

Figure 1.

Figure 1.

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Particle characterization and the catalytic effect of Pyro-Fe on DHA. (a) Schematic illustration showing the construction of the core-shell structure of ZnP@DHA/Pyro-Fe nanoparticles. (b-d) TEM images of ZnP (b), ZnP@Pyro-Fe (c), and ZnP@DHA/Pyro-Fe (d). Scale bar = 50 nm. (e) Number-average diameter of ZnP@DHA, ZnP@Pyro-Fe, and ZnP@DHA/Pyro-Fe characterized by DLS. (f) Decomposition of Chol-DHA in ZnP@DHA/Pyro-Fe after incubation with cysteine in the presence of 0.5% Triton X-100. (g) ROS generation of ZnP@DHA catalyzed by ZnP@Pyro-Fe in the presence of cysteine under O2-free condition.

Pyro-Fe catalyzes ROS production from DHA

We investigated the catalytic effect of reduced Pyro-Fe on DHA degradation by determining the remaining Chol-DHA content after incubation with either ZnP@DHA or ZnP@DHA/Pyro-Fe with or without cysteine in the presence of 0.5% Triton X-100, which was used to disrupt the lipid bilayer and release Chol-DHA. As shown in Figure 1f, the Chol-DHA in ZnP@DHA hydrolyzed over time and decreased by ~50% in 2 h. The addition of Pyro-FeIII had no effect on DHA degradation, but the reduction of Pyro-FeIII to Pyro-FeII with cysteine further decreased the Chol-DHA content to 15%. Pyro-FeII appeared to contribute more to the degradation of Chol-DHA than cysteine (to 36%).

We then tested if Pyro-Fe could enhance ROS production from DHA. In the presence of O2, ZnP@Pyro-FeII catalyzed the generation of ROS directly from O2, with approximately 8-fold increase in the ROS amount. The addition of ZnP@DHA further increased ROS production to approximately 10-fold (Figure S6). However, in the absence of O2, ZnP@Pyro-FeII could not generate ROS, while ZnP@DHA/Pyro-FeII increased ROS amount to approximately 6-fold (Figure 1g). This result indicates that Pyro-FeII catalyzes the decomposition of DHA to produce ROS in an O2-independent manner, which is consistent with a previous report on FeII-catalyzed DHA decomposition to produce carbon-center radicals.41

Pyro-Fe enhances the cytotoxicity of DHA

After incubation with ZnP@Pyro-Fe for 24 h, the iron concentration in cells significantly increased by 24.8-fold compared to the PBS control group. Incubation with ZnP@DHA/Pyro-Fe notably increased the intracellular iron content by 41.6-fold (Figure S7). ZnP@Pyro-Fe itself showed no cytotoxicity at a concentration of 50 μM on both MC38 and CT26 cells, but significantly enhanced the cytotoxicity of ZnP@DHA, decreasing its IC50 value from 16.5 ± 1.2 to 8.0 ± 1.2 μM on CT26 cells and 14.8 ± 1.6 to 5.2 ± 0.9 μM on MC38 cells, respectively (Figure S8 and Table S1). We also performed MTS assay on normal HEK293T cells and observed similar cytotoxicity, suggesting that ferroptosis-induced cell killing effect is not cell type dependent (Figure S9). We then studied the mechanism for ZnP@DHA/Pyro-Fe induced cytotoxicity. Cell EM first confirmed the ferroptosis induced by ZnP@DHA/Pyro-Fe (Figure 2a). Cells treated with ZnP@DHA/Pyro-Fe showed smaller, ruptured mitochondria with increased mitochondrial membrane intensities, a typical pathological morphology of mitochondria observed in artemisinin derivative triggered ferroptosis.42 Mitochondrial membrane potential change probed by tetraethylbenzimidazolylcarbocyanine iodide (JC-1) further confirmed that ZnP@DHA or ZnP@DHA/Pyro-Fe-induced ferroptosis effectively depolarized mitochondrial membrane potential (Figure S10).Additionally, flow cytometry assay showed that treatment with ZnP@DHA/Pyro-Fe induced more apoptotic cells (42.4%) than ZnP@DHA alone (32.7%) (Figure S11), suggesting that the enhancement of ROS generation induced by ZnP@DHA/Pyro-Fe mediates both ferroptosis and apoptosis for cytotoxicity. Furthermore, CT26 cells exposed to 10 μM ZnP@DHA showed significant decreases of cells in the S phase (32.5%) and G0/G1 phase (32.2%), and a corresponding increase of cells in the G2/M phase (35.4%) when compared to the control group (48.0%, 40.1%, and 11.1% for S, G0/G1, and G2/M phases, respectively). In the presence of Pyro-Fe, ZnP@DHA induced the arrest of cell growth in the G2/M phase (42.9%) more effectively, directly correlating with the inhibition of cell proliferation (Figure 2b). The enhancement of DHA antitumor activity by ferrous iron suggests that simultaneous administration of artemisinin-like drugs and intracellular iron compounds may present a simple and effective way to treat cancer.

Figure 2.

Figure 2.

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ZnP@DHA/Pyro-Fe enhances ferroptosis and kills cancer cells. (a) EM images of ZnP@DHA/Pyro-Fe treated cells showing smaller, ruptured mitochondria with increased mitochondrial membrane densities caused by ferroptosis. (b) Pyro-Fe enhances cell cycle arrest induced by ZnP@DHA. (c-f) Pyro-Fe enhances intracellular ROS generation to cause DNA double strand breaks and lipid peroxidation, as shown by γ-H2AX foci formation (c,d) and increased oxidized BODIPY (e,f) (n = 3). (g-i) The effect of ferroptosis inhibitors on cytotoxicity (g), ROS production (h), and lipid peroxidation (i) caused by ZnP@DHA and ZnP@Pyro-Fe (n = 3). Scale bars = 200 nm (a) and 20 μm (c and e). * p < 0.05, ** p < 0.01, *** p < 0.001.

Pyro-Fe facilitates intracellular ROS generation

Next, we determined whether cell death was dependent on DHA-induced ROS generation. A cell-permeable fluorogenic 2′,7′-Dichlorodihydrofluorescin diacetate (H2-DCFDA) probe that can be oxidized by different ROS and converted to a highly fluorescent dye (DCF) was used to determine intracellular ROS levels. Cells treated with ZnP@DHA alone or with ZnP@DHA/Pyro-Fe showed stronger DCF staining (Figure 2c). Quantitative analysis by flow cytometry revealed stronger DCF fluorescence intensity and more DCF positive cells after incubation with ZnP@DHA/Pyro-Fe (Figure 2d), which correlates with cell death.

Intracellular ROS have multiple effects as they can cause damage to the lipid membrane, DNAs, proteins, and subcellular organelles by direct oxidation reactions or through secondary products of these reactions. ROS can also mediate numerous physiological processes, including innate immune response and cell signaling.43, 44 DNA double-strand break (DSB), one of the most critical DNA lesions related to cell-death, induces phosphorylation of the histone, H2AX. This newly phosphorylated protein, γ-H2AX, initiates the recruitment and localization of DNA repair proteins, therefore acting as an early marker for the identification of DNA DSB.45 ZnP@DHA significantly increased the number of γ-H2AX foci in CT26 cells, suggesting DHA damage via oxidation by the generated ROS. Due to the enhancement on ROS production, Pyro-Fe further enhanced the DNA damage effect of DHA, as evidenced by increased γ-H2AX foci formation (Figure 2e).

The effect of DHA on lipid peroxidation was investigated using BODIPY C11 (581/591), a specific sensor for intracellular lipid peroxidation that shifts the fluorescence emission from red to green. Quantitative single-cell analysis revealed an increase in the intensity of oxidized BODIPY C11 (581/591) and the number of oxidized BODIPY C11 (581/591) positive cells (45.7%) in response to ZnP@DHA, indicating increased lipid peroxidation. ZnP@DHA/Pyro-Fe led to more pronounced increases in the number of cells with lipid peroxidation (65.0%) and the intensity of oxidized BODIPY C11 (581/591), confirming the enhancement of DHA-based ROS generation by Pyro-Fe (Figure 2f). DNA damage and lipid peroxidation results indicate that DHA treatment elicits a multitude of subcellular changes to cause cancer cell death.

Pyro-Fe enhances DHA-induced ferroptosis

Iron-dependent ROS generation and lipid peroxidation have been described as the driving forces for ferroptosis, although their underlying molecular mechanism remains unclear.46, 47 Since both Pyro-Fe and Chol-DHA are very hydrophobic due to their conjugation to phospholipid and cholesterol, we hypothesized that they would prefer lipid-rich areas in cells to cause strong lipid oxidation and ferroptosis. Furthermore, as the iron is chelated to the tetradentate pyropheophorbide ring, the catalytic effect of Pyro-Fe should not be affected by iron chelators. We tested whether lipid ROS scavenger, ferrostatin 1 (Fer-1), or iron chelator, deferoxamine (DFO), could block cell death via scavenging the Fe pool. Co-addition of Fer-1 (20 μM) or DFO (0.1 μM) reduced but did not fully block the cytotoxicity of ZnP@DHA and rescued the viability of CT26 cells to approximately 75% from 48% for the ZnP@DHA treatment only, indicating than labile Fe is partially responsible for DHA-induced cell death through ferroptosis (Figure 2g). Total ROS accumulation and lipid peroxidation induced by ZnP@DHA were also suppressed by co-treatment with Fer-1 or DFO, as indicated by decreased DCF signal and BODIPY C11 (581/591) oxidation (Figure 2h, i). However, the addition of DFO to the ZnP@DHA/Pyro-Fe treatment did not rescue CT26 cells from cell death, ROS accumulation, or lipid peroxidation (Figure 2g–i), indicating significant enhancement of DHA-induced ferroptosis by co-delivered Pyro-Fe.

Pyro-Fe enhances the antitumor efficacy of DHA

Pharmacokinetic and biodistribution studies were carried out on SD rats and CT26-tumor bearing mice with ZnP@DHA/Pyro-Fe doped with 20 mol% Pyrolipid (relative to the sum of Pyro-Fe and Pyrolipid), respectively. The particle showed a prolonged blood circulation with a half-life of 3.5 h and an accumulation of 5%ID/g in tumors 24 h post injection (Figure S12 and Table S2), indicating that the particle escapes form MPS uptake and enriches in tumors through the enhanced permeability and retention (EPR) effect.

Next we examined if Pyro-Fe could enhance the antitumor activity of DHA on CT26 and MC38 colorectal tumor models established by subcutaneous injection of cancer cells into the right flank regions of BALB/c and C57BL/6 mice, respectively. When the tumors reached ~100 mm3, mice were intraperitoneally (i.p.) injected with ZnP@DHA, ZnP@Pyro-Fe or ZnP@DHA/Pyro-Fe at a DHA dose of 10 mg/kg and/or Fe dose of 1.96 mg/kg every 3 days. Compared to the rapidly increasing tumor volumes in the PBS group, ZnP@DHA moderately inhibited CT26 tumor growth, resulting in a tumor growth inhibition index (TGI, defined as [1-(mean volume of treated tumors/mean volume of control tumors)] ×100%) of 28.8%. In contrast, the treatment of ZnP@DHA/Pyro-Fe significantly inhibited tumor growth to afford a TGI of 57.9% (Figure 3a), indicating the enhancement of DHA antitumor activity by Pyro-Fe. The average tumor weights were also reduced significantly, with 65.1% and 32.5 % reduction in tumor weights over the PBS control for ZnP@DHA/Pyro-Fe and ZnP@DHA groups, respectively (Figure 3b).

Figure 3.

Figure 3.

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Pyro-Fe enhances the antitumor efficacy of ZnP@DHA. (a-d) Growth curves (a,c) and final weights (b,d) of CT26 (a,b) and MC38 (c,d) tumors treated with combination of ZnP@DHA and ZnP@Pyro-Fe (n = 6). (e) Representative CLSM images of TUNEL assays of MC38 tumor tissues. DNA fragment in apoptotic cells was stained with fluorescein-conjugated deoxynucleotides (green) and the nuclei were stained with DAPI (blue). Scale bars: 50 μm. (f) Histological analysis of MC38 tumor tissues. Scale bars: 100 μm. Red arrows refer to injection of different particles. Statistical significance: *P < 0.05, **P < 0.01 and ***P < 0.001.

We observed a similar tumor growth inhibitory effect on the MC38 tumor model with TGI values of 56.1% and 24.3% for ZnP@DHA/Pyro-Fe and ZnP@DHA groups, respectively (Figure 3c). The weights of tumors treated with PBS, ZnP@/Pyro-Fe, ZnP@DHA, and ZnP@DHA/Pyro-Fe were 1.61± 0.10 g, 1.59 ± 0.12 g, 1.21 ± 0.16 g, and 0.55 ± 0.09 g, respectively (Figure 3d). These results confirm the ability of Pyro-Fe to enhance the antitumor efficacy of DHA. To ascertain ZnP@DHA/Pyro-Fe induced ferroptosis as the major mechanism for antitumor efficacy, we immunostained tumor slides after treatments to probe SLC7A11, a glutamate-cystine exchanger, as one of the negative regulators for ferroptosis. The ZnP@DHA group showed modest green fluorescence, indicating moderate ferroptotic cell death (Figure 3e). In comparison, ZnP@DHA/Pyro-Fe treatment showed completely depleted SLC7A11 with no green fluorescence signals, suggesting stronger ROS generation abrogated the negative regulator for ferroptosis.48 The down-regulation of SLC7A11 might be mediated by the p53 pathway, which was activated by ROS to reduce the expression of SLC7A11.49, 50 Similarly, we observed SLC7A11 downregulation on CT26 cells after treated with ZnP@DHA or ZnP@DHA/Pyro-Fe by western blot (Figure S13). The TdT-mediated dUTP nick end labeling (TUNEL) assay showed that the ZnP@DHA/Pyro-Fe group had higher fluorescence intensity of DNA fragmentation and percentage of dead cells (Figure S14). Histological analysis of tumor tissues also showed that the ZnP@DHA/Pyro-Fe group induced more cell death than ZnP@DHA and PBS groups (Figure 3f). Furthermore, no obvious body weight loss and abnormal histopathology were observed in major organs for all treated groups (Figure S15–16), suggesting the lack of general toxicity for these treatments.

Pyro-Fe potentiates immunostimulatory effects of DHA

Endoplasmic reticulum (ER) stress and ROS production are two essential intracellular pathways for immunogenic cell death (ICD).51, 52 When occurring simultaneously, they activate danger signaling pathways to allow the trafficking of damage associated molecular patterns (DAMPs) to extracellular space. ICD releases tumor-associated antigens, DAMPs, and pro-inflammatory cytokines to facilitate the presentation of antigens to naïve T cells for an antigen-specific immune response against solid tumors.53 Calreticulin (CRT), an indicator of ICD, is transported to the cell surface in response to ER stress and serve as an “eat-me” signal to enhance the engulfment of dying tumor cells and their debris by immature dendritic cells (DCs) and macrophages.54, 55 We demonstrated that DHA effectively induced immunogenic phenotypes of tumor cells, as evidenced by CRT exposure on the surface of 30.6% cells (Figure 4a,c). The translocation of CRT biomarker to tumor cell surface was confirmed by CLSM imaging (Figure 4b). In the presence of Pyro-Fe, CRT exposure was further enhanced as shown by the increases in CRT fluorescence intensity and CRT-positive cells. We also quantified the release of HMGB-1, another ICD marker that stimulates antigen presentation to T cells, from treated cells by enzyme-linked immunosorbent assay (ELISA). As shown in Figure 4d, incubation with ZnP@DHA increased HMGB-1 release (12.30 ± 0.90 ng/mL) from cancer cells, which was further increased upon treatment with ZnP@DHA/Pyro-Fe (15.22 ± 0.36 ng/mL) compared to PBS (2.51 ± 0.25 ng/mL). To validate the immunogenicity beyond induction of ICD, we co-cultured as-treated cells with bone marrow-derived dendritic cells (BMDCs) and characterized the expression of functional markers for antigen presentation. As shown in Figure S17, BMDCs incubated with cancer cells that had been treated with ZnP@DHA or ZnP@DHA/Pyro-Fe presented higher expression of co-stimulatory markers CD80 and CD86 as well as MHC-II, indicating that ferroptosis-induced ICD promotes antigen presentation.

Figure 4.

Figure 4.

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Pyro-Fe potentiates immunostimulatory effects of ZnP@DHA. (a-c) CRT exposure on the cell surface of CT26 cells after treatment with ZnP@DHA and/or ZnP@Pyro-Fe, assessed by flow cytometry analysis (a,c) and CLSM (b), respectively (n = 3). Scale bars: 20 μm. (d) HMGB-1 release from tumor cells treated with ZnP@DHA and/or ZnP@Pyro-Fe (n = 3). (e) Priming of T-cell responses trigged by ZnP@DHA and/or ZnP@Pyro-Fe. Dead and dying MC38 tumor cells treated with ZnP@DHA and/or ZnP@Pyro-Fe were injected into the right footpads of C57BL/6 mice to determine the capacity of draining lymph node cells in producing IFN-γ in response to MC38 lysates (n = 3). Statistical significance: *P < 0.05, **P < 0.01 and ***P < 0.001.

We also determined the ability of MC38 cells treated with ZnP@DHA/Pyro-Fe to prime T cells in vivo by inoculating these dead and/or dying cells into the footpads of C57BL/6 mice. Six days later, popliteal lymph node cells were harvested from these mice and re-stimulated with MC38 lysates. IFN-γ production by these activated T cells was then detected by an ELISA kit. As shown in Figure 4e, cells treated with ZnP@DHA (3.44 ± 0.42 ng/mL) and ZnP@Pyro-Fe (0.98 ± 0.18) primed T cells for IFN-γ production compared to PBS (0.43 ± 0.16 ng/mL), but the combination treatment further enhanced T cell priming for IFN-γ production (5.42 ± 0.81 ng/mL).

ZnP@DHA/Pyro-Fe enhances intratumoral immune cell infiltration

Given DHA induces immunogenicity in tumor cells, we investigated whether ZnP@DHA/Pyro-Fe could elicit antitumor immunity in vivo by determining the changes of intratumoral immune cell populations 12 days after i.p. injection of ZnP@DHA or ZnP@DHA/Pyro-Fe. Compared to PBS, ZnP@DHA treatment showed significantly higher infiltration of CD4+ T cells (8.6 ± 2.1 %), NK cells (3.5 ± 2.0 %), and B cells (1.6 ± 0.8 %) in tumors over PBS treatment (4.1 ± 1.9 %, 1.1 ± 0.8 %, 0.3 ± 0.2 %, respectively) (Figure 5a–e). ZnP@DHA/Pyro-Fe treatment significantly increased the infiltration of CD8+ T cells (5.9 ± 1.8 %) in addition to CD4+ T cells (6.7 ± 1.9 %), NK cells (2.4 ± 1.3 %), and B cells (2.1 ± 1.3 %) into tumors over the PBS control (Figure 5a). The increased CD8+ T cells in tumors were confirmed by immunofluorescence assay. ZnP@DHA/Pyro-Fe treatment instigated more CD3ɛ+ cell infiltration in the tumor tissues than ZnP@DHA treatment and most of the tumor-infiltrating CD3ɛ+ cells were CD8+ (Figure 5f). These results show the ability of ZnP@DHA/Pyro-Fe in promoting CD8+ T-cell infiltration into tumors. As CD8+ T cells are the predominant T-cell subset responsible for antitumor immune response, the increased infiltration of CD8+ T cells suggests the potential of generating potent immune response against tumors. This conjecture was supported by the observation that treatment with ZnP@DHA/Pyro-Fe significantly increased the number of tumor-specific IFN-γ producing CD8+ T cells as determined by an IFN-γ enzyme-linked immunospot (ELISpot) assay (Figure 5g).

Figure 5.

Figure 5.

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ZnP@DHA/Pyro-Fe enhances intratumoral immune cell infiltration. (a-d) The infiltration of macrophages and dendritic cells in tumors 12 days after the first treatment by immunofluorescence and flow cytometry (n=5). (e) The infiltration of CD8+ T cells, CD4+ T cells, NK cells, and B cells in tumors 12 days after the first treatment by flow cytometry of cell-surface staining (n = 5). (f) Immunofluorescence analysis showing the infiltration of CD8+ T cells 12 days after the first treatment. (g) Antigen-specific IFN-γ producing T cells detected by ELISpot assay 12 days after the first treatment (n = 5). Statistical significance: *P < 0.05, **P < 0.01 and ***P < 0.001.

ZnP@DHA/Pyro-Fe sensitizes tumors to checkpoint blockade immunotherapy

To determine whether the immune response triggered by ZnP@DHA/Pyro-Fe could sensitize tumors to checkpoint blockade therapy, we first evaluated the PD-L1 expression after treatment in vitro. Flow cytometric analysis showed that ZnP@DHA or ZnP@DHA/Pyro-Fe upregulated PD-L1 expression on CT26 cells (Figure S18), indicating the potential synergy for combination with an anti-PD-L1 antibody. We then investigated the antitumor activity of ZnP@DHA/Pyro-Fe plus anti-PD-L1 (α-PD-L1, Clone: 10F.9G2, Catalog No. BE0101, BioXCell) on both CT26 and MC38 colorectal cancers. Anti-PD-L1 showed a slight inhibitory effect on CT26 tumors at a dose of 75 μg/mouse, but no effect on MC38 tumors. This difference can be attributed to relatively immunogenic CT26 tumors with large amounts of granzyme B+ NK cells and reinvigorated CD8+ T cells and more immunosuppressive MC38 tumors with a large amount of monocytic myeloidderived suppressor cells (mMDSCs).56 Combination treatment with ZnP@DHA/Pyro-Fe and anti-PD-L1 significantly retarded the growth on both CT26 and MC38 tumors, leading to TGI values of 85% and 82.3%, respectively (Figure 6a–d).

Figure 6.

Figure 6.

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ZnP@DHA/Pyro-Fe treatment sensitizes tumors to α-PD-L1. (a-d) Growth curves (a,c) and final weights (b,d) of CT26 (a,b) and MC38 (c,d) tumors treated with ZnP@DHA/Pyro-Fe with or without α-PD-L1 (n = 6). (e,f) Therapeutic effect of the combination with anti-PD-L1 on Rag2−/− mice (e) and on CD8+ T cells depleted C57BL/6 mice (f) (n = 5). Red arrows refer to nanoparticle and/or anti-PD-L1 injections while green arrows refer to injection of anti-CD8 antibody or isotype control.

To confirm the contribution of the immune system to tumor control, we tested the therapeutic effect of ZnP@DHA/Pyro-Fe plus α-PD-L1 on MC38 tumors implanted in immunodeficient Rag2−/− mice which lack mature lymphocytes, including both T and B cells. As expected, the effect of α-PD-L1 in the combination therapy was completely abrogated on Rag2−/− mice with ZnP@DHA/Pyro-Fe plus α-PD-L1 treatment showing similar antitumor efficacy as ZnP@DHA/Pyro-Fe treatment (Figure 6e). It is worth noting that the therapeutic effect of ZnP@DHA/Pyro-Fe against MC38 tumors in immunodeficient Rag2−/− mice was much worse than that against MC38 tumors in immunocompetent C57BL/6 mice, underscoring the important role of the immune system on the therapeutic response. To further determine whether the antitumor effect depends on the host CD8+ T cells, we depleted CD8+ T cells on MC38 tumors-bearing mice via i.p. administration of an anti-CD8 monoclonal antibody. The therapeutic effect of ZnP@DHA/Pyro-Fe plus anti-PD-L1was significantly weakened with the depletion of CD8+ T cells (Figure 6f). Taken together, these findings support our hypothesis that involvement of immune system is crucial to the therapeutic responses mediated by ZnP@DHA/Pyro-Fe.

Conclusion

We developed Zn-pyrophosphate core-shell nanoparticles for the co-delivery of Chol-DHA and Pyro-Fe to colorectal tumors in mouse models. Reduced Pyro-Fe can catalyze the decomposition of DHA to produce ROS in O2-independent manner, causing ferroptosis of cancer cells. ZnP@DHA/Pyro-Fe particles prolong circulation half-lives of Chol-DHA and Pyro-Fe to increase their uptake in tumors. The enhanced delivery of Chol-DHA and Pyro-Fe to tumors induces more ROS production and elicits significant tumor inhibition in vivo. Importantly, reduced Pyro-Fe enhances the immunostimulatory effect of DHA, as evidenced by more CRT exposure and higher HMGB-1 release. The combination of DHA and Pyro-Fe enhances intratumoral CD8+ T cell infiltration and potentiates anti-PD-L1 immune checkpoint blockade in non-immunogenic colorectal tumors. With broad efficacy, low toxicity, and high selectivity, DHA delivered by nanoparticles is a promising candidate for cancer treatment and for combination therapy with immune checkpoint blockade.

Methods

Materials, cell Lines, and animals.

All of the chemicals and reagents were purchased from Sigma-Aldrich and Fisher (USA) unless otherwise noted. 1,2-dioleoyl-snglycero-3-phosphate (DOPA), 1,2-dioleyl-sn-glycero-3-phosphocholine (DOPC), cholesterol, and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino(polyethylene glycol)2000] (DSPE-PEG2k) were purchased from Avanti Polar Lipids (USA). Murine colon adenocarcinoma cell CT26 and MC38 cells were purchased from the American Type Culture Collection (ATCC, Rockville, MD) and cultured in RPMI 1640 and Dulbecco’s Modified Eagle’s Medium (DMEM), respectively, supplemented with 10% FBS, 100 U/mL penicillin G sodium and 100 μg/mL streptomycin sulfate in a humidified atmosphere containing 5% CO2 at 37°C. BALB/c female mice (6 weeks, 18–22 g), C57BL/6 female mice (6 weeks, 18–22 g), and Rag2−/− female mice (6 weeks, 18–22 g) were provided by Harlan-Envigo Laboratories, Inc (USA). The study protocol was reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Chicago.

Synthesis and characterization of ZnP@DHA, ZnP@Pyro-Fe and ZnP@DHA/Pyro-Fe.

ZnP core was synthesized according to our previously reported method.34 ZnP was then mixed with DOPC, cholesterol, DSPE-PEG2k (2:1:1:1 in molar ratio) in THF, added to 500 μL of 30% (v/v) ethanol/water at 50 °C in the presence of Chol-DHA and/or Pyro-Fe, respectively, and stirred at 1,700 rpm for 1 min. After complete evaporation of THF and ethanol, ZnP@DHA, ZnP@Pyro-Fe, and ZnP@DHA/Pyro-Fe were subjected to particle size and zeta potential determination by dynamic light scattering (DLS) using a Zetasizer (Nano ZS, Malvern, UK) and transmission electron microscopy (Tecnai Spirit, FEI, USA) imaging to observe the morphology of nanoparticles. In order to determine the drug loadings, nanoparticles were centrifuged at 13,000 rpm for 30 min and resuspended in THF to dissolve the lipid layer. The amounts of released Chol-DHA and Pryo-Fe were then determined by liquid chromatography-mass spectrometry (6540 Q-Tof MS-MS, Agilent, USA) and inductively coupled plasma-mass spectrometry (Masshunter 7700, Agilent, USA), respectively. To test colloidal stability, nanoparticles diluted in phosphate buffered solution containing 5 mg/mL BSA were incubated at 37 °C, and the changes in particle size and PDI over time were determined by DLS.

ROS generation in solution.

ZnP@DHA, ZnP@Pyro-Fe, or ZnP@DHA/Pyro-Fe with or without 5 mM cysteine was dispersed in 0.5% Triton X-100 that contains 10 μM H2-DCFDA (Thermo Fisher Scientific, USA). The ROS generation in the presence or absence of O2 was investigated by determining the change of DCF fluorescence over time.

Cell EM.

CT26 cells were treated with ZnP@DHA, ZnP@Pyro-Fe or ZnP@DHA/Pyro-Fe at a dose of 10 μM Pyro-Fe or/and 10 μM DHA for 24 h. Cells were then collected and fixed with OsO4 staining solution for observation under Tecnai Spirit TEM.

Cell cycle assay.

CT26 cells seeded in six-well plates (5 × 104 cells/well) were treated with ZnP@DHA, ZnP@Pyro-Fe or ZnP@DHA/Pyro-Fe at a dose of 10 μM Pyro-Fe or/and 10 μM DHA for 24 h. Treated cells were then collected, washed with PBS, fixed with 70% ethanol at 4°C overnight. After treatment with RNase A for 45 min and PI staining for 30 min, cells were subjected to cytometric analysis to determine the alteration of cell cycle.

Intracellular ROS generation.

CT26 cells were treated with ZnP@Pyro-Fe, ZnP@DHA, or ZnP@DHA/Pyro-Fe at a concentration of 10 μM Pyro-Fe or/and 10 μM DHA for 24 h. After incubation with 10 μM H2-DCFDA (Thermo Fisher) for further 1 h, cells were collected, washed with PBS, and then analyzed by flow cytometry.

DNA double strand break.

After treatment with ZnP@Pyro-Fe, ZnP@DHA, or ZnP@DHA/Pyro-Fe at a concentration of 10 μM Pyro-Fe or/and 10 μM DHA for 24 h, CT26 cells were fixed with 4% paraformaldehyde at room temperature for 10 min, permeabilized with 0.1% Trition X-100 at room temperature for 10 min, blocked with 2% BSA at room temperature for 1 h, and stained with FCS DNA damage kit (Invitrogen) for the oxidative DNA damage detection by γ-H2AX foci. After staining with DAPI for another 10 min, the cells were washed twice with PBS and observed under CLSM (Olympus, FV1000).

CRT exposure analysis.

CT26 cells were incubated with ZnP@Pyro-Fe, ZnP@DHA, or ZnP@DHA/Pyro-Fe at a dose of 10 μM Pyro-Fe and/or 10 μM DHA for 24 h, collected, incubated with Alexa Fluor 488-CRT antibody (Enzo Life Sciences, USA) for 2 h, stained with PI for 10 min, and analyzed by flow cytometer to determine CRT exposure. The fluorescence intensity of stained cells was gated on PI− cells. For surface detection of CRT, CT26 cells seeded on 10 mm2 glass coverslips placed in 6-well plates were treated with ZnP@Pyro-Fe, ZnP@DHA, or ZnP@DHA/Pyro-Fe at a dose of 10 μM Pyro-Fe and/or 10 μM DHA for 24 h, washed with PBS three times, incubated with Alexa Fluor 488-CRT antibody for 2 h, stained with DAPI for 10 min, and observed under CLSM for CRT expression on the cell membrane.

HMGB-1 release.

CT26 cells were treated with ZnP@Pyro-Fe, ZnP@DHA, or ZnP@DHA/Pyro-Fe at a dose of 10 μM Pyro-Fe and/or 10 μM DHA for 24 h. After that, the medium was collected, and the release of HMGB-1 was detected by ELISA according to manufacturer instructions (Chondrex, USA).

Priming assay.

After treatment with ZnP@Pyro-Fe, ZnP@DHA, or ZnP@DHA/Pyro-Fe at a dose of 10 μM Pyro-Fe and/or 10 μM DHA for 24 h, 1 × 106 MC38 cells were injected into the footpad of C57Bl/6 mice. Six days later, popliteal lymph node cells were collected and prepared into single-cell suspension by homogenizing and filtering through a sterile cell strainer (70 μm; Becton Dickinson, USA). 1 × 105 lymph node cells were incubated in 200 μL medium in 96-well plates in the presence of MC38 cell lysates killed by freeze-thaw cycle for 3 days, and then IFN-γ secreted to the supernatants was determined by ELISA (eBioscience, USA).

Pharmacokinetics and Biodistribution.

ZnP@DHA/Pyro-Fe was doped with 20% Pyrolipid (relative to the sum of Pyro-Fe and Pyrolipid) and used for pharmacokinetic (PK) and biodistribution (BD) analysis. Particles were dosed intravenously to SD rats (at a dose of 4 mg/kg Pyro-Fe) for PK and to CT26-bearing mice (at a dose of 10 mg/kg Pyro-Fe) for BD studies. Blood samples were collected at 5 min, 0.5 h, 1 h, 3 h, 5 h, 8 h and 24 h from rats post injection and tumors were collected from mice 24 h post injection. Blood samples were centrifuged at 3000 rpm for 5 min to obtain plasma and then measured by UV-Vis. Tumors were homogenized with 0.5% triton X-100 and then measured by IVIS for their fluorescence signals. UV-Vis and fluorescence data were compared to calibration curves in order to obtain plasma Pyrolipid concentrations.

In vivo antitumor efficacy.

1 × 106 CT26 cells were subcutaneously injected into the right flank region of 6-week-old BALB/c mice to establish CT26-bearing mice model. Similarly, MC38-bearing mice model were established by subcutaneously injecting 1 × 106 MC38 cells into the right flank region of 6-week-old C57Bl/6 wild-type or Rag2−/− mice. 12 days later, mice were intraperitoneally (i.p.) injected with ZnP@DHA/Pyro-Fe particles or controls with equivalent doses of 10 mg/kg DHA, 1.96 mg/kg Fe, and/or 75 μg PD-L1 antibody once every 3 days for 3 doses for CT26 tumors and 4 doses for MC38 tumors. To evaluate the therapeutic efficacy, tumor growth and body weight evolution were monitored. Tumor size was measured with a digital caliper every day and calculated as follows: (width2 × length)/2. All mice were euthanized when the tumor size of the control group exceeded 2 cm3 and the excised tumors were photographed and weighed. The tumors were embedded in optimal cutting temperature medium, sectioned at 5 μm thickness and subjected to haematoxylin and eosin (H&E) stain for histopathological analysis, SLC7A11 (Cell Signaling, USA) immunostaining for ferroptosis, and TUNEL (Invitrogen, USA) assay for apoptosis. Hearts, livers, lungs, spleens, and kidneys were also excised and sectioned at 5 μm thickness, stained with H&E, and observed under a light microscope (Pannoramic Scan Whole Slide Scanner, Perkin Elmer, USA).

Depletion of T cells.

In addition to treatment with ZnP@DHA/Pyro-Fe plus anti-PD-L1, tumor-bearing C57Bl/6 mice were treated with anti-CD8 (0.2 mg/mouse, i.p., OKT-8, BioXCell, USA) at day 0 and day 5 to deplete CD8+ T cells. Treatment with IgG2a (0.1 mg, C1.18.4, BioXCell, USA) was used as a control. The tumor volumes were recorded as above.

ELISpot assay.

ELISPOT assay (Mouse IFN-γ ELISPOT Ready-SET-Go!®; Cat. No. 88-7384-88; eBioscience) was performed to determine tumor-specific immune responses to IFN-γ in vitro. A Millipore Multiscreen HTS-IP plate was coated with anti-Mouse IFN-γ capture antibodies overnight at 4 °C. Single-cell suspensions of splenocytes obtained from MC38 tumor-bearing mice 12 days after the first treatment were seeded onto the antibody-coated plate at a concentration of 2 × 105 cells/well, incubated with or without KSPWFTTL (10 mg/ml; PEPTIDE 2.0) for 48 h at 37°C. Biotin-conjugated anti-IFN-γ detection antibody was then added into the plate and incubated at room temperature for 2 h, followed by Avidin-HRP incubation for another 2 h at room temperature. After addition of 3-amino-9-ethylcarbazole (AEC) substrate solution (Sigma, Cat. AEC101), IFN-γ spots were detected by ELISpot reader.

Flow cytometry assay for immune response.

Twelve days after the first treatment, tumors were harvested, incubated with 1 mg/mL collagenase I (Gibco™, USA) at 37 °C for 1 h, ground with the rubber end of a syringe, filtered through nylon mesh filters, and washed with PBS. The single-cell suspension was incubated with anti-CD16/32 (clone 93; eBiosciences) for 30 min to reduce nonspecific binding to FcRs, and further stained with the following fluorochrome-conjugated antibodies for another 30 min: CD45 (30-F11), CD3ɛ (145–2C11), CD4 (GK1.5), CD8 (53–6.7), B220 (RA3–6B2), NKp46 (29A1.4). After that, cells were subjected to flow cytometry by LSR FORTESSA (BD Biosciences) and data was analyzed by FlowJo software (TreeStar, Ashland, OR).

Immunofluorescence assay.

Twelve days after the first treatment, tumors were harvested, sectioned at 5 μm thickness, fixed in acetone at −20°C for 10 min, blocked with 2% BSA for 1 h, and incubated with primary antibodies against CD3ɛ (Santa Cruz) and CD8 (Thermo Scientific) at 4 °C overnight, followed by incubation with dye-conjugated secondary antibodies at room temperature for another 1 h. After DAPI staining, the sections were washed with PBS and observed using CLSM (Olympus, FV1000).

Supplementary Material

1

Acknowledgements

We thank Ziwan Xu for experimental help. We acknowledge the National Cancer Institute (1R01CA216436 and 1R01CA223184) and the University of Chicago Medicine Comprehensive Cancer Center (NIH CCSG: P30 CA014599) for funding support.

Footnotes

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Author Credit Statement:

W.H., X.D., K.N., and W.L. conceived and designed the experiments. W.H., X.D., K.N., Y.L., and C.C. performed experiments. W.H., X.D., K.N., Y.L., C.C., and W.L. analyzed the data. W.H., X.D., K.N., Y.L., C.C., and W.L. wrote the manuscript.

Supplementary information is available in the online version of the paper.

Declaration of interests

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.

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