Anti PD-1 Induces M1 Polarization in the Glioma ... · CD11b þand Iba1 monocytes ... study...

CLINICAL CANCER RESEARCH | TRANSLATIONAL CANCER MECHANISMS AND THERAPY

Anti–PD-1 Induces M1 Polarization in the GliomaMicroenvironment and Exerts Therapeutic Efficacy in theAbsence of CD8 Cytotoxic T Cells A CGanesh Rao1, Khatri Latha1, Martina Ott1, Aria Sabbagh1, Anantha Marisetty1, Xiaoyang Ling1,Daniel Zamler2, Tiffany A. Doucette1, Yuhui Yang1, Ling-Yuan Kong1, Jun Wei1, Gregory N. Fuller3,Fernando Benavides4, Adam M. Sonabend5, James Long6, Shulin Li7, Michael Curran8, andAmy B. Heimberger1

ABSTRACT◥

Purpose: Anti-programmed cell death protein 1 (PD-1) therapyhas demonstrated inconsistent therapeutic results in patients withglioblastoma (GBM) including those with profound impairments inCD8 T-cell effector responses.

Experimental Design: We ablated the CD8a gene in BL6 miceand intercrossed them with Ntv-a mice to determine how CD8 Tcells affect malignant progression in forming endogenous gliomas.Tumor-bearing mice were treated with PD-1 to determine theefficacy of this treatment in the absence of T cells. The tumormicroenvironment of treated and controlmice was analyzed by IHCand FACS.

Results: We observed a survival benefit in immunocompetentmice with endogenously arising intracranial glioblastomas afterintravenous administration of anti–PD-1. The therapeutic effectof PD-1 administration persisted inmice even after genetic ablation

of the CD8 gene (CD8�/�). CD11bþ and Iba1þ monocytes andmacrophages were enriched in the gliomamicroenvironment of theCD8�/� mice. The macrophages and microglia assumed a proin-flammatory M1 response signature in the setting of anti–PD-1blockade through the elimination of PD-1–expressingmacrophagesand microglia in the tumor microenvironment. Anti–PD-1 caninhibit the proliferation of and induce apoptosis of microgliathrough antibody-dependent cellular cytotoxicity, as fluorescentlylabeled anti–PD-1 was shown to gain direct access to the gliomamicroenvironment.

Conclusions:Our results show that the therapeutic effect of anti–PD-1 blockade in GBM may be mediated by the innate immunesystem, rather than by CD8 T cells. Anti–PD-1 immunologicallymodulates innate immunity in the glioma microenvironment—likely a key mode of activity.

IntroductionImmunotherapy has revolutionized the treatment of cancer. This

has generated interest in harnessing the immune system as a treatmentfor glioma, the most common primary brain tumor in humans (1–5).However, the effectiveness of immunotherapy against glioma is atten-uated by the immunosuppressive tumor microenvironment (6). In thecontext of metastatic cancer to the brain, immunotherapy has dem-onstrated significant efficacy, suggesting that this treatment is not

impeded by the blood–brain tumor barrier (7). Treatment of patientswith glioblastoma (GBM) with immune checkpoint inhibitors maybenefit a select patient subset (8, 9). However, these patients are knownto be profoundly immunosuppressed (10) and, in particular, lympho-penic (11). The number of cytotoxic CD8þ T cells, thought to becritically important to mediate the effects of immunotherapy, is verylow in subsets of patients with GBM (12), in part, related to theirsequestration in the bone marrow (13). In patients with GBM whodemonstrate a response to anti–PD-1 antibody (Ab), it is unclear whatimmune cell ismediating the antitumor effect because the CD8T cell ispresumed to be completely refractory to immune modulation (10).

The role of the T cell in the process of gliomagenesis is also unclear.GBMs frequently arise de novo butmay also originate froma low-gradeglioma precursor. Despite an initially indolent course, during whichsurvival time may be many years, low-grade gliomas almost inevitablyprogress to GBM (14–16). After this malignant transformation, sur-vival rates drop precipitously to 12–15 months. We have previouslyshown a direct correlation between an immune-suppressive micro-environment and malignant progression (17). As the immune systemrecognizes and eradicates tumor cells, some tumor cells evade theimmune system by avoiding detection or by becoming immunesuppressive to diminish the tumoricidal effects of CD8 T cells (18–20).Thus, by the time of diagnosis, GBM has already been subject toimmunoediting by T cells andmight not be susceptible to this immunecell population, even in the presence of immunotherapies that enhanceT-cell activity.

Here, we show that PD-1 Ab delivered intravenously significantlyincreases survival in immunocompetent mice with endogenouslyforming tumors (21, 22). To model the lack of CD8 T-cell effectorsobserved in human patients, we genetically modifiedmice to eliminate

1Department of Neurosurgery, Baylor College of Medicine, The University ofTexas MD Anderson Cancer Center, Houston, Texas. 2Department of GenomicMedicine and Cancer Biology, The University of Texas MD Anderson CancerCenter, Houston, Texas. 3Department of Pathology, The University of Texas MDAnderson Cancer Center, Houston, Texas. 4Department of Epigenetics andMolecular Carcinogenesis, The University of Texas MD Anderson Cancer Center,Houston, Texas. 5Department of Neurosurgery, Feinberg School of Medicine,Robert H Lurie Comprehensive Cancer Center, Northwestern University,Chicago, Illinois. 6Department of Biostatistics, The University of Texas MDAnderson Cancer Center, Houston, Texas. 7Department of Pediatrics, TheUniversity of Texas MD Anderson Cancer Center, Houston, Texas. 8Departmentof Immunology, The University of Texas MD Anderson Cancer Center, Houston,Texas.

Corresponding Authors: Amy B. Heimberger, The University of Texas MDAnderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030. Phone:713-792-2400; Fax: 713-794-4950; E-mail: [email protected]; andGanesh Rao, Department of Neurosurgery, Baylor College of Medicine, 7200Cambridge, Houston, TX 77030. E-mail: [email protected]

Clin Cancer Res 2020;26:4699–712

doi: 10.1158/1078-0432.CCR-19-4110

�2020 American Association for Cancer Research.

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the CD8 T cells. We hypothesized that the absence of the CD8 effectorresponse might promote malignant progression. Alternatively, selec-tive pressure of the immune effector response might induce the tumorto become more malignant and immunosuppressive through geneticalterations and instability of the tumor. We show that the CD8 T-cellpopulation does not influence glioma formation rates, tumor-freesurvival times, or malignant progression and that the innate immunesystem compensates for CD8 T-cell loss primarily through theinflux of immune-reactive macrophages and microglia. Even in theabsence of CD8 T cells, we observed a significant therapeutic effectfrom the administration of intravenous anti–PD-1 antibodies. Com-mensurate with this effect was a significant decrease in immune-suppressive PD-1þ macrophages within the tumor microenviron-ment, possibly allowing for greater tumor clearance by the proin-flammatory M1 macrophage population.

Materials and MethodsStudy designSample size and rules for stopping data collection

For the binary comparison, we set the control rate at 95%, for ax2 test with two-sided 5% alpha and 80% power, with 30 mice pergroup, we could detect a proportion in the experimental group of 67%as being significantly different from the control rate. Because apredefined interim analysis demonstrated marked differences in sur-vival after we used 10 animals per group, further data collection wasnot needed.

Data inclusion/exclusion criteriaProspectively, mice were replaced who died within three weeks after

injection of the RCAS vector or that did not receive at least three dosesof either anti–PD-1 Ab or IgG.

OutliersNo outliers were excluded from analysis.

Selection of endpointsThe primary prospective endpoint of the study was survival.

ReplicatesExperimental replicates are designated in each legend.

RandomizationMice were randomly sorted into treatment arms starting on day 21

after the initiation of gliomagenesis when thymic output is at amaximum (23) and after maturation of the immune system (24, 25).

BlindingThe investigatorswho assessed,measured, andquantified the results

were blinded to the experimental conditions and outcomes. Theprimary endpoint was death, which was recorded by the animalcaretakers and then associated with the treatment group.

Cell linesThe murine microglia cell line EOC-20 was purchased from the

ATCC and was cultured in DMEM (Corning Inc), supplemented with10% FBS and 1% penicillin/streptomycin at 37�C in a humidifiedatmosphere of 5% CO2 and 95% air. These cells were maintained bytrypsin passage every 2–3 days. The cell line was confirmed to be free ofMycoplasma.

AnimalsAll mice were housed in the MD Anderson Isolation facility in

accordance with Laboratory Animal Resources Commission stan-dards, and all work was supervised by the Institutional Animal Careand Use Committee at MD Anderson Cancer Center (Houston, TX;Protocol 00000900-RN01).

Generation of a C57BL/6J congenic strain carrying a null alleleof CD8 and the Ntv-a transgene

The Ntv-a transgene [avian cell surface receptor (TVA) for sub-group A avian leukosis virus under the control of a glial progenitor-specific promoter derived from the human nestin (NES) gene] and theCD8a-targeted allele were moved from their respective genetic back-grounds onto the C57BL/6 background by marker-assisted backcross-ing (26) to yield Ntv-a/CD8�/� and Ntv-a/CD8þ/þ mice.

Vector constructsThe RCAS/Ntv-amodel was described previously (21). Both RCAS-

PDGFB (27) and RCAS-STAT3 (17) vectors have been described. Thevector constructs are propagated in DF-1 chicken fibroblasts. Livevirus was produced by transfecting plasmid versions of RCAS vectorsinto DF-1 cells using FuGene6 (Roche). DF-1 cells senesce 1–2 daysafter injection. In mice injected with DF-1 cells with nontumor-inducing vectors, no long-term inflammatory response is observedin the brain or the brains appear histologically normal consistent withprior reports using this model (28, 29).

In vivo somatic cell transfer in transgenic miceTo transfer genes via RCAS vectors, 5 � 104 DF-1 producer cells

transfected with the RCAS vectors in 1–2 mL of PBS were injected intothe frontal lobes of mice using a 10-mL gastight Hamilton syringe (30).Micewere injectedwithin 24–48 hours after birth. Themicewere killed90 days after injection or sooner if they demonstrated morbidity. TheRCAS-PDGFB and RCAS-STAT3 model has been found to recapit-ulate many of the key immune features of human gliomas includingmacrophage infiltration and have been used to study antitumorimmunity for a variety of immune therapeutic strategies (31, 32).

Quantitative real time-PCRIn addition to IHC methods for detecting expression of CD8a in

tumor-bearing tissue, we also performed RT-PCR assay on the brainsof mice (n ¼ 3). After the mice were killed, their forebrains were

Translational Relevance

Immune checkpoint inhibition (ICI) has largely been ineffectiveagainst glioblastoma (GBM), likely due to the uniquely immuno-suppressed microenvironment of this primary brain tumor. Thepaucity of CD8þ T cells in GBM has long been considered thereason for the failure of ICI. However, the population of anti-programmed cell death protein 1 (PD-1þ) macrophages is veryrobust in GBM and may be targeted by ICI. We show that ICItargeting PD-1 results in significant survival gains in glioma-bearing immunocompetentmice evenwhenCD8T cells are absent.Treatment with anti–PD-1 antibody shifts the polarization ofremaining macrophages to the inflammatory (and antitumor)M1 phenotype. In humans, this strategy may not have directantitumor effect, but may be useful to reverse immunosuppressionby the resident macrophage population. Thus, combinatorialimmune strategies which include ICI, may be a rational next stepin the treatment of GBM.

Rao et al.

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removed and frozen in liquid nitrogen. Tissue specimens were homog-enized and RNAwas extracted using Qiagen RNeasyMini Kit. Reversetranscription was performed using Bio-Rad iScript cDNA SynthesisKit. Quantitative RT-PCR was performed using FastStart SYBR GreenMaster reagent (Roche) with the following primers: CD8a forward (F):CAAGCCCAGACCTTCAGAGA; CD8a reverse (R): TCCCCATCA-CACCCCTACTA. Data were normalized to internal GAPDH and b2-microglobulin.

Phenotypic characterization of tumorWe dichotomized tumors as either low-grade or high-grade on

hematoxylin and eosin (H&E)–stained tumor sections, depending onthe presence of histologic features including neovascularization andnecrosis. Tumor gradingwas performed by the study neuropathologist(G.N. Fuller).

Syngeneic clonotypic intracranial glioma modelTo induce intracerebral tumors in C57BL/6J mice, GL261 cells were

injected at a dose of 5 � 104 cells/mL as described previously (33).

Flow cytometry analysisPeripheral blood (50 mL) was collected and placed in 4mL ofmouse

RBC lysis buffer (eBioscience). Splenic single-cell suspensions wereprepared by mechanical dissociation of the spleen, followed by filtra-tion, and lysis of the RBCs. Cells were stained with anti-mouse CD4,CD8a, CD19, CD3e, NK1.1, ls-T, FoxP3, and granzyme B antibodies(eBioscience). For granzyme B measurements, T-cell activation wasperformed before antibody staining. Splenic single-cell suspensionswere prepared and cultured in RPMI1640 (containing 10% FBS) plusIL2. T cells were activated for 7 days using a Dynabead Mouse TActivator CD3/CD28 Kit. Antibodies used for flow cytometry in thisstudy (anti-CD11c, anti-CD11b, anti-CD19, anti-NK1.1, anti-CD3,and anti-MHCII) were from eBioscience, and anti-TNF-a was fromBecton Dickinson. Single-cell suspensions were prepared with aNeural Tissue Dissociation Kit from Miltenyi Biotec. For CD11b andTNF-a analysis, cells were treatedwith Cell Stimulation Cocktail and aprotein transport inhibitor (eBioscience) overnight, stained for surfaceCD11b, and fixed and permeabilized for TNF-a detection (using flowcytometry). TMEM 119 was used to stain microglia via ex vivo flowcytometry and by IHC (34).

IHC and immunofluorescence analysesMouse brains were fixed in 10% buffered formalin and were

paraffin-embedded, with 4-mm sections being used for IHC analysis.To detect Iba1 (1:1,000), the anti-Iba1 antibody fromWako was used.To quantify Iba1-positive cells, we counted the total number of cellsand the number of positively stained cells in the areas of highest tumor-cell density in 10 nonoverlapping microscopic fields (400X magnifi-cation) in tumor-bearing brains taken from mice in each group. Todetect CD8þ T cells in brain sections, we performed immunofluo-rescence using Fluor-conjugated antibodies from eBioscience (1:50).PD-1 for immunofluorescence was detected using antibodies fromAbcam (ab214421) or from R&D Systems (AF1021) and TMEM119using Abcam (ab209064). Alexa Fluor–conjugated secondary antibo-dies were used from Invitrogen Thermo Scientific. ProLong GoldAntifade Mountant with 40-diamidino-2-phenylindole (DAPI;Thermo Fisher Scientific) was used as the mounting medium. Slideswere further processed for imaging and confocal analysis using anOlympus Fluoview FV1000microscope.We quantified the percentageof positive cells by counting the number of cells that stained for both

Iba1 and TMEM119 and PD-1 in at least five nonoverlapping micro-scopic fields (magnification, 400� and/or 600�) from each genotype.The number of positive cells was divided by the total number ofDAPIþ

cells.

Ex vivo flow cytometry of intracerebral gliomasWhen themice started to show signs of neurologic deficit, they were

euthanized and the brains were collected after cardiac perfusion withPBS. To isolate immune cells from the brains, the brainsweremanuallydissected, filtered through a 70-mm cell strainer (BD Biosciences) andthe myelin was depleted from the single-cell suspension with Percollgradient centrifugation or magnetic bead separation (MACS MiltenyiBiotec) according to the manufacturer instructions. Next, cells wereincubated with Protein Transport Inhibitor Cocktail 500x (ThermoFisher Scientific) for 4 to 5 hours at 37�C. To prevent nonspecificbinding, cells were incubated with Fc-Block (CD16/32, Biolegend) andthen stained with fixable viability dye eFluor 780 to exclude dead cells(Thermo Fisher Scientific). To determine the different immune cellsubsets, the following antibodies were used: anti-mouse CD45 BV510,anti-mouse CD11b PerCP/Cy5.5, anti-mouse PD-1 BV421, anti-mouse CD3 PerCP/Cy5.5, anti-mouse CD4 FITC, anti-mouse IFNgPE/Cy7, anti-mouse CD49b – PE/Cy7, anti-mouse CD4 – BV510, (allBioLegend), anti-mouse CD25 BV510 (BD Biosciences), anti-mouseFoxp3 PE (Thermo Fisher Scientific), anti-mouse TMEM119(Abcam), goat anti-rabbit AlexaFluor 488 (highly cross-absorbed; LifeTechnologies). For fixation and permeabilization of the cells, theeBioscience Foxp3/Transcription factor Fixation/Permeabilization Kit(Thermo Fisher Scientific) was used according to the manufacturer’sinstructions. The cells were measured using FACS Celesta (BDBiosciences) and the data analysis was done with FlowJo software.

NanoString assayRNA (200 ng) at a concentration of 40 ng/mL in a total volume of

5 mL was prepared for NanoString assay analysis with the immune-specific gene array kit (NanoString Technologies, Inc). Sample prep-aration and hybridization were performed for the assay according tothe manufacturer’s instructions. Briefly, RNA samples were preparedby ligating a specific DNA tag (mRNA-tag) onto the 30 end of eachmature mRNA, and excess tags were removed via restriction enzymedigestion at 37�C. After processing with the mRNA sample prepara-tion kit, the entire 10-mL reaction volume containing mRNA andtaggedmRNAswas hybridizedwith a 10-mL reporter CodeSet, 10mL ofhybridization buffer, and a 5-mL capture ProbeSet (for a total reactionvolume of 35mL) at 65�C for 16–20 hours. Excess probeswere removedusing two-step magnetic bead-based purification with an nCounterPrep Station. The specific target molecules were quantified using annCounter Digital Analyzer by counting the individual fluorescent barcodes and assessing targetmolecules. The data were collected using thenCounter Digital Analyzer after obtaining images of the immobilizedfluorescent reporters in the sample cartridge using a charge-coupleddevice camera. These data were then normalized to mRNA geneexpression data for the GSE5099 Classical M1 VS Alternative M2macrophage gene panel (35). The cluster analyses were used todetermine deregulated genes between the anti–PD-1 and the IgGisotype control group by multigroup comparison using Qlucoresoftware. Gene counts were loaded into GSEA 4.0.1 as a tab-delimited text expression matrix, with each row representing a geneand its expression across the samples. Analyses were run with thedefault settings and 100 permutations. Dataset labels were created togroup the columns that were either anti–PD-1 antibody or IgG treated.

Immunologic Modulation of Gliomagenesis

AACRJournals.org Clin Cancer Res; 26(17) September 1, 2020 4701




Microglia assaysCell viability was assessed using the Presto-blue assay according to

the manufacturer’s recommendations (Invitrogen, A13261). Briefly,3,000 cells were seeded overnight into a 96-well plate containing100 mL/well of cell culture medium. Cells were treated with variousconcentrations of IgG (BioXcell; BE0089) and the anti–PD-1 antibody(BioXcell; BE0146) for 24, 48, 72, 96, and 120 hours. Next, 10 mL of cellviability reagent was added at the end of the incubation periods andincubated for 10 minutes in the dark (protected from light). Fluores-cencewasmeasured at 560 nm.Cell proliferationwasmeasured using aBrdU incorporation assay according to the manufacturer’s recom-mendations (Cell Signaling Technology; catalog No. 6813). Briefly,3,000 cells were seeded overnight and treated with various concentra-tions of IgG or the anti–PD-1 Ab for 24 and 48 hours. Twenty-fourhours before the proliferation measurement, the 10� BrdU solutionwas added at a concentration of 1� and the cells were incubated at37�C in an atmosphere of 95%air and 5%CO2. The cells werefixed andincubated with the detection antibody for an hour followed byincubation with HRP secondary antibody and the TMB substrate.Absorbance was measured at 450 nm. For assessment of the effects oncell cycle, the EOC-20 cells were harvested, fixed, stained with a PIantibody, and analyzed by flow cytometry after incubating with125 mg/mL of either the anti–PD-1 antibody or IgG.

Antibody-dependent cell cytotoxicity assayIn a typical antibody-dependent cell cytotoxicity (ADCC) assay, an

antibody binds on the surface of target cells, the effector cell FCreceptor recognizes cell-bound antibodies, and the cross-linkingresults in apoptosis of target cells. To investigate whether PD-1signaling was mechanistically required, we used two different blockingantibodies [RPM1-14 (BioXcell, BE0146) and 29F.1A12 (BioXcell,BE0273)] and a nonblocking antibody [RPM1-30 (eBioscience, 14-9981-82)]. RPM1-14 (anti–PD-1) is produced in rats and as such, theFc gamma R IV receptor of the mouse does not recognize/react withthe rat IgG2a (36). Repeat administration of rat anti-mouse antibodieswill trigger in vivo generation of mouse anti-rat responses. To reca-pitulate the in vivo vaccination response to RPM1-14, mouse anti-ratIgG antibodies were added to experimental arms. TheADCCassaywasperformed using standardmurine FcgRIIIADCCeffector cells accord-ing to manufacturer’s recommendations (Promega, G7015). Briefly, aday before the assay, target cells (EOC20) were plated in freshmediumin a 96-well plate and incubated in a CO2 incubator at 37�C. On thefollowing day, either IgG or anti–PD-1 or secondary antibodies werediluted in medium and added to the target cells, followed by theaddition of effector cells. The ratio of Effector:Target cells is main-tained at 20:1 in triplicate for 24 hours. The Bio-Glo luciferase reagentwas added and the luminescence (in relative light units, RLUs) wasdetermined using a SynergyHTX multimode plate reader.

Treatment with anti–PD-1 monoclonal Ab and in vivo depletionsWe used RMP1-14, a murine mAb against PD-1 (BioXCell). Mice

were injected with RCAS-PDGFBþ RCAS-STAT3 to induce primarilyhigh-grade tumors, consistent with glioblastoma. Mice were random-ized between the two treatment arms beginning 3 weeks after genetransfer in the perinatal period (approximates adolescence in humans)and were treated with either RPM1-14 (200 mg) or an IgG isotypecontrol (BioXCell) intravenously (via tail vein injection) three timesweekly for up to 5 weeks. At three weeks, thymic output is at its maxi-mum (23) and the immune system is fully mature (24, 25). Animalsthat died from tumor progression prior to the initiation of at least threedoses of anti–PD-1 antibody were replaced. Mice were monitored as

described previously and killed when they exhibited neurologicmorbidity. Their brains were removed and fixed in 4% formalin.

Anti–PD-1 fluorescent tagging and in vivo biodistributionanalysis

The PD-1 antibody was fluorescently tagged with Alexa Fluor 647using the SAIVI Rapid Ab Labeling Kit according to the manufac-turer’s recommendations (S30044, Invitrogen). Briefly, 1 mg of theantibody at a concentration of 2 mg/mL was incubated with the Alexa647 dye for 1 hour at room temperature with gentle stirring. A 3-cmcolumn was prepared by using the resin in the kit and washed twicewith elution buffer before loading the sample. The labeled antibodywas loaded onto the column, and all the eluted fractions were collected.The first-eluted colored bands contained the labeled antibody. Absor-bance of the purified conjugated antibody was measured at both A280and 650 nm and protein concentration was calculated using a Nano-drop 1000 spectrophotometer (Thermo Fisher Scientific). To evaluatewhether the anti–PD-1 antibody was able to infiltrate brain tumors,C57BL/6micewith intracerebral GL261 tumors established for 20 daysor mice injected with RCAS-PDGFBþRCAS-STAT3 in the Ntv-aþwild-type or in the homozygous CD8�/� background that wereneurologically symptomatic were injected intravenously with 200 mgof Alexa Fluor 647–conjugated anti–PD-1 antibody. The mice werekilled and their organs were harvested after 3 hours and imaged usingan IVIS 200 fluorescence imager.

Statistical analysisTheCochran–Mantel–Haenszel test was used to compare the tumor

incidence between different injection sets. Student two-sample t testwas applied to compare immune cell compositions between differentgroups. Kaplan–Meier survival curves were used to estimate unad-justed tumor latency. To compare the time-to-event variables betweengroups, the log-rank test was used to compare distributions. All testswere two-sided, and P < 0.05 was considered statistically significant.Statistical analysis was carried out using R version 3.1.2 software(R Core Team) and Graphpad Prism version 6.01 software (GraphpadSoftware, Inc).

ResultsEnhancement of innate immunity in the gliomamicroenvironment in the absence of CD8 T cells

To analyze the immune gliomamicroenvironment in the absence ofCD8 T cells, Ntv-a mice were backcrossed into a BL6 background andintercrossed with CD8�/� mice to create Ntv-a/CD8�/� mice. Toascertain the CD8 T-cell composition in the CD8�/, heterozygous(CD8�/þ), and wild-type mice (CD8þ/þ), their peripheral blood, bonemarrow, and spleens were analyzed. No CD8þ cells were detected inthese tissues of CD8�/�mice (Supplementary Fig. S1A). HeterozygousCD8 mice (Supplementary Fig. S1B) displayed a level of CD8 T cellssimilar to that in wild-type mice (Supplementary Fig. S1C and S1D;P > 0.05). The flow cytometry data were consistent with the geneticanalysis of CD8 T-cell loss achieved in the CD8 KO background. Weidentified intratumoral infiltration of CD8þ T cells in the CD8þ/þ

mice, but as expected, these cells were absent in the CD8�/� mice(Supplementary Fig. S1E). To test the hypothesis that the CD8 effectorresponse facilitates evasion of immune detection, thereby decreasinganimal survival and promoting malignant progression, gliomas wereinducedwithRCAS-PDGFBþRCAS-STAT3 inNtv-amice in either theCD8þ/þ or CD8�/� background. CD8þ/þ wild-type mice survivedlonger (49.5 days, 95%CI: 27–69) than the CD8�/�mice (27 days, 95%

Rao et al.





CI: 21–46) although this was not statistically significant (Fig. 1A,P ¼ 0.2281, H.R.1.354). In addition, there was no difference in theincidence of high-grade gliomas, regardless of CD8 status (Fig. 1B).

CD11b is a marker of myeloid cells of the innate immune systemincluding macrophages. In the tumor-bearing mice we found anincrease in the CD11bþ cells in the CD8�/� mice relative to CD8þ/þ

mice in both the spleen and blood (Fig. 1C; P ¼ 0.02 for blood andP < 0.0001 for spleen). There was also a significant difference in themacrophage and microglia marker Iba1þ in tumors induced inCD8�/� mice compared with CD8þ/þ mice (Student t test,

P ¼ 0.0009; Fig. 1D). MHCþCD11bþ cells demonstrated a >2-foldincrease in the gliomas of CD8�/� mice relative to CD8þ/þ mice(Fig. 1E). This expansion was not an intrinsic property of the CD8�/�

mice, as shown by the lack of an increase in the percentage of CD11bþ

cells in the brains of nontumor-bearing control mice. These macro-phages possess features that reflect a proinflammatory immune pro-pensity based on TNF-a expression (Fig. 1F; P ¼ 0.001). Increases ininnate immunity in the gliomamicroenvironment were also found in asecond glioma model in which retrovirus induced PGFRþPten�/�murine gliomas were depleted of CD8 T cells (37). Cumulatively, these

Figure 1.

CD8 knockout (CD8�/�) does not have an impact on survival in a genetically engineered murine model of glioma and demonstrate a compensatory increase inmacrophages in the glioma microenvironment. A, As shown by the Kaplan–Meier curves, survival was not affected when 5 � 104 DF-1 cells/mouse containing theRCAS-PDGFBþRCAS-STAT3 geneswere injected bilaterally into the frontal brain lobes of CD8�/�mice.B,At the time of death or when the animal wasmoribund, anautopsywas performed, andmicroscope slides containing sections of the central nervous system (CNS)were stainedwith H&E for tumor grading. TheCD8 status didnot influence the glioma grade.C,CD11bþmonocytes andmacrophageswere found to occurmore frequently in the blood and spleens of tumor-bearing CD8�/�micethan in those of CD8þ/þmice (� , P¼ 0.0202 for blood; �� , P < 0.0001 for spleen). The analysis was conducted using ex vivo flow cytometry and the percentage ofCD11bþ cells was calculated on the basis of the total alive cells with 30,000 total events analyzed.D,CD11bþMHC IIþ cells were found by ex vivo flow cytometry to beenriched in the CD8�/� group relative to the wild-type CD8þ/þ group (�� , P¼ 0.0001). The percentage of dual expressing CD11bþMHC IIþ cells was calculated onthe basis of the total alive cells with 30,000 total events analyzed. E, Dot plot summarizing a significantly higher percentage of Iba1þ cells in gliomas in the CD8�/�

mice than in CD8þ/þ mice. P ¼ 0.001. F, Functional analysis by detecting intracellular TNF-a expression with flow cytometry demonstrated increased numbers ofTNF-aþ-expressing CD11bþ cells in glioma-bearing CD8�/�mice (�� , P¼ 0.0001). The analysis was conducted using ex vivo flow cytometry and the percentage ofCD11bþ cells that had intracellular expression of TNF-awas calculated on the basis of the total alive cells with 30,000 total events analyzed. G, CD8 knockout micedemonstrate a compensatory increase in the peripheral CD4 compartment. The CD4þ T-cell percentage in spleens was increased in CD8�/�mice (��P¼ 0.0002).H, Because there was an increase in the frequency of CD4þ T cells in the CD8�/� mice, the fraction of Tregs within the CD4 compartment was assessed, butno differences were observed in this in either the blood or spleen compartments, regardless of CD8 status. I, The percentage of CD4þ T cells producing granzymeB was found to be increased only in CD8�/� mice harboring intracranial tumors (�� , P < 0.0001).






data indicate an association of the innate immune system withgliomagenesis and antitumor immune reactivity.

To ascertain whether there was a compensatory increase in CD4T cells in the absence of CD8T cells in thesemodels, we analyzed bloodand spleens for the percentage of CD4 T cells in the CD8�/� and wild-type mice. The frequency of CD4 T cells was elevated in the spleens ofCD8�/� mice relative to that in the CD8þ/þ mice (P ¼ 0.0002).However, there was no difference in the frequency of the CD4 T cellsin the blood of the CD8�/� mice relative to the wild-type mice

(Fig. 1G). Previously, an increased regulatory T-cell fraction wasshown to be present within the CD4 compartment in patients withhigh-grade gliomas (11). To determine whether there was an alterationof glioma-induced immune suppression between the CD8�/� andCD8þ/þ mice, both their blood and spleens were analyzed for thefraction of CD4þ and FoxP3þ T cells. We found no difference in thefraction of CD4þ FoxP3þT cells between glioma-bearing CD8�/� andCD8þ/þ mice (Fig. 1H), indicating that Tregs are not differentiallymodulated. Because some CD4 T cells are cytotoxic cells (38, 39),

Figure 2.

CD8 knockout mice (CD8�/�) demonstrate a compensatory increase inmultiple peripheral immune compartments.A,CD4þ T cells were not detected in the gliomasof either CD8�/�mice or wild-type mice (CD8þ/þ) by IHC staining. CD19þ cells (B) and NK1.1þ (C) populations in the blood and spleen of glioma-bearing mice werefound to be significantly higher in the CD8�/� group relative to the CD8þ/þ group (�� , P ¼ 0.0004 and ��, P < 0.0001, respectively). However, these cellpopulations were not detected within the brain tumors. Spleen staining is the positive control. Representative IHC-stained images at 200�magnification for A andB (left, bar¼ 100 mm); A and Bmiddle and right at 400�magnification, scale bar, 50 mm. D, Kaplan–Meier estimates of tumor-free survival in glioblastoma-bearingNtv-amice in thewild type (CD8þ/þ) or CD8�/� background treatedwith anti–PD-1 or IgG isotype control (n¼ 11–13 per group). Themedian overall survival timewas68 days in the anti–PD-1 antibody-treated wild-type mice and 40 days in the control group (log-rank test, P ¼ 0.0002; left). The median overall survival time was61 days in the anti–PD-1 antibody-treated CD8�/� mice and 39 days in the control group (log-rank test, P < 0.0001).

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including those directed against malignancies (refs. 40, 41; a per-forin- and granzyme B-mediated process; ref. 42), we exploredwhether the increase in CD4 frequency was tied to these specificfunctional activities. In mice harboring gliomas, there was a 2.4-foldincrease of granzyme bþ CD4 T cells in the spleens of CD8�/� micerelative to that in CD8þ/þ mice (Fig. 1I). However, this expansionwas not an intrinsic property of the CD8�/� background, as shownby the absence of an increase in the percentage of granzyme bþCD4 T cells in nontumor-bearing control mice (not injected withDF-1 cells) possibly related to the immunologic recognition of theintracranial tumor. Further analysis of the CD8�/� mice harboringgliomas demonstrated no expansion of this population in theglioma microenvironment (Fig. 2A), indicating that the CD4cytotoxic T cell does not have a dominant role in the immunologiccontrol of gliomagenesis.

Compensatory immune cell expansionoccurs peripherally in theabsence of CD8 T cells

Because we observed expansion of the CD4 T-cell population inthe spleens of mice with the CD8�/� background (Fig. 2A), weinvestigated whether this occurred with other immune cell popula-tions such as CD19þ B cells (Fig. 2B) and NK1.1þ cells (Fig. 2C). Inthe CD8�/� glioma-bearing mice, there was a compensatoryincrease in both B cells and NK cells, in the blood and spleen.However, we saw no CD19þ B cells in gliomas arising in either theCD8�/� (n ¼ 8) or CD8þ/þ wild-type mice (n ¼ 9). Similar findingswere obtained for the NK1.1 population in the CD8�/� (n ¼ 8) andwild-type backgrounds (n ¼ 9), again indicating that although theremay be expansion of these populations in the periphery as a reactionto the tumor but there was no evidence of their involvement intrafficking to and exerting an effector response in the tumormicroenvironment.

Anti–PD-1 exerts a therapeutic effect in the absence of CD8T cells

To investigate the role of anti–PD-1 in the absence of CD8 T cells,we induced high-grade gliomas in Ntvaþ/BL6 mice of the CD8þ/þ

background by coinjecting RCAS-PDGFBþRCAS-STAT3, as has beendescribed previously (17). Mice were observed for 3 weeks and treatedwith anti–PD-1 intravenously through tail-vein injection. Animalswere treated with 200 mg of either rat IgG (control) or anti–PD-1administered intravenously every week for up to 5 weeks. Mice areweaned at 3 weeks after birth and they reach maturity (adulthood) by8–10 weeks (43). On the basis of this report, the mice used in theseexperiments started treatment as teenagers with the vast majority oftreatments occurring in adulthood. The median survival time for theanti–PD-1–treated group was 68 days, and in the IgG control group itwas 40 days; (log-rank test, P ¼ 0.0002; Fig. 2D), demonstratingtherapeutic activity of anti–PD-1 in this model and similar to theGL261 model (44–46). To investigate whether anti–PD-1 would exerta therapeutic effect in the CD8�/� mice, we treated a cohort of thesemice. Compared with the anti–PD-1–treated groups, median survivaltimes for the IgG-treated controls were 40 days (relative to the PD-1Ab–treated group of 68 days; log-rank test, P ¼ 0.0002) and 39 days(relative to the PD-1Ab-treated control group of 61 days; log-rank test,P < 0.0001) in the CD8þ/þ and the CD8�/� mice, respectively(Fig. 2D). The difference in the survival of the CD8þ/þ mice and theCD8�/� mice treated with anti–PD-1 (Fig. 2D) was not statisticallysignificantly different (68 vs 61 days, respectively, log-rank test P ¼0.0727). These results demonstrate that a similar therapeutic effect wasobtained for anti–PD-1 regardless of the presence of CD8 T cells.

CD11bþ myeloid cells are the predominant immune cellspopulation in the brains of tumor-bearing Ntv-a/CD8�/�

and Ntv-a/CD8þ/þ miceTo investigate which immune cells subsets could mediate the anti–

PD-1–mediated survival benefit in the absence of CD8 T cells, weisolated immune cells from the whole brains of tumor-bearing Ntv-a/CD8�/� and Ntv-a/CD8þ/þ mice treated with PD-1 antibody or IgGcontrol for multicolor flow cytometry (Supplementary Fig. S2). First,we quantified the percentage of myeloid cells (defined as CD11bþ

cells), CD4T cells (defined asCD3þCD4þ cells), andNK cells (definedas CD3� CD49þ cells) because those immune cells have already beenreported to express PD-1 (Fig. 3A). With an average of around 70%,CD11bþ myeloid cells (Fig. 3A, left) represented the majority ofimmune cells in all groups, whereas the contribution of CD4 T cells(middle), was less than 5%, and NK cells (right), were less than 1%.There was no statistical significant difference in the percentage of thedifferent immune cells subsets between any of treatment groups withintheir genotype. Next, we determined the PD-1 expression levels oneach of these cell subsets. The CD11bþ myeloid cell subset showed anaverage PD-1 expression between 10% and 20% (Fig. 3B, left) with nosignificant differences between the treatment groups. CD11bþmyeloidcells, but not CD4 T cells, are present throughout the brain paren-chyma. Notably, myeloid cells only showed PD-1 expression in thepresence of glioma (Supplementary Fig. S3). Thus, the flow cytometryanalysis, which analyzed the whole brain including the glioma, under-estimates themodulation of this population in the setting of anti–PD-1treatment within the glioma microenvironment. In the CD4 T-cellsubset (Fig. 3B, middle), there was much more variability in thepercentage of PD-1 expression within the different groups (Fig. 3B,middle) with expression levels ranging from11.5% to 89.4%. Therewasa statistically significant difference detected between the anti-IgG andthe anti–PD-1 in the Ntv-a CD8þ/þ mice (P ¼ 0.0425) but not inthe Ntv-a/CD8�/� mice. In the NK-cell subset, only 2% to 4%showed PD-1 expression and there was no difference between thedifferent groups (Fig. 3B, right). Because the CD4 T cells showed quitehigh PD-1 expression, even though they are not very frequent andcontribute only to very small extent to the immune cell composi-tion, we examined whether the anti–PD-1 treatment influencestheir functional status. Therefore, we stained the CD4 T cells forIFN-g (Fig. 3C, left), TNF-a (Fig. 3C, middle), and for regulatoryT cells (Fig. 3C, right), but we could not detect any significantchanges, indicating the CD4 T cells are not the main contributors ofour observed effects.

Anti–PD-1 recalibrates the glioma-infiltrating macrophages/microglia to an M1 phenotype

Because PD-1 expression has been previously described for tumor-associated macrophages (47), we evaluated whether PD-1 expressionwas present within gliomas and coexpressed with macrophages withinthe CNS. Using dual immunofluorescence, clusters of PD-1- and Iba1-positive cells were detected in both the CD8þ/þ and CD8�/� mice(Supplementary Fig. S4 and S5). Throughout the glioma microenvi-ronment, there was heterogeneous expression of Iba1þPD-1�,Iba1�PD-1þ, and Iba1þPD-1þ cells. Given that the tumor-associated macrophages express PD-1, and that there are scant CD8T cells in gliomas at diagnosis, we hypothesized that PD1 blockademight act independently of CD8 T-cell–based immune responses. Toassess whether PD-1 blockade has any role in the regulation ofmacrophages/microglia in the tumor microenvironment we per-formed multi-immunofluorescent staining for the microglia-specificmarker TMEM119, Iba1, and PD-1 (Fig. 4A). We noticed that in the






anti–PD-1–treated group relative to the IgG control during thetherapeutic window, the microglia/macrophage expression wasdecreased in both the CD8þ/þ and the CD8�/� backgrounds.Notably, relative to the IgG control group, we detected a markeddecrease in the populations expressing Iba1þ TMEM119þ

PD1þcells within the tumor in both CD8þ/þ and CD8�/� mice(P < 0.0001; Fig. 4B). The decreased expression of macrophage/microglia markers Iba1 and TMEM119 (Supplementary Fig. S6)suggest that the innate immune system may be mediating the effectsof the anti–PD-1 antibody.

Given that the PD-1–expressingmacrophage/microglial populationwas reduced in the setting of anti–PD-1 therapy, we evaluated whetherthe remainingmacrophage/microglia populationwas exerting a proin-flammatory or M1 classical skewed phenotype, as these cells can existeither in a nonpolarized state or along a continuum that is eitherpro- or anti-inflammatory (48). Gliomas in either the CD8þ/þ or theCD8�/� backgrounds treated with anti–PD-1 demonstrated a signi-ficant skewing in genes associated with the proinflammatory M1phenotype relative to the IgG isotype control–treated mice based onNanoString profiling (Fig. 4C). The upregulated genes were related to

Figure 3.

Flow cytometric analysis of immune cell subsets isolated from the whole brain of glioblastoma-bearing Ntv-amice in the wild type (CD8þ/þ) or CD8�/� backgroundtreated with PD-1 antibody or IgG isotype control.A, Percentage of CD11bþmyeloid (left; CD8þ/þ IgG vs anti–PD-1 P¼ 0.7933; CD8�/� IgG vs anti–PD-1 P¼ 0.6872),CD3þ CD4þ T cells (middle; CD8þ/þ IgG vs anti–PD-1 P¼ 0.9975; CD8�/� IgG vs anti–PD-1 P¼ 0.5572), and cells NK cells (right; CD8þ/þ IgG vs anti–PD-1 P¼ 0.7146;CD8�/� IgG vs anti–PD-1 P¼0.3732) of all immune cells isolated from thewhole brains of tumor-bearingNtv-amice.B, PD-1 expression onCD11bþmyeloid cells (left;CD8þ/þ IgG vs anti–PD-1 P¼0.1551; CD8�/� IgG vs anti–PD-1 P¼0.8539), CD3þCD4þ T cells (middle; CD8þ/þ IgG vs anti–PD-1 P¼0.0425; CD8�/� IgG vs anti–PD-1,P¼0.2647), andNK cells (right; CD8þ/þ IgG vs anti–PD-1 P¼0.2294; CD8�/� IgG vs anti–PD-1 P¼0.5275). C,Quantification of functional CD3þCD4þ T-cell subsetsin the brains of tumor-bearingmice, percentage of IFNg (left; CD8þ/þ IgG vs anti–PD-1, P¼0.9441; CD8�/� IgG vs anti–PD-1, P¼0.5492) and TNF-a (middle; CD8þ/þIgG vs anti–PD-1, P¼0.8415; CD8�/� IgG vs anti–PD-1 P¼0.2869) expressing cytotoxic CD3þCD4þ T cells, and percentage of regulatory T cells (Treg; right; CD8þ/þIgG vs anti–PD-1 P¼0.5431; CD8�/� IgG vs anti–PD-1 P¼0.2994). Two-sided unpaired t test was performed to compare the treatment groupswithin the genotypes,only the statistical significant values (P ≤ 0.05) are indicated in the figure.

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class I MHC–mediated antigen processing and presentation(eg, TRIM69), skewing to or maintaining the M1 phenotype(eg, LP1N1, EPST1 ZNF229, HDAC3), macrophage differentiation(eg, ERMAP), toll-like receptor transcription (eg, HCFC2), phagocy-tosis (eg, ATP10A), and survival (eg, E4F1; Supplementary Fig. S7).Together, these results indicate that anti–PD-1 enriches for a classicalM1 skewed macrophage phenotype in the glioma microenvironment.

Anti–PD-1 has direct effects on PD-1–expressing macrophage/microglia expressing cells, but also triggers ADCC

To define the direct effects of the anti–PD-1 antibody on macro-phages/microglia, we used EOC-20, a murine microglia cell line thatexpresses PD-1 (Fig. 5A) to a similar extent as tumor-infiltratingmyeloid cells (defined as CD45þ CD11bþ cells; Fig. 5B), to assay cellviability, proliferation, and cell cycle. At a concentration of 125 mg/mLof anti–PD-1, there was direct loss of cell viability after one day ofcoincubation that was further enhanced by extended exposure

(Fig. 5C). The effect of the anti–PD-1 antibody was partially mediatedby direct inhibition of cellular proliferation (Fig. 5D). After treatmentwith the anti–PD-1 antibody for 48 hours, EOC20 cells were analyzedfor cell-cycle distribution with flow cytometry, and there was no dif-ference upon exposure to anti–PD-1 (Fig. 5E). Given that the max-imum circulating concentration of anti–PD-1 was approximately3mg/mL (200-mg infusion in a 60-mLbloodvolume),we testedwhetherthe anti–PD-1 antibody could be further potentiated secondary to theinvolvement of ADCC. There was no substantial increase in cytotox-icity as detected by luminescence released from the microglia targetcells in the presence of the isotype IgG control and effector cells, but thepresence of anti–PD-1 increased this by 113% (Fig. 5F). Becauserepeated administration of the anti–PD-1 rat anti-mouse antibodyin vivo would trigger an anti-rat humoral response, a secondaryantibody (anti-rat IgG) was added to the ADCC assay, which furtherenhanced luminescence release by 233%. To further investigate wheth-er PD-1 signaling ismechanistically required, both blocking antibodies

Figure 4.

Anti–PD-1 recalibrates the glioma-infiltrating macrophages/microglia to an M1 phenotype. A, Representative immunofluorescent images of Iba1þTMEM119þPD-1þ

microglia infiltration in a glioblastoma-bearing mouse (CD8þ/þ background, with similar staining in the CD8�/� background) after IgG or anti–PD-1 Ab treatment,400� magnification, scale bar ¼ 50 mm. B, Scatter plot demonstrating difference in PD-1–expressing Iba1þTMEM119þ expression in the wild-type CD8þ/þ and KOCD8�/�backgroundmice demonstrating a significant reduction in the number of Iba1þTMEM119þmicroglia after anti–PD-1 antibody treatment (P

(RMP1-14 and 29F.1A12) and a nonblocking antibody (RMP1-30)were evaluated in the ADCC assay. All three antibodies triggeredADCC elimination of PD-1–expressing microglia, indicating thatanti–PD-1 triggers the elimination of PD-1–expressing immune-suppressive innate immune cells in the CNS through an ADCC-mediated mechanism independent of PD-1 blockade.

Anti–PD-1 antibodies can be detected in CNS gliomasTo ascertain whether anti–PD-1 was able to penetrate CNS gliomas,

the anti–PD-1 antibody was fluorescently labeled and subsequentlyinjected intravenously into mice. Nonglioma–bearing, untreatedC57BL/6 mice showed no baseline fluorescence, but mice treated withthe labeled anti–PD-1 antibody demonstrated fluorescence in thebrain and systemic organs (Fig. 6A). In tumor-bearing mice treatedwith labeled anti–PD-1 antibody, ex vivo analysis of the entire braindemonstrated elevated fluorescence in the right frontal lobe where the

GL261 glioma cells were implanted, but no focal fluorescence wasdetected in nontumor-bearing mice (Fig. 6B). Increased fluorescencewas detected in the cerebellum (ie, posterior fossa) including innonglioma-bearingmice. Subsequent sequential coronal sections fromanterior to posterior directly confirmed the presence of tumor in theright frontal lobe that correlated with this increased fluorescence(Fig. 6C). Because GL261 is surgically implanted and grows in a focalmanner, this increased fluorescence could be an artifact of the model.As such, we evaluated the penetration of fluorescently labeled anti–PD-1 in the Ntv-a model in which gliomagenesis is triggered in theneonatal period. In addition, gliomas generated in the Ntv-a model arediffusely infiltrative and can be multifocal, more closely recapitulatinghuman gliomas. In the Ntv-a CD8�/� mice bearing gliomas treatedwith labeled anti–PD-1, increased fluorescence was detected in mul-tiple areas of the CNS relative tomicewithout gliomas (Fig. 6D). Theseareas were confirmed to histologically contain glioma (Fig. 6E).

Figure 5.

Direct and ADCC activity of anti–PD-1 Ab on PD-1–expressing macrophage/microglia. A, PD-1 expression of EOC-20 cells, gated on CD45þ CD11bþ cells. B, PD-1expression on myeloid cells (CD45þ, CD11bþ) isolated from tumor-bearing Ntv-a mice. C, Coincubation of only the anti–PD-1 antibody at the designatedconcentrations with PD-1 expressing EOC-20 microglia resulted in diminished cellular viability starting 1 day after exposure, which was further enhanced withincreased exposure time. D, Coincubation of BrdU-labeled EOC-20 microglia with increasing concentrations of anti–PD-1 relative to the IgG control demonstrateddecreased proliferative capacity. E, Cell-cycle analysis of EOC-20 cells exposed to IgG control or anti–PD-1 demonstrating that these antibodies do not affect cellularproliferation. F,ADCC assay detecting lactic dehydrogenase leakage [luminescence; relative light units (RLU)] from target EOC-20microglia cells upon exposure toanti–PD-1 and in the presence of effector cells capable of mediating ADCC. Mouse microglial target cells, EOC20, were incubated with control antibody or anti–PD-1antibody at a concentration of 125 mg/mL or 12.5 mg/mL, followed by the addition of effector cells. The E:T ratio was 20:1. After 8 hours of induction at 37�C, Bio-Gloluciferase reagentwas added, and luminescence (RLU)was determined. TheADCCwas further potentiatedby the presence of a secondary antibody (mouse anti-rat)that could be generated by repeat administration of a rat anti-mouse antibody (anti–PD-1) in vivo. M, media; T, target; E, effector cell. When the anti–PD-1 antibodywas decreased to 12.5 mg/mL, similar results were obtained.

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Cumulatively, these data indicate that the anti–PD-1 can pass into theCNS, especially within gliomas in which there is breakdown of theblood–brain barrier (BBB; such as GL261) and in gliomas that haveevolved intrinsically within the CNS that are more diffusely infiltrat-ing, such as those induced in Ntv-a mice.

DiscussionIn our previous clinical trial of patients with recurrent glioblastoma

that were treated with anti–PD-1 prior to surgery, immune profiling ofthe tumor microenvironment revealed a marked paucity of effectorT cells but a profound predominance of macrophages displayingheterogeneous immune-stimulatory and immune-suppressive pheno-types. In our clinical trial (49) and in another (8), patients withglioblastoma treated in an adjuvant setting had better than expectedoutcomes. Thus, there arises the paradox of an anti–PD-1 agent havingtherapeutic activity in an oncologic setting that is notable for T cells

that are sequestered in the bone marrow and completely refractory tobeing reinvigorated with immune checkpoint inhibitors (10). It islikely that anti–PD-1 can switch its therapeutic effect between variousimmune populations given their relative frequencies. In malignanciesenriched in T-cell infiltration, anti–PD-1 likely exerts most of itstherapeutic activity through direct T-cell–ligand interactions. In con-trast, in malignancies such as glioblastoma that are devoid of T cells,anti–PD-1 activity may exert a therapeutic effect through alternativeimmune populations such asmacrophages andmicroglia. In this latterscenario, the therapeutic activity is mediated through the eliminationof an immune-suppressive, tumor-supportive PD-1þ macrophage/microglia population by myeloid-to-myeloid ADCC-mediated fratri-cide mechanisms and/or to M1 macrophage polarization. PD-1 haspreviously been shown to be expressed by macrophages, which limitstheir phagocytic capacity (47) and as such, treatment with anti–PD-1may be enhancing this activity in vivo. Proinflammatory M1 macro-phages mediate direct tumor killing through secreted products like

Figure 6.

Anti–PD-1 in vivo biodistribution analysis. A, C57BL/6 mice were either untreated (control) or injected with 200 mg of fluorescently labeled anti–PD-1. After 3 hours,their organswere harvested, rinsed in PBS, positioned on a petri dish, and then imaged using the IVIS 200 Fluorescence Imager. The organswere then photographed.B, Thebrain fromanontumor-bearingC57BL/6mouse treatedwithfluorescently labeled anti–PD-1 (top) or bearing intracerebral GL261 (bottom). The arrowdenotesthe location of the GL261 implantation. C, The brains from Bwere then sequentially coronally sectioned, with the non-tumor–bearing brain on the left and the GL261-implanted brain on the right. The sections were positioned anterior to posterior on the petri dish and imaged. The horizontal arrow denotes the location of theimplanted GL261 cells in the right frontal lobe. Increased fluorescent intensity is detected in the posterior cerebellum. All brain sections were imaged for the sameexposure time. D, Nonglioma–bearing (control) or glioma-bearing Ntv-a mice in the knockout (CD8�/�) background were treated with 200 mg of fluorescentlylabeled anti–PD-1. After 3 hours, their brains were harvested, rinsed in PBS, coronally sectioned, positioned on a petri dish, and then imaged using the IVIS 200Fluorescence Imager. The brains were then photographed. Increasing fluorescence intensity is seen to correlate with increasing concentration of anti–PD-1. E, H&E-stained coronal sections of GL261 (top) and Ntv-a CD8�/� (bottom) from C and D. Magnification 20�; scale bar, 100 mm. Arrows indicate the region of the tumor.






nitric oxide or tumor necrosis factor (50). This alternative mechanismof anti–PD-1 activity also provides an explanation for the failure ofbiomarkers to predict clinical responses in cancers such as glioblas-toma, as these markers are focused on the immune functional featuresof the adaptive immune system such as the abundance of antigens(ie, mutational burden) and the presence of T-cell infiltration andligand frequency (ie, PD-1þ TILs, PD-L1), which are unlikely to be thedominant mechanism of therapeutic activity of anti–PD-1 in thesetting of myeloid-enriched malignancies. Because PD-1 is notexpressed in gliomas, it is unlikely that the anti–PD-1 is exerting adirect effect on the tumor cell. We are unable to exclude the possibilitythat other immune populations are also contributing to the therapeuticeffect of the anti–PD-1 and it is likely that a combination of immunepopulations play a role in activity. Cumulatively, this is the first studyto demonstrate that the anti–PD-1 antibody may promote a proin-flammatory M1 signature within the glioma microenvironment.

The purpose of this study was not to suggest that monotherapywith anti–PD-1 in glioblastoma is effective for all patients withGBM (49). Rather, the purpose of our study was to more fullyunderstand the mechanisms of action of this agent in GBM so thatwemightmore appropriately select rational combinations and identifypotential response biomarkers. To date, immune checkpoint inhibitorresponse biomarkers are surrogates for antitumor T-cell responseswhich include T-cell counts in the tumor microenvironment; tumormutational burden, microsatellite instability, and POLE mutationsas T-cell targets; IFN signatures; and the inhibitory ligand forPD-1–PD-L1. We are also not attempting to refute the contributionof CD4 cytotoxic T cells to these responses (51–53), but are ratherexpanding consideration to PD-1–expressing macrophages andmicroglia as also being involved in the antitumor immune responseswith these agents. As such, future clinical trials using ICIs shouldalso consider including the immune phenotype and function ofmacrophages which may be all the more relevant in glioblastomagiven the predominance of this immune population. Combinatorialstrategies, such as macrophage polarization therapeutics, are beingactively considered with anti–PD-1. However, our data indicatesthis may not be necessary because anti–PD-1 is already eliminatingPD-1–expressing microglia and driving M1 polarization and assuch should be deprioritized.

Because CNS macrophages arise from peripherally derived mono-cytes, the anti–PD-1 antibody need not have penetrated into the CNSto eliminate PD-1þ immune-suppressive macrophages in the gliomamicroenvironment. Most immune cells, including the monocyte-derived macrophages in the tumor microenvironment, arise from theperiphery where they can interact with the anti–PD-1 antibody.However, the elimination of PD-1–expressing microglia, which orig-inate in the CNS during embryogenesis, implies that the anti–PD-1antibody is capable of some degree of CNS penetration, especially intumor regions in which there is breakdown of the BBB. Fluorescently-tagged anti–PD-1 accumulated in the glioma relative to the surround-ing brain, confirming that the anti–PD-1 antibody can gain access tothe glioma microenvironment. Future studies will be directed atassessing the concentrations of the anti–PD-1 antibody in infiltratingnoncontrast-enhancing regions of the brain that do not have appreci-able breakdown of the BBB.

Eliminating the CD8 T cell from the microenvironment of anevolving glioblastoma enabled us to isolate the macrophage as thecell mediating the therapeutic effect of checkpoint inhibition.We werealso able to study how the absence of CD8þ T cells affects gliomadevelopment, malignancy, and survival. A previous study showed thatthe CD8þ T cell is defective and nonoperational in murine models of

spontaneously-arising astrocytomas, even at early stages of tumordevelopment (54). However, this contrasts with recent reports thatlevels of CD8þ tumor-infiltrating lymphocytes are inversely correlatedwith glioma grade and are associated with long-term survival (55).Alternatively, the CD8þT-cell populationmay only have a role duringthe early stages of tumor development (56, 57). Our primary hypoth-esis was that the elimination of the CD8 immune effector populationwould produce an increased incidence of high-grade gliomas resultingfrom the lack of immunologic recognition and control. Because thepresumptive antitumor immune effector population is not present inthe CD8�/� genetic background relative to wild-type mice, the tumor-bearing CD8�/� mice should have had an environment favoringgrowth of tumor cells. Alternatively, without the selective pressure ofCD8 T cells culling tumor cells sensitive to elimination, the moreresistant cells escaping immune surveillance might not proliferate,resulting in some degree of quiescence. If this hypothesis were correct,then there would be increased survival in mice with the CD8�/�

genetic background associated with less malignant progression.Surprisingly, we found that there was no difference in survivaltime and no difference in the incidence of glioma grade between thetwo genotypes. These results show that CD8 T cells do not influencesurvival of mice that undergo de novo glioma formation. Yet, Kaneand colleagues report that while CD8 T cells do not influencesurvival similar to our glioma model, the presence of CD8 T cellsdo influence the tumor genome, phenotype (oncogenes/tumorsuppressors, MAPK activation), and composition of the immuno-logic microenvironment (37).

Another observation from our study was that there was elevatedMHC-II and Iba1 expression associated with the monocyte/macrophage population present in the glioma in the absence of CD8T cells. These cells demonstrated a proinflammatory activated profile.Activated macrophages have been shown to be capable of tumoricidalactivity (58). Similar compensatory findings have been reported in aGL261 glioblastoma model treated with a tumor lysate vaccine withOX40L-Fc stimulant in which the vaccine efficacy was independent ofthe CD8þ T-cell population, but dependent on CD4þ T cells, NK cells,and B cells (59); however, this study did not specifically delve into therole of the proinflammatory macrophage. The macrophages alsodemonstrated immune-suppressive features, including PD-1 expres-sion and elaboration of arginase. Consistent with our findings fromhuman GBM specimens (48), the glioma-infiltrating macrophages inour models demonstrated marked immunologic heterogeneity ofphenotype and function.

A confounder of our study is the possibility that the CD19þ andNK1.1þ populations play a role in the earlier stages of gliomagenesisand the CD8 KO model may have an impact on the development ofother immune cells. However, given the sustained survival in thismodel and the fact that gliomagenesis is induced in the postnatalperiod, it is not technically feasible to perform in vivo depletions inneonatal mice. This study also underscores the importance of genet-ically engineered mouse models in which gliomas are formed de novoin the brain for the study of the immune system, as there was asignificant difference in the infiltration of T cells and macrophagesbetween endogenously forming tumors and orthotopic xenografts.These results and the studies by others (37) indicate that CD8 T cellsmight influence features of glioma development such as genotype,phenotype, immunogenicity, and the microenvironment throughimmunoediting, and that these effector cells might not be responsiblefor responses to anti–PD-1 blockade. This may be related to anunappreciated role of the innate immune system in modulatingmalignant degeneration, as has been previously suggested (60).

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Disclosure of Potential Conflicts of InterestA.M. Sonabend reports personal fees from Abbvie (consulting fees) outside the

submitted work; in addition, A.M. Sonabend is listed as a coinventor of a patentregarding predicting response to immunotherapy in gliomas that is owned byColumbia University. J.P. Long reports grants from Brockman Foundation duringthe conduct of the study. S. Li reports grants from NIH during the conduct of thestudy. M.A. Curran reports personal fees from ImmunoGenesis, ImmunOS, Agenus,Alligator, Aptevo,Mabimmune,Oncoresponse, Pieris, Xencor, andMerck outside thesubmitted work; in addition, M.A. Curran is listed as a co-inventor on four patentslicensed to ImmunoGenesis Inc and owned by MD Anderson Cancer Center relatedto PD-L1 antibodies, PD-L2 antibodies, PD-L1/PD-L2 dual specific antibodies, andPD-L1/PD-L2 bispecific antibodies. No potential conflicts of interest were disclosedby the other authors.

Authors’ ContributionsG. Rao: Conceptualization, resources, data curation, formal analysis, supervision,

funding acquisition, writing-original draft, project administration, writing-reviewand editing. K. Latha: Data curation, writing-review and editing. M. Ott: Datacuration, writing-review and editing. A. Sabbagh: Data curation, writing-review andediting. A. Marisetty: Data curation, writing-review and editing. X. Ling: Datacuration, formal analysis, writing-review and editing. D. Zamler: Data curation,writing-review and editing. T.A. Doucette: Data curation, writing-review andediting. Y. Yang: Data curation, writing-review and editing. L.-Y. Kong: Datacuration, writing-review and editing. J. Wei: Data curation, writing-review and

editing. G.N. Fuller: Data curation, writing-review and editing. F. Benavides:Data curation, writing-review and editing. A.M. Sonabend: Data curation,writing-review and editing. J. Long: Formal analysis, writing-review and editing.S. Li: Writing-review and editing. M. Curran: Data curation, writing-review andediting. A.B. Heimberger: Conceptualization, resources, data curation, supervision,funding acquisition, writing-original draft, writing-review and editing.

AcknowledgmentsWe thank David M. Wildrick, Ph.D., for scientific editing of the manuscript

and Audria Patrick for administrative support. This work was supported by grantsfrom the Brockman Foundation, the Dr. Marnie Rose Foundation, the Ben andCatherine Ivy Foundation, The University of Texas MD Anderson Cancer CenterGBM Moonshot program, and the NIH CA120813, P50 CA127001, and NS094615.This study made use of the Research Animal Support Facility-Smithville (LaboratoryAnimal Genetic Services), which is supported by P30 CA016672 DHHS/NCI CancerCenter Support grant toTheUniversity of TexasMDAndersonCancerCenter and theTLC Foundation.

The costs of publication of this article were defrayed in part by the payment of pagecharges. This article must therefore be hereby marked advertisement in accordancewith 18 U.S.C. Section 1734 solely to indicate this fact.

Received December 19, 2019; revised April 16, 2020; accepted June 11, 2020;published first June 18, 2020.

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Rao et al.




2020;26:4699-4712. Published OnlineFirst June 18, 2020.Clin Cancer Res Ganesh Rao, Khatri Latha, Martina Ott, et al. Cellsand Exerts Therapeutic Efficacy in the Absence of CD8 Cytotoxic T

PD-1 Induces M1 Polarization in the Glioma Microenvironment−Anti

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