Abstract
This protocol describes how to induce large numbers of tumor-specific cytotoxic T cells (CTLs) in the spleens and lymph nodes of mice receiving dendritic cell (DC) vaccines and how to modulate tumor microenvironments (TMEs) to ensure effective homing of the vaccination-induced CTLs to tumor tissues. We also describe how to evaluate the numbers of tumor-specific CTLs within tumors. The protocol contains detailed information describing how to generate a specialized DC vaccine with augmented ability to induce tumor-specific CTLs. We also describe methods to modulate the production of chemokines in the TME and show how to quantify tumor-specific CTLs in the lymphoid organs and tumor tissues of mice receiving different treatments. The combined experimental procedure, including tumor implantation, DC vaccine generation, chemokine-modulating (CKM) approaches, and the analyses of tumor-specific systemic and intratumoral immunity is performed over 30–40 d. The presented ELISpot-based ex vivo CTL assay takes 6 h to set up and 5 h to develop. In contrast to other methods of evaluating tumor-specific immunity in tumor tissues, our approach allows detection of intratumoral T-cell responses to nonmanipulated weakly immunogenic cancers. This detection method can be performed using basic laboratory skills, and facilitates the development and preclinical evaluation of new immunotherapies.
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Introduction
Recent studies involving cellular immunotherapies and inhibitors of immune checkpoint molecules (checkpoint blockade) have demonstrated the ability of the immune system to control tumor growth1,2,3,4,5,6. To date, immunotherapies have been particularly effective in the therapy of tumors that are already infiltrated with CTLs6,7,8,9,10,11,12.
Here, we describe a protocol to enhance the numbers of tumor-specific type-1 cells in cancer-bearing mice and present strategies to reprogram TMEs for enhanced CTL attraction. The methods include the generation of murine type-1-polarized DCs13,14 loaded with autologous tumor material14,15, the induction of tumor-specific CTLs in murine lymphoid organs, and different forms of TME modulation to promote effective CTL entry into tumor lesions16,17,18. We further outline methods to evaluate the changes in the numbers of tumor-specific CTLs in tumor lesions19 during immunotherapy.
Development of the protocol
Intratumoral accumulation of CTLs is an independent prognostic factor for survival of patients with different cancer types20,21,22,23,24,25,26 and is required for the clinical effectiveness of checkpoint blockade therapies6,7,8,9,10,11,27,28. By contrast, multiple studies have shown that increases in circulating tumor-specific T cells in the course of immunotherapy do not predict clinical responses29,30. These observations highlight the importance of tumor-specific T cell entry into the TME as the key limiting factor in the effectiveness of anticancer immunity during natural immune surveillance and cancer immunotherapy31,32,33,34.
New strategies to promote the accumulation of CTLs in tumor lesions. Several strategies have been shown to activate endogenous antigen-presenting cells (APCs) and other TME components, resulting in enhanced cross-priming of tumor-antigen-specific CD8+ T cells, augmented local chemokine production, and enhanced CTL accumulation34,35. Productive cross-priming of CD8+ T cells in vivo involves early innate immune recognition of cancer cells and induction of type-1 interferons (IFNs) in DCs36. The stimulator of interferon genes (STING) pathway detects the presence of a tumor and can drive DC activation and induction of T-cell immunity against tumor-associated antigens (TAAs) in vivo37,38. Alternative strategies involve targeting exogenous antigens and adjuvants to DCs (reviewed in Kreutz et al.39), or exposing tumor tissues to combinatorial CKM adjuvants involving exogenous type I IFNs40 in combination with Toll-like receptor (TLR) ligands or interleukin (IL)-18 (refs. 16, 18, 41, 42, 43, 44), or targeted delivery of CTL-attracting chemokine genes to tumors45,46.
Dendritic cell therapies. To bypass dysfunction of endogenous DCs in cancer-bearing individuals, ex vivo-generated and antigen-loaded DCs can be used as therapeutic agents. The ability of DC vaccines to promote intratumoral CTL infiltration was seen in patients with hormone-refractory prostate cancer receiving Sipuleucel-T (Provenge)47, the first FDA-approved APC-based cancer vaccine48.
Modes of DC generation and maturation. Mouse and human DCs can be generated ex vivo from bone marrow or blood precursors using granulocyte/monocyte colony-stimulating factor (GM-CSF), alone or in combination with IL-4 or tumor necrosis factor (TNF)-α49,50,51,52,53,54,55. Mouse DCs can also be generated from thymic or splenic progenitor cells56,57. Pretreatment of mice with fms-like tyrosine kinase 3 ligand (Flt3L), a factor promoting DC development, can be used to increase the numbers of DC progenitors and DCs in bone marrow, spleen, lymph nodes, and other tissues58,59.
DC maturation is needed for effective induction of T-cell immunity60. Factors that can induce maturation of DCs include TLR ligands (e.g., microbial components such as lipopolysaccharide (LPS), peptidoglycan, cholera toxin, filamentous hemagglutinin, inactivated Bacillus Calmette–Guérin (BCG), or double-stranded RNA)61, cytokines (including type I interferons, IFN-γ, TNF-α, and IL-1β), prostaglandin E2 (PGE2), damage-associated molecular pattern molecules released by dying or damaged cells (apoptotic bodies, heat shock proteins, and urate crystals)62, or signals provided by other immune cells (T, natural killer (NK), NKT, or γδT cells)63. A maturation cocktail used to generate DCs for early clinical trials involved TNF-α, IL-1β, IL-6, and PGE2 (refs. 64, 65). However, observations that PGE2-matured DCs produce only very low amounts of IL-12p70 (refs. 13, 66, 67, 68, 69) prompted a search for alternative DC maturation factors. The ability of DC vaccines to produce IL-12p70 is critical for the induction of Th1/CTL/NK-cell-dominated type-1 immunity70,71,72,73,74 and is predictive of the positive clinical outcomes in DC-vaccinated cancer patients75,76,77. Among others, we proposed the combination of LPS with IFN-γ to induce highly immunogenic, high-IL-12-producing DCs66,70,71,78,79,80. LPS may be replaced with cytosine-phosphorothioate-guanine (CpG), polyinosinic:polycytidylic acid (poly-IC), or mixtures of more than one TLR ligand to generate DCs with high levels of co-stimulatory molecules necessary for T-cell activation and high production of IL-12, necessary for type-1 immune responses13,61,81,82,83.
Selection of a relevant antigen source is another aspect critical for the therapeutic activity of DC vaccines. One commonly used approach is the use of peptides or recombinant proteins representing defined TAAs84,85,86. An alternative approach is the use of tumor lysates or tumor apoptotic bodies as a source of DC-presented antigens87,88,89. Compared with peptide approaches, whole-tumor-based vaccines can provide a range of antigens and induce polyclonal immune responses90. Especially for vaccination against tumor types that do not have well-characterized common tumor-rejection antigens, UV beta and gamma (UVBγ)-irradiated tumor (UVBγ-Tu) cells15,91 or freeze–thaw tumor lysates14 are a convenient source of multiple TAAs, including patient-specific unique tumor antigens. DCs can also be transfected with total mRNA or DNA from tumors to induce CTLs against unique tumor antigens92.
Routes of DC administration. DC entry into lymph nodes is important for their ability to effectively activate tumor-specific CTLs93,94. Although the intravenous route is the standard mode of administration of the APC-based Sipuleucel-T (Provenge)47, DCs administered intradermally have been shown to migrate to the lymph nodes better than intravenously or intraperitoneally injected DCs95,96,97. Ultrasound-guided intranodal injection or intralymphatic administration can also be used to deliver DCs to lymph nodes97,98,99. Alternatively, intratumoral injection of DCs can be used for the induction of antitumor immune responses97,100,101. Although intratumorally injected immature DCs take up tumor antigens and migrate to the draining lymph nodes, intratumorally injected mature DCs induce local priming of CD8+ T cells in the tumors, rather than in lymphoid tissues101,102.
Approaches to enhance local effectiveness of tumor-infiltrating CTLs. In addition to inducing high numbers of tumor-specific T cells and promoting their entry into tumors, the effectiveness of anti-tumor immunity benefits from counteracting tumor-associated immunosuppression and blocking immune checkpoints103,104. Preclinical models have shown that immune checkpoint blockade (anti-CTLA-4 and anti-PD-1/PD-L1 mAbs) can synergize with vaccines105,106,107,108. T-cell co-stimulatory factors109 (e.g., CD40, CD70, LIGHT, OX40L, B7H3, and 4-1BBL), functional inhibition of T regulatory (Treg) cells, and/or myeloid-derived suppressor cell (MDSC) activity by cyclooxygenase-2 (COX-2) (refs. 110, 111, 112) and indoleamine 2,3-dioxigenase 1 (IDO1) (ref. 113) inhibitors also predispose the TME toward effective antitumor immunity. The combinatorial approaches combining DC or viral therapies with COX-2 blocker described in this protocol have a dual role, promoting both CTL influx into tumor lesions and their local antitumor activity.
Applications of the current protocol
This protocol is designed to enhance the numbers of tumor-specific CTLs within tumor lesions and to evaluate their changing numbers in the course of immunotherapy. The approaches involve systemic immunization using DC vaccines and different forms of systemically or locally applied modulation of TME, such as systemic or local application of combinatorial adjuvants, intratumoral injection of DCs, or viral therapies.
The presented DC-based approaches can be used to efficiently induce CTLs recognizing different tumor-relevant antigens and their entry into tumors. We anticipate that they may be combined with the blockade of tumor survival pathways114, oncolytic virus therapies115, TNF superfamily ligands116, and blockers of inhibitory factors in the TME (e.g., immune checkpoints, IDO or COX2 blockers). The application of combinatorial adjuvants involving type I IFNs in combination with TLR ligands to promote the entry of the CTLs into the tumor lesions can be used as self-standing therapies following tumor resection, or can be combined with vaccines, checkpoint blockade, or adoptive T-cell therapies to enhance their effectiveness against advanced unresectable tumors.
Comparisons with alternative techniques
Common methods of evaluating the numbers of tumor-specific T cells in tumor tissues involve the use of T cells from genetically manipulated mice with defined T-cell receptors (e.g., OT-I or OT-II), and genetically modified tumor cell lines that uniformly express highly immunogenic model antigens, such as ovalbumin (OVA) (reviewed in Dranoff117). As the artificially high immunogenicity of such genetically manipulated tumor cells is known to substantially affect both the magnitude and character of immune responses (reviewed in Ngiow et al.118), such models may not be fully relevant to the immunotherapy of human tumors, which rarely express strong antigens and are antigenically heterogenous.
Advantages of the current protocol
The ability to evaluate polyclonal T-cell responses to multiple tumor-related weak antigens (mutated or overexpressed 'self' antigens) provides a more direct translatable readout, by eliminating the need for surrogate antigens or genetically manipulated T cells with artificial T-cell receptors (TCRs) and allowing the evaluation of T cells recognizing different TAAs expressed by cancer cells. It eliminates the need for known cancer antigens and an arbitrarily selected threshold of antigen specificity, reflected in the design of tetramers or defined α/β TCR combinations. Our protocol allows for enumeration of spontaneously arising and immunotherapy-induced T cells that are able to recognize any cancer cell. Thus, the key advantages of the current technique are its ability to evaluate responses to multiple TAAs without the need for prior identification of the relevant immunogenic epitopes, alleviating the bias related to the narrow focus on high-affinity TCR-antigen interactions, and its immediate general applicability to multiple tumor models and mouse strains.
Limitations of the current protocol and comparison with other methods
Our method relies on the detection of cytokine-producing cells (IFN-γ and potentially other mediators of effector and suppressive T cells) upon stimulation of tumor-isolated lymphocytes with relevant cancer cells. Cytokine-producing cells can be detected by flow cytometry or ELISpot analysis, and potentially by single-cell analyses of CD8+ T-cell degranulation (such as CD107 assay)119 and target cell killing120.
Our approach allows detection of antigen-specific cells that are capable of infiltrating tumor lesions in vivo and that specifically recognize cancer cells. However, isolated tumor-infiltrated lymphocytes (TILs) and/or tumor-associated lymphocytes (TALs) must be rechallenged in vitro with the original cancer cells (nonprofessional APCs) to detect tumor-specific responses, which may result in apoptosis of a proportion of antigen-specific cells.
In terms of the ability to detect very low cell numbers, ELISpot assay and detection by tetramers are very similar (1 in 3–10 tetramer-positive cells are positive in ELISpot analysis)121. The advantage of detection by tetramers is that cells can be additionally stained for various markers and subsets of tetramer-positive cells can be analyzed. However, as T-cell responses to tumor epitopes are known to be weaker than antiviral responses or responses to model antigens, such as OVA, ex vivo analysis by tetramers allows for detection of only a relatively narrow range of the potential tumor epitopes available.
Overview of the protocol
The three main parts of the protocol are generation of DCs (Fig. 1, Steps 1–7), quality control of the DCs (QC: Fig. 1, Step 9), and monitoring of the therapeutic and immunomodulatory effect of DCs in mice inoculated with tumor cells (Fig. 1, Steps 10–24). Each of these parts can be performed at separate time points, starting with DC generation, and followed by QC and monitoring of the effectiveness of DC vaccine on tumor progression. DC generation starts with administration of Flt3L to C57BL/6J mice on day −11. It is followed by CD11c+ cell isolation and induction of mature type-1 polarized DCs on day 0, including optional cryopreservation of DCs. QC involves confirming the mature DC phenotype, detecting expression of co-stimulatory molecules (Fig. 2a), and determining the ability to produce IL-12 upon CD40L stimulation (Fig. 2b). Additional analyses include in vitro induction of type-1 immune responses by DCs (Fig. 2c), assessment of the capacity of DCs to migrate to lymph nodes (Fig. 2d), and in vivo induction of type-1 immune responses (Fig. 2e). Our tumor model involves intraperitoneal injection of cancer cells into experimental mice. A day before DC vaccination, mice are imaged to determine the baseline intensity of the bioluminescence signal. Mice receive vaccination with tumor-loaded DCs at day 4 after cancer cell inoculation. 3 d after DC vaccination, CKM adjuvants are applied (e.g., intratumoral DC injections; combination of IFN-α, Ampligen, and COX2 blockers; or chemokine-expressing vaccinia virus) to modulate the TME. The cycles of DC therapy and CKM regimen should be repeated weekly to prolong the survival of the tumor-bearing mice. Separate sets of experimental mice are used for tumor progression/survival monitoring and immune monitoring. Enumeration of tumor-specific type-1 immune cells is typically performed 4–5 d after CKM adjuvant application. Although the current protocol has been developed using the C57BL/6 mice, other mouse strains in combination with the relevant MHC-compatible tumor models may be used instead.
Experimental design
DCs should be generated from mice of the same genetic background as that of the experimental mice to avoid allogeneic immune responses. In this protocol, we use CD11c+ splenocytes isolated from Flt3L-expressing B16 tumor-bearing donor mice (Box 1) to generate DCs. During the maturation, we expose the isolated CD11c+ cells to UVBγTu cells (e.g., MC38 or ID8A cells) and use the generated DCs to induce tumor-specific TILs into the experimental mice injected with matching tumor cells.
DC generation. In this immunotherapy protocol, we replaced the in vitro generation14,82 of bone marrow-derived DC with in vivo induction of intrasplenic DC accumulation after injection of B16-Flt3L cells to enhance the efficiency and reduce the costs115. We inject Flt3L-expressing B16 cells into subcutaneous tissue in the right flank of C57BL/6 mice115 to avoid the need for daily injections and the cost of recombinant Flt3L. Alternatively, daily injection of 10 μg of Flt3L subcutaneously to mice for 9–11 consecutive days can be done instead. Administration of either recombinant Flt3L or Flt3L-expressing B16 cells results in expansion of both CD8α+ and CD8α− DC subsets (CD11c+ class II+)58,122,123. CD11c+ immature DCs can then be isolated from the spleens of mice injected with Flt3L-B16 cells or the recombinant Flt3L-treated animals and exposed to cytokines and/or TLR ligands ex vivo to generate type-1 polarized DCs.
Vaccine generation. To effectively promote anti-tumor immunity, DCs must be mature and produce high levels of IL-12 (refs. 13, 61, 70, 71, 72, 73, 74, 75, 76, 77, 81, 82, 83). The manner in which DCs are matured will affect the phenotype and functional activity (i.e., IL-12-producing capacity) of the mature DCs. To achieve translational analogy to our clinical DC vaccines76, the LPS-based maturation method82 was replaced with the mouse variant of our previously developed maturation cocktail13. Our DC maturation cocktail contains the recombinant murine cytokines rmGM-CSF, rmIL-1β, rmIFNα, rmIFNγ, rmTNFα, and poly-IC to convert DCs into type-1 polarized DCs.
We observed that, compared with our human protocol13,66,124,125, the maturation period of mouse DCs needed to be reduced from 16–48 to 4 h for optimal DC stimulatory capacity. The 4 h–matured DCs can secrete IL-12p70 while showing high levels of expression of key co-stimulatory and maturation markers (e.g., CD40, CD80, CD86, and CD83) and the lymph node-homing chemokine receptor CCR7 (Giermasz et al.14; Fig. 2a,b).
We use a UVBγ irradiation-based technique to prepare apoptotic tumor cells (UVBγ-Tu cells) for antigen loading15,91,99,126 (Box 2). The ability to take up UVBγ-Tu cells is reduced in DCs that have completed the maturation process127. Thus, we load DCs with UVBγ-Tu cells during their maturation to generate type-1 polarized DCs that produce high levels of IL-12p70 and cross-present tumor antigens15,91.
In a single process using ten donor mice, we can generate up to 150 million DCs, which, after QC testing, will suffice for ∼500 doses of a single batch of vaccine.
QC of tumor-loaded DCs. Before vaccine administration in tumor-bearing mice, we test each batch of DC vaccines for consistency in phenotypic and functional characteristics. We perform flow-cytometry analysis for expression of markers such as IA-Eb, H2Kb, CCR7, CD14, CD40, CD80, CD83, and CD86 in DCs that have been matured in vitro with Th1-inducing cocktail. Apart from expressing co-stimulatory molecules and CCR7 (Fig. 2a), type-1 polarized DCs produce IL-12p70 upon stimulation with CD40L128 (Fig. 2b). Consistent with the key role of IL-12p70 in the induction of type-1 immunity, tumor-loaded DCs matured in the presence of type-1-inducing cocktail induce high numbers of tumor-specific T cells129 in in vitro cultures of syngeneic splenocytes (Fig. 2c). Another important functional characteristic of DCs is their ability to migrate to draining lymph nodes (Box 3). In vivo migration of DCs generated from firefly luciferase (CAG-luc-eGFP) transgenic mice can be traced by bioluminescence imaging (BLI) after injection. In vivo migration of DCs from B6 CD45.1 mice or carboxyfluorescein succinimidyl ester (CFSE)-labeled DCs from C57BL/6J mice130 can be detected by flow cytometry of lymph node-infiltrating cells. The advantage of using CAG-luc-eGFP DCs is that migration can be monitored in the same mice over time (Fig. 2d). Generation of CD45.1+ DCs avoids the need for DC labeling, as the cells can be detected by staining for the CD45.1 marker. Furthermore, the functional activity of DCs can most comprehensively be evaluated by the in vivo induction of tumor-specific type-1 immune response assay (Fig. 2e).
Mouse tumor models. The evolving role of animal models in cancer vaccine development has been recently reviewed118,131. The assessment of immunotherapy using inducible tumor models in genetically manipulated mice or carcinogen-induced tumor models is particularly relevant to the physiological situation, as they involve chronic inflammation, development of immune tolerance, tumor immune editing, and immunosuppressive events. At the same time, transplantable models (we have used ovarian cancer ID8A and colorectal cancer MC38 cells) offer multiple advantages, especially regarding their costs and feasibility to address specific mechanistic questions (e.g., manipulation of antigenicity, suppressive activity, resistance to apoptosis, chemokine production), as recently summarized by Mac Keon et al.132. In orthotopic tumor models, the tumor cell lines are implanted into the relevant organ and are allowed to form metastases, reflecting the human disease133. The ID8A tumor-bearing mice develop massive malignant ascites in the late stages, mirroring the characteristics of human epithelial ovarian cancer. As different sites of tumor inoculation can give rise to different immune microenvironments, these can affect tumor progression and therapeutic outcomes134. Therefore, our protocol involves two alternative sites of MC38 tumor implantation (subcutaneous and intraperitoneal) in syngeneic C57BL/6J mice, which are both similarly susceptible to the described immunotherapies.
Promoting the infiltration of tumor-specific T cells into tumor tissues. Homing of effector type-1 immune cells into the TME is critical for effective immunotherapy. To promote the migration of CTLs to tumors, we developed two strategies of TME modulation (Fig. 3a). Our basic protocol combines two individual doses of DC injection. In the first subcutaneous vaccine is followed 3 d later by a second intratumoral booster (Fig. 3b,c). Alternatively, the vaccination may be combined with the selective induction of effector-attracting chemokines using combinatorial CKM adjuvants16. Multiple chemokines can be targeted to manipulate and alter the TME, in order to correct the local balance of immunosuppressive and effector cells (reviewed in Devaud et al.135). A potent way to manipulate TME is the use of TLR ligands (TLR-Ls). TLR-Ls trigger broad inflammatory responses that elicit rapid innate immunity and promote the activation of the adaptive immune reaction136. Examples of commonly used TLR-Ls are poly-IC (TLR3 agonist) and CpG (TLR9 agonist)137,138. Although systemic application of the individual TLR-Ls can induce a substantial nonselective systemic response composed of both the effector cell-attracting chemokines and Treg/MDSC attractants, their combination with cytokines and blockers of the COX2 pathway allows the focus of their activity on tumor tissues and selectively induces desirable chemokines16,17,18. Therefore, we use the combination of type-1 interferons, TLR ligands (such as poly-IC), and COX2 inhibitors (such as celecoxib or aspirin) (Fig. 3). Another approach to selectively target tumor lesions139 is the introduction of effector cell–attracting chemokines by administration of genetically manipulated viruses136,140. Oncolytic viruses are known to stimulate multiple TLRs141 and predominantly infect cancer cells139. Directing cytokines (e.g., IFN-α and IL-12) specifically to tumors, using intratumoral administration of viral vectors or DC vaccines, has been shown to reprogram the TME and increase efficacy of additional immunotherapeutic agents76,142,143,144.
The presented DC vaccination and CKM approaches may be used as individual modalities or in combinatorial approaches to enhance efficacy of intratumoral tumor-specific T cells to achieve optimal therapeutic outcomes145,146.
Detection and quantification of tumor-specific immune responses in the tumor tissues compared with evaluation of systemic immunization. The in vivo tumor-specific cytotoxicity assay using i.v. injection of a 1:1 mixture of nonloaded CFSEnegative/dim and antigen-loaded CFSEhigh splenocytes115 (Boxes 4 and 5) is a straightforward method to determine the magnitude of the systemic tumor-specific immune response in tumor-bearing mice (Box 6 and Fig. 2e). However, this strategy does not allow evaluation of the effectiveness of new cancer therapies to promote the increase in the numbers of tumor-specific T cells within tumor tissues. To address this issue, we developed a highly sensitive ex vivo IFN-γ ELISpot-based protocol allowing us to compare the numbers of tumor-specific cells in splenocytes19 and TILs. Our protocol involves the isolation of immune cells from cancerous tissues (e.g., solid primary tissues and malignant ascites) using tissue digestion and subsequent Percoll gradient centrifugation, followed by ex vivo exposure of the isolated immune cells to the relevant antigens, such as whole cancer cells (Fig. 4) or, potentially, defined peptide-loaded target cells.
Materials
REAGENTS
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RPMI-1640 medium (Gibco, Invitrogen, cat. no. 61870-010)
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Recombinant mouse IL-1β (research grade; Miltenyi, cat. no. 130-094-053)
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Recombinant mouse TNF-α (research grade; Miltenyi, cat. no. 130-094-085)
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Recombinant mouse IFN-γ (research grade; Miltenyi, cat. no.130-094-048)
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Recombinant mouse GM-CSF (research grade; Miltenyi, cat. no. 130-094-043)
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Recombinant mouse IFN-α (research grade; Miltenyi, cat. no.130-093-131)
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Recombinant mouse IL-2 (research grade; Miltenyi, cat. no.130-094-055)
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Recombinant mouse IL-7 (research grade; Miltenyi, cat. no. 130-094-636)
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Poly-IC (Sigma-Aldrich, cat. no. P9582-5MG)
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Ampligen (polyI:polyC12U; Hemispherx Biopharma)
Critical
Ampligen is safer and more specific in comparison with poly-IC.
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Celecoxib (Biovision, cat. no. 1574100)
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Aspirin (Sigma-Aldrich, cat. no. A5376-100G)
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Trypan blue (0.4% (wt/vol); Corning Cellgro, cat. no. MT-25-900-CI)
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L-Glutamine (L-glu; Gibco; Invitrogen, cat. no. 25030-024)
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Penicillin–streptomycin (10,000 U/ml; Gibco; Invitrogen, cat. no. 15140-114)
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Non-essential amino acids (NEAAs; Gibco; Invitrogen, cat. no. 11140-035)
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Sodium pyruvate (Gibco; Invitrogen, cat. no. 11360-039)
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2β-mercaptoethanol (2βME; Sigma-Aldrich, cat. no. M3148)
Caution
β-mercaptoethanol is toxic upon inhalation, upon contact with skin, and if swallowed, and it is hazardous to the aquatic environment. Avoid contact with skin, eyes, and clothing, and handle it with gloves in a chemical fume hood.
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FBS (Gemini Foundation B, cat. no. 900-208)
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ACK lysis buffer (Life Technologies, cat. no. A10492-01)
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Mouse CD11c microbeads (Miltenyi Biotec, cat. no. 130-052-001)
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Collagenase (10% (wt/vol) stock; Sigma-Aldrich, cat. no. C0130-100MG)
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Tumor Dissociation Kit (enzymes D, R, and A; Miltenyi Biotec, cat. no. 130-096-730)
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Percoll (Sigma-Aldrich, cat. no. P1644)
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PBS (Cellgro Cell Culture PBS (1×); Corning, cat. no. MT-21-040-CV)
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EDTA (500 mM; Life Technologies, cat. no. 15575-020)
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Carboxyfluorescein succinimidyl ester (CFSE; Life Technologies, cat. no. C34554)
Critical
Protect the reagent from light.
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Ethanol (diluted 70% (vol/vol); Pitt Pharmco, cat. no. 111000200CS)
Caution
Ethanol is flammable, and it may cause skin and eye irritation. Avoid contact with skin, eyes, and clothing, and handle it with gloves in a chemical fume hood.
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Luciferin (Gold Biotechnology, cat. no. Luck-1g)
Critical
Protect the luciferin from light.
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OVA peptide SIINFEKL (Genescript, cat. no. rp10611)
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R4-6A2 anti-IFN-γ biotinylated detecting antibody (Mabtech, cat. no. 3321-6-250)
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AN18 anti-IFNγ-coating antibody (Mabtech, cat. no. 3321-3-1000)
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ABC reagent (Vector Laboratories, cat. no. PK-6100)
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ELISpot AEC substrate kit (Vector Laboratories, cat. no. SK-4200) (Vector Laboratories, cat. no. PK6100)
Caution
ELISpot Substrate reagent is flammable and is harmful upon contact with skin, of if swallowed or inhaled. Avoid contact with skin and eyes and handle it with gloves.
Antibodies for flow cytometry
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CD80-APC (Armenian hamster IgG, 0.2 mg/ml; BioLegend, cat. no. 104714)
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CD40-APC (IgG2a, 0.2 mg/ml; BioLegend, cat. no. 124612)
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CD86-APC (IgG2a, 0.2 mg/ml; BioLegend, cat. no. 105012)
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CD14-FITC (IgG1, 0.5 mg/ml; BD Biosciences, cat. no. 553739)
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CCR7-PE (IgG2a, 0.2 mg/ml; BioLegend, cat. no.120106)
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IA/Eb-FITC (IgG2a, 0.5 mg/ml; BioLegend, cat. no.114406)
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H2Kb-FITC (IgG2a, 0.5 mg/ml; BioLegend, cat. no. 116505)
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IgG1-FITC (IgG1, 0.5 mg/ml; BD Biosciences, cat. no. 553953)
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IgG2a-FITC (IgG2a, 0.5 mg/ml; BD Biosciences, cat. no. 554688)
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IgG-PE (Armenian hamster IgG, 0.2 mg/ml; BioLegend, cat. no. 400908)
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IgG2a-APC (IgG2a, 0.2 mg/ml; BioLegend, cat. no. 400219)
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IgG1-APC (IgG1, BD Biosciences, cat. no. 555751)
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CD45.1-APC (IgG2a, 0.2 mg/ml; BioLegend, cat. no. 110714)
Critical
Protect the antibodies from light.
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IL-12 ELISA Kit (R&D, cat. no. DY419)
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BSA (Sigma-Aldrich, cat. no. A9647)
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dH2O (Life Technologies, cat. no. 10977)
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Tween 20 (Fisher Scientific, cat. no. BP337-500)
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10× PBS (Cellgro PBS (1×); Corning, cat. no. MT-46-013-CM)
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NaCl (Sigma-Aldrich, cat. no. S7653)
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Na2HPO4·2H2O (Sigma-Aldrich, cat. no. 71643)
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KH2PO4 (Sigma-Aldrich, cat. no. P5655)
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ELISA substrate (Fisher Scientific, cat. no. ENN301)
Caution
ELISA substrate is flammable and is harmful upon contact with skin, or if swallowed or inhaled. Avoid contact with skin and eyes, and handle it with gloves.
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H2SO4 (Sigma-Aldrich, cat. no. 339741)
Caution
Sulfuric acid is highly flammable, and it may cause severe skin burns and eye damage. Avoid contact with skin, eyes, and clothing, and handle it with gloves in a chemical fume hood.
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Paraformaldehyde, 32% (wt/vol) solution (PFA; Electron Microscopy Sciences, cat. no. 15714-S)
Caution
PFA is a hazardous reagent; use a chemical fume hood and wear protective gloves and a mask when you are working with it.
Cells
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Flt3L-expressing B16 melanoma cells147 (C57BL/6J mouse strain), a gift from M. Kronenberg, La Jolla Institute for Allergy and Immunology
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Critical
The cell lines should be regularly checked to ensure they are authentic and are not infected with mycoplasma.
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Critical
B16-Flt3L cells can be replaced with recombinant Flt3L (Invitrogen, cat. no. 14-8001-80) as described in the Experimental design section.
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CD40L-transfected J558 (J558-CD40L) (Balb/c mouse strain) cells148, a gift from P. Lane, University of Birmingham. Alternatively, CD40L-expressing EL-4-B5 cells are available from Kerafast (cat. no. EVU301); however, we have never used them.
Critical
Soluble CD40L (MEGACD40L; Enzo Life Sciences, cat. no. ALX-522-120-C010) can be substituted for the CD40L-expressing cells, as described in Step 9B(iv).
Critical
The cell lines should be regularly checked to ensure they are authentic and are not infected with mycoplasma.
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MC38 (C57BL/6J mouse strain) colorectal cancer cells (luciferase-expressing)149 (available from Kerafast, cat. no. ENH204)
Critical
The cell lines should be regularly checked to ensure they are authentic and are not infected with mycoplasma.
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ID8A (luciferase-expressing), a cell line derived from spontaneous in vitro malignant transformation of C57BL/6 mouse ovarian surface epithelial cells, was a gift from T.J. Curiel, University of Texas Health Science Center at San Antonio
Critical
The cell lines should be regularly checked to ensure they are authentic and are not infected with mycoplasma.
Critical
Cancer cell lines should be of the same genetic background as the mice into which the cells will be injected, to avoid any allogeneic immune responses and tumor rejection.
Mice
-
Critical
Donor and experimental mice used in separate steps of the protocol are indicated in Table 1 and listed below.
C57BL/6J (The Jackson Laboratory, cat. no. 000664)
Caution
All procedures involving animal experiments should follow approved institutional and governmental animal protocols and comply with the relevant guidelines and regulations of the local animal ethics committee. The animal studies reported here were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Pittsburgh Cancer Institute (protocol 14063339).
-
FVB-Tg(CAG-luc,-GFP)L2G85Chco/J—backcrossed on the C57BL/6 line for >14 generations150—can be obtained from A. Beilhack, University Hospital of Wurzburg, or from the Jackson Laboratories (cat. no. 008450)
Caution
All procedures involving animal experiments should follow approved institutional and governmental animal protocols and comply with the relevant guidelines and regulations of the local animal ethics committee. The animal studies reported here were approved by the IACUC at the University of Pittsburgh Cancer Institute (protocol 14063339).
-
B6.SJL-PtprcaPepcb/BoyJ (C57BL/6 CD45.1; The Jackson Laboratory, cat. no. 002014)
Caution
All procedures involving animal experiments should follow approved institutional and governmental animal protocols and comply with the relevant guidelines and regulations of the local animal ethics committee. The animal studies reported here were approved by the IACUC at the University of Pittsburgh Cancer Institute (protocol 14063339).
EQUIPMENT
-
2-ml Pipettes (sterile, individually wrapped; Denville, cat. no. p7132)
-
5-ml Pipettes (sterile, individually wrapped; Denville, cat. no. p7133)
-
10-ml Pipettes (sterile, individually wrapped; Denville, cat. no. p7134)
-
50-ml Pipettes (sterile, individually wrapped; Denville, cat. no. p7135)
-
Alcohol prep pads (Fisher Scientific, cat. no. 22-363-750)
-
12 × 75-mm Polystyrene tubes (Falcon, cat. no. 14-959-11B)
-
Sterile cell culture plates (12 well; Denville, cat. no. t1012)
-
Sterile cell culture flasks (75 mm2; Falcon, cat. no. 13-680-65)
-
Sterile cell culture plates (96 well, flat-bottom; Denville, cat. no. t1096)
-
Sterile cell culture plates (96 well, V-bottom; Corning Costar, cat. no. CLS3894)
-
High-binding 96-well plates for ELISA (Costar 96-well EIA/RIA plates; Corning, cat. no. 07-200-35)
-
Cell strainers (100 μm; Falcon, cat. no. 08-771-19)
-
Cell strainers (70 μm; Falcon, cat. no. 08-771-2)
-
Conical centrifuge tubes (50 ml; Denville, cat. no. C1060-p)
-
Conical centrifuge tubes (15 ml; Denville, cat. no. c1018-p)
-
Eppendorf centrifugation tubes (2 ml; Sigma-Aldrich, cat. no. T2795-1000EA)
-
ELISpot plates (Millipore, cat. no. MAHAS4510)
-
Forceps (Fisher Scientific, cat. no. 08-880)
-
Scissors (Fisher Scientific, cat. no. 08-951-20)
Caution
Scissors are sharp. Handle them with care.
-
Needles, 18 gauge, 22 gauge, 27 gauge (BD Biosciences, cat. nos. 305195, 305155, 305109)
Caution
Needles are sharp. Handle them with care.
-
0.5-ml Syringes (BD Biosciences, cat. no. 305620)
-
3-ml Syringe (BD Biosciences, cat. no. 309585)
-
10-ml Syringe (BD Biosciences, cat. no. 309604)
-
Pipette tips (10, 200, 1,000 μl; MidSci, cat. nos. AV10R-H, AV200, AV1000)
-
Cryovials (Thermo Scientific, cat. no. 12-565-164N)
-
Sterile beakers (Pyrex, cat. no. 02-540M)
-
Pasteur pipettes (Denville, cat. no. p0458-9)
-
Countess cell counter (Invitrogen) or Neubauer hemocytometer (Fisher Scientific, cat. no. 02-671-10)
-
Microscope (Carl Zeiss, model no. Axiovert 200)
-
FACS instrument (Accuri or Fortessa; BD Biosciences)
-
MACS MultiStand (Miltenyi Biotec, cat. no. 130-042-303)
-
QuadroMACS separator (Miltenyi Biotec, cat. no. 130-090-976)
-
Cell incubator (temperature = 37 °C, CO2 level = 5% (vol/vol); Thermo/Kendro, model no. Heracell 150)
-
Gamma irradiator (Nordion International)
Caution
Gamma irradiation is hazardous. Appropriate safety measures should be taken while working with a gamma irradiator.
-
UVBγ light (Spectroline, model no. EB-160C)
Caution
UV light is hazardous. Appropriate personal protective equipment should be worn while using a UV source.
-
ELISpot reader (ImmunoSpot, Cellular Technology)
-
ELISA reader (Perkin Elmer, model no. Victor 2)
-
In vivo bioluminescence imager (IVIS; Perkin Elmer)
-
Temperature-controlled freezing container (Mr Frosty; Thermo Scientific, cat. no. 51000001)
-
Centrifuge (Eppendorf, model no. 5810 R)
-
Pipetman (P10-P100; Gilson, cat. no. F123615G)
-
Pipetman (P100-P1000; Gilson, cat. no. F123602G)
-
Pipet-Aid (Drummond Scientific)
-
Pipettes (0.5–10 μl, 20–200 μl, 100–1,000 μl; Eppendorf, cat. nos. 3121.000.020, 3121.000.054, 3121.000.062)
-
Multichannel pipette (20–300 μl; Thermo Scientific, cat. no. 4661140)
REAGENT SETUP
FBS
-
We purchase the serum in frozen 500-ml bottles and store them directly at −20 °C. Individual bottles should be slowly thawed (at 4 °C overnight) and heat-inactivated for 30 min at 56 °C, and then serum aliquots should be prepared in 50-ml tubes and frozen again. Freeze–thaw cycles should be avoided. The serum can be stored for up to 2 years at −20 °C.
Standard isotonic Percoll solution
-
Prepare 10× acidic (pH: 4.6, 1.051 g/ml) PBS from 13.5 g NaCl, 0.125 g of Na2HPO4·2H2O, 2.1 g KH2PO4, and 200 ml dH2O. Sterilize acidic PBS by 0.22-μm filtration, and store at 4 °C in 4-ml aliquots. The maximum recommended storage time is 3 months. Percoll must be mixed with acidic PBS 10:1 and further diluted with culture medium to prepare 60% (vol/vol), 45% (vol/vol), and 34% (vol/vol) standard isotonic Percoll solution (SIP). Freshly prepare SIP before use and keep it at 37 °C.
Cell culture medium
-
To prepare cell culture medium, supplement RPMI-1640 medium with 1% (vol/vol) (100 U/ml) penicillin–streptomycin, 1% (vol/vol) (1 mM) sodium pyruvate, 1% (vol/vol) NEAAs, 0.1% (vol/vol) (14.3 mM) 2βME, 1% (vol/vol) (2 mM) L-glu, and 10% (vol/vol) FBS and store at 4 °C. The maximum recommended storage time is 1 month.
Cytokines
-
Many cytokine concentrations are indicated in IU/ml, which is the correct form for comparison on the basis of functional activity. However, for some cytokines, this information is not available. Values can differ greatly from company to company, suggesting that measurement of activity may be challenging. We provide information referring to specific cytokines from specific companies. Adjustments must be made when cytokines from different vendors are used. We prepare all cytokines as 100× or 1,000× concentrated stock solutions and store the aliquots at −20 °C. The maximum recommended storage time is 6 months. The concentrations vary, as indicated in Table 2.
DC maturation medium
-
To generate effector type-1 polarized DCs, prepare maturation medium before setting up the DC maturation. The volume needed depends on the number of isolated CD11c+ cells—∼10 ml of maturation medium is required for maturation of DCs isolated from one spleen. Supplement culture medium with cytokines as indicated in Table 2. The cytokines are stored at −20 °C and should be completely thawed at RT before adding to the medium. Allow the prepared medium to equilibrate to 37 °C before adding to the cultures.
Critical
Freshly prepare the DC maturation medium before use.
Cell-freezing medium
-
For freezing medium 1 (FM1), supplement RPMI-1640 medium with 50% (vol/vol) FBS. Freezing medium 2 (FM2) contains RPMI-1640 medium (40% (vol/vol)), FBS (40% (vol/vol)), and DMSO (20% (vol/vol)). Store both media at 4 °C. The maximum recommended storage time is 1 month.
CKM adjuvants for modulation of the TME
-
We prepare a solution of 5 × 104 IU/ml IFN-α and 0.25 mg/ml Ampligen in PBS for injecting 200 μl i.p. per mouse (for i.p. tumor models) and 2 × 105 IU/ml IFN-α and 1.0 mg/ml Ampligen in PBS for injecting 50 μl intratumorally per mouse (for s.c. MC38 tumor models).
Critical
Freshly prepare the CMK adjuvant solutions before use.
COX-2 inhibitors
-
We prepare a 4 mg/ml stock solution of celecoxib and a 200 mg/ml stock solution of aspirin in DMSO and store aliquots at −20 °C. The maximum recommended storage time is 6 months. We make 500 μg/ml (12.5% (vol/vol) DMSO, celecoxib) and 2.5 mg/ml (1.25% (vol/vol) DMSO, aspirin) working solutions in PBS immediately before administration (for injecting 200 μl per mouse).
Critical
Freshly prepare the COX-2 inhibitor solutions before use.
CFSE reagent
-
Prepare CFSE reagent in the dark by adding 18 μl of DMSO to the vial of lyophilized CFSE. Store CFSE reagent at −20 °C. The maximum recommended storage time is 1 month.
EDTA in PBS
-
For detachment of tumor cells, prepare 1 mM EDTA in PBS by adding 1 ml of 500 mM EDTA to 500 ml PBS and sterile-filter the solution through a 0.22-μm filter. Store at 4 °C. The maximum recommended storage time is 3 months.
MACS buffer
-
For MACS buffer (0.5% (wt/vol) BSA and 2 mM EDTA in PBS), supplement 500 ml of PBS with 2.5 g of BSA and 2 ml of 500 mM EDTA and sterile-filter the solution through a 0.22-μm filter. Store at 4 °C. The maximum recommended storage time is 3 months.
Blocking buffer
-
Prepare blocking buffer (3.0% (wt/vol) BSA in PBS) by adding 1.5 g of BSA to 50 ml of PBS and sterile-filter the solution through a 0.22-μm filter. Store at 4 °C. The maximum recommended storage time is 1 month.
Wash buffer
-
We prepare 10 liters of wash buffer (0.05% (vol/vol) Tween 20 in PBS) by diluting 1 liter of 10× PBS with 9 liters of dH2O and adding 5 ml of Tween 20. Store at room temperature (20–23 °C). The maximum recommended storage time is 3 months.
ABC reagent
-
Prepare avidin–peroxidase complex (Vector Laboratories) by adding one drop each of reagents A and B to 10 ml of wash buffer.
Critical
Freshly prepare ABC reagent before use.
Digestion buffer
-
Thaw digestion enzymes D, R, and A from the Tumor Dissociation Kit at room temperature (see manufacturer's instructions for reconstitution information). Per 2.5 ml of digestion medium, 100 μl of enzyme D, 50 μl of enzyme R, and 12.5 μl of enzyme A are required.
Critical
Freshly prepare the digestion buffer before use.
Luciferin
-
Thaw luciferin at room temperature. Reconstitute 1 g of luciferin in the dark with 33.3 ml of cold PBS and freeze at −80 °C in 1-ml aliquots. The maximum recommended storage time is 6 months.
Paraformaldehyde 4% (wt/vol) solution
-
Dilute 5 ml of paraformaldehyde 32% (wt/vol) solution with 35 ml of PBS. Store at 4 °C. The maximum recommended storage time is 6 months.
Procedure
Generation of type-1 polarized mouse DCs
Timing 4–5 h
-
1
Resuspend the CD11c+ cells (see Box 1 for CD11c+ cell isolation) in cell culture medium at 4 × 106 per ml and add 1 ml of cell suspension to each well of a 12-well culture plate. Add 1 ml of DC maturation medium to each well. Examine the DC cultures for the density and general appearance of the cells, using an inverted microscope. Immediately after addition of DC maturation medium, add 100 μl of 1 × 106 UVBγ-Tu cells per ml per well (40:1 CD11c+ cells (DCs)/tumor cell ratio).
Critical Step
Ensure that the same cancer cell line that will be injected into the experimental mice is used for DC loading (see Box 2 for preparation of UVBγTu cells). To generate control unloaded type-1 polarized DCs, do not add UVBγ-Tu cells to the maturing DCs.
Critical Step
Add UVBγ-Tu cells within 10 min after addition of DC maturation medium to ensure optimal take up of tumor antigens by maturing DCs.
-
2
Gently rock the plate back and forth to ensure even distribution of cells. Place the culture plates in the cell incubator at 37 °C and 5% (vol/vol) CO2 for 4 h. The target maturation time is 4 h before harvesting, but the maturation period can be extended to 4.5 h in the case that many plates must be harvested.
-
3
Transfer the plates to a refrigerator for 5–10 min. Depending on the number of plates, you may perform harvesting of the DCs in batches so that the plates are not kept at 4 °C for longer than 30 min, as this might affect DC function.
Critical Step
In contrast to the classic cytokine-based maturation cocktail (supplemented with PGE2), type-1 polarized mature DCs are more adherent to the plastic, displaying more elongated morphology. Incubation of the plates at 4 °C is important to reduce the adherence to the plastic and recover high numbers of viable DCs.
Collection of mature type-1 polarized mouse DCs
Timing 30–60 min
-
4
Detach DCs by vigorous pipetting and collect them in a 15-ml conical centrifuge tube. Refill each well with 1 ml of cold PBS and check for remaining adherent cells using a microscope. If some cells remained attached, vigorously pipette again and collect the cells in the 15-ml tube. Repeat until all cells are collected.
-
5
Centrifuge the cells at 400g for 5 min at 4 °C. Resuspend cell pellets in all tubes with 1-ml of cold PBS and combine the UVBγ-Tu-loaded DCs (or unloaded DCs) into one tube. Assess viability of the cells by Trypan blue exclusion. If the viability is below 50–70%, we recommend discarding the batch of DCs and repeating Steps 1–5.
Cryopreservation and thawing of type-1 polarized mature DCs
Timing 40–50 min
-
6
Keep the harvested type-1 polarized mature DCs at 4 °C while preparing for cryopreservation. The number of DCs frozen per cryovial depends on the number of planned vaccine doses and the number of mice receiving the vaccine. Prepare to freeze the DCs in at least as many cryovials as the number of planned vaccine doses plus an additional vial for QC. For example, if mice need to receive three cycles of vaccination and two vaccine doses per cycle, then freeze the DCs in at least 7 (2 × 3 + 1) cryovials. Freeze at least 1 × 106 DCs per cryovial.
-
7
Centrifuge the type-1 polarized mature DCs at 400g for 5 min at 4 °C and resuspend in FM1 (0.5 ml per cryovial). Place the cells on ice for 10 min and add the same volume of FM2. Place the cells in a temperature-controlled freezing container (e.g., Nalgene, Mr Frosty) and store at −80 °C overnight before their transfer to liquid nitrogen.
Pause point
DCs can be frozen for up to 1 year of cryopreservation, after which the viability will decrease.
-
8
To thaw the cells, place the vials in a water bath at 37 °C and transfer the cells to cold PBS (20 ml) immediately after thawing. Spin down the DCs (5 min, 400g, 4 °C), resuspend them in 20 ml of fresh PBS and centrifuge again (5 min, 400g, 4 °C). Resuspend the cells in 1 ml of culture medium and count the cells.
Quality control of the generated DCs
-
9
To check the quality of the DCs, follow option A for phenotype analysis of the DCs, option B for analysis of IL-12 production, option C for in vitro evaluation of effector type-1 immune cell induction, option D for in vivo evaluation of DC migration, and option E for in vivo evaluation of systemic tumor-specific immunity. Although the in vivo induction of tumor-specific type-1 immunity represents the most definitive test of DC function, the analysis of IL-12 production is a very strong predictor of their overall functional status.
-
A
Phenotype analysis of the DCs • TIMING 40–50 min for staining and ∼20 min for flow analysis
-
i
Adjust the cell density of the type-1 polarized mature DCs to 1 × 106 cell per ml in culture medium.
-
ii
Prepare five staining mixes by adding the following antibodies to 100 μl of blocking buffer: 1 μl of CD14-FITC, 5 μl of CCR7-PE, and 2.5 μl of CD80-APC (mix 1);
1 μl of IA/Eb-FITC and 2.5 μl of CD40-APC (mix 2);
1 μl of H2Kb-FITC and 1 μl of CD86-APC (mix 3);
1 μl of IgG1-FITC, 5 μl of IgG-PE, 2.5 μl of IgG2a-APC (mix 4); and
1 μl of IgG2a-FITC and 10 μl of IgG1-APC (mix 5).
-
iii
Add 200 μl of cell suspension (2.0 × 105 cells per well) to 5 wells of a V-bottom 96-well plate and centrifuge at 400g for 5 min at 4 °C.
-
iv
Discard the supernatant, add 200 μl of blocking buffer, and centrifuge at 400g for 5 min and at 4 °C.
-
v
Discard the supernatant and add 100 μl of each staining mix prepared in the previous step to one well with cells in a 96-well plate and mix. After 20–30 min incubation at 4 °C in the dark, wash the cells with 100–200 μl of blocking buffer and resuspend in 100 μl of 4% (wt/vol) PFA.
-
vi
Perform flow analysis following the instructions of the flow cytometer manufacturer (Fig. 2a).
-
i
-
B
Analysis of IL-12 production • TIMING 20–30 min for culture set-up and ∼6 h for ELISA
-
i
Start cultures of J558-CD40L cells in cell culture medium at 2 × 105 cells per ml and maintain between 1 × 105 and 1 × 106 cells per ml for at least 3–5 d after thawing before their use, to ensure their high viability and potency.
-
ii
Adjust the cell density of the type-1 polarized mature DCs to 1 × 106 cell per ml in culture medium.
-
iii
Add 100 μl of type-1 polarized mature DCs to 6 wells in a sterile 96-well flat-bottom plate.
-
iv
Prepare 5 × 105 cells per ml of J558-CD40L cells in culture medium and add 100 μl of cell suspension to the 3 wells with DCs. Add 100 μl of culture medium to the remaining 3 wells for controls. Place the culture plate in an incubator (37 °C, 5% (vol/vol) CO2) and collect the media from the cells after 24 h. Alternatively, soluble CD40L with enhancer (MEGACD40L, 1 μg/ml; Enzo Life Sciences, cat. no. ALX-522–120-C010) (ref. 151) can be used instead of J558-CD40L cells.
Pause point
Supernatants may be stored at −20 °C until analysis. The maximum recommended storage time is 1 month.
-
v
Perform IL-12 ELISA following the manufacturer's instructions (Fig. 2b).
-
i
-
C
In vitro evaluation of effector type-1 immune cell induction • TIMING 1–2 h for setting up the IVS culture, ∼2 h for setting up the ELISpot plate, and ∼5 h for developing the ELISpot plate
-
i
Adjust the cell density of the type-1 polarized mature DCs to 1 × 106 cell per ml in culture medium.
-
ii
Add 0.25 ml of tumor-loaded DCs in cell culture medium to each well of a 48-well plate (4.0 × 105 cells per well).
-
iii
Prepare the splenocyte suspension (Box 3) and resuspend the splenocytes in cell culture medium at a density of 4.0 × 106 cells per ml. Add 0.25 ml of splenocytes to DC cultures to induce in vitro stimulation (IVS) and incubate at 37° C and 5% (vol/vol) CO2 for 7–10 d.
-
iv
On day 4 of culture, add IL-2 (25 U/ml) and IL-7 (0.5 ng/ml) to the DC–splenocyte co-cultures.
-
v
Prepare the ELISpot plate. To do this, add 100 μl of 15 μg/ml AN18 anti-IFNγ-coating antibody to each well of an ELISpot plate in PBS, wrap the plate with Parafilm, and incubate overnight at 4° C.
-
vi
Wash the wells by adding 200 μl of RPMI-1640 and wait 3–5 min before removing the media. Repeat the wash three times.
-
vii
Block the wells by adding 200 μl of culture medium for 30 min at 37 °C.
Critical Step
The IFNγ ELISpot plate must be pre-coated with IFNγ capture antibody at least 1 d before harvesting the splenocytes in the next step to ensure maximal efficiency. Alternatively, the plate may be coated for 2 h at 37° C; however, such incubation is less efficient.
Pause point
The maximum recommended storage time for an anti-IFN-γ antibody precoated and blocked ELISpot plate is 72 h at 4° C.
-
viii
Harvest the DC-stimulated splenocyte co-cultures on days 7–10 in a 15-ml conical centrifuge tube. Centrifuge the cells at 400g at room temperature for 3–5 min, discard the supernatant, and resuspend the cells in 5 ml of PBS. Centrifuge the cells at 400g and room temperature for 3 min, then resuspend the cells in 0.5 ml of cell culture medium and count the cells using a hemocytometer.
-
ix
Discard the culture medium in the anti-IFN-γ antibody precoated and blocked plate (from Step 9C(vii)) and plate 100 μl of 1 × 105 splenocytes in six wells of the plate.
-
x
Harvest the tumor cells (use the tumor cell line that was used for loading of DCs) in culture with 1 mM EDTA in PBS as described in step 1 of Box 2 and centrifuge at 400g and room temperature for 5 min.
-
xi
Resuspend the tumor cells at 1 × 106 cells per ml in culture medium and irradiate with 20 Gy. Centrifuge at 400g and room temperature for 5 min and resuspend the irradiated tumor cells at 4 × 105 cells per ml.
-
xii
Add 100 μl of irradiated tumor cells (4 × 104 cells) to 3 wells with DC-stimulated splenocytes. Add 100 μl of culture medium to 3 wells of control DC splenocytes. Incubate the cultures at 37° C, 5% (vol/vol) CO2 for 24–48 h.
-
xiii
Develop the ELISpot plate. To do this, first wash the plate five times by soaking 3–5 min in 200–300 μl of wash buffer (PBS/0.05% (vol/vol) Tween 20). After the washes, add 100 μl of 1 μg/ml R4-6A2 biotinylated secondary antibody in PBS/0.5% (wt/vol) BSA and incubate at room temperature for 2 h. 10 min before the incubation is finished, prepare ABC reagent according to the manufacturer's instructions.
-
xiv
Mix the ABC reagent and let it stand at room temperature for 30 min.
-
xv
Wash the plate five times as in Step 9C(xiii), add 100 μl of ABC reagent, and incubate the plate at room temperature for 1 h.
-
xvi
Wash the plate three times by soaking 3–5 min in wash buffer, followed by an additional two washes in PBS. While washing the plate, prepare AEC substrate according to the manufacturer's instructions.
-
xvii
Add 100 μl of AEC reagent per well. Spots appear after 5–20 min, depending on the assay. Once spots are clearly visible, but before background appears, rinse the plate thoroughly in dH2O. Remove the plastic lid from the plate and allow the plate to dry. Read the plate using an ELISpot reader (Fig. 2c).
-
i
-
D
In vivo evaluation of DC migration • TIMING 10–20 min for DC injection and ∼15 min for imaging per five mice
-
i
Adjust the cell density of UVBγ-Tu-loaded DCs to 1 × 106 cell per ml in PBS.
-
ii
Inject 50 μl of DCs (5 × 105) in PBS with a 1-cc syringe and a 27-gauge or 30-gauge needle into the footpad of the C57BL/6J mouse.
-
iii
Image mice by IVIS after 1, 12, 24, and 48 h (Fig. 2d). To do this, inject luciferin (100 μl of 30 mg/ml i.p. with a 1-cc syringe and a 27-gauge or 30-gauge needle) before imaging. Follow the instructions outlined in Step 12 for animal handling and care to image the mice.
Critical Step
To conduct this analysis, generate the DCs from firefly luciferase (CAG-luc-eGFP) transgenic mice (FVB-Tg(CAG-luc,-GFP)L2G85Chco/J; The Jackson Laboratory, cat. no. 008450) (ref. 150). If you generate the DCs from B6 Cd45.1 mice, or from C57BL/6J mice, you must perform evaluation of DC migration by flow cytometric analysis instead of imaging (Box 4).
-
i
-
E
In vivo evaluation of effector type-1 immune cell induction • TIMING 8–10 h for preparation and injection of OVA-loaded DCs, 3–4 h for preparation of target cells, and 3–4 h for flow analysis of harvested splenocytes
-
i
Resuspend the unloaded DC cells (from Step 8) in culture medium at 1 × 107 per ml and add OVA peptide to cells at 1.0 μg per 106 cells. Incubate in a 50-ml conical centrifugation tube at 37 °C for 1.5–2.0 h. Fill the tube with PBS to 35–40 ml and centrifuge at 400g for 5 min at room temperature.
-
ii
Resuspend the cells in PBS at 3 × 106 cells per ml in PBS and inject 3–4 × 105 OVA-loaded DCs/0.1 ml of PBS s.c. with a 1-cc syringe and a 27-gauge or 30-gauge needle into the flank of naive mice.
-
iii
Antigen loading of target cells. 3–5 d after vaccination with OVA-loaded DCs, isolate splenocytes from a B6 Cd45.1 mouse as described in Box 3. Resuspend splenocytes at 2 × 107 cells per ml in culture medium and divide into two 15-ml conical centrifugation tubes. Add OVA peptide to splenocytes in one of the tubes to load at 0.5 μg per 106 cells. Add nothing or a control peptide to the splenocytes in the other tube. Incubate both tubes at 37 °C for 1.5–2.0 h. Fill the tubes with PBS to 12–15 ml and centrifuge at 400g for 5 min at room temperature.
-
iv
Label loaded splenocytes with CFSE and unloaded splenocytes with 20× diluted CFSE as described in Box 5. Mix unloaded CFSElow- and loaded CFSEhigh-labeled splenocytes at a 1:1 ratio in PBS to 2.5 × 107 cells per ml and inject 200 μl i.v. into DC-vaccinated mice (3–5 d after vaccination with OVA-loaded DCs) and into at least two naive (unvaccinated) mice.
Critical Step
For the i.v. injections, keep the mouse warm under a heat lamp (or other heating device), making sure not to overheat the animal. After placing the animal in a restraint device, hold the tail between the thumb and forefinger and clean it with 70% (vol/vol) ethanol. Keep the tail under pressure and place the needle with the bevel facing upward at the middle or slightly distal part of the tail. Insert the needle at least 3 mm into the tail, while keeping the needle parallel to the vein. Inject the material in a slow, smooth motion, without aspirating. If swelling occurs or if there is resistance to the injection, remove the needle from the tail and repeat the procedure slightly above the initial site of injection.
Critical Step
The number of injected cells is not critical, although more cells are better, as the cells will be detected by flow cytometry. We aim for 5 × 106 cells per mouse.
-
v
Harvest spleens 16 h afterward and isolate splenocytes as described in Box 3.
-
vi
Flow analysis. Determine the specific killing by the comparative analysis of live CFSElow and CFSEhigh cells as presented in Figure 2e.
-
vii
(Optional) Stain the cells for CD45.1 as described in step 9 of Box 4. Make a histogram of live CD45.1+ cells and gate on CFSElow and CFSEhigh cells.
-
i
-
A
Inoculation of mice with tumor cells
Timing 5–10 min per five mice
-
10
Culture the relevant tumor cells for at least two passages after thawing before harvesting. Harvest the tumor cells as described in step 1 of Box 2. Wash the cells twice with 10–20 ml of PBS and resuspend in PBS for subsequent injection (5 × 106 per ml for MC38Luc or 20 × 106 per ml for ID8A cells).
-
11
Inject the tumor cells. Follow option A to perform i.p. injection (disseminated peritoneal tumor model) or option B for s.c. injection (localized/solid tumor).
-
A
Intraperitoneal injection
-
i
Use a 1-cc syringe and a 27-gauge needle to inject 1 × 106 MC38Luc or 4 × 106 ID8A cells in 200 μl of PBS i.p. into 6–8-week-old B6 mice for colorectal or ovarian cancer models, respectively.
Critical Step
To make the injection into lower left abdominal quadrant (the preferred injection site for i.p. injections), manually restrain the mice with dorsal recumbence (abdomen side up) and the cranial end of the animal pointed down. Enter the abdominal cavity with the needle bevel-side up and slightly angled (15–20 degrees). After penetrating the abdominal wall (∼4–5 mm), attempt to aspirate (pulling back slightly on the needle) to ensure that abdominal organs, such as the bladder or colon, have not been penetrated.
-
i
-
B
Subcutaneous injection
-
i
Inject 0.5 × 106 MC38Luc cells in 100 μl PBS s.c. into the lower right flank.
Critical Step
For s.c. injections, shave the right flank of the mice and then manually restrain the mice with the right side pointed up. When penetrating the skin, hold the needle bevel-side up and angled 15–25 degrees. Advance the needle ∼2–3 mm below the skin surface so that the tip is visible under the skin. When injecting, a bleb should develop, otherwise withdraw the needle slightly.
-
i
-
A
Monitoring of tumor progression
Timing ∼15 min per five mice
-
12
Inject mice i.p. with luciferin (100 μl of 30 mg/ml using a 1-cc syringe and a 27-gauge or 30-gauge needle) before imaging. Wait 8 min after the luciferin injection before transferring the mice to a 5% (vol/vol) isoflurane chamber before imaging (firefly luciferase signal peaks after 10 min and then declines).
Critical Step
Optimal in vivo imaging time should be determined by taking images over incremental periods of time. We image mice every 5–7 d.
-
13
Use the control panel to manually set the imaging parameters or use the software's Imaging Wizard (under 'sequence setup') to automatically set the imaging parameters. Adjust the field of view depending on number of mice imaged. The field of view size affects the sensitivity. Note that longer exposure time, higher binning, and a lower F/Stop (a wider aperture allows more light to reach the charge-coupled device) increase sensitivity. Images should take no longer than 5 min of exposure time.
Critical Step
Luciferase signal can decrease at later time points (owing to ascites accumulation in the ID8A model or because of ulceration in the MC38 s.c. model), therefore, tumor progression should be concomitantly monitored by increase in mouse weight or circumference (for i.p. tumors) or by assessing tumor size with a caliper (for MC38 s.c. tumors).
Induction of tumor-infiltrating specific type-1 immune cells
Timing 20–30 min for vaccine preparation and ∼10 min for injection into five mice
Critical Step
We recommend imaging and weighing mice 1 d before therapy initiation and grouping mice randomly into untreated and treated sets.
-
14
Thaw DCs as described in Step 8. Refer to Table 1 to determine the number of vials needed. Dilute the cells to 3 × 106 cells per ml in PBS and inject 3–4 × 105 DCs per 0.1 ml of PBS s.c. into the left flank of tumor-bearing mice (as in Step 10B(i)), on days 3–7 after injecting tumor cells as a repeat injection two to four times, with a 1-week interval between treatments.
Critical Step
If a s.c. tumor model is investigated, DCs should be injected s.c. into the opposite flank.
-
15
Perform TME modulation 3 d after each s.c. DC vaccination. Follow option A to modulate the TME by i.p. injection of the type-1 polarized mature DCs or option B for CKM adjuvant-mediated modulation.
-
A
DC injection
-
i
Inject 3–4 × 105 type-1 polarized mature DCs in 0.2 ml of PBS i.p. (for the i.p. model) or 3–4 × 105 type-1 polarized mature DCs in 0.05 ml intratumorally (for the s.c. model).
-
i
-
B
CKM adjuvant injection
-
i
Inject 1 × 104 IU of IFN-α and 50 μg Ampligen (Fig. 3b) in 0.2 ml of PBS i.p. (for the i.p. model) or 1 × 104 IU of IFN-α and 50 μg Ampligen in 0.05 ml intratumorally (for the s.c. model).
-
i
-
A
-
16
Inject 0.2 ml of 500 μg/ml celecoxib (in 12.5% (vol/vol) DMSO) or 0.2 ml of 2.5 mg/ml (1.25% (vol/vol) DMSO) aspirin i.p. every 2–3 d starting as early as day 4 after tumor inoculation.
Ex vivo immunomonitoring
Timing 8–10 h for setting up an ELISpot plate and ∼6 h for developing the ELISpot plate
Critical Step
Initiate Step 17 before the mice from Step 18 approach the relevant time point.
-
17
Add 100 μl of 15 μg/ml AN18 anti-IFNγ coating antibody in PBS to each well of an ELISpot plate and wrap the plate with Parafilm to precoat the plate. Incubate overnight at 4 °C. Alternatively, the plate may be coated for 2 h at 37° C; however, such incubation is less efficient.
-
18
Monitor animals in accordance with institutional guidelines. Kill mice by CO2 inhalation, followed by cervical dislocation when one of the following end points is met: weight exceeds 140% of initial body weight, mice appear to be in distress (anorexic, emaciated, lethargic), changes in behavior occur (immobile and/or non-responsive), tumor size exceeds 2 cm in diameter, or ulceration occurs (for s.c. tumors).
Critical Step
Do not allow mice to die due to high tumor burden. Typically, the untreated mice must be euthanized before day 30 (MC38 model) or day 40 (ID8A model).
-
19
Isolate the TILs/TALs from the tumors (in the case of the s.c. tumor model; option A) or peritoneal washes/cancer ascites (in the case of the i.p. tumor model; option B).
-
A
Harvesting TILs from tumors
-
i
Make a cut to enter the peritoneal cavity and collect the tumor tissue into 15-ml conical tubes filled with 2-ml of PBS.
-
ii
Add 2 ml of PBS to a Petri dish to cover the bottom. Transfer the harvested tumor tissue and mince it with two scalpels to <0.5-mm pieces. Transfer the tumor pieces to a new 15-ml conical centrifugation tube and add enough digestion buffer to cover the tumor pieces.
Critical Step
If the tumor pieces are very small, transfer them directly to a 2-ml Eppendorf centrifugation tube, add 500 μl of digestion buffer, and then mash with the rubber piston of the syringe.
-
iii
Incubate tumors at 37 °C for 20–45 min on a rotator. After the tissue digestion, transfer the tumors to a pre-wetted 100-μm strainer placed on top of a 50-ml conical centrifuge tube and mash with the rubber piston of the syringe and continuously wash with PBS to a final volume of 10–20 ml.
-
iv
Centrifuge the cells at 400g for 3–5 min at room temperature, discard the supernatant, and resuspend the cells in 1 ml ACK lysis buffer to remove the red blood cells. Add an additional 3 ml of ACK lysis buffer on top and rotate the conical centrifuge tube to mix. Incubate the cells at room temperature for 4 min.
-
v
Add 30 ml of PBS and centrifuge the cells at 400g for 3 min at room temperature, then resuspend them in 20 ml of PBS and transfer the mixture to a 70-μm strainer.
-
vi
Count the cells using a hemocytometer.
-
vii
Centrifuge the cells at 400g for 3–5 min at room temperature, resuspend the cells in 3 ml of 60% (vol/vol) SIP, and transfer them to 15-ml conical tube.
-
i
-
B
Harvesting TALs from peritoneal washes/cancer ascites
-
i
Use a 10-ml syringe and an 18-gauge needle to obtain the peritoneal washes/cancer ascites. Fill the syringe with 3–5 ml of PBS. After penetrating the abdominal wall (∼4–5 mm) into the lower left abdominal quadrant, inject PBS into the i.p. cavity and then aspirate as much peritoneal fluid as possible. Ensure that the abdominal organs, such as the bladder or colon, have not been penetrated. In such case, try to readjust the needle by pulling it backward for a few millimeters and changing the angle. Blood or fecal material may indicate that the intestine has been ruptured. If fecal material or blood is aspirated, discard the collected sample. Transfer the peritoneal fluid to a 50-ml conical tube.
-
ii
Centrifuge the cells at 400g for 3–5 min at room temperature, discard the supernatant, and resuspend the cells in 1-ml of ACK lysis buffer to remove the red blood cells. Add an additional 3 ml of ACK lysis buffer on top and rotate the conical centrifuge tube to mix. Incubate the cells at room temperature for 4 min.
-
iii
Add 30 ml of PBS and centrifuge the cells at 400g for 3 min at room temperature, then resuspend them in 20-ml of PBS, and transfer the mixture to a 70-μm strainer.
-
iv
Count the cells using a hemocytometer.
-
v
Centrifuge the cells at 400g for 3 min at room temperature, resuspend the cells in 3 ml of 60% (vol/vol) SIP, and transfer them to a 15-ml conical tube.
-
vi
Layer 3 ml of 45% (vol/vol) SIP slowly on top of the 60% (vol/vol) SIP layer by expelling the 45% (vol/vol) SIP at a shallow angle down the side of the centrifuge tube. Take care not to mix with the bottom layer. Repeat with a similar technique to layer 3 ml of 34% (vol/vol) SIP on top of the 45% (vol/vol) SIP layer.
-
vii
Centrifuge the tubes at 2,400g for 30 min at room temperature without applying the brake.
-
viii
Collect the bottom interface (Fig. 4a) (∼2–3 ml) into a new 15-ml conical centrifugation tube, fill the tube to 15 ml with PBS, and centrifuge at 600g for 10 min at room temperature. Resuspend the cell pellet in 0.5 ml of the cell culture medium.
Critical Step
High centrifugation speed and dilution of transferred TILs/TALs with PBS is necessary to allow effective centrifugation.
-
i
-
A
-
20
Adjust the cell concentration to 1 × 106 per ml with cell culture medium. If the cell number is <1 × 106 per ml, resuspend the samples in 0.7 ml of the cell culture medium.
Critical Step
When analyzing the ELISpot data, normalize the number of spots to the number of cells loaded into the wells.
-
21
Isolate splenocytes as described in Box 3 4–5 d after TME modulation (Step 15) and resuspend to 5 × 106 per ml in culture medium.
-
22
Wash the ELISpot plate from Step 17 three times by adding 200 μl of RPMI-1640 and waiting 3–5 min before removing media. Block the plate by adding 200 μl of culture medium for 30 min at 37° C.
-
23
Discard the culture medium from the plate and dispense 100 μl (5 × 105) splenocytes and 1 × 105 TILs/TALs into six wells of the ELISpot plate. Harvest the tumor cells in culture with 1 mM EDTA in PBS and centrifuge at room temperature as described in step 1 of Box 2. Resuspend the tumor cells at 8 × 105 cells per ml of culture medium and irradiate with 20 Gy. Add 100 μl of (4 × 104) tumor cells to three wells with splenocytes and TIL/TALs (Fig. 4b). Add 100 μl of culture medium to make three control wells of splenocytes and TIL/TALs (Fig. 4b). Incubate the cultures at 37° C, 5% (vol/vol) CO2 for 24–48 h.
-
24
Develop the ELISpot plate as in Step 9C(xiii).
Troubleshooting
Troubleshooting advice can be found in Table 3.
Timing
This protocol covers a minimum of 30 d. A workflow schema is provided in Figure 1
Generation of tumor-loaded DCs
Box 1, Flt3L-B16 cell injection into donor mice, day −11 ± 1: 1 h
Box 2, preparation of UVBγ-Tu cells, day −1: 1.5 h
Box 3, preparation of the splenocyte suspension, day 0: 1 h
Box 4, extraction of cells from the lymph nodes of mice, days 1–3: 2 h
Box 5, labeling of the cells with CFSE (several time points): 30–45 min
Box 6, flow analysis of the spleens for in vivo specific cytotoxicity assay, days 13–15: 2–3 h
Steps 1–8, generation, collection, cryopreservation, and thawing of type-1 polarized DCs from donor mice, day 0: 10 h
QC of DCs
Step 9, phenotype analysis, in vitro evaluation of type-1 immune cell induction, and J558-CD40L restimulation of DCs from donor mice, day 1: 6 h
Step 9A, phenotype analysis of the DCs, day 2: 40–50 min for staining and ∼20 min for flow analysis
Step 9B, ELISA to evaluate the capacity of DCs from donor mice to produce IL-12, day 2: 20–30 min for culture set-up and ∼6 h for ELISA
Step 9C, in vitro evaluation of effector type-1 immune cell induction, day 2: 1–2 h for setup of the IVS culture, ∼2 h for setup of the ELISpot plate, and ∼5 h for developing the ELISpot plate
Step 9D, in vivo evaluation of DC migration, day 2: 10–20 min for DC injection and ∼15 min for imaging per five mice
Step 9E, in vivo evaluation of effector type-1 immune cell induction, day 2: 8–10 h for preparation and injection of OVA-loaded DCs, 3–4 h for preparation of target cells, and 3–4 h for flow analysis of harvested splenocytes
Steps 10 and 11, inoculation of experimental mice with tumor cells, day −4: 5–10 min per five mice
Steps 12 and 13, monitoring of tumor progression (BLI) of experimental mice, weekly from day −1: ∼15 min per five mice
Steps 14–16, induction of tumor-infiltrating-specific type-1 immune responses in experimental mice with type-1 polarized DCs (and CKM), weekly from days 0 and 3 : 20–30 min for vaccine preparation and ∼10 min for injection into five mice
Monitoring of the therapeutic and immunomodulatory effects
Step 17, preparation of ELISpot plate to detect intratumoral type-1 immune cells from experimental mice, day 12: 10 min
Steps 18–23, isolation of splenocytes and TILs/TALs from experimental mice and setup of the ELISpot assay, days 13–15: 8–10 h
Step 24, developing of ELISpot plate to detect intratumoral type-1 immune cells from experimental mice, days 16/17: 6 h
Anticipated results
Typical results of immunotherapy-induced changes (following DC therapy and combinatorial approaches with TME modulation) in animal survival are shown in Figure 3c,d. It may be expected that the s.c. injections of tumor-loaded type-1 polarized DCs (3 × 105 cells on days 5 and 12), used as a single treatment, will not significantly prolong the survival of MC38 intraperitoneal tumor–bearing mice. The data from this therapeutic model are in contrast to prevention models14 and the therapy of genetically manipulated tumors with DCs loaded with strong antigens81,152. However, additional intraperitoneal administration of DCs (on days 8 and 15) significantly prolongs survival of the mice (median survival = 29 d versus 23 d; P = 0.0021; n = 10 mice per group) (Fig. 3c). Furthermore, we evaluated the effect of substituting additional intraperitoneal administration of DCs with intraperitoneal injection of CKM (IFN-α and poly-IC) and COX-2 inhibitor (celecoxib or aspirin) on prolongation of mice survival (Steps 15 and 16). Typically, mice treated with s.c. injection of DCs only- (Fig. 3d) or a CKM regimen alone (Fig. 3e) do not demonstrate an advantage in survival. However, the combinatorial approach shows significant prolongation of mouse survival (median survival = 45 d (celecoxib); P < 0.0001 and 42 (aspirin); P = 0.0014 versus 35 d for control; n = 10 mice per group)) (Fig. 3d).
The current protocol is designed to enhance the infiltration of tumor-specific CTLs into the TME (Fig. 4). High tumor production of CCL5/RANTES (ligand for CCR5) and CXCL9/MIG, CXCL10/IP10, and CXCL11/ITAC (three known ligands for CXCR3) is associated with high CTL infiltration in cancers153,154,155. Our own studies show strong correlations between the intratumoral production of CCL5, CXCL9, and CXCL10 and local infiltration of CD8+ granzyme B+ CTLs16. To achieve the antitumor effectiveness of immune responses in the cancer microenvironment, TME modulation is critical and can be induced by administration of a CKM regimen, e.g., IL-12-producing DCs; combination of IFN-α, poly-IC and COX2 blockers; and chemokine-expressing Vaccinia viruses. However, additional combinatorial approaches are likely to be effective for selective attraction of specific subsets of immune cells. In this regard, the currently presented protocol can be modified to include additional elements and readouts, for example, ELISpot assays that detect T cells that produce alternative cytokines, and thus are able to detect T cells with different functions.
References
Johnson, L.A. & June, C.H. Driving gene-engineered T cell immunotherapy of cancer. Cell Res. 27, 38–58 (2017).
Kalos, M. & June, C.H. Adoptive T cell transfer for cancer immunotherapy in the era of synthetic biology. Immunity 39, 49–60 (2013).
Maus, M.V. et al. Adoptive immunotherapy for cancer or viruses. Annu. Rev. Immunol. 32, 189–225 (2014).
Palucka, K. & Banchereau, J. Cancer immunotherapy via dendritic cells. Nat. Rev. Cancer 12, 265–277 (2012).
Topalian, S.L. et al. Immunotherapy: the path to win the war on cancer? Cell 161, 185–186 (2015).
Tumeh, P.C. et al. PD-1 blockade induces responses by inhibiting adaptive immune resistance. Nature 515, 568–571 (2014).
Herbst, R.S. et al. Predictive correlates of response to the anti-PD-L1 antibody MPDL3280A in cancer patients. Nature 515, 563–567 (2014).
Brahmer, J.R. et al. Safety and activity of anti-PD-L1 antibody in patients with advanced cancer. N. Engl. J. Med. 366, 2455–2465 (2012).
Taube, J.M. et al. Association of PD-1, PD-1 ligands, and other features of the tumor immune microenvironment with response to anti-PD-1 therapy. Clin. Cancer Res. 20, 5064–5074 (2014).
Topalian, S.L. et al. Safety, activity, and immune correlates of anti-PD-1 antibody in cancer. N. Engl. J. Med. 366, 2443–2454 (2012).
Ji, R.R. et al. An immune-active tumor microenvironment favors clinical response to ipilimumab. Cancer Immunol. Immunother. 61, 1019–1031 (2012).
Gajewski, T.F., Schreiber, H. & Fu, Y.X. Innate and adaptive immune cells in the tumor microenvironment. Nat. Immunol. 14, 1014–1022 (2013).
Mailliard, R.B. et al. Alpha-type-1 polarized dendritic cells: a novel immunization tool with optimized CTL-inducing activity. Cancer Res. 64, 5934–5937 (2004).
Giermasz, A.S. et al. Type-1 polarized dendritic cells primed for high IL-12 production show enhanced activity as cancer vaccines. Cancer Immunol. Immunother. 58, 1329–1336 (2009).
Lee, J.J., Foon, K.A., Mailliard, R.B., Muthuswamy, R. & Kalinski, P. Type 1-polarized dendritic cells loaded with autologous tumor are a potent immunogen against chronic lymphocytic leukemia. J. Leukoc. Biol. 84, 319–325 (2008).
Muthuswamy, R. et al. NF-kappaB hyperactivation in tumor tissues allows tumor-selective reprogramming of the chemokine microenvironment to enhance the recruitment of cytolytic T effector cells. Cancer Res. 72, 3735–3743 (2012).
Muthuswamy, R., Corman, J.M., Dahl, K., Chatta, G.S. & Kalinski, P. Functional reprogramming of human prostate cancer to promote local attraction of effector CD8(+) T cells. Prostate 76, 1095–1105 (2016).
Muthuswamy, R., Wang, L., Pitteroff, J., Gingrich, J.R. & Kalinski, P. Combination of IFNα and poly-I:C reprograms bladder cancer microenvironment for enhanced CTL attraction. J. Immunother. Cancer 3, 6 (2015).
Downs-Canner, S. et al. Complement inhibition: a novel form of immunotherapy for colon cancer. Ann. Surg. Oncol. 23, 655–662 (2016).
Bach, S. et al. A yeast-based assay to isolate drugs active against mammalian prions. Methods 39, 72–77 (2006).
Galon, J., Fridman, W.H. & Pages, F. The adaptive immunologic microenvironment in colorectal cancer: a novel perspective. Cancer Res. 67, 1883–1886 (2007).
Zhang, L. et al. Intratumoral T cells, recurrence, and survival in epithelial ovarian cancer. N. Engl. J. Med. 348, 203–213 (2003).
Sato, E. et al. Intraepithelial CD8+ tumor-infiltrating lymphocytes and a high CD8+/regulatory T cell ratio are associated with favorable prognosis in ovarian cancer. Proc. Natl. Acad. Sci. USA 102, 18538–18543 (2005).
Fridman, W.H. et al. Prognostic and predictive impact of intra- and peritumoral immune infiltrates. Cancer Res. 71, 5601–5605 (2011).
Fridman, W.H., Pages, F., Sautes-Fridman, C. & Galon, J. The immune contexture in human tumours: impact on clinical outcome. Nat. Rev. Cancer 12, 298–306 (2012).
Pages, F. et al. Effector memory T cells, early metastasis, and survival in colorectal cancer. N. Engl. J. Med. 353, 2654–2666 (2005).
Llosa, N.J. et al. The vigorous immune microenvironment of microsatellite instable colon cancer is balanced by multiple counter-inhibitory checkpoints. Cancer Discov. 5, 43–51 (2015).
Le, D.T. et al. PD-1 blockade in tumors with mismatch-repair deficiency. N. Engl. J. Med. 372, 2509–2520 (2015).
Astsaturov, I. et al. Amplification of virus-induced antimelanoma T-cell reactivity by high-dose interferon-alpha2b: implications for cancer vaccines. Clin. Cancer Res. 9, 4347–4355 (2003).
Rosenberg, S.A. et al. Tumor progression can occur despite the induction of very high levels of self/tumor antigen-specific CD8+ T cells in patients with melanoma. J. Immunol. 175, 6169–6176 (2005).
Fox, B.A. et al. Defining the critical hurdles in cancer immunotherapy. J. Transl. Med. 9, 214 (2012).
Apetoh, L. et al. Consensus nomenclature for CD8+ T cell phenotypes in cancer. Oncoimmunology 4, e998538 (2015).
Kalinski, P., Muthuswamy, R. & Urban, J. Dendritic cells in cancer immunotherapy: vaccines and combination immunotherapies. Expert Rev. Vaccines 12, 285–295 (2013).
Gajewski, T.F. The next hurdle in cancer immunotherapy: overcoming the non-T-cell-inflamed tumor microenvironment. Semin. Oncol. 42, 663–671 (2015).
Tang, H. et al. Facilitating T cell infiltration in tumor microenvironment overcomes resistance to PD-L1 blockade. Cancer Cell 29, 285–296 (2016).
Fuertes, M.B., Woo, S.R., Burnett, B., Fu, Y.X. & Gajewski, T.F. Type I interferon response and innate immune sensing of cancer. Trends Immunol. 34, 67–73 (2013).
Woo, S.R. et al. STING-dependent cytosolic DNA sensing mediates innate immune recognition of immunogenic tumors. Immunity 41, 830–842 (2014).
Ohkuri, T. et al. STING contributes to antiglioma immunity via triggering type I IFN signals in the tumor microenvironment. Cancer Immunol. Res. 2, 1199–1208 (2014).
Kreutz, M., Tacken, P.J. & Figdor, C.G. Targeting dendritic cells—why bother? Blood 121, 2836–2844 (2013).
Yang, X. et al. Targeting the tumor microenvironment with interferon-β bridges innate and adaptive immune responses. Cancer Cell 25, 37–48 (2014).
Wong, J.L., Muthuswamy, R., Bartlett, D.L. & Kalinski, P. IL-18-based combinatorial adjuvants promote the intranodal production of CCL19 by NK cells and dendritic cells of cancer patients. Oncoimmunology 2, e26245 (2013).
Wong, J.L., Berk, E., Edwards, R.P. & Kalinski, P. IL-18-primed helper NK cells collaborate with dendritic cells to promote recruitment of effector CD8+ T cells to the tumor microenvironment. Cancer Res. 73, 4653–4662 (2013).
Kalinski, P. & Gingrich, J.R. Toward improved effectiveness of bladder cancer immunotherapy. Immunotherapy 7, 1039–1042 (2015).
Lu, H. TLR agonists for cancer immunotherapy: tipping the balance between the immune stimulatory and inhibitory effects. Front. Immunol. 5, 83 (2014).
Liu, Z. et al. CXCL11-armed oncolytic poxvirus elicits potent antitumor immunity and shows enhanced therapeutic efficacy. Oncoimmunology 5, e1091554 (2016).
Francis, L. et al. Modulation of chemokines in the tumor microenvironment enhances oncolytic virotherapy for colorectal cancer. Oncotarget 7, 22174–22185 (2016).
Kantoff, P.W. et al. Sipuleucel-T immunotherapy for castration-resistant prostate cancer. N. Engl. J. Med. 363, 411–422 (2010).
Fong, L. et al. Activated lymphocyte recruitment into the tumor microenvironment following preoperative sipuleucel-T for localized prostate cancer. J. Natl. Cancer Inst. 106 http://dx.doi.org/10.1093/jnci/dju268 (2014).
Inaba, K., Young, J.W. & Steinman, R.M. Direct activation of CD8+ cytotoxic T lymphocytes by dendritic cells. J. Exp. Med. 166, 182–194 (1987).
Caux, C., Dezutter-Dambuyant, C., Schmitt, D. & Banchereau, J. GM-CSF and TNF-alpha cooperate in the generation of dendritic Langerhans cells. Nature 360, 258–261 (1992).
Inaba, K. et al. Identification of proliferating dendritic cell precursors in mouse blood. J. Exp. Med. 175, 1157–1167 (1992).
Bender, A., Sapp, M., Schuler, G., Steinman, R.M. & Bhardwaj, N. Improved methods for the generation of dendritic cells from nonproliferating progenitors in human blood. J. Immunol. Methods 196, 121–135 (1996).
Schuler, G. & Steinman, R.M. Dendritic cells as adjuvants for immune-mediated resistance to tumors. J. Exp. Med. 186, 1183–1187 (1997).
Sallusto, F. & Lanzavecchia, A. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and downregulated by tumor necrosis factor alpha. J. Exp. Med. 179, 1109–1118 (1994).
Peters, J.H. et al. Signals required for differentiating dendritic cells from human monocytes in vitro. Adv. Exp. Med. Biol. 329, 275–280 (1993).
Brasel, K., De Smedt, T., Smith, J.L. & Maliszewski, C.R. Generation of murine dendritic cells from flt3-ligand-supplemented bone marrow cultures. Blood 96, 3029–3039 (2000).
Saunders, D. et al. Dendritic cell development in culture from thymic precursor cells in the absence of granulocyte/macrophage colony-stimulating factor. J. Exp. Med. 184, 2185–2196 (1996).
Maraskovsky, E. et al. Dramatic increase in the numbers of functionally mature dendritic cells in Flt3 ligand-treated mice: multiple dendritic cell subpopulations identified. J. Exp. Med. 184, 1953–1962 (1996).
Brasel, K. et al. Hematologic effects of flt3 ligand in vivo in mice. Blood 88, 2004–2012 (1996).
Albert, M.L., Jegathesan, M. & Darnell, R.B. Dendritic cell maturation is required for the cross-tolerization of CD8+ T cells. Nat. Immunol. 2, 1010–1017 (2001).
Sparwasser, T. et al. Bacterial DNA and immunostimulatory CpG oligonucleotides trigger maturation and activation of murine dendritic cells. Eur. J. Immunol. 28, 2045–2054 (1998).
Binder, R.J., Anderson, K.M., Basu, S. & Srivastava, P.K. Cutting edge: heat shock protein gp96 induces maturation and migration of CD11c+ cells in vivo. J. Immunol. 165, 6029–6035 (2000).
Kalinski, P. et al. Natural killer-dendritic cell cross-talk in cancer immunotherapy. Exp. Opin. Biol. Ther. 5, 1303–1315 (2005).
Jonuleit, H. et al. Pro-inflammatory cytokines and prostaglandins induce maturation of potent immunostimulatory dendritic cells under fetal calf serum-free conditions. Eur. J. Immunol. 27, 3135–3142 (1997).
Anguille, S., Smits, E.L., Lion, E., van Tendeloo, V.F. & Berneman, Z.N. Clinical use of dendritic cells for cancer therapy. Lancet Oncol. 15, e257–e267 (2014).
Vieira, P.L., de Jong, E.C., Wierenga, E.A., Kapsenberg, M.L. & Kalinski, P. Development of Th1-inducing capacity in myeloid dendritic cells requires environmental instruction. J. Immunol. 164, 4507–4512 (2000).
Kalinski, P., Schuitemaker, J.H., Hilkens, C.M. & Kapsenberg, M.L. Prostaglandin E2 induces the final maturation of IL-12-deficient CD1a+CD83+ dendritic cells: the levels of IL-12 are determined during the final dendritic cell maturation and are resistant to further modulation. J. Immunol. 161, 2804–2809 (1998).
Kalinski, P., Vieira, P.L., Schuitemaker, J.H., de Jong, E.C. & Kapsenberg, M.L. Prostaglandin E(2) is a selective inducer of interleukin-12 p40 (IL-12p40) production and an inhibitor of bioactive IL-12p70 heterodimer. Blood 97, 3466–3469 (2001).
Whittaker, D.S., Bahjat, K.S., Moldawer, L.L. & Clare-Salzler, M.J. Autoregulation of human monocyte-derived dendritic cell maturation and IL-12 production by cyclooxygenase-2-mediated prostanoid production. J. Immunol. 165, 4298–4304 (2000).
Hilkens, C.M., Kalinski, P., de Boer, M. & Kapsenberg, M.L. Human dendritic cells require exogenous interleukin-12-inducing factors to direct the development of naive T-helper cells toward the Th1 phenotype. Blood 90, 1920–1926 (1997).
Kalinski, P., Hilkens, C.M., Snijders, A., Snijdewint, F.G. & Kapsenberg, M.L. IL-12-deficient dendritic cells, generated in the presence of prostaglandin E2, promote type 2 cytokine production in maturing human naive T helper cells. J. Immunol. 159, 28–35 (1997).
Butterfield, L.H., Gooding, W. & Whiteside, T.L. Development of a potency assay for human dendritic cells: IL-12p70 production. J. Immunother. 31, 89–100 (2008).
Gustafsson, K. et al. Recruitment and activation of natural killer cells in vitro by a human dendritic cell vaccine. Cancer Res. 68, 5965–5971 (2008).
Wesa, A., Kalinski, P., Kirkwood, J.M., Tatsumi, T. & Storkus, W.J. Polarized type-1 dendritic cells (DC1) producing high levels of IL-12 family members rescue patient TH1-type antimelanoma CD4+ T cell responses in vitro. J. Immunother. 30, 75–82 (2007).
Carreno, B.M. et al. IL-12p70-producing patient DC vaccine elicits Tc1-polarized immunity. J. Clin. Invest. 123, 3383–3394 (2013).
Okada, H. et al. Induction of CD8+ T-cell responses against novel glioma-associated antigen peptides and clinical activity by vaccinations with α-type 1 polarized dendritic cells and polyinosinic-polycytidylic acid stabilized by lysine and carboxymethylcellulose in patients with recurrent malignant glioma. J. Clin. Oncol. 29, 330–336 (2011).
Zitvogel, L. et al. IL-12-engineered dendritic cells serve as effective tumor vaccine adjuvants in vivo. Ann. N. Y. Acad. Sci. 795, 284–293 (1996).
Xu, S. et al. Rapid high efficiency sensitization of CD8+ T cells to tumor antigens by dendritic cells leads to enhanced functional avidity and direct tumor recognition through an IL-12-dependent mechanism. J. Immunol. 171, 2251–2261 (2003).
Ten Brinke, A., Karsten, M.L., Dieker, M.C., Zwaginga, J.J. & van Ham, S.M. The clinical grade maturation cocktail monophosphoryl lipid A plus IFNgamma generates monocyte-derived dendritic cells with the capacity to migrate and induce Th1 polarization. Vaccine 25, 7145–7152 (2007).
Chiang, C.L. et al. Optimizing parameters for clinical-scale production of high IL-12 secreting dendritic cells pulsed with oxidized whole tumor cell lysate. J. Transl. Med. 9, 198 (2011).
Hokey, D.A., Larregina, A.T., Erdos, G., Watkins, S.C. & Falo, L.D. Jr. Tumor cell loaded type-1 polarized dendritic cells induce Th1-mediated tumor immunity. Cancer Res. 65, 10059–10067 (2005).
Budiu, R.A. et al. Immunobiology of human mucin 1 in a preclinical ovarian tumor model. Oncogene 32, 3664–3675 (2013).
Schreibelt, G. et al. Commonly used prophylactic vaccines as an alternative for synthetically produced TLR ligands to mature monocyte-derived dendritic cells. Blood 116, 564–574 (2010).
Zitvogel, L. et al. Therapy of murine tumors with tumor peptide-pulsed dendritic cells: dependence on T cells, B7 costimulation, and T helper cell 1-associated cytokines. J. Exp. Med. 183, 87–97 (1996).
Mayordomo, J.I. et al. Bone marrow-derived dendritic cells pulsed with synthetic tumour peptides elicit protective and therapeutic antitumour immunity. Nat. Med. 1, 1297–1302 (1995).
Paglia, P., Chiodoni, C., Rodolfo, M. & Colombo, M.P. Murine dendritic cells loaded in vitro with soluble protein prime cytotoxic T lymphocytes against tumor antigen in vivo. J. Exp. Med. 183, 317–322 (1996).
Fields, R.C., Shimizu, K. & Mule, J.J. Murine dendritic cells pulsed with whole tumor lysates mediate potent antitumor immune responses in vitro and in vivo. Proc. Natl. Acad. Sci. USA 95, 9482–9487 (1998).
Schnurr, M. et al. Apoptotic pancreatic tumor cells are superior to cell lysates in promoting cross-priming of cytotoxic T cells and activate NK and gammadelta T cells. Cancer Res. 62, 2347–2352 (2002).
Strome, S.E. et al. Strategies for antigen loading of dendritic cells to enhance the antitumor immune response. Cancer Res. 62, 1884–1889 (2002).
Chiang, C.L., Coukos, G. & Kandalaft, L.E. Whole tumor antigen vaccines: where are we? Vaccines 3, 344–372 (2015).
Wieckowski, E. et al. Type-1 polarized dendritic cells loaded with apoptotic prostate cancer cells are potent inducers of CD8(+) T cells against prostate cancer cells and defined prostate cancer-specific epitopes. Prostate 71, 125–133 (2010).
Specht, J.M. et al. Dendritic cells retrovirally transduced with a model antigen gene are therapeutically effective against established pulmonary metastases. J. Exp. Med. 186, 1213–1221 (1997).
Mullins, D.W. et al. Route of immunization with peptide-pulsed dendritic cells controls the distribution of memory and effector T cells in lymphoid tissues and determines the pattern of regional tumor control. J. Exp. Med. 198, 1023–1034 (2003).
Okada, N. et al. Administration route-dependent vaccine efficiency of murine dendritic cells pulsed with antigens. Br. J. Cancer 84, 1564–1570 (2001).
Engleman, E.G. & Fong, L. Induction of immunity to tumor-associated antigens following dendritic cell vaccination of cancer patients. Clin. Immunol. 106, 10–15 (2003).
Fong, L., Brockstedt, D., Benike, C., Wu, L. & Engleman, E.G. Dendritic cells injected via different routes induce immunity in cancer patients. J. Immunol. 166, 4254–4259 (2001).
Bedrosian, I. et al. Intranodal administration of peptide-pulsed mature dendritic cell vaccines results in superior CD8+ T-cell function in melanoma patients. J. Clin. Oncol. 21, 3826–3835 (2003).
Grover, A. et al. Intralymphatic dendritic cell vaccination induces tumor antigen-specific, skin-homing T lymphocytes. Clin. Cancer Res. 12, 5801–5808 (2006).
Radomski, M. et al. Prolonged intralymphatic delivery of dendritic cells through implantable lymphatic ports in patients with advanced cancer. J. Immunother. Cancer 4, 24 (2016).
Hu, J. et al. Induction of potent antitumor immunity by intratumoral injection of interleukin 23-transduced dendritic cells. Cancer Res. 66, 8887–8896 (2006).
Nishioka, Y., Hirao, M., Robbins, P.D., Lotze, M.T. & Tahara, H. Induction of systemic and therapeutic antitumor immunity using intratumoral injection of dendritic cells genetically modified to express interleukin 12. Cancer Res. 59, 4035–4041 (1999).
Thompson, E.D., Enriquez, H.L., Fu, Y.X. & Engelhard, V.H. Tumor masses support naive T cell infiltration, activation, and differentiation into effectors. J. Exp. Med. 207, 1791–1804 (2010).
Melero, I. et al. Evolving synergistic combinations of targeted immunotherapies to combat cancer. Nat. Rev. Cancer 15, 457–472 (2015).
Vasaturo, A., Verdoes, M., de Vries, J., Torensma, R. & Figdor, C.G. Restoring immunosurveillance by dendritic cell vaccines and manipulation of the tumor microenvironment. Immunobiology 220, 243–248 (2015).
Pardoll, D.M. The blockade of immune checkpoints in cancer immunotherapy. Nat. Rev. Cancer 12, 252–264 (2012).
Ribas, A. et al. Dendritic cell vaccination combined with CTLA4 blockade in patients with metastatic melanoma. Clin. Cancer Res. 15, 6267–6276 (2009).
Rosenblatt, J. et al. PD-1 blockade by CT-011, anti-PD-1 antibody, enhances ex vivo T-cell responses to autologous dendritic cell/myeloma fusion vaccine. J. Immunother. 34, 409–418 (2011).
Wilgenhof, S. et al. Phase II study of autologous monocyte-derived mRNA electroporated dendritic cells (TriMixDC-MEL) plus ipilimumab in patients with pretreated advanced melanoma. J. Clin. Oncol. 34, 1330–1338 (2016).
Melero, I., Hervas-Stubbs, S., Glennie, M., Pardoll, D.M. & Chen, L. Immunostimulatory monoclonal antibodies for cancer therapy. Nat. Rev. Cancer 7, 95–106 (2007).
Fujita, M. et al. COX-2 blockade suppresses gliomagenesis by inhibiting myeloid-derived suppressor cells. Cancer Res. 71, 2664–2674 (2011).
Wong, J.L., Obermajer, N., Odunsi, K., Edwards, R.P. & Kalinski, P. Synergistic COX2 induction by IFNγ and TNFα self-limits type-1 immunity in the human tumor microenvironment. Cancer Immunol. Res. 4, 303–311 (2016).
Obermajer, N., Muthuswamy, R., Lesnock, J., Edwards, R.P. & Kalinski, P. Positive feedback between PGE2 and COX2 redirects the differentiation of human dendritic cells toward stable myeloid-derived suppressor cells. Blood 118, 5498–5505 (2011).
Vacchelli, E. et al. Trial watch: IDO inhibitors in cancer therapy. Oncoimmunology 3, e957994 (2014).
Vanneman, M. & Dranoff, G. Combining immunotherapy and targeted therapies in cancer treatment. Nat. Rev. Cancer 12, 237–251 (2012).
Li, J. et al. Chemokine expression from oncolytic Vaccinia virus enhances vaccine therapies of cancer. Mol. Ther. 19, 650–657 (2011).
Jensen, S.M. et al. Signaling through OX40 enhances antitumor immunity. Semin. Oncol. 37, 524–532 (2010).
Dranoff, G. Experimental mouse tumour models: what can be learnt about human cancer immunology? Nat. Rev. Immunol. 12, 61–66 (2011).
Ngiow, S.F., Loi, S., Thomas, D. & Smyth, M.J. Mouse models of tumor immunotherapy. Adv. Immunol. 130, 1–24 (2016).
Betts, M.R. et al. Sensitive and viable identification of antigen-specific CD8+ T cells by a flow cytometric assay for degranulation. J. Immunol. Methods 281, 65–78 (2003).
Zaritskaya, L., Shurin, M.R., Sayers, T.J. & Malyguine, A.M. New flow cytometric assays for monitoring cell-mediated cytotoxicity. Expert Rev. Vaccines 9, 601–616 (2010).
Klenerman, P., Cerundolo, V. & Dunbar, P.R. Tracking T cells with tetramers: new tales from new tools. Nat. Rev. Immunol. 2, 263–272 (2002).
Karsunky, H., Merad, M., Cozzio, A., Weissman, I.L. & Manz, M.G. Flt3 ligand regulates dendritic cell development from Flt3+ lymphoid and myeloid-committed progenitors to Flt3+ dendritic cells in vivo. J. Exp. Med. 198, 305–313 (2003).
Pulendran, B. et al. Distinct dendritic cell subsets differentially regulate the class of immune response in vivo. Proc. Natl. Acad. Sci. USA 96, 1036–1041 (1999).
Kalinski, P., Schuitemaker, J.H., Hilkens, C.M., Wierenga, E.A. & Kapsenberg, M.L. Final maturation of dendritic cells is associated with impaired responsiveness to IFN-gamma and to bacterial IL-12 inducers: decreased ability of mature dendritic cells to produce IL-12 during the interaction with Th cells. J. Immunol. 162, 3231–3236 (1999).
Watchmaker, P.B. et al. Independent regulation of chemokine responsiveness and cytolytic function versus CD8+ T cell expansion by dendritic cells. J. Immunol. 184, 591–597 (2010).
Hoffmann, T.K., Meidenbauer, N., Dworacki, G., Kanaya, H. & Whiteside, T.L. Generation of tumor-specific T-lymphocytes by cross-priming with human dendritic cells ingesting apoptotic tumor cells. Cancer Res. 60, 3542–3549 (2000).
Albert, M.L. et al. Immature dendritic cells phagocytose apoptotic cells via alphavbeta5 and CD36, and cross-present antigens to cytotoxic T lymphocytes. J. Exp. Med. 188, 1359–1368 (1998).
Snijders, A., Kalinski, P., Hilkens, C.M. & Kapsenberg, M.L. High-level IL-12 production by human dendritic cells requires two signals. Int. Immunol. 10, 1593–1598 (1998).
Wieckowski, E. et al. Type-1 polarized dendritic cells loaded with apoptotic prostate cancer cells are potent inducers of CD8(+) T cells against prostate cancer cells and defined prostate cancer-specific epitopes. Prostate 71, 125–133 (2011).
MartIn-Fontecha, A. et al. Regulation of dendritic cell migration to the draining lymph node: impact on T lymphocyte traffic and priming. J. Exp. Med. 198, 615–621 (2003).
Wei, W.Z., Jones, R.F., Juhasz, C., Gibson, H. & Veenstra, J. Evolution of animal models in cancer vaccine development. Vaccine 33, 7401–7407 (2015).
Mac Keon, S., Ruiz, M.S., Gazzaniga, S. & Wainstok, R. Dendritic cell-based vaccination in cancer: therapeutic implications emerging from murine models. Front. Immunol. 6, 243 (2015).
Westwood, J.A., Darcy, P.K. & Kershaw, M.H. The potential impact of mouse model selection in preclinical evaluation of cancer immunotherapy. Oncoimmunology 3, e946361 (2014).
Devaud, C. et al. Tissues in different anatomical sites can sculpt and vary the tumor microenvironment to affect responses to therapy. Mol. Ther. 22, 18–27 (2014).
Devaud, C., John, L.B., Westwood, J.A., Darcy, P.K. & Kershaw, M.H. Immune modulation of the tumor microenvironment for enhancing cancer immunotherapy. Oncoimmunology 2, e25961 (2013).
Rakoff-Nahoum, S. & Medzhitov, R. Toll-like receptors and cancer. Nat. Rev. Cancer 9, 57–63 (2009).
Lou, Y. et al. Antitumor activity mediated by CpG: the route of administration is critical. J. Immunother. 34, 279–288 (2011).
Amos, S.M. et al. Adoptive immunotherapy combined with intratumoral TLR agonist delivery eradicates established melanoma in mice. Cancer Immunol. Immunother. 60, 671–683 (2011).
John, L.B. et al. Oncolytic virus and anti-4-1BB combination therapy elicits strong antitumor immunity against established cancer. Cancer Res. 72, 1651–1660 (2012).
Bartlett, D.L. et al. Oncolytic viruses as therapeutic cancer vaccines. Mol. Cancer 12, 103 (2013).
Kirn, D.H. & Thorne, S.H. Targeted and armed oncolytic poxviruses: a novel multi-mechanistic therapeutic class for cancer. Nat. Rev. Cancer 9, 64–71 (2009).
Kerkar, S.P. et al. IL-12 triggers a programmatic change in dysfunctional myeloid-derived cells within mouse tumors. J. Clin. Invest. 121, 4746–4757 (2011).
Zhang, L. et al. Improving adoptive T cell therapy by targeting and controlling IL-12 expression to the tumor environment. Mol. Ther. 19, 751–759 (2011).
Lotze, M.T. et al. Cytokine gene therapy of cancer using interleukin-12: murine and clinical trials. Ann. N. Y. Acad. Sci. 795, 440–454 (1996).
Schlom, J. Therapeutic cancer vaccines: current status and moving forward. J. Natl. Cancer Inst. 104, 599–613 (2012).
Datta, J. et al. Rationale for a multimodality strategy to enhance the efficacy of dendritic cell-based cancer immunotherapy. Front. Immunol. 6, 271 (2015).
Mach, N. et al. Differences in dendritic cells stimulated in vivo by tumors engineered to secrete granulocyte-macrophage colony-stimulating factor or Flt3-ligand. Cancer Res. 60, 3239–3246 (2000).
Lane, P., Burdet, C., McConnell, F., Lanzavecchia, A. & Padovan, E. CD40 ligand-independent B cell activation revealed by CD40 ligand-deficient T cell clones: evidence for distinct activation requirements for antibody formation and B cell proliferation. Eur. J. Immunol. 25, 1788–1793 (1995).
Thirunavukarasu, P. et al. A rationally designed A34R mutant oncolytic poxvirus: improved efficacy in peritoneal carcinomatosis. Mol. Ther. 21, 1024–1033 (2013).
Wilhelm, K. et al. Graft-versus-host disease is enhanced by extracellular ATP activating P2X7R. Nat. Med. 16, 1434–1438 (2010).
Kaka, A.S., Foster, A.E., Weiss, H.L., Rooney, C.M. & Leen, A.M. Using dendritic cell maturation and IL-12 producing capacity as markers of function: a cautionary tale. J. Immunother. 31, 359–369 (2008).
Nakamura, Y. et al. Helper function of memory CD8+ T cells: heterologous CD8+ T cells support the induction of therapeutic cancer immunity. Cancer Res. 67, 10012–10018 (2007).
Musha, H. et al. Selective infiltration of CCR5(+)CXCR3(+) T lymphocytes in human colorectal carcinoma. Int. J. Cancer 116, 949–956 (2005).
Kunz, M. et al. Strong expression of the lymphoattractant C-X-C chemokine Mig is associated with heavy infiltration of T cells in human malignant melanoma. J. Pathol. 189, 552–558 (1999).
Ohtani, H., Jin, Z., Takegawa, S., Nakayama, T. & Yoshie, O. Abundant expression of CXCL9 (MIG) by stromal cells that include dendritic cells and accumulation of CXCR3+ T cells in lymphocyte-rich gastric carcinoma. J. Pathol. 217, 21–31 (2009).
Sampath, P. et al. Crosstalk between immune cell and oncolytic vaccinia therapy enhances tumor trafficking and antitumor effects. Mol. Ther. 21, 620–628 (2013).
Obermajer, N., Muthuswamy, R., Odunsi, K., Edwards, R.P. & Kalinski, P. PGE(2)-induced CXCL12 production and CXCR4 expression controls the accumulation of human MDSCs in ovarian cancer environment. Cancer Res. 71, 7463–7470 (2011).
Acknowledgements
We dedicate this article to Eva Wieckowski (who died unexpectedly on 14 December 2015). We thank T.J. Curiel for providing the ID8A ovarian cancer cell line, P. Lane for CD40L-expressing J558 cells, and M. Kronenberg for Flt3-L-expressing B16 cells. We thank K. Lemon for assistance with monitoring the mice and BLI, and P. Bailey for critical review and editorial comments. This work was supported by NIH grant CA132714 (to P.K.), by a 2015 CRI Clinical Strategy Team Grant (to P.K.), and by the David C. Koch Regional Therapy Cancer Center (D.L.B.).
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N.O., J.U., and R.R. participated in data generation; N.O. and J.U. selected and evaluated the experimental data; N.O., J.U., R.M., R.R., E.W., P.K., and D.L.B. participated in the design of the protocol; N.O. and P.K. wrote the manuscript.
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Methods of production and clinical use of human variants of type-1 polarized DCs are a subject of two US patents (to P.K.). The remaining authors declare no competing financial interests.
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Obermajer, N., Urban, J., Wieckowski, E. et al. Promoting the accumulation of tumor-specific T cells in tumor tissues by dendritic cell vaccines and chemokine-modulating agents. Nat Protoc 13, 335–357 (2018). https://doi.org/10.1038/nprot.2017.130
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DOI: https://doi.org/10.1038/nprot.2017.130
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