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Abstract
Background Despite its potential utility in delivering direct tumor killing and in situ whole-cell tumor vaccination, tumor cryoablation produces highly variable and unpredictable clinical response, limiting its clinical utility. The mechanism(s) driving cryoablation-induced local antitumor immunity and the associated abscopal effect is not well understood.
Methods The aim of this study was to identify and explore a mechanism of action by which cryoablation enhances the therapeutic efficacy in metastatic tumor models. We used the subcutaneous mouse model of the rhabdomyosarcoma (RMS) cell lines RMS 76-9STINGwt or RMS 76-9STING-/-, along with other murine tumor models, in C57BL/6 or STING-/- (TMEM173-/-) mice to evaluate local tumor changes, lung metastasis, abscopal effect on distant tumors, and immune cell dynamics in the tumor microenvironment (TME).
Results The results show that cryoablation efficacy is dependent on both adaptive immunity and the STING signaling pathway. Contrary to current literature dictating an essential role of host-derived STING activation as a driver of antitumor immunity in vivo, we show that local tumor control, lung metastasis, and the abscopal effect on distant tumor are all critically dependent on a functioning tumor cell-intrinsic STING signaling pathway, which induces inflammatory chemokine and cytokine responses in the cryoablated TME. This reliance extends beyond cryoablation to include intratumoral STING agonist therapy. Additionally, surveys of gene expression databases and tissue microarrays of clinical tumor samples revealed a wide spectrum of expressions among STING-related signaling components.
Conclusions Tumor cell-intrinsic STING pathway is a critical component underlying the effectiveness of cryoablation and suggests that expression of STING-related signaling components may serve as a potential therapy response biomarker. Our data also highlight an urgent need to further characterize tumor cell-intrinsic STING pathways and the associated downstream inflammatory response evoked by cryoablation and other STING-dependent therapy approaches.
- Adaptive Immunity
- Immunomodulation
- Sarcoma
- Tumor Microenvironment
- Translational Medical Research
Data availability statement
All data relevant to the study are included in the article or uploaded as supplementary information. Not applicable.
This is an open access article distributed in accordance with the Creative Commons Attribution Non Commercial (CC BY-NC 4.0) license, which permits others to distribute, remix, adapt, build upon this work non-commercially, and license their derivative works on different terms, provided the original work is properly cited, appropriate credit is given, any changes made indicated, and the use is non-commercial. See http://creativecommons.org/licenses/by-nc/4.0/.
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WHAT IS ALREADY KNOWN ON THIS TOPIC
Cryotherapy has been applied clinically, mostly for tumor debulking and pain control.
Although anecdotal evidence of the abscopal effects has been observed in patients with advanced stage cancer, the essential immunological mechanisms and drivers of this clinical response are not well understood.
Conventional wisdom suggests cryoablation induces immunogenic cell death thereby activating the cGAS-STING pathway in host myeloid cells.
WHAT THIS STUDY ADDS
We demonstrate an unequivocal role of an ongoing and functional STING pathway within the residual, live tumor cells as a major driver of local control and the systemic abscopal effects on advanced tumors following cryoablation, further supported by profiling variable STING pathway components among sarcomas.
HOW THIS STUDY MIGHT AFFECT RESEARCH, PRACTICE OR POLICY
Our new knowledge provides the foundation to further study the role of the STING-IFNβ signaling axis and its downstream chemokine and cytokine responses in improving cryoablation efficacy and as biomarkers of clinical cryoablation therapy response.
Background
Cryoablation1 is an established clinical intervention in which freezing of tissues is achieved by passing argon gas or liquid nitrogen via a probe under direct CT/MRI/ultrasound-guided visualization with repeated freeze-and-thaw cycles, reaching an ultra-cold temperature of −40°C to −70°C during the freeze cycles and −20°C to −40°C during the thaw cycles.2–4 Cryoablation has been previously used to treat a diverse group of cancers, often as a palliative debulking measure for unresectable tumors.2–6 The ice ball created by cryoablation results in direct mechanical destruction of tissues and cells due to osmotic shock, ruptured cell membranes from ice crystal formation, and ischemia and reperfusion injury secondary to disruption of blood supply from vascular stasis. With the rising interests in cancer immunotherapy, cryoablation has shown synergy with host immune activation under certain circumstances. Preclinical studies demonstrated that cryoablation-mediated tumor destruction induces the activation and maturation of immune cells critical for an effective antitumor immune response, and this immune activation can synergize further with the administration of a checkpoint blocking antibody such as anti-CTLA-4 Ig.7 The release of tumor antigens from dying tumor cells can activate tumor-specific immune responses through antigen presentation by dendritic cells (DCs) to T cells, thereby allowing these adaptive cells to survey and destroy both ablated and non-ablated tumors at distant sites; this phenomenon is known as the ‘abscopal effect’.8 9 In addition to shedding tumor antigens, cryoablated tumor tissues predominantly undergo immunogenic cell death (ICD),10–12 which preserves intact tumor cell-derived antigens and releases damage-associated molecular patterns (DAMPs) such as double-stranded DNA (dsDNA), ATP, and heat shock proteins (HSP70 and HMGB1), which promotes innate immune cell maturation including DC activation and effective cross-presentation of tumor cell-derived antigens to T cells.13 14 In support of this observation, studies have shown that variable duration and tissue applications of cryoablation resulted in differing necrotic and apoptotic cell death cores as well as varying immune activation landscapes in the treated area.12 Despite these theoretical advantages, clinical response to cryoablation is highly variable and suffers from the absence of a reliable biomarker that predicts a beneficial therapeutic outcome of this approach.15
To this end, mounting evidence suggests that activation of the cyclic GMP-AMP synthase–stimulator of interferon genes (cGAS-STING) innate immune sensing pathway in myeloid cells may be a critical link to ICD.16 17 STING is an essential protein for the innate immune defense against a wide array of microbial pathogens. STING-related contributions to homeostasis and disease pathogenesis have been described in innate myeloid immune cells,18 as well as T,19–21 non-hematopoietic stromal22 and endothelial23–25 cells. Detection of cytosolic dsDNA in the context of viral and bacterial infections,26–29 aging and damaged cells,30 or cancer,31–33 is performed by the cytoplasmic DNA sensor, cGAS. cGAS catalyzes the formation of the second messenger cyclic dinucleotide (CDN), 2′3′-cyclic-GMP-AMP (cGAMP).34 Accumulation of cGAMP in the cytoplasm activates the cytosolic domain of its ER membrane receptor, STING, via phosphorylation of the serine-365 (S365) residue in the C-terminal tail of STING,20 21 which initiates the translocation of this scaffolding polymer to the Golgi, and the recruitment of TANK-binding kinase 1 (TBK1) and interferon (IFN) regulator factor 3 (IRF3), on which the phosphorylation of all three components occurs, ultimately leading to the production of type I IFN, in particular, IFN-β that further activates antigen presenting cells (APCs) to enhance cross-presentation of tumor antigens to CD8+ T cells.16 17 26 35–37
Within tumor cells, the cGAS-STING pathway can be activated by cytosolic dsDNA originating from micronuclei secondary to the genomic instability of tumors, particularly those with mismatch repair deficiencies,33 or after DNA-damaging therapies.32 38–40 The cytosolic dsDNA activates cGAS in tumor cells, leading to cGAMP production41 42 which is exported as a paracrine immunotransmitter, subjected to degradation by extracellular hydrolase, ENPP1,43 and imported into host myeloid cells through the SLC19A1 transporter,41 44 thereby triggering a type I IFN response and subsequent induction of antitumor immunity. Considering all available literature together, it is widely accepted that while cGAMP could come from either tumor cells or myeloid cells, activation of the host cell–derived, in particular, myeloid cell–derived STING pathway is the critical and converging determinant of antitumor immune activation.23 41 Based on this, clinical approaches using direct intratumoral injection of a STING agonist (SNX281) have been tried (NCT04609579).45 46
Our current study directly investigates the functional role of STING signaling in therapeutic response to cryoablation by determining whether there is a positive correlation between tumor ICD post-cryoablation and STING pathway activation. As recent evidence suggests that tumor cell-intrinsic STING is critical to the production of type I IFNs and promotion of CD8+ T cell activation via enhancing the function of APCs,33 it has been proposed that tumor cell-intrinsic STING enhances tumor immunogenicity by increasing immune cell activation and triggering tumor cell regression.47–49 Therefore, we dissect the functional contribution of tumor cell-intrinsic STING pathway activation versus that derived from host tissues in response to cryoablation. Our results challenge the current paradigm of antitumor immunity activation resulting from myeloid STING activation in the tumor microenvironment (TME) and provide evidence that tumor cell-derived STING signaling plays an indispensable role in determining the clinical efficacy of not only cryoablation but also STING agonistic therapies.
Methods
Mice
As rhabdomyosarcoma (RMS) cell line RMS 76-9 originated from male mice, female mice spontaneously reject inoculated tumor growth at a high frequency. To eliminate confounding issues related to H-Y dependent tumor rejection, only male mice were used in this study. Male C57BL/6J and C57BL/6J-Sting1gt/J (STING-/-) mice were obtained from The Jackson Laboratory (Bar Harbor, Maine, USA). Male B6 nude mice (B6.Cg/NTac-Foxn1nu NE10) were obtained from Taconic. Animals were housed, bred, and handled in the Animal Resource Center facilities at Case Western Reserve University according to approved protocols.
Cell lines
Parental mouse sarcoma cell lines RMS 76-9 and M3-9-M (embryonal RMS derived from C57BL/6 transgenic for hepatocyte growth factor and heterozygous for mutated P53) were generously provided by C. Mackall.50 51 Murine pancreatic cancer cell line, KPC1242, was kindly provided by Dr Stanley Huang (Department of Pathology, Case Western Reserve University) and was originally derived by the Tuveson laboratory (Cold Spring Harbor, New York, USA) from a spontaneously arising tumor from KRasLSL.G12D/+-p53R172H/+-Pdx-Cre (KPC) transgenic mice on the C57BL/6 background.52 All cells were grown in RPMI 1640 supplemented with 10% Fetal Bovine Serum, 1% 4-(2-hydroxyethyl)-1-piperazine ethanesulfonic acid (HEPES), 1% non-essential amino acids, and 1% penicillin and streptomycin, 1% sodium pyruvate, and 1% L-glutamine. RMS 76-9STING-/- knockout cells were generated via CRISPR using guide RNA (gRNA) 5′-TACTTGCGGTTGATCTTACC-3′. gRNA guide was complexed with TracrRNA, then added to S.P.Cas9 (QB3 MacroLab) to form a ribonucleoprotein. Electroporation was performed using the Lonza Nucleofector (SE Cell Line 96-well Nucleofector Kit). Single-cell colonies were isolated (clones 2, 7 and 8), and gene knockout efficiency was confirmed by western blot. RMS 76-9STINGwt mNeonGreen and RMS 76-9STING-/- E2-Crimson cells were generated by transfection of RMS 76-9 with pNCS-mNeonGreen53 plasmid (purchased from Allele Biotechnology) and CSII-prEF1a-E2-Crimson-NLS plasmid (Addgene plasmid number 125263) using Lipofectamine 3000 Transfection Reagent (Thermo Fisher Scientific) following the manufacturers’ protocols. Bright cells were sorted by flow cytometry using a BD FACS ARIA flow cytometer.
Cryoablation of primary tumors
To explore the effects of cryoablation on tumor growth, C57BL/6, Nude and STING-/- mice were inoculated subcutaneously in the back with 1×106 RMS 76-9 cells, which were resuspended in 50 μL of Hanks’ Balanced Salt Solution (HBSS). To investigate the role of host STING in tumor growth, C57BL/6 and STING-/- mice were inoculated subcutaneously in the back with 1×106 RMS 76-9 cells resuspended in 50 μL of HBSS. To investigate the role of tumor-intrinsic STING in tumor growth, wild-type (WT) and STING-/- mice were inoculated subcutaneously in the back with 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- cells, or their M3-9-M and KPC1242 counterparts, resuspended in 50 μL of HBSS.
Abscopal effects of secondary tumors following cryoablation of primary tumor
To determine the extent of the abscopal effect from tumor cell-intrinsic STING post local cryoablation, C57BL/6 and STING-/- mice were inoculated subcutaneously with 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- cells resuspended in 50 μL of HBSS on the left flank in the back and with 5×105 RMS 76-9STINGwt cells resuspended in 50 μL of HBSS on the corresponding opposite (right) flank in the back of the same mice. Tumors on the left flank were treated with local cryoablation and then both flanks were monitored twice a week for tumor growth. Tumor sizes were calculated as follows: volume = (d1*d2∧2)/2, where d1 is the long measurement and d2 is the shorter measurement.
Tumor cryoablation procedure
All cryoablation procedures were performed using the ProSense Cryosurgical System (IceCure Medical). This device uses liquid nitrogen to reach cooling temperatures (–196°C). When tumor sizes became approximately 100 mm3, mice were anesthetized with 1%–3% isoflurane and the cryoprobe was placed directly on the primary tumor mass, and cryoablation was performed at a high rate of freeze with each tumor undergoing three cycles of freezing (1 min freeze and 3 min thaw).
In vivo depletion of CD4+ and CD8+ T cells
As previously addressed, mice were inoculated subcutaneously in the back with 1×106 RMS 76-9 cells, and primary tumors were cryoablated 10 days post inoculation. CD4+ or CD8+ T cell subsets were depleted in C57BL/6 tumor-bearing mice by administering 100 µg (in 100 µL of HBSS) of either anti-CD4 antibody (GK1.5) or anti-CD8 antibody (2.43) via intraperitoneal injection 1 day before cryoablation (9 days post inoculation) of tumor, as well as 3 and 6 days after cryoablation treatment (13 and 16 days post inoculation). The tumor volume was monitored every 3 days.
Bone marrow-derived dendritic cell (BMDC) culture
Single-cell suspensions of bone marrow cells were obtained from C57BL/6J and STING−/− mice. The cells were placed in 6-well plates and cultured in RPMI-1640 medium containing 10% FBS, 1% HEPES, 1% non-essential amino acids, 1% sodium pyruvate, 1% pen/strep, and 1% L-glutamine and supplemented with 15 ng/mL granulocyte macrophage-colony stimulating factor (GM-CSF) and 10 ng/mL interleukin-4 (IL-4). Fresh media with GM-CSF and IL-4 was added into culture on days 0, 3 and 5. BMDCs were harvested for co-culture on day 7. RMS 76-9 cells were treated with cryoablation and incubated for 6 hours. BMDCs were co-cultured with cryo-ablated RMS 76-9 cells or treated with 100 μg/mL 5,6-dimethylxanthenone-4-acetic acid (DMXAA) in 24-well plates at the ratio of 1:1 for an additional 6 hours for quantitative PCR (qPCR) assay and 12 hours for flow cytometric analysis. ELISA and qPCR assay were performed on purified CD11c+ cells, which were isolated with EasySep Mouse CD11c Positive Selection Kit II (STEMCELL).
Bone marrow-derived macrophage (BMDM) culture
Single-cell suspensions of bone marrow cells were resuspended at 1×106 per 1 mL in Dulbecco’s Modified Eagle Medium (DMEM) media containing 10% FBS, 1% HEPES, 1% non-essential amino acids, 1% sodium pyruvate, 1% Pen/Strep, and 1% L-glutamine and supplemented with 20%–25% LADMAC (mouse bone marrow-derived cell line producing colony stimulating factor-1) supernatant (10 mL/100 mm dish). Supernatant was removed and fresh media (10 mL 20%–25% LADMAC in 10% FBS) was added on days 5 and 8. On day 10, macrophages were harvested and cultured in 6-well plates (1×106/well) in 3 mL of LADMAC media/well. Spent supernatant was then removed and 3 mL of complete media was added on day 11. On day 12, macrophages were harvested and stimulated with 100 ng/mL LPS and 20 ng/mL IFN-γ (M1 polarization condition), or 20 ng/mL IL-4 (M2 polarization condition). Macrophages were harvested after 48 hours and analyzed by flow cytometry for the expression of M2 or M1 activation markers. For tumor co-culture experiments, macrophages were co-cultured with cryoablated or sham-ablated RMS 76-9 cells in 24-well plates at 1:1 ratio and analyzed by flow cytometry after 48 hours.
Tumor isolation and tissue preparation
Tumors were harvested either 3 days (qPCR and ELISA) or 7 days post cryoablation (flow cytometry), weighed for comparison against volume measurements, and then finely chopped with a razor blade. The tumor was then placed into a conical tube containing RPMI without supplement, collagenase D and DNase-1, then incubated at 37°C in a mixer for 1 hour. The mixture was then passed through a 40 µM strainer twice to obtain a single cell suspension. Purified CD11c+ cells were then isolated from the tumor suspension using an EasySep Mouse CD11c Positive Selection Kit II (STEMCELL) prior to qPCR. For ELISA, tumors were excised and homogenized in 1 mL phosphate-buffered saline (PBS). After homogenization, the concentration of IFN-β was measured with the Mouse Interferon Beta Serum Quantikine ELISA kit (R&D Systems) in accordance with the manufacturer’s instructions.
Flow cytometry and t-distributed stochastic neighbor embedding (t-SNE) analysis
For surface staining, 1×106 cells in 100 µL of Fluorescent Activated Cell Sorting (FACS) buffer were incubated with unlabeled rat anti-mouse blocking Fc antibody for 30 minutes (min) on ice followed by primary antibody staining or isotype control antibodies on ice in the dark for 45 min. After washing twice, cells were resuspended in 100 μL of FACS buffer and incubated with 7-AAD for 10 min. Cells were then washed twice with ice-cold FACS buffer. For intracellular flow cytometric analysis, 96-well plates were coated with 1 µg/mL unlabeled anti-CD3 for 90 min at 37°C. Harvested tumor cell suspensions were then plated with 1 μL/mL of GolgiStop (eBioscience) for 6 hours on the coated 96-well plates. Tumor cells were then fixed and permeabilization using the Cytofix/Cytoperm kit (BD Biosciences), then intracellularly stained with anti-granzyme B (GZMB) or anti-IFN-γ antibodies according to manufacturer’s instructions. Flow cytometry was performed on a Beckman Coulter CytoFLEX Flow Cytometer or BD Accuri C6 flow cytometer, and acquired data were then analyzed using the FlowJo analysis software. In addition to conventional FACS analysis, high-dimensional clustering using tSNE was performed on the same number of live CD45+ events from each sample, as indicated, which was exported and clustered based on the mean fluorescent intensity (MFI) of each marker of interest by the Rphenograph package. Antibodies used are as follows: anti-CD45.2-FITC (Clone: 104, Cat: 109806), anti-CD3-PE (Clone: 145-2 C11, Cat: 100308), anti-CD3-PEDazzle594 (Clone: 17A2, Cat: 100246), anti-CD4-PE-Cy7 (Clone: RM4-5, Cat: 100528), anti-CD8-PerCP/Cy5.5 (Clone: 53-6.7, Cat: 100734), anti-IFN-gamma-APC (Clone: XMG1.2, Cat: 505810), anti-Granzyme B-AF700 (Clone: QA16A02, Cat: 372221), anti-NK-1.1-FITC (Clone: PK136, Cat: 108706) anti-CD11c-FITC (Clone: N418, Cat: 117306), anti-F4/80-FITC (Clone: BM8, Cat: 123108), anti-CD206-PE (Clone: C068C2, Cat: 141706), anti-I-Ab-APC (Clone: AF6-120.1, Cat: 116418), anti-CD80-APC (Clone: 16-10 A1, Cat: 104714), anti-CD80-PE (Clone: 16-10 A1, Cat: 104708), anti-CD86-APC (Clone: GL-1, Cat: 105012) from BioLegend, anti-CD86-PE (Clone: GL1, Cat: 12-0862-82) from eBioscience, anti-CD45.2-PE-Cy5 (Clone: 30-F11, Cat: 553082) and anti-CD40-PE (Clone: 3/23, Cat: 553791) from BD Pharmingen. All flow cytometry analyses of the TME immune composition were performed with freshly isolated single cell suspensions obtained on the same day of tumor harvesting without freeze-thaw manipulations.
Quantitative PCR assay
TRIZOL Reagent or RNAspin Mini kit (GE Healthcare, UK) was used for RNA extraction and isolation, followed by DNAseI treatment to eliminate genomic DNA. RNA was quantified using the NanoDrop2000 spectrophotometer. Reverse transcription (RT) of 1 µg of total RNA was performed using the SuperScript First Strand Synthesis System for RT-PCR kit (Life Technologies, Carlsbad, California, USA). Real-time quantitative PCR was carried out on the resulting cDNA using SYBR Green PCR Master Mix (Life Technologies) on the ABI 7300 real-time PCR machine (Life Technologies) to determine the transcript expression of IL-12β and IFN-β; cytochrome C served as a reference gene. Analysis was performed using the delta-delta CT (2−ΔΔCT) method. The primer sequences were as follows: cytochrome c forward: CTGCCACAGCATGGATTATG; cytochrome c reverse: CATCATCATTAGGGCCATCC; IL-12β forward: CATCTGCTGCTCCACAAGAA; IL-12β reverse: TTGGTGCTTCACACTTCAGG; IFN-β forward: ACTTGAAGAGCTATTACTGGAGGG; IFN-β reverse: TTCCTGAAGATCTCTGCTCGG; Tmem173 forward: TGTCTCCCCATTCAGAAGCC; Tmem173 reverse: CATCTTCTGCTTCCTAGACCGG.
Western blot analysis
Cells were washed with PBS and resuspended in RIPA lysis buffer (Sigma-Aldrich) containing Halt protease and phosphatase inhibitors (ThermoFisher) for 30 min on ice. Cell debris was removed by centrifugation at 12,000 × g for 15 min at 4°C. Total protein lysate was quantified by using the Bio-Rad DC protein assay kit (Bio-Rad Cat. No. 500-0116). About 30 µg of protein was boiled for 5 min at 95°C in 4X LDS sample buffer containing β-ME, then separated on Bolt 4–12% Tris-Glycine gels (ThermoFisher), and transferred onto 0.2 µm nitrocellulose membranes (ThermoFisher). Membranes were blocked for 30 min at room temperature in Pierce StartingBlock Blocking buffer (ThermoFisher) followed by incubation of membranes overnight at 4°C with primary antibodies against STING (D2P2F/D1V5L), Phospho-STING (Ser365) (D8F4W), IRF-3 (D83B9), Phospho-IRF-3 (Ser396) (D6O1M), TBK1 (D1B4), Phospho-TBK1 (Ser173) (D52C2), cGAS (D3O8O), MLKL (D6W1K) or Phospho-MLKL (Ser345) (D6E3G) (Cell Signaling Technology) at 1:1000 dilution in 5% BSA–0.02% Tween-20–1XTBS (TBST). β-ACTIN (13E5) or GAPDH (D16H11) served as the protein loading control. Membranes were then washed three times for 5 min in TBST and incubated for 2 hours at room temperature with a 1:2000 dilution of relevant secondary antibodies, washed an additional three times, then incubated in Advansta WesternBright ECL-HRP Substrate. Bands were visualized using ChemiDoc (Bio-Rad). After visualization, membranes were stripped for 5 min in Restore Plus Western Blot Stripping Buffer (ThermoFisher), washed three times in TBST and reblocked before incubating in the next antibody.
Chemokine and cytokine arrays
1×106 RMS STINGwt and RMS STING-/- tumors were injected subcutaneously in the flank of C57BL/6 mice on day 0. Tumors were cryoablated on day 10 when tumor size reached 100 mm3. Five days later (day 15), tumors were harvested, weighed, minced, then resuspended in 300 µL PBS, vortexed and pelleted at 500 g for 5 min at 4°C. Then, 100 µL of supernatant from each mouse was pooled by groups. 200 µL of pooled supernatant from each group was then used to perform the Proteome Profiler Mouse XL Cytokine Array (R&D Systems; Cat. No. ARY028) and 175 µL of pooled supernatant from each group was used to perform the Proteome Profiler Mouse Chemokine Array Kit (R&D Systems; Cat. No. ARY020), each according to manufacturer’s instructions. Densitometry was then performed on the dot blot arrays using ImageJ with the Analyze/Measure command. Density was calculated by subtracting the background and then normalized based on average weight (in grams) of tumor per group.
Cell proliferation and cell death analysis
To assess tumor cell proliferation and confluency in vitro, tumor cells were treated with vehicle, 50 µg/mL, 100 µg/mL, 200 µg/mL or 400 µg/mL DMXAA or cryoablation for one cycle of 10, 20, 30 or 60 s in quadruplicate. Live cell phase contrast imaging was then performed every 4 hours up to 96 hours to assess confluency by the Sartorius IncuCyte Live-Cell Analysis System. For cell death analysis, tumor cells were treated in vitro with 100 µg/mL DMXAA or cryoablation for one cycle of 10, 20, 30 or 60 sec and incubated for 24, 48 and 72 hours in triplicate. The frequency of live cells (Annexin-V-7-AAD-), early apoptotic cells (Annexin-V+7-AAD-) and late apoptotic cells (Annexin-V+7-AAD+) was then analyzed by flow cytometry.
Immunohistochemistry
For Caspase-3 staining and terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) assay, formalin-fixed paraffin-embedded (FFPE) preserved tumor blocks obtained 3 days following cryoablation were sectioned, and a minimum of 4 tissue sections per condition were sent to the Tissue Histology Core at University Hospitals for antibody staining under standard protocols. The stained sections were then scored for Caspase-3+ tumor cells within the live tumor core per high power field (Casp-3) or overall tumor areas (TUNEL). The RMS tissue microarray (TMA) was purchased from US Biomax (Cat. No. SO2082A). Tumor and lung tissue sections were deparaffinized and hydrated through xylenes, graded alcohol series, and finally washed for 5 min in tap water. Heat-induced epitope retrieval was performed using citrate buffer pH 6.0 (Thermo Scientific AP9003-500). Endogenous peroxidase activity was blocked by incubating the sections in 3% hydrogen peroxide in water. Sections were blocked with 2.5% normal horse serum, then incubated with polyclonal anti-TMEM173 antibody (Millipore Sigma HPA038534). ImmPRESS Excel Amplified Polymer Staining Kit, Anti-Rabbit IgG and Peroxidase (Vector Laboratories MP-7601) were used for amplification and visualization of signals according to manufacturer’s protocol. The intensity of STING staining among RMS tumor cells and mononuclear cells within each tissue core was visually determined by an independent sarcoma pathologist (STS) using a scale of 0–3.
Confocal imaging
Mouse lung tissues were filled with Optimal Cutting Temperature (OCT) and fixed with periodate-lysine-paraformaldehyde (PLP) medium (0.625 mL PFA 16%, 3.75 mL L-lysin 0.2 M pH 7.4, 5.65 mL P-buffer 0.1 M pH 7.4, 0.0212 g NaIO4) overnight at 4o C. They were then washed in P-buffer (81 mL Na2HPO4 0.2 M and 19 mL Na H2PO4 0.2M with 100 mL of water) and incubated overnight in sucrose solution (30% in P-buffer) at 4°C. The lungs were then embedded in OCT, snap frozen, and store at −20°C. Tissues were sectioned and treated with ProLong Diamond Antifade Mountant with DAPI from (Thermo Fisher Scientific). Imaging of slides was performed using an inverted Leica SP5 confocal microscope system using an Argon Laser (488 nm) for the mNeonGreen and HeNe (633 nm) for the E2-crimson. Images were analyzed using Imaris software from Bitplane.
Statistical analysis
Analyses were performed using GraphPad Prism software V.8. All data are shown as mean ± SEM. Statistical analyses include two-tailed Student’s t-test with unequal variance, one-way analysis of variance (ANOVA) or two-way ANOVA with Bonferroni’s multiple comparisons test. We defined statistical significance as follows: *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001; NS indicates no significant difference. The analysis and p value reported are presented in each figure panel’s corresponding legend.
Results
Cryoablation promotes RMS regression through adaptive immunity
To investigate whether cryoablation can effectively control established RMS 76-9 tumor burden in vivo, we subcutaneously inoculated RMS 76-9 cells50 51 into naïve C57BL/6 mice. Ten days later, when the tumor volume reached approximately 100 mm3, three cycles of percutaneous cryoablation were applied directly on the tumor in a single treatment session. The tumor volume was then assessed two to three times a week. Cryoablation effectively reduced the size of the established tumor mass while suppressing the rate of local recurrence over the following 2 weeks (figure 1A). Next, we explored whether the robust local tumor regression following in vivo percutaneous cryoablation was the direct result of tumor cell death caused by freezing, or possibly contributed by the host immune activation in response to cryoablation-induced cell death of RMS 76-9. To examine the latter possibility, we repeated in vivo tumor cryoablation in nude mice. Interestingly, the effect of cryoablation was greatly reduced in nude mice which all succumb to disease days after the last tumor measurement on day 21, indicating that the adaptive T-cell response was required for the observed therapeutic efficacy of cryoablation in vivo (figure 1B). To further validate this observation, we depleted either CD4+ T cells or CD8+ T cells in tumor-bearing C57BL/6 mice during the peri-cryoablation period. Indeed, depletion of CD4+ T cells resulted in partial abrogation of the observed cryoablation-induced tumor regression (figure 1C, online supplemental figure S1), whereas removal of CD8+ T cells resulted in tumor growth that was not significantly different from untreated tumors (figure 1D, online supplemental figure S1). Taken together, these data underscore the importance of host adaptive T-cell immunity in mediating a seemingly curative antitumor effect using a cryoablation procedure that proved to be only sublethal against the tumor without a functional adaptive T-cell immunity in this model.
Supplemental material
Local therapeutic effect of rhabdomyosarcoma (RMS) 76-9 cryoablation is dependent on CD8+ T cells and partially dependent on CD4+ T cell. (A) C57BL/6 mice or (B) nude mice were inoculated subcutaneously with 1×106 RMS 76-9 cells, on days 9–10 (indicated by arrow) primary tumors underwent cryoablation (cryo) or were left untreated (non-cryo). Tumor growth was measured two to three times a week. (C, D) C57BL/6 mice were inoculated with 1×106 RMS 76-9 cells and treated with cryoablation on day 10. About 100 µg of anti-CD4 (C) or anti-CD8 (D) mAb was then administrated on days 10, 13 and 16. Data are presented as mean ± SEM (n=5 mice per group). Statistical analysis was performed by two-way analysis of variance with Bonferroni’s multiple comparisons test (B–E) or by two-tailed unpaired t-test (A, F). *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.
Tumor cryoablation induces DC maturation and infiltration of functional T cells into TME
APCs induce antitumor immunity by engulfing and presenting tumor antigens to initiate adaptive T-cell responses.13 14 Therefore, we sought to determine if tumor cryoablation increased APC stimulation and maturation. To test the impact of tumor cryoablation on DC maturation in vitro, we cryoablated in vitro cultured RMS 76-9 cells, co-cultured them with bone marrow-derived dendritic cells (BMDCs), and examined the expression of various stimulatory markers on APCs, including CD80, CD86, CD40 and major histocompatibility complex II (MHC II) using flow cytometry. Cell lysates from cryoablated RMS 76-9 cells were able to stimulate the maturation of CD11c+ DCs, as evidenced by a significant increase in the frequency of co-cultured BMDCs expressing CD86, CD40 and MHC II, as compared with BMDCs co-cultured with non-cryoablated RMS 76-9 cells (figure 2A). Other forms of RMS 76-9 cell death, such as those induced by radiation and chemotherapy, resulted in similar levels of MHC II and CD86 expression on co-cultured BMDCs as compared with non-cryoablated RMS76-9 cells (online supplemental figure S2). Similar effects were observed in freshly isolated DCs from the TME of tumor-bearing C57BL/6 mice 3 days after percutaneous tumor cryoablation in vivo. A higher percent of MHC II and CD80 expressing DCs were found in cryoablated TME as compared with those in the non-cryoablated group (figure 2B). Similarly, cryotherapy of RMS 76-9 increases Il-12β transcription by co-cultured BMDCs in vitro (figure 2C) as well as in DCs isolated from cryoablated TME in vivo (figure 2D). In addition to inducing immune-stimulating BMDCs, cryoablated RMS also decreased bone marrow-derived macrophage (BMDM) expression of CD206+ under similar co-culture conditions (figure 2E). Our results suggest that tumor cryoablation induces APC maturation toward immune stimulating and antitumor phenotypes. We next investigated changes in the immune cell infiltration within the TME following cryoablation. Seven days after cryoablation, residual tumor mass was harvested and analyzed for changes in tumor-infiltrating lymphocytes (TILs) using multiparametric flow cytometry (online supplemental figure S3A). Cryoablated tumor TME harbored a significant increase in the frequency of CD45+ cells (figure 2F). These results suggest that cryoablation may activate antitumor immune responses by supporting DC maturation and recruitment of hematopoietic cells to the TME.
In vivo cryoablation induces dendritic cell (DC) maturation and infiltration of functional lymphocytes into rhabdomyosarcoma (RMS) 76-9 tumor microenvironment (TME). (A) RMS 76-9 cells were untreated (non-cryo) or treated with cryoablation and incubated at 37°C for 6 hours in vitro. The cells were then co-cultured with bone marrow-derived dendritic cells (BMDCs) for 12 hours in vitro. CD11c+ DCs were then assessed for the maturation markers I-Ab, CD80, CD86 and CD40 by flow cytometry. (B) C57BL/6 mice were inoculated with 1×106 RMS 76-9 cells and treated with cryoablation on day 10. Three days later, tumor tissue was analyzed by flow cytometry for the frequency of live CD11c+I-Ab+ and CD11c+CD80+ cells in the TME (n=4). (C, D) DCs were isolated by using positive CD11c selection following co-culture of cryoablated RMS 76-9 lysate and BMDC in vitro (with lipopolysaccharide (LPS) treatment as a positive control) (C), or from bulk tumors 3 days post cryoablation in vivo (D) and evaluated for Il-12β expression by qPCR (n=3). (E) Bone-marrow derived macrophages (BMDM) co-cultured with in vitro cryoablated or non-ablated RMS 76-9 were analyzed for the frequency of F4/80 and CD206 by flow cytometry. BMDM grown in the presence of IL-4 was used as a control. (F) Flow cytometric analysis of bulk RMS 76-9 tumors taken 7 days after cryoablation (17 days post inoculation) was performed to detect live, CD45+ cells (n=5). Data are presented as mean ± SEM. Statistical analysis was performed by two-tailed unpaired t-test (A–F). *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.
Host STING contributes to the in vivo efficacy of cryoablation
Based on the observed antitumor immune responses following cryoablation, we hypothesize that dsDNA released from cryoablated RMS 76-9 tumors triggers the STING pathway in APCs and other cellular elements in the tumor-bearing host to elicit immune cell recruitment, which effectively suppresses local RMS 76-9 recurrence. Previous studies have shown that dsDNA derived from tumor cells as a result of cytotoxic agents or stress can trigger STING pathway activation in nearby host APCs, resulting in the release of IFN-β, chemokines CXCL9/10, and enhanced cross-presentation of tumor antigens to CD8+ T cells.17 35 36 54–58 To test whether this paradigm applies to tumor cryoablation, we assessed and observed a significant increase in the amount of IFN-β production in the TME 3 days after cryoablation as compared with non-ablated TME (figure 3A). To examine whether cryotherapy of RMS tumor was able to activate STING signaling pathway in APCs, we measured IFN-β production by qPCR in DCs directly isolated from tumor TME 3 days after in vivo tumor cryoablation of RMS. Tumors treated with cryoablation induced a significant increase in Ifn-β mRNA transcript (figure 3B) and protein secretion (figure 3C) in tumor-infiltrating DCs. Since multiple signaling pathways can trigger type I IFN production, including Toll-like receptor (TLR) signaling in APCs,59 next we examined the specific contribution of STING pathway in mediating type I IFN production in cryoablation. We co-cultured in vitro cryo-treated RMS 76-9 cells with BMDCs derived from STING-/- mice, and observed a complete absence of Ifn-β transcription as compared with BMDCs from wild-type mice (figure 3D), suggesting that the STING signaling pathway is essential for tumor cryoablation-induced type I IFN production in DCs. As prior reports have described an essential role for host STING pathway in cancer therapy,17 35 36 54 we next investigated how systemic host STING pathway contributes to the cryoablation-mediated tumor control in vivo. We inoculated RMS 76-9 cells into STING-/- and C57BL/6 mice. Surprisingly, while RMS 76-9 grew aggressively in both groups without cryoablation therapy, cryoablation of established RMS tumor in STING-/- mice resulted in a significant partial reduction in tumor size over the subsequent 2 weeks, although there was not as robust of a response as seen in wild-type mice (figure 3E). Interestingly, IFN-β production in the TME of tumor-bearing STING-/- mice were similarly enhanced 3 days following cryoablation as in wild-type mice (figure 3A, online supplemental figure S4F). These results suggest that, while host STING signaling activation and associated type I IFN production can provide partial local tumor control following cryoablation, they cannot fully account for the complete in vivo therapeutic effects observed in wild-type mice.
Host STING partially contributes to local efficacy of cryoablation in vivo. (A) C57BL/6 mice were inoculated with 1×106 RMS 76-9 cells and tumors were treated with cryoablation on day 10. Three days later, interferon-β (IFN-β) expression in bulk tumor tissues was assessed by ELISA (n=8–7). (B, C) Dendritic cells (DCs) were enriched from bulk tumor tissues using a positive CD11c selection kit and assessed for Ifn-β mRNA by quantitative PCR (qPCR) (B) and IFN-β protein secretion by ELISA (C). (D) RMS 76-9 cells were treated with cryoablation and incubated for 6 hours in vitro, followed by co-culturing for an additional 6 hours with bone marrow-derived dendritic cells (BMDCs) from C57BL/6 (wild type (WT); white bars) or STING-/- mice (STING-/-; black bars). Untreated BMDCs served as control. Ifn-β mRNA expression was then measured by qPCR. (E) C57BL/6 mice or STING-/- mice inoculated with RMS 76-9 cells were monitored for tumor growth following treatment with cryoablation on day 10 (arrow; n=5). Data are presented as mean ± SEM and statistical analysis was performed by two-tailed unpaired t-test (A–E) or two-way analysis of variance with Bonferroni’s multiple comparisons test (F). *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.
Tumor-intrinsic STING mediates antitumor immune responses following cryoablation
Thus far, our data indicate that the host STING pathway plays a partial role in regulating RMS 76-9 tumor growth, as the antitumor immune response of cryoablation was impaired but not completely abrogated in STING-/- mice (figure 3E). This exciting observation raises the possibility that tumor cell-intrinsic STING may contribute to cryoablation-induced therapeutic efficacy. Previous studies have suggested that the tumor cell-intrinsic STING pathway may be essential in triggering antitumor immunity in other contexts.47–49 We set out to test whether tumor cell-intrinsic STING also impacts cryoablation efficacy. First, we validated that there was an intact STING signaling pathway in RMS 76-9 and then ablated Tmem173 expression via CRISPR-Cas9 genome editing (RMS 76-9STING-/-), which was confirmed by both western blot (online supplemental figure S4A) and qPCR (online supplemental figure S4B). We also tested whether the STING pathway was functional in RMS 76-9STINGwt and RMS 76-9STING-/- cells by treating with STING agonist, DMXAA. Both the upregulation of IFN-β (online supplemental figure S4C) and the phosphorylation of IRF3 (online supplemental figure S4D) were decreased in DMXAA-treated RMS 76-9STING-/- as compared with wild-type cells. Next, we examined whether the STING pathway can be activated in live RMS 76-9STINGwt or RMS 76-9STING-/- cells upon co-culture with cryo-treated RMS 76-9STINGwt lysates in vitro. On exposure to dead tumor lysate, Ifn-β transcription was upregulated only in the live RMS 76-9STINGwt, while live RMS 76-9STING-/- cells remained unaffected (online supplemental figure S4E). To determine whether tumor cell-intrinsic STING signaling influences the effectiveness of cryoablation, we inoculated RMS 76-9STINGwt or RMS 76-9STING-/- cells into either C57BL/6 or STING-/- mice. Although RMS 76-9STING-/- exhibit faster growth kinetics in vitro as compared with RMS 76-9STINGwt (online supplemental figure S8A), both cell lines exhibit similar tumor growth kinetics in C57BL/6 mice in the absence of cryoablation (figure 4A). As expected, cryoablation of RMS 76-9STINGwt tumor in wild-type mice not only induced sustained local tumor control (figure 4A), but also produced a robust systemic immunity to suppress distant lung metastasis (figure 4B,C). To our surprise, however, cryoablation of RMS 76-9STING-/- tumors in C57BL/6 mice was rendered completely ineffective, despite the presence of functionally intact STING signaling in the host (figure 4A). Concurrently, we also observed a significant increase in RMS lung metastases in the same group (figure 4B,C), suggesting ineffective systemic immunity. To test the possibility that this observation was not due to selection bias of the specific RMS 76-9STING-/- tumor clone, we tested two additional RMS 76-9STING-/- tumor clones, Clones 7 and 8, either alone or in combination, and observed the same phenomenon of a complete loss of antitumor efficacy following cryoablation as seen in figure 4A, online supplemental figure S5).
Tumor cell-intrinsic STING mediates local antitumor immune response following cryoablation in the presence of host STING. (A) Tumor growth was monitored in C57BL/6 mice inoculated with either 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- cells and cryoablated (arrow) on day 10. (B, C) Lung metastases were assessed on day 35 by serial immunohistochemical (IHC) sections. Scale bar: 5 mm (n=5). (D–P) Flow cytometric analysis was performed on bulk tumors 17 days post inoculation and 7 days following cryoablation. (D) t-distributed stochastic neighbor embedding (t-SNE) analysis of lymphocyte profiling on live CD45+ cells was performed. t-SNE heatmap plots show the mean fluorescent intensity (MFI) expression level and the distribution of each indicated marker. Low to high levels of protein expression are depicted in the gradient from blue (low) to red (high). (E) t-SNE density plots of live CD45+ cells in each treatment group were shown. (F–P) Conventional flow cytometric analysis of CD45+, CD3+, CD3+CD4+ and CD3+CD8+ lymphocytes that express intracellular IFNγ+ (F–I) or GZMB (J–L). CD4+ lymphocytes expressing Foxp3 (M), NK cells (N), myeloid cells expressing CD11c (O) or CD11b (P) were also analyzed. Data are presented as two-way analysis of variance with Bonferroni’s multiple comparisons test (A, C), and two-tailed unpaired t-test (F–P). *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001. WT, wild-type.
To further understand how tumor cell-intrinsic STING signaling changes immune cell components in the post-cryoablation TME, we profiled the immune landscape of the TME using t-SNE on multiparameter flow cytometric analyses of all freshly isolated, live CD45+ lymphocyte subsets 7 days after cryoblation. The heatmap t-SNE plots show the distribution of mean fluorescent intensity (MFI) for each specific marker per cluster (figure 4D). There were clear differences in the TME expression profiles for each treatment group as shown by the number of events per cluster in each of the density plots (figure 4E). Therefore, TILs were further analyzed by conventional flow cytometric analysis (online supplemental figure S3B). Interestingly, the basal level of various lymphocyte compartments was significantly reduced or trending toward a reduction in C57BL/6 mice bearing RMS 76-9STING-/- tumors as compared with RMS 76-9STINGwt. Corroborating with gross local tumor burden (figure 4A), we observed a significant increase in the total CD4+ T cells expressing IFN-γ (figure 4H) and granzyme B (GZMB; figure 4J),60–65 NK cells (figure 4N), and the amount of GZMB expression in CD8+ T cells (figure 4L) in the post-cryoablated RMS 76-9STINGwt TME as compared with RMS 76-9STING-/- TME in wild-type mice, as was Foxp3+ CD4+ cells (figure 4M). There was also a trend showing higher levels of total CD45+, CD3+, or CD8+ T cells expressing IFN-γ (figure 4F, G and I) and frequency of CD8+ T cells expressing GZMB in cryoablated RMS 76-9STINGwt compared with RMS 76-9STING-/- (figure 4K). We did not observe any significant changes in the myeloid cell populations within the cryoablated TME of either tumors (figure 4O,P).
To further isolate the direct impact of tumor cell-intrinsic STING signaling on host antitumor immunity, we inoculated RMS 76-9STINGwt and RMS 76-9STING-/- tumors into STING-/- mice. In this model, the only source of intact STING signaling pathway comes solely from RMS 76-9STINGwt, while mice bearing RMS 76-9STING-/- tumors completely lack STING in both host and tumor. Validating the importance of tumor cell-derived STING signaling in mediating cryoablation-induced antitumor effect in wild-type mice (figure 4A), a significant tumor growth inhibition was again observed when cryoablation was carried out in STING-/- mice bearing RMS 76-9STINGwt tumor (figure 5A). We also observed a significant inhibition in RMS 76-9 lung metastases in the same experimental group (figure 5B,C). To further understand how tumor cell-intrinsic STING changes immune cell components in the TME post cryoablation, TILs were analyzed 7 days after cryoablation. Unique clusters identified by t-SNE were again generated similar to figure 4D (figure 5D). Conventional flow cytometry analysis confirmed a significant increase in IFN-γ-expressing CD45+/CD3+/CD8+ TILs (figure 5E, F and H), and CD3+ TILs post cryoablation of RMS 76-9STINGwt tumor as compared with RMS 76-9STING-/- tumor in the setting of STING-/- mice, thereby providing a cellular mechanism of partial cryoablation efficacy afforded by cryoablation of RMS 76-9STINGwt (figure 5A). In contrast to wild-type mice, the significant changes in frequency of CD4+ T cells expressing IFN-γ (figure 5G) or GZMB (figure 5I), and the frequency and intensity of GZMB-expressing CD8+ T cells (figure 5J,K) were lost. The differences among Foxp3+ CD4+ cells, natural killer (NK) cells, CD11c+ myeloid cells and CD11b+ myeloid cells were also lost in the context of STING-/- mice.
Tumor cell-intrinsic STING mediates antitumor immune response following cryoablation in the absence of host STING. (A) STING-/- mice inoculated with either 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- tumor cells were treated with cryoablation (arrow) on day 10 and monitored for tumor growth. (B, C) Lung metastases were assessed by serial immunohistochemical (IHC) sections on day 35. Scale bar: 5 mm (n=5). (D) t-distributed stochastic neighbor embedding (t-SNE) density plots of lymphocyte profiling on live CD45+ cells from bulk tumor 17 days post inoculation were shown for each treatment group. (E–O) Conventional flow cytometric analysis of CD45+, CD3+, CD3+CD4+ and CD3+CD8+ lymphocytes that express intracellular IFN-γ+(E–H) or GZMB (I–K). CD4+ lymphocytes expressing Foxp3 (L), NK cells (M), myeloid cells expressing CD11c (N) or CD11b (O) were also analyzed. Data are presented as mean ± SEM and statistical analysis was performed by two-way analysis of variance with Bonferroni’s multiple comparisons test (A, C), and two-tailed unpaired t-test (E–O). *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001. RMS, rhabdomyosarcoma; WT, wild-type.
As different tumor models have shown variability in their responses to STING signaling,66 we examined the applicability of our finding regarding tumor-intrinsic STING signaling to cryoablation efficacy by testing two additional syngeneic tumor cell lines, M3-9-M, another embryonal RMS derived from transgenic mice expressing the hepatocyte growth factor and heterozygous for mutated P53,50 67 and KPC1242, a spontaneously arising pancreatic cancer cell line isolated from KRasLSL.G12D/+-p53R172H/+-Pdx-Cre (KPC) transgenic mice on a homogenous C57BL/6 background.52 In both tumor cells we ablated STING expression via CRISPR-Cas9 genome editing (M3-9-MSTING-/- and KPC1242STING-/-). We confirmed STING protein expression by western blot (online supplemental figure S6). We also tested whether the STING pathway was functional in these cells by treating with DMXAA (data not shown).56 Similar to RMS 76-9, cryoablation of M3-9-MSTINGwt and KPC1242STINGwt resulted in significantly reduced tumor burden in C57BL/6 mice mice, while the same intervention was ineffectual against M3-9-MSTING-/- and KPC1242STING-/- tumors, even in C57BL/6 mice with intact host STING signaling pathways (figure 6).
Tumor cell-intrinsic STING mediates local antitumor immune response following cryoablation of different tumor types. C57BL/6 (A) or STING-/- (B) mice inoculated with either 1×106 M3-9-MSTINGwt or M3-9-MSTING-/- tumor cells were treated with cryoablation on day 12 when the tumors reached 100 mm3 and then monitored for tumor growth. (C) C57BL/6 mice inoculated with either 1×106 KPC1242STINGwt or KPC1242STING-/- were treated with cryoablation on day 15 when the tumors reached 100 mm3 and then monitored for tumor growth. WT, wild-type.
Our tumor models showed that cryoablation efficacy was dependent on tumor-intrinsic STING and associated with increases in functional CD8+IFN-γ+ T cells, an effector population shown to be a target of downstream STING signaling involving the IFN-β-CXCL9/10 axis.54–58 Therefore, we interrogated IFN-β, CXCL9/10 and other chemokine and cytokine production within various TME following cryoablation (online supplemental figure S7). Total IFN-β increased in both RMS 76-9STINGwt (figure 3F) and M3-9-MSTINGwt TME (figure 7A) as compared with the STING-/- tumor TME following cryoablation. Cryoablation of STINGwt tumor dramatically increases a variety of chemokines and cytokines, including the canonical chemokines, CXCL9 and CXCL10, as well as a variety of others (figure 7B–D). Most significantly, chemokine and cytokine protein production was not significantly changed before and after cryoablation in STING-/- tumor TME, even though the host stroma contains a functional STING signaling pathway (figure 7B–D).
Preferential inflammatory chemokine and cytokine production in STINGwt RMS tumor microenvironment (TME) following cryoablation. C57BL/6 mice inoculated with 1×106 M3-9-MSTINGwt or M3-9-MSTING-/- were non-treated (non-cryo) or treated with cryoablation on day 10 when tumors reached 100 mm3. Five days later, tumors were harvested, weighed, minced, and suspended in PBS. (A) The supernatants were analyzed for IFN-β protein concentration by ELISA and normalized to tumor weight. For the chemokine and cytokine arrays, supernatants from each mouse were pooled by group, and the pooled supernatant for each group was then used to analyze chemokines (B) and cytokines (C, D). Densitometry was performed on the dot blot arrays (online supplemental figure S7) and normalized to average tumor weight, and results are displayed based on a relative intensity scale.
A potential alternative explanation for the observed lack of cryoablation efficacy involving STING-/- tumors irrespective of host STING status was the reported protective role of STING deficiency in tumor cell death.68 69 Indeed, at baseline we saw a growth advantage of RMS 76-9STING-/- tumor as compared with RMS 76-9STINGwt tumor (online supplemental figure S8A) in vitro. Treatment with DMXAA impaired RMS 76-9STINGwt growth in a dose-dependent manner, while 76-9STING-/- was not affected until a high DMXAA dose range (online supplemental figure S8B). However, both tumors showed similar growth suppression when subjected to various cryoablation treatment regimens in vitro (online supplemental figure S8C). In fact, analysis of different phases of apoptotic cell death over a 3-day time span revealed that RMS 76-9STINGwt have a higher percentage of live tumor cells, while RMS 76-9STING-/- exhibited more early and late apoptotic markers across time and cryoablation regimens (online supplemental figure S8D-F). Furthermore, we did not observe any significant differences in necroptosis resulting from cryoablation between the two cell lines as assessed by phosphorylation of MLKL (online supplemental figure S9A). Lastly, we assessed the level of Caspase-3+ and TUNEL+ tumor cells in vivo within the non-necrotic tumor zone 3 days following cryoablation, and found no significant differences in the level of cell death following cryoablation (online supplemental figure S9D,E). In fact, our Caspase-3 staining data suggest that there appears to be a slightly higher basal level of Caspase-3+ tumor cells among RMS 76-9STING-/- tumors (online supplemental figure S9E), although we failed to observe any significant differences in cleaved Caspase-3 and Gasdermin D activation following cryoablation in vitro by western blot analysis (data not shown). Our data suggest that intrinsic differences of cell proliferation or various forms of cell death between STINGwt and STING-/- tumors cannot explain our observed therapeutic efficacy of cryoablation.
To further elucidate whether tumor cell-intrinsic STING signaling with cryoablation affects systemic immunity to influence lung metastatic incidence, or if RMS 76-9STING-/- are more metastatic prone,70 less sensitive to immune attack, or both (figures 4B, C, 5B,C), we generated fluorescently labeled RMS 76-9 STINGwt mNeonGreen and RMS 76-9 STING-/- E2-Crimson cells, respectively. Equal numbers of both cell lines were mixed, inoculated subcutaneously into C57BL/6 mice, and applied percutaneous cryoablation therapy to the primary tumor 10 days after inoculation (online supplemental figure S10A-G). On day 35, lung metastasis was analyzed by both flow cytometry (online supplemental figure S10B, S10C) and tissue fluorescent microscopy (online supplemental figure S10G). A significant decrease in the percentage of RMS 76–9 STINGwt mNeonGreen cell subpopulation was observed in the lungs of mice treated with cryoablation as compared with sham-ablated control mouse lung (online supplemental figure S10D), while cryoablation had no effect on the proportion of RMS 76-9 STING-/- E2-Crimson tumor cell metastases in the lung tissue compared with non-cryoablated controls (online supplemental figure S10E). Comparing lung metastatic lesions in the lungs following cryoablation, the percentage of RMS 76-9 STING-/- E2-Crimson cells in the lung was proportionally higher than that of RMS 76-9 STINGwt mNeonGreen cells (online supplemental figure S10F), suggesting a correlation between the lack of tumor cell-intrinsic STING and tumor metastatic tendency or resistance to antitumor immunity.
Tumor-intrinsic STING pathway promotes the abscopal effect post-local cryoablation
To further pinpoint the impact of tumor cell-intrinsic STING pathway on systemic antitumor immune activation rather than tumor cell-intrinsic metastatic or immunogenicity tendency, we directly tested the effect of cryoablation of primary tumor on the growth of a concurrently established secondary tumor, a phenomenon called the ‘abscopal effect’.8 9 To do this, we implanted two separate tumors into the same mouse, with one tumor inoculated in the left flank (primary tumor) and the other inoculated simultaneously in the contralateral flank (secondary tumor). Either RMS 76-9STINGwt or RMS 76-9STING-/- tumor was inoculated subcutaneously in the left flank of C57BL/6 mice as the primary tumor, and only RMS 76-9STINGwt was inoculated concurrently in the contralateral flank as the secondary tumor in all recipient mice (figure 8A). This approach eliminated the discrepancy in the inherent immunogenicity of the abscopal tumor as seen in the previous lung metastasis models between RMS 76-9STINGwt or RMS 76-9STING-/- (figures 4 and 5). Primary tumors were treated with percutaneous cryoablation on day 10 after tumor inoculation, and both the primary and the secondary tumor growths were then monitored. The tumor reduction efficacy of cryoablation in the primary tumor was completely lost in the context of RMS 76-9STING-/- tumor (figure 8B, black), while RMS 76-9STINGwt primary tumor exhibited a robust response (figure 8B, yellow), consistent with our previous observations (figures 3E and 4A). Evaluating the abscopal effect, we observed that cryoablation of primary RMS 76-9STINGwt tumor led to a significant reduction of the advanced, untreated secondary tumor (figure 8C, yellow). However, this abscopal effect on secondary RMS 76-9STINGwt was lost on cryoablation of RMS 76-9STING-/- primary tumor (figure 8C, black). To further investigate the sole impact of tumor-intrinsic STING signaling pathway on the observed abscopal effect, we used the same concomitant dual tumor injection model in STING-/- mice. Again, RMS 76-9STINGwt or RMS 76-9STING-/- RMS tumor was inoculated subcutaneously in the left flank of STING-/- mice as the primary tumor, and only RMS 76-9STINGwt was inoculated concurrently in the contralateral flank as the secondary tumor in all recipient STING-/- mice (figure 8D). Primary tumors were treated with percutaneous cryoablation on day 10 after tumor inoculation, and both the primary and the secondary tumor growths were then monitored. The tumor reduction efficacy of cryoablation in the primary tumor was completely lost in the context of RMS 76-9STING-/- tumor (figure 8E, black). However, RMS 76-9STINGwt primary tumor again exhibited a robust response to cryoablation despite the absence of host STING signaling (figure 8E, yellow), consistent with our previous observations (figure 5A). Importantly, we observed that cryoablation of primary RMS 76-9STINGwt tumor led to a significant reduction of the advanced, untreated secondary tumor (figure 8F, yellow) in the complete absence of host STING signaling. However, this abscopal effect on secondary RMS 76-9STINGwt was completely abrogated on cryoablation of RMS 76-9STING-/- primary tumor (figure 8F, black). Taken together, tumor cell-intrinsic STING signaling critically impacts the abscopal effect of the non-cryoablated advanced distant tumors as induced by cryoablation of the primary tumor.
Rhabdomyosarcoma (RMS) 76-9-intrinsic STING dictates systemic immunity to control distant tumor burden following local cryoablation of the primary tumor. (A–C) C57BL/6 mice were inoculated with 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- tumor in the left flank (primary tumor) and 5×105 RMS 76-9STINGwt was inoculated concurrently in the contralateral flank (secondary tumor) (A). Three cycles of cryoablation (arrows) were applied to the primary tumor when it reached ~100 mm3, 10 days later. The primary (B) and secondary tumors (C) were then monitored until day 30. Inset to the right of (C) shows individual tumor measurements of the abscopal tumors in various groups. (D–F) STING-/- mice were inoculated with 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- tumor in the left flank (primary tumor) and 5×105 RMS 76-9STINGwt was inoculated concurrently in the contralateral flank (secondary tumor) (D). Three cycles of cryoablation were applied to the primary tumor when it reached ~100 mm3, 10 days later. The primary (E) and secondary tumors (F) were then monitored until day 30. Inset to the right of (F) shows individual tumor measurements of the abscopal tumors in various groups. Data are presented as mean ± SEM (n=5 mice per group). Statistical analysis was performed by two-way analysis of variance with Bonferroni’s multiple comparisons test (A–F). *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001. WT, wild-type.
Tumor-intrinsic STING contributes to an in vivo therapeutic response to STING agonism
Thus far, our data support a critical contribution of tumor cell-intrinsic STING pathway in mediating immune-mediated RMS 76-9 tumor rejection in vivo. To address whether this is solely a cryoablation-specific phenomenon, or if the presence of an intact tumor cell-intrinsic STING signaling pathway contributes to effective immune-mediated tumor rejection in general, we administered the STING agonist DMXAA intratumorally into wild-type or STING-/- mice harboring established RMS 76-9STINGwt or RMS 76-9STING-/- tumors 10 days following tumor inoculation (figure 9). Interestingly, while DMXAA administration effectively reduced RMS 76-9STINGwt growth in C57BL/6 mice (figure 9A, yellow), the absence of STING in either the tumor cells (figure 9A, black) or the host (figure 9B, yellow) alone partially suppressed tumor growth, while a complete absence of STING in both tumor cells and the host completely abolished in vivo antitumor efficacy of STING agonism (figure 9B, black). These data suggest that the presence of an intact tumor cell-derived STING signaling is critical for in vivo tumor rejection not only in the setting of cryoablation (figures 4 and 5) but also is applicable in the setting of therapies involving direct STING agonist administration.
Tumor cell-intrinsic STING mediates antitumor immune response to intratumoral STING agonist therapy in vivo. (A) C57BL/6 mice inoculated with either 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- tumor cells. DMXAA (250 µg) was injected intratumorally on day 10 (arrow) and tumor growth was monitored. (B) STING-/- mice inoculated with either 1×106 RMS 76-9STINGwt or RMS 76-9STING-/- tumor cells. DMXAA (250 µg) was injected intratumorally on day 10 (arrow) and tumor growth was monitored. Data are presented as mean ± SEM (n=4–5 mice per group). Statistical analysis was performed by two-way analysis of variance with Bonferroni’s multiple comparisons test. *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001. WT, wild-type.
STING pathway expression in clinical sarcoma
Given the importance of STING signaling in dictating therapeutic efficacy, we wish to determine the status of STING signaling in various human cancers as an explanation for the inconsistent outcomes of clinical bulk tumor cryoablation. To investigate the clinical correlation between STING pathway and clinical tumor samples, Tmem173 transcript expression was examined from RNA-seq data from 2142 samples obtained from the St. Jude Cloud PeCan database, spanning all brain, hematopoietic and solid tumor types available on the database71 72 (online supplemental figure S11A). We also examined other components of the STING pathway including Cgas, Tbk1 and Irf3 (online supplemental figure S11A). These patient-derived data showed a wide variation in the expression level of STING pathway components in pediatric and adolescent and young adult (AYA) cancers. We then analyzed the expression of STING pathway transcripts in pediatric and AYA RMS patients. Interestingly, Tmem173 and Tbk1 transcript expression in RMS were lower in metastatic and relapsed patients when compared with primary tumor at diagnosis (online supplemental figure S11B), supporting our observation in the RMS 76-9 model that tumor cell-intrinsic STING is critical for antitumor immunity. Furthermore, we performed immunohistochemistry (IHC) to detect STING expression on an RMS tissue microarray (TMA) containing alveolar, embryonal, pleomorphic and spindle cell and sclerosing RMS samples from 104 different patients (online supplemental figure S11G). Both the RMS tumor cells and mononuclear cells within each tissue core were then scored for STING intensity (online supplemental figure S11C). Although there was a wide range of STING expression across each RMS subtypes (online supplemental figure S11D), we found that mononuclear cells expressed higher levels of STING in the TME than the tumor (online supplemental figure S11E). However, this differential STING expression between tumor and mononuclear cells was lost in the higher grade and more advanced stage tumors (online supplemental figure S11F), reflecting a decrease in overall STING expression, which is supported by the Tmem173 transcript expression profile of the PeCan dataset.
Discussion
Cryoablation, along with other thermal or tissue injury ablation techniques, including laser ablation, radio-ablation, alcohol ablation and high-temperature infrared ablation, has been used clinically for non-invasive debulking of large, unresectable tumors. As an established alternative to surgical debulking of inoperable tumors, cryoablation has been demonstrated to be safe and effective in advanced tumors of the kidney, liver, adrenal, lung.3–6 It has also received Unite States Food & Drug Administration (U.S. FDA) breakthrough status in 2021 for the treatment of early-stage T1 invasive breast cancer or non-operable lesions.73 It is well tolerated with minimal side effects even in metastatic lung and bone lesions in advanced cancers.4 74 Cryoablation has also been applied in musculoskeletal cancers over the years, mostly in the palliative setting.75 Recently, an ongoing clinical study (NCT05302921) is currently assessing cryoablation in combination with dual checkpoint inhibition for relapsed/refractory pediatric solid tumors including RMS.
Tumors induced to undergo ICD can serve as ‘in situ tumor vaccines’ to activate DC and T cells, leading to the generation of an immunogenic hot TME and a robust systemic antitumor immune response.13 14 Cryoablation is a direct tumor-targeting approach that promises these unique benefits. This is evident from the more than 30 clinical trials that have been conducted.76–78 A distinct theoretical advantage of cryoablation over high-temperature or radio-ablation techniques is its ability to preserve intact tumor antigens and provide ICD-related DAMPs. Despite these theoretical benefits, clinical outcomes of tumor cryoablation—both obtaining local control and achieving an abscopal effects—are highly unpredictable with variable magnitudes of response and frequent failure of local recurrence. The molecular determinants driving these clinical outcomes are not well understood. Remarkably, our current data argue for a heretofore unappreciated but critical role of functional tumor cell-intrinsic STING signaling as a major determinant in cryoablation-induced antitumor immunity activation and clinical responses, a finding that is in sharp contrast to the current dogma which dictates that host-derived STING pathway is the converging critical responder.79–81 Instead, our data suggest that STING pathway activation in myeloid cells only partially contributed to primary tumor control and systemic abscopal effect, while a functional tumor STING signaling pathway serves as the dominant determinant of therapy response.
Using the RMS 76-9 sarcoma tumor model, we demonstrated that successful cryoablation therapies against advanced tumor growth requires the adaptive immune response, as cryoablation failed to control primary tumor growth in immune-deficient mice resulting from rapid local recurrence and metastasis following cryoablation. While CD4+ T cells partially contributed to this immune-mediated tumor regression following cryoablation, CD8+ T cells were absolutely essential for the in vivo antitumor efficacy. Cryoablation enhanced the infiltration of total CD45+ cells and functional CD8+ T cells82 and increased the expression of co-stimulatory molecules on DCs. These findings support the view that cryoablation boosted tumor antigen presentation by DCs and T cell activation and recruitment through an ICD-mediated in situ vaccine effect.
Previous studies have reported that tumor cells undergoing ICD actively export DNA fragments to initiate an innate immune response.83 84 Consequently, the presence of cytosolic dsDNA in APCs triggers host STING-mediated type I IFN activation and adaptive immunity.17 32 35 36 Here, we showed increased IFN-β in the TME following tumor cryoablation, and observed increased production of IFN-β by DCs after tumor cryoablation in vitro and in vivo. Since APCs can be activated to express type I IFN via STING-dependent and STING-independent pathways that lead to IFN-β production,50 we used BMDCs from STING-/- mice to demonstrate the STING dependency of IFN-β production by APCs in our model. Although these findings support the involvement of a myeloid cell-mediated mechanism that contributes to the therapeutic effects of cryoablation and reaffirmed prior studies showing the requirement of the myeloid STING pathway for the antitumor immunity effects associated with multiple cancer therapeutics,17 35 36 we consistently observed a partial blunting of tumor growth in the complete absence of host STING signaling. This finding suggests either an alternative source of STING signaling, or that STING signaling does not completely account for the therapeutic efficacy of cryoablation.
In addition to being a potential source of CDN/cGAMP, live RMS 76-9 maintains a functional STING signaling pathway and serves as a major source of IFN-β after STING activation either by co-culturing with cryoablated tumor or DMXAA. Genetic deletion of Tmem173 within RMS 76-9 completely abrogated cryoablation-induced IFN-β upregulation, indicating a STING dependency of this process within RMS 76-9. Surprisingly, cryoablation of RMS 76-9STINGwt was efficacious in suppressing primary tumor growth, as well as preventing lung metastasis regardless of whether or not the tumor-bearing hosts express functional STING in all tissue compartments. Beyond RMS 76-9-intrinsic STING signaling directly contributing to the clinical efficacy of cryoablation observed in STING-/- mice, a contributing factor may be the enhanced T cell survival in tumor-bearing STING-/- mice, as recent evidence demonstrated that STING activity induces T cell apoptosis,85–87 and gain-of-function STING-N153S mutation resulted in T cell deficiency.85 In sharp contrast to RMS 76-9STINGwt tumor, however, neither tumor-bearing C57BL/6 nor STING-/- mice benefited from cryoablation when treating RMS 76-9STING-/- tumor, and experienced rampant lung metastasis. These results provide strong evidence that tumor-intrinsic STING signaling drives both the local response to cryoablation and the generation of an effective systemic antitumor immunity. This conclusion was further supported by the observed abscopal effect in the co-injection model. Importantly, the vital contribution of tumor-intrinsic STING signaling to generate effective local immune TME extends beyond cryoablation into other therapeutic modalities including direct intratumoral injection of the STING agonist, DMXAA, suggesting a generalized vital role of tumor cell-intrinsic STING in generating effective antitumor immunity in other ICD-inducing cancer therapies.
Recent reports suggest a critical role for STING in tumor cell death, dormancy and metastasis. Although we did observe a slight baseline proliferative advantage of STING-/- tumors, we also observed a slightly enhanced susceptibility of these cells to cell death both in vitro and in vivo. We did not observe any significant difference in the growth of STINGwt and STING-/- tumors in vivo at baseline. These data led us to believe that the loss of cryoablation efficacy in STING-/- tumors is not due to intrinsic survival advantage of the STING-/- tumor following cryoablation; rather, it is the lack of antitumor immunity in these tumors following the intervention that resulted in the lack of therapeutic efficacy.
Our investigation into tumor-intrinsic STING signaling pathway kinetics suggests that downstream signaling molecule activation requires time (up to 4 hours) and the effect can be transient. These data lend an explanation to the anecdotal clinical observation that partial and sublethal cryoablation rather than complete cryoablation results in more robust response since leaving live, bulky tumor after the procedure to respond to cryoablated tumors may provide a more robust STING signaling in tumors with a functional STING signaling pathway. Furthermore, our current discovery offers an explanation as to why clinical application of cryoablation in advanced tumors has been met with varied and unreliable responses thus far. Our survey of available pediatric tumor databases revealed a heterogeneous expression patterns among pediatric cancers, of which recurrent, refractory and metastatic RMS represents one of the persistent clinical therapeutic challenges and unmet clinical need,88–90 with improvement in 5-year survival experiencing one of the smallest increases of all pediatric and AYA cancers.91–95 Although outside the scope of the current study, investigations are sorely needed to further understand the status of STING and related signaling molecules in clinical tumor samples, and epigenetic or other methodologies to reactivate epigenetically silenced STING-related pathways as a means to enhance their responsiveness to cryoablation and other immune-activating therapeutic agents.96 97 For example, the regulation of STING activity is not solely dependent on cGAMP binding and activation of STING. In the basal state, STING is bound to its negative regulator, NLRC3.98 99 On exposure to ATP and stress, NLRC3 releases STING which allows binding of cGAMP and subsequent phosphorylation of TBK1 and IRF3 for the initiation of transcriptional programming including IFN-β production. These and other additional regulations of STING-related pathways require a careful investigation to fully capture the potential translational utility of our finding.
We provided preclinical proof of concept that tumor-intrinsic STING signaling contributes critically to cryoablation-mediated tumor suppression in vivo in at least three different tumor models, although with varying degree of efficacy. However, our current observations were made with a single cryoablation intervention in order to standardize cryoablation parameter in the experimental approach. Whether employing different treatment timing, duration and frequency would further enhance the antitumor immunity remains to be determined. Moreover, the potential direct effects of tumor cell-intrinsic STING on metastatic potential versus immunogenicity should be further investigated. Despite the overall improvement in the abscopal efficacy in our co-implantation tumor model, the abscopal tumor remained sizeable and many progressed over time, suggesting the need for the addition of combination therapy, such as immune checkpoint blockade approaches (eg, NCT05302921) to improve overall systemic response. Lastly, it remains to be determined whether the observed effect of tumor cell-intrinsic STING contribution to local and systemic immunity converges on IFN-β, chemokines or other as-yet undiscovered effector molecules. Although IFN response is the most recognized signaling activity of STING and such IFN signaling is widely believed to be the major contributor to STING-mediated antiviral and antitumor responses, recent evidence suggests the existence of IFN-independent activities of STING in hematopoietic-derived cells including T cells,18 20 21 as mice harboring the functionally deficient STINGS365A/365A point mutant are still capable of exhibiting STING-related IFN-independent activities. Importantly, when analyzing chemokine and cytokine productions within TME, we found elevated IFN-β levels following cryoablation of RMS STINGwt irrespective of host STING status, whereas INF-β levels remained at baseline in cryoablated STING-/- RMS TME even in wild-type mice. IFN-β was not the only cytokine behaving this way. Indeed, chemokine and cytokine arrays of various TME revealed increases in several chemokines and cytokines (eg, CXCL9, CXCL10, CCL6, CCL8, CCL9/10, IL-16, and CXCL2) in the RMS STINGwt TME following cryoablation even within the context of a wild-type mouse. In contrast, little change in the cytokine and chemokine arrays were observed in the RMS STING-/- TME following cryoablation, despite the host being fully capable of responding to STING activation (figure 7). This intriguing interplay between tumor STING signaling and host response warrants further rigorous investigations.
In conclusion, we identified the tumor cell-intrinsic STING pathway as a critical contributor to both local response and the abscopal effect in cryoablation and local STING agonist therapy. Our results highlight the tumor cell-intrinsic STING-related signaling expression as a biomarker of potential therapy responses in cryoablation, and argues for the overexpression and/or reactivation of STING related pathway in tumor cells as a strategy for developing immunotherapeutic approaches and increasing the potential therapeutic benefits against solid tumors.
Data availability statement
All data relevant to the study are included in the article or uploaded as supplementary information. Not applicable.
Ethics statements
Patient consent for publication
Ethics approval
All animal experiments were performed in animals 8–12 weeks old with strict adherence to the active experimental protocols approved by the Institutional Animal Care and Use Committee at CWRU (protocol number 2015-0018) and performed in accordance with the guidelines of the American Association for Accreditation of Laboratory Animal Care and the NIH.
Acknowledgments
The authors would like to thank Dr Jason Williams as an expert consultant in cryoablation, IceCure Medical, Ltd. for making the ProSense CryoSurgical System available for this project; Adam Kresak for the help with tissue histology; and Jan Skotheim for providing the CSII-prEF1a-E2-Crimson-NLS plasmid (Addgene plasmid # 125263). IHC images were obtained with a Nanozoomer S60 slide scanner funded by NIH shared instrumentation grant (ORIP S10OD024981). The authors also acknowledge support of the Center for Pediatric Immunotherapy at the Angie Fowler AYA Cancer Institute/UH Rainbow Babies & Children’s Hospital. MA is supported by a scholarship from Qassim University, Buraydah, Saudi Arabia. AY-CH is supported as a fellow of the Harrington Discovery Institute and an endowed chair from the Theresia G. & Stuart F. Kline Family Foundation.
References
Supplementary materials
Supplementary Data
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Footnotes
MA and SLT contributed equally.
Contributors Conceptualization: MA and AY-CH. Formal analysis: MA, SLT, MB, STS, SHC. Resources: AY-CH. Data curation: MA, SLT, JM, DK, SE, MA, SHC. Writing—original draft preparation: MA, SLT. Writing—review and editing: MA, SLT, MB, SHC, AY-CH. Visualization: MA, SLT, AY-CH. Supervision: AY-CH. Project administration: MA, SLT, AY-CH. Funding acquisition: AY-CH. All authors have read and agreed to the published version of the manuscript. MA and SLT contributed equally to this work. AY-CH is responsible for the overall content of this work as the guarantor.
Funding This work was funded by the St. Baldrick’s Foundation, Pediatric Cancer Research Foundation, Alex’s Lemonade Stand Foundation for Childhood Cancer, MIB Agents, Children’s Cancer Research Fund, Sarcoma Foundation of America, the Char & Chuck Fowler Family Foundation, the I’m Not Done Yet Foundation, Case Comprehensive Cancer Center AYA Oncology Pilot Grant, Case Comprehensive Cancer Center-VeloSano Bike to Cure Pilot Award, the Risman Family Philanthropic Funds, the Cleveland Foundation, and the Alan & Karen Krause Family Foundation. M. Alshebremi is supported by Qassim University, Qassim, Saudi Arabia for the PhD scholarship. M. Abiff is supported by NIH/NCI NRSA F31 award (F31CA254259) and NIH/NIGMS T32 award (T32GM007250). MB is supported by NIH/NIGMS T32 award (T32GM007250) and NIH/NCI T32 award (T32CA059366).
Competing interests None declared.
Provenance and peer review Not commissioned; externally peer reviewed.
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