Polypharmacologic Reprogramming of Tumor-Associated Macrophages toward an Inflammatory Phenotype

Solid tumors comprise heterogeneous populations of cancer and noncancerous cells that interact through direct contact and secreted factors, thus establishing the tumor microenvironment (TME). TME alters the behavior of noncancerous cells, resulting in phenotypes with tumor-supportive properties. For example, tumor cells and immune cells that reside within TME release a number of anti-inflammatory and proangiogenic factors that result in protumoral polarization of tumor-associated macrophages (TAM), one of the abundant types of immune cells found in TME. Thus, polarized TAMs can further promote tumor progression by stimulating proliferation, invasion, angiogenesis, immunosuppression, metastasis, endowing resistance against chemo- and radiotherapy, and decreased efficacy of immunotherapy (1, 2). Furthermore, clinical evidence supports that patients with higher TAM infiltration have poor prognoses in several cancers (3). Given their role in tumor progression and the clinical evidence, TAMs have been proposed as a promising therapeutic target (4–6). However, physiologic TAMs are scarce and difficult to study using existing experimental strategies. Therefore, mechanisms that drive TAM protumor polarization, as well as potential targeting opportunities for drug development, remain incompletely understood.

Accumulated evidence suggests that TAMs are highly plastic and dynamic cells with complex signaling networks optimized to rapidly adjust the phenotype in response to external stimuli (7). Therefore, TAMs exhibit heterogenous (8) and continuum phenotypes between inflammatory and anti-inflammatory states (9). One strategy to model TME in vitro is to employ tumor conditioned medium (CM) of cancer cells, which has been shown to contain a near-complete set of secreted factors and metabolites found in TME and has been used in several TAM studies aimed at improving our understanding of TAM biology (10–14). However, quantitative, phenotypic drug screening assays aimed at TAMs have not been described before, thus limiting the ability to discover compounds with potential therapeutic benefits in this context. To address this need, we established an in vitro quantitative TAM polarization model using human monocyte cell line THP-1 and CM from multiple cancer models. Using cellular elongation measurement as a quantitative measure of TAM polarization, we screened a library of 85 kinase inhibitors targeting most of the human kinome. We identified BMS-7948933, a dual-targeting inhibitor of c-MET and VEGFR2, as the most potent blocker of cellular elongation and protumoral function of TAMs. BMS-794833 suppressed tumor growth on mouse triple-negative breast cancer (TNBC) models. Surprisingly, we found the TAM-inhibitory function of BMS-794833 does not involve c-MET or VEGFR2 but a range of other targets that include focal adhesion kinases (FAK) and cytoskeletal-related proteins, SRC family kinases, STAT3, and p38 MAPKs. We also observed that targeting a single pathway exhibited little to no effect on TAM polarization, whereas BMS-794833–mediated targeting of multiple signaling pathways exhibited the most potent effect on TAM polarization. Our study underlines the complex regulation of signaling pathways during TAM polarization and the necessity of concomitant blocking of multiple signaling pathways to reprogram protumoral TAM function.

Cell lines including 4T1, CT26, Py8119, ACHN, THP-1, and Jurkat cells were purchased from ATCC. HCC70, HCC1419, and HCC1937 were gifts from Dr. Peggy Porter, Fred Hutchinson Cancer Research Center (FHCRC). 4T1, CT26, HCC70, HCC1419, HCC1937 cells were maintained in RPMI1640 media (Gibco by Thermo Fisher Scientific) supplemented with 10% (v/v) FBS, 1% penicillin–streptomycin (P/S, final concentration: 100 U/mL of penicillin, 100 μg/mL of streptomycin; Gibco). Py8119 cells were maintained in F-12K medium (ATCC) with 5% FBS. THP-1 cells were cultured in RPMI1640 media supplemented with 10% FBS, 1% P/S, 1 mmol/L sodium pyruvate (Lonza), and 55 nmol/L β-mercaptoethanol (Gibco). Jurkat cells were maintained in RPMI1640 media with 10% FBS, 1% P/S, 10 mmol/L HEPES (Santa Cruz Biotechnology), and 1 mmol/L sodium pyruvate. All cells were cultured in humidified 37°C incubators with 5% CO₂ atmosphere.

For the collection of CM from tumor tissue and cancer cells, serum-free tumor slice culture (TSC) medium (15, 16) was used. Cancer cells grown to subconfluent were washed with PBS twice and incubated in TSC medium for 1–2 days. For the collection of CM from tumor tissues, tumors were cut into 4–5 mm cubes, washed with PBS twice, and incubated with 10 mL of TSC medium per 1 g tumor for one day. After collecting the spent medium, fresh TSC medium was added to the tumor pieces to collect the second pool. The collected medium was centrifuged at 2,000 × g for 10 minutes, and the supernatant was used as CM. TSC medium without conditioning was used as an experimental control. For the collection of TAM CM, TAMs polarized for 3 days were washed with PBS twice, then incubated for 24 hours in RPMI1640 medium for THP-1 maintenance. The supernatant was centrifuged at 2,000 × g for 10 minutes, to remove cell debris. RPMI1640 medium for THP-1 culture without conditioning was used for experimental control.

THP-1 cells were differentiated into macrophages by inducing with phorbol 12-myristate 13-acetate (PMA, LC Laboratories) at 25 ng/mL for one day. The cell densities used were; 1.3–1.5 × 10⁴/well for 96-well plate, 4 × 10⁵ cells/well for 6-well plate, 2.4 × 10⁶ cells for 10-cm dish. The medium was replaced with medium containing cancer CM at 25%–50% (v/v) with volume adjustment to 50% with TSC medium, and 50% (v/v) RPMI1640 for THP-1 cell culture. Cellular elongation was measured with NeuroTrack analysis software accompanied with IncuCyte Zoom Live Imaging system (Sartorius).

THP-1 cells were seeded at 1.5 × 10⁴ cells per well of 96-well plates with PMA at 25 ng/mL. After a day, the medium was replaced to CM mix (25% 4T1 cell CM, 25% TSC medium, with 50% RPMI1640 for THP-1 culture, YOYO-3 at 1:10,000 dilution) together with kinase inhibitors at 8 serial concentrations of 10, 3.3, 1.1, 0.37, 0.12, 0.04, 0.01, and 0 μmol/L with triplicates. The 85 inhibitors tested are listed in Supplementary Table S1. All small molecules were constituted in DMSO for the stock solution, and DMSO (0.1%, up to 1% based on the volume of inhibitor solution) was supplemented as vehicle control. A red fluorescent viability dye, YOYO-3 (Thermo Fisher Scientific) was supplemented to the culture to detect cellular death. The phase contrast and red fluorescent images were taken every 2 hours over 3–5 days using IncuCyte Zoom instrument. The target kinase profiles of BMS-794833 in the acellular system were described in Rata and colleagues (17).

THP-1 cells at 1.5 × 10⁴ cells per well of 96-well plate were induced with PMA at 25 ng/mL and polarized for 3 days with the indicated tumor CM with or without inhibitors. The TAMs were washed with PBS twice, and 4T1-nucGFP cells were seeded at 3 × 10³ cells per well density with RPMI1640 supplemented with 0.5% FBS. The cell numbers of 4T1 were counted on the basis of nuclear GFP using the software accompanied with IncuCyte Zoom.

Tube formation assay of endothelial cells was performed using IncuCyte Angiogenesis 96-well PrimeKit Assay (Sartorius) with modifications. For the angiogenesis assay of TAM CM, the cells were seeded following the manufacturer's protocol. Three days postseeding, TAM CM was supplemented at 50% concentration diluted in the assay medium included in the kit. The cells were incubated for 6 days with a medium replacement on day 2 and 5 during the culture. The plate was scanned and analyzed with IncuCyte Zoom. For assay of BMS-794833–treated TAM CM, human primary fibroblasts (NHDF) and GFP-expressing human umbilical vein endothelial cells (GFP-HUVEC) were minimally expanded. NHDFs were seeded at 1 × 10⁴ cells per well of 96-well plate with DMEM with 10% FBS and 1% P/S and incubated for 2 days till confluent. GFP-HUVECs were seeded at 2 × 10³ cells per well in EGM-2 on top of the NHDF layer. On the next day, the medium was replaced with 50% TAM CM diluted in EGM-2, and cells were incubated for 6 days. A half volume of medium was replated every 2 days. The plate was scanned and analyzed with IncuCyte S3.

Jurkat cells were washed with PBS and seeded onto a 24-well transwell culture insert with 3-μm pores (Celltreat Scientific Products) at 5 × 10⁵ cells/well with RPMI1640 supplemented with 2.5% FBS. The inserts were placed onto wells containing TAM CM or control RPMI1640 medium for THP-1, and incubated for 19 hours. The relative number of cells migrated to the bottom chamber was quantified using Celltiter Glo (Promega). The remaining cells in the inserts were analyzed with Caspase-Glo 3/7 assay (Promega) to confirm that Jurkat cells incubated with TAM CM did not induce apoptosis compared with the control medium.

All animal studies were approved by the Institutional Animal Care and Use Committee of FHCRC. BALB/c and C57BL/6J females were sourced from The Jackson Laboratory. NSG female mice were inbred in-house by Comparative Medicine of FHCRC. 4T1 cells or Py8119 cells suspended at 1–2 million/150 μL of 33% (v/v) Matrigel (Thermo Fisher Scientific) were transplanted subcutaneously to the right flank of 6–8 week-old BALB/c and NSG, or C57BL/6J female mice, respectively. After tumors reached 50–100 mm³, mice were randomized into treatment and untreated groups. The mice received an intratumoral injection of BMS-794833 at 25 mg/kg dose, BMS-5 at 30 mg/kg dose, or vehicle control twice weekly. BMS-794833 (Selleckchem) was dissolved into 4% (v/v) DMSO (Sigma-Aldrich), 45% (v/v) PEG300 (Sigma-Aldrich), and 5% (v/v) Tween 80 (Fisher Scientific). BMS-5 (Medchem Express) was dissolved into 5% DMSO, 30% PEG400, 5% propylene glycol, and 0.5% Tween 80. The tumor size was measured by caliper twice weekly and by weight at the experimental endpoint. The tumor volumes were calculated with the following equation: V = 4/3π×(L/2)×(W/2)×(H/2), (L = length, W = width, H = height). HCI010 model was kindly gifted from Dr. Alana Welm (University of Utah, Salt Lake City, UT) and expanded orthotopically in NSG female mice as described before (18). Py8119 tumors used for CM collection used in THP-1–derived TAM polarization were prepared by injecting Py8119 cells subcutaneously to NSG female mice.

At the end of the animal study, tumors and lungs were harvested from mice and fixed in 10% neutral-buffered formalin (Millipore Sigma) for 7 days. The tissues were embedded into paraffin blocks and cut into sections. Hematoxylin and eosin staining of lung sections was performed by the Histopathology core at the Fred Hutchinson Cancer Research Center (Seattle, WA). The entire tissue was scanned with TissueFAXS (TissueGnostics). The number and area of metastatic foci were evaluated using QuPath software (19). For detection and characterization of macrophages, the staining and analyses were performed by the Histopathology core at the Fred Hutchinson Cancer Research Center using ARG1, F4/80, and iNOS as primary antibodies.

All the statistical analyses were performed using Graphpad Prism 8.0.1 software. The statistical analyses used for each data are indicated in the figure legends (*, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001). Heat maps were prepared using pheatmap package in R Studio.

For more detailed methods, see Supplementary Materials and Methods.

To develop a physiologically relevant in vitro TAM polarization model, we exposed macrophages differentiated from THP-1 cells, a human monocyte cell line, to tumor CM collected from multiple tumor models (Fig. 1A). CM from the mouse TNBC cell line 4T1 induced protrusions in the THP-1–derived macrophages, causing them to exhibit drastically elongated cellular morphology starting from day 2 with peaking at around day 4 (Fig. 1B). Given that previous studies have shown that TAM cellular elongation correlates strongly with protumor phenotypes (10, 12–14, 20–23), we adopted cellular elongation quantified by live-cell imaging as an indicator of TAM polarization (Fig. 1A). We then assessed whether the TAM polarization model reproduces the molecular characteristics of TAMs in vivo using a gene set panel adapted from ref. 8. Gene expression profiles of macrophages induced by the CM from 4T1 cells upregulated TAM-associated genes and temporally induced inflammatory marker genes compared with macrophages cultured without 4T1 CM (Fig. 1C). The TAM-related secretory proteins were also detected from the 4T1 CM–induced TAMs (Supplementary Fig. S1A). Consistent with previous reports (24), the TAM model induced both inflammatory (CCL1, Groα, IL1β, IL6, IL8, IL12, MIP-1α, MIP-1β, CCL5, TNFα, sICAM) and anti-inflammatory cytokines (IL1RA, IL4, IL10, SerpinE1) to 1.4–13.2 and 1.2–1.8 folds, respectively, over control macrophages (Supplementary Fig. S1A). Similar cellular elongation during TAM polarization was observed with CM collected from other breast cancer models (4T1 tumor, Py8119 tumor, a Ras-expressing human mammary epithelial cells HMLE-Ras, breast cancer PDX models, human breast cancer cells: HCC70, HCC1419, HCC1937, HCC70 tumor; Fig. 1D, top), melanoma (B16.F10.Ova), pancreas cancer (Panc02), colon cancer (MC38 and CT26), and human renal carcinoma (ACHN; Fig. 1D, bottom). In contrast, CM from untransformed HMLE cells, NHDF, mouse hepatocytes (AML12), and mouse fibroblast (3T3) did not polarize TAMs as effectively as cancer CM (Fig. 1D, top), except for human embryonic kidney cells (HEK293T) that partially elongated cells. These results provide validation that our tumor CM-induced TAM polarization system is reliable in vitro model for studying TAM function.

To further investigate the protumoral behavior of our TAM model and further validate that this model recapitulates in vivo physiology of TAMs, we performed a series of coculture experiments. Coculture of 4T1 breast cancer cells with breast cancer CM-induced TAMs stimulated cancer cell proliferation, recapitulating previously reported growth stimulus effects of TAMs (Fig. 2A). Similar cancer cell growth stimulation by TAM was observed with TAMs induced by CM from other cancer models (melanoma, pancreas, and colon cancer; Fig. 2B). To assess proangiogenesis ability of TAM, we treated HUVECs with TAM CM that was enriched with secretion factors from polarized TAMs, and monitored the ability of HUVECs forming tube-like structures. Our data show that TAM CM exposure stimulates both length and network branching of endothelial cells (Fig. 2C). It is known that TAMs impede T-cell infiltration into the tumor by secretory factors (25–27). Consistently, TAM CM suppressed Jurkat cell migration in a transwell migration assay compared with macrophage CM (Fig. 2D). Together, these results further validate our in vitro TAM polarization model as the system recapitulates in vivo ability of TAMs to promote cell proliferation and angiogenesis and suppress T-cell chemotaxis. Therefore, this model offers an opportunity to conduct screening campaigns to identify compounds that reprogram TAMs with protumoral traits into antitumoral traits.

Protein kinases (kinases) are critical components of cellular signaling networks that transmit extra/intracellular stimuli to cellular responses. Given that TAM polarization is an external stimuli–regulated process, we hypothesized that kinases will play a major role in this process, and therefore, kinase inhibitors may represent useful pharmacologic agents to inhibit protumoral TAM polarization. To examine this further, we used our model system and cellular elongation as an indicator of TAM polarization levels to screen a curated collection of 85 kinase inhibitors (Fig. 3A) that achieves broad kinome coverage (289 kinases out of 298 kinases measured), with lower than 50% residual activities at 0.5 μmol/L (Fig. 3B; ref. 28). We tested this inhibitor collection at eight serially diluted concentrations (0–10 μmol/L) on our TAM model using THP-1 cells induced with CM from 4T1 cultured cells (Fig. 3A). The efficacy of each inhibitor on TAM polarization was quantified as inhibition of cellular elongation compared with vehicle control. To omit inhibitors causing cellular death, we supplemented the assay with YOYO-3, a red fluorescent dye that stains membrane-compromised cells. The response to each inhibitor was evaluated at the highest concentration ranging from 0 to 10 μmol/L that did not exhibit cellular death. As a result, of the 85 tested inhibitors, 33 inhibitors caused a reduction in elongation by 3%–93%, 26 inhibitors showed no change and/or induced cellular death, and surprisingly, 26 caused an increase in elongation by 14%–225% (Fig. 3C; Supplementary Table S1). Sorafenib, a multikinase inhibitor of Raf, VEGFR, and PDGFR, has been previously shown to inhibit M2 (anti-inflammatory) macrophage phenotype and restore inflammatory cytokine expression (29–31). In our assay, sorafenib efficiently suppressed cellular elongation at 3.3 μmol/L or higher, with 81.5% inhibition at 10 μmol/L (Fig. 3A) and was the fourth-most effective inhibitor (Fig. 3C), validating the reliability of the screening system.

Among the tested inhibitors, BMS-7948333, a dual inhibitor of c-MET and VEGFR2 (32), was the most potent inhibitor of cellular elongation during TAM polarization, with 93% inhibition at 10 μmol/L with no evidence of cell death (Fig. 3D; Supplementary Fig. S1B). We confirmed the inhibitory effect of BMS-794833 on cellular elongation in our other TAM polarization models using breast cancer CM (4T1 tumor, Py8119 tumor, and PDX) as well as CM from other cancer models (Fig. 3D; Supplementary Fig. S1C). Time-course transcriptional analyses of BMS-794833–treated cells showed a marked decrease in expression of the anti-inflammatory macrophage marker (MRC1/CD206), and enhanced expressions of inflammatory cytokine-coding genes, such as IL1B, CCL2, CCL4, IL12A, IL6, and TNFA (Fig. 3E). Cytokines secreted from BMS-794833–treated TAMs were also measured using protein arrays and absolute quantification using Luminex (Supplementary Fig. S1D). BMS-794833 treatment suppressed secretion of anti-inflammatory cytokines IL4, IL13, and IL10, and enhanced inflammatory cytokines such as IL6, IL8, IL27, MIP-1α/β, CCL2, TNFα, and IL1β. These results demonstrate that BMS-794833 could abrogate TAM polarization and reprogram TAMs to proinflammatory phenotypes.

We next investigated the effect of BMS-794833 on the protumoral activities of TAMs. BMS-7949833 treatment decreased the growth-stimulatory effect on 4T1 cells following coculture with 4T1 or Py8119 tumor CM–induced TAMs (Fig. 3F), as well as other cancer models (Supplementary Fig. S1E). In addition, CM collected from BMS-794833–treated TAMs reduced angiogenesis stimulation potential and increased T-cell chemotaxis ability compared with untreated TAMs (Fig. 3G and H). Coculturing with BMS-794833–treated TAMs increased expression of PDCD1 (PD-1–coding gene) and Jurkat cell adhesion, indicating stimulation of inflammatory response (Supplementary Fig. S1F). We further validated the inhibitory effect of BMS-794833 on TAM polarization models prepared using bone marrow–derived cells (BMDC) from BALB/c and C57BL/6J mice induced with CM from 4T1 and Py8119 tumors, respectively. In agreement with our previous results, BMDC-derived TAMs exhibit elongated morphology and induced expression of TAM-related anti-inflammatory genes, and BMS-794833 treatment suppressed both cellular elongation and expression of these genes (Supplementary Fig. S1G and S1H). Taken together, these results demonstrate that BMS-794833 identified through the phenotypic screen can impede TAMs from enacting their protumoral functions in vitro and reprogram them into more proinflammatory phenotypes.

We next investigated the effect of BMS-794833 on primary breast tumor growth in mice. First, we evaluated the effect of BMS-794833 on the proliferation of breast cancer cells in monoculture. BMS-794833 inhibited cellular growth of 4T1 breast cancer cells by 28% at 10 μmol/L (Fig. 4A). Py8119 cells were slightly more sensitive, with IC₅₀ at 4.1 μmol/L and inhibition by 87% at 10 μmol/L. These results showed that BMS-794833 has a modest inhibitory effect on breast cancer cell growth. Next, we administered BMS-794833 to 4T1 tumor transplanted subcutaneously in BALB/c mice. Biweekly treatment of BMS-794833 at 25 mg/kg dose intratumorally significantly reduced tumor weight by 52% compared with vehicle control without a decrease of body weight (Fig. 4B; Supplementary Fig. S2A). The tumor-suppressive effect of BMS-794833 on 4T1 was similar in immunodeficient NSG mice (Supplementary Fig. S2B). Reduction of tumor growth was also observed in Py8119 tumors grown subcutaneously in C57BL/6J mice with no obvious systemic toxicity (Fig. 4C; Supplementary Fig. S2A). IHC of cleaved caspase-3 in BMS-794833–treated tumors revealed that BMS-794833 treatment caused an increased trend of apoptosis (Supplementary Fig. S2C). In addition, BMS-794833–treated 4T1 tumors developed fewer microvessels (Supplementary Fig. S2D). The BMS-794833 treatment also decreased lung metastasis in the 4T1 tumor model (Fig. 4D). Py8119-bearing mice did not develop visible lung metastatic foci in either control or treated tumors within the experimental period. These results demonstrate that BMS-794833 suppresses tumor growth and metastasis, and tumor suppression is likely due to its effect on cancer and stromal cells.

To investigate whether BMS-794833 treatment impacts the profiles and characteristics of TAMs and other immune cells, we analyzed 4T1 and Py8119 tumors treated or untreated with BMS-794833 by flow cytometry (Supplementary Fig. S3A). BMS-794833 caused a significant decrease in live cells (Supplementary Fig. S3B), consistent with cleaved caspase-3 staining observed in tumor sections (Supplementary Fig. S2C). In both tumor models, BMS-794833 treatment did not change the percentage of overall immune populations (CD45⁺ cells; Supplementary Fig. S3C), myeloid populations (CD45⁺CD11b⁺ cells), and their most marker expressions (Supplementary Fig. S3D–S3H), and most lymphoid populations (Supplementary Fig. S3I). TAM population (CD45⁺CD11b⁺Ly6G⁻Ly6C⁻F4/80⁺) exhibited a slight decrease under treatment but was not statistically significant (Supplementary Fig. S3E). The most prominent change in population was in monocytes, defined by Ly6C immature marker, which was significantly increased in 4T1 and less significantly in the Py8119 model (Supplementary Fig. S3E). Blood-derived monocytes are a major source of TAMs, continuously replenishing TAM in tumor tissues (33). An increase in monocytes may be due to enhanced monocyte recruitment by tissue-regenerative responses induced by cellular death, increased chemoattractants in cancer and stromal cells (such as CCL2 and MIP-1α/β increased in THP-1–derived TAM by BMS-794833 treatment; Fig. 3E; Supplementary Fig. S1D), or inhibition of polarization into TAMs. To investigate whether BMS-794833 causes incomplete TAM polarization, surface markers of THP-1–derived TAMs treated or untreated with BMS-7948333 were analyzed. Untreated TAMs exhibited increased expression of CD11b (a myeloid cell marker), CD68 (a pan-macrophage marker), and CD14 (monocytes/macrophage marker) compared with the parental THP-1 cells (Supplementary Fig. S2E). BMS-794833–treated TAMs retained a similar level of CD11b⁺ cells, but decreased CD68 and CD14 expression at the cell surface. These results suggest that the BMS-794833 has inhibitory roles on complete differentiation/polarization of monocytes to TAM.

Next, we investigated gene expression of TAMs and their precursor cells, monocytes, in BMS-794833–treated tumors. The TAM and Ly6C⁺ monocyte populations of 4T1 and Py8119 tumors were isolated through FACS (Supplementary Fig. S2F and S2G). Gene expression analyses of TAMs isolated from BMS-794833–treated 4T1 tumors significantly upregulated inflammatory marker genes (Cxcl10, Tnf), and suppressed protumoral gene expression (Chi3l3; Fig. 4E). Ly6C⁺ monocytes from BMS-794833–treated 4T1 tumors exhibited a similar decrease in the anti-inflammatory Chi3l3 and increase in inflammatory markers, but to a lesser extent (Supplementary Fig. S2H). In the Py8119 tumors, the gene expression of TAM and Ly6C⁺ monocytes were less significant; neither TAM nor monocytes of Py8119 tumor induce TNFα, but notably, TAM increased expression of Nos gene, which encodes inducible NO synthase causing cytotoxicity (Supplementary Fig. S2H). These results demonstrate that BMS-794833 induced inflammatory phenotypes in intratumoral TAMs and Ly6C⁺ monocytes. We further analyzed the distribution and activation status of TAMs by multiplex IHC (Fig. 4F). TAMs in both treated and untreated tumors were distributed throughout the tissues as clusters. Total TAMs (F4/80⁺), anti-inflammatory (F4/80⁺Arg1⁺), and inflammatory (F4/80⁺iNOS⁺) TAMs were quantified within the intact tissue area. The total TAM distribution remained at the same level, and inflammatory TAMs were scarce, but anti-inflammatory TAMs showed a decreasing trend in BMS-794833–treated tumors (Fig. 4F). These results further support that BMS-794833 suppressed anti-inflammatory phenotypes of TAMs in vivo.

We sought to investigate the mechanism by which BMS-794833 impairs TAM polarization. Although c-MET and VEGFR2 are considered to be two main targets of BMS-794833, in vitro target profiling revealed that BMS-794833 inhibited 29 kinases to below 50% residual activity at 0.5 μmol/L (Fig. 5A; Supplementary Fig. S4A; Supplementary Table S2). To examine whether c-MET and VEGFR2 are responsible for inhibition in TAM polarization, we first tested selective inhibitors for c-MET and VEGFR2, JNJ-38877605 and cediranib, respectively, on the TAM model. Neither compound inhibited TAM elongation (Supplementary Fig. S4B), suggesting that the inhibitory effect of BMS-794833 results from inhibition of either another single target or a polypharmacologic effect on multiple targets. To test this further, we also inhibited the top 18 kinases targeted by BMS-794833 using selective inhibitors, but a majority of these inhibitors were not as potent as BMS-794833 (Fig. 5B). Only inhibitors chemically-analogous to BMS-794833 showed a similar level (cabozantinib) or less (BMS-777607) inhibition on cellular elongation during TAM polarization (Supplementary Fig. S4C). Cabozantinib is newly recommended as standard-of-care combined with nivolumab for renal cell carcinoma (34). We validated that CM from human renal carcinoma, ACHN cell line, polarized THP-1–derived TAMs (Fig. 1D), and cabozantinib inhibited cellular elongation but was less potent than BMS-794833 (Supplementary Fig. S4D). These results suggest that targeting a single kinase is insufficient to suppress TAM elongation, and that BMS-794833 was the most potent inhibitor to inhibit multiple signaling pathways to effectively block TAM polarization.

To characterize the effect of BMS-794833 further, we performed a phosphorylated tyrosine kinase array (Supplementary Fig. S4E and S4F) and reverse-phase protein array (RPPA) screening followed by Western blot validation (Fig. 5C). TAMs polarized with 4T1 CM were treated with BMS-794833 for full time (3 days) or the last 30 minutes of the polarization period. BMS-794833 treatment decreased phosphorylation of proteins related to focal adhesion–cytoskeleton regulation, SRC family kinases, JAK-STAT3, p38 MAPK, AKT, and NFκB (Fig. 5D–F). The results also revealed differential response at the time of inhibition by BMS-794833; phosphorylation of FAK, PYK, cofilin, and p38 MAPK can be inhibited at the last 30 minutes of the polarization period, whereas total LIMK1, phosphorylated STAT3, AKT, and NFκB were downregulated only upon full-time treatment. In detail, phosphorylated FAK, PYK2, and cofilin decreased within 30 minutes and remained low after 3 days of BMS-794833 treatment, while the total protein level of LIMK1, an upstream kinase that phosphorylates cofilin (35), decreased after 3 days (Fig. 5D). Among the 9 SRC family kinases (36), THP-1–derived TAMs expressed HCK, FGR, LYN, FYN, and a low amount of c-SRC (Fig. 5E). Phosphorylation of total SRC family kinases was induced in TAM polarization and decreased by 3-day BMS-794833 treatment (Fig. 5E). Phosphorylated STAT3, a transcription regulator of immunosuppressive cytokines, MMPs, and angiogenesis factors (7) was decreased significantly after long-term BMS-794833 treatment (Fig. 5F). The phosphorylation level of p38 MAPK, a conditional regulator of pro- or anti-inflammatory response (37), dropped within 30 minutes and also decreased total protein amount after 3 days. Phosphorylated ERK1/2 was increased by BMS-794833 treatment within 30 minutes and remain high on 3 days (Fig. 5F). However, in the validation experiment with mouse BMDC-derived TAMs, phosphorylation of Erk1/2 and Mek1/2 decreased, indicating that the increase of phosphorylated ERK is a THP-1–specific observation (Fig. 5G). Downregulation of phosphorylated Src family, Pyk2, Cofilin, Stat3, and Akt by BMS-794833 were validated in the mouse BMDC-derived TAM model (Fig. 5G).

As TAMs dynamically regulate signaling activation over time, we further investigated time-dependent activation of signaling pathways during TAM polarization, leveraging the advantage of our in vitro system (Supplementary Fig. S4G). We observed an increase in the phosphorylation of AKT, ERK1/2, and p38 MAPK within 1 hour in response to 4T1 tumor CM, followed by an increase in the phosphorylation of NFκB, which peaked at 24 hours, indicating activation of these proteins is an early event in TAM polarization. In contrast, phosphorylation of FAK, cofilin, SRC family, and STAT3 was increased from day 2, indicating that these proteins are activated in a late event. Importantly, BMS-794833 treatment suppressed induction of p38 MAPK, NFκB, FAK, Cofilin, SRC family, and STAT3, suggesting that BMS-794833 targets both early and late-stage protein signaling during TAM polarization. Together, these data highlight temporal activation of multiple pathways during TAM polarization, and the effect of BMS-794833 on these signal cascades.

TAMs exhibit dynamic changes in gene expression and protein phosphorylation during the polarization period (Figs. 1, 3, 5). To evaluate relative contributions of each change to TAM polarization and to the response to BMS-794833, we performed partial least square regression (PLS-R) analysis of time-course gene expression and protein phosphorylation of TAMs treated or untreated with BMS-794833, with cellular elongation as a response (Supplementary Fig. S5). Component 1 (40% explained variance) delineated time-dependent TAM polarization, and component 2 (19% explained variance) discriminated BMS-794833 treated and untreated groups (Supplementary Fig. S5A). The top 30 standardized coefficient of component 2 includes positive coefficients (toward BMS-794833–treated group) of inflammatory cytokine-coding genes and negative coefficients (toward untreated group) of phosphorylated STAT3, cofilin, p38 MAPK, and total LIMK1 (Supplementary Fig. S5B and S5C). These results indicate that the effect of BMS-794833 can be explained by both increases in inflammatory cytokine expression and a decrease in phosphorylated/total proteins involve in TAM polarization.

To investigate the contribution of the individual signaling pathway in TAM polarization, we tested selective inhibitors for FAK, LIMK1, SRC family, and STAT3 (Fig. 6A). Two potent FAK inhibitors (PF00562271 and NVP-TAE226) showed an inconsistent effect on TAM elongation and did not provide conclusive evidence for the importance of FAK. A highly selective LIMK inhibitor BMS-5 inhibited cell elongation, consistent with the role of LIMK1 in actin polymerization. LIMK1 is predominantly expressed in the TAM population of breast cancer patient tumors (Supplementary Fig. S6A). BMS-5 treatment upregulated secretion of inflammatory cytokines (Supplementary Fig. S6B) and lowered the growth stimulation effect of TAMs (Supplementary Fig. S6C). However, BMS-5 did not show any tumor-suppressive effect on 4T1 tumor–bearing mice (Supplementary Fig. S6D). Western blot analyses revealed that BMS-5 suppresses phosphorylation of FAK, cofilin, and SRC family kinases, but phosphorylation of STAT3 remained unchanged (Fig. 6B). A STAT3 inhibitor, Stattic (38), had a modest effect on TAM elongation at 10 μmol/L (Fig. 6A). The immunoblot confirmed suppressed phosphorylation of STAT3 (Fig. 6B), and also showed decreased phosphorylated SRC. However, phosphorylation of FAK, cofilin, and total LIMK remained unchanged. Src inhibitor-1, which inhibits most SRC family kinase, stimulated cellular elongation (Fig. 6A). Immunoblot showed that Src inhibitor-1 induces phosphorylated SRC, FAK, and STAT3 within 30 minutes, and increased LIMK1 expression after 3 days (Fig. 6B). The results indicate that SRC signal blockade results in compensatory induction of SRC and other TAM-related signals.

As both BMS-5 and Stattic could inhibit TAM elongation with complementary inhibition of phosphorylated cofilin and STAT3, we tested whether the combination treatment of BMS-5 and Stattic potentiate inhibitor effect on TAM polarization. A combination of BMS-5 and Stattic suppressed TAM elongation more potently than either BMS-5 or Stattic alone at the same dose (5 μmol/L; Supplementary Fig. S6E). The Bliss combination index (39) was 1.02, suggesting that combination treatment resulted in an additive effect. These results indicate that targeting both LIMK1actin-related signal and STAT3 better suppress TAM polarization.

Overall, the results above suggest that in vitro TAM polarization assay system enables investigations of the state of signaling pathways during polarization, and our findings indicate targeting a single kinase/pathway is not sufficient to block TAM polarization. The results reiterate that the potency of BMS-794833 is due to its multitargeting of SRC, actin regulation, and STAT3, inactivating surrogate pathways that support the plastic nature of TAMs.

TAMs receive aggregated signals in the form of growth factors, cytokines, chemokines, extracellular vesicles, and metabolites accumulated in TME. How TAMs process this multitude of signals to alter their morphology and function to support the tumor growth and spread remains poorly understood. Current in vitro model systems that rely on cytokine-supplemented cultures, such as IL4 and IL13, do not adequately recapitulate the complex nature of TME to study TAM function (40). To overcome this limitation, we used media collected from tumor tissues, tumor CM, to induce a complex signaling and phenotypic response through a mixture of secreted factors contained in the CM. Although our in vitro system does not model specific cellular interactions that may impact TAM polarization, we confirmed that tumor CM was sufficient to induce protumoral phenotypes known in TAMs. The monocyte cell line and cancer CM-based system provided a relatively uniform and accessible source for TAMs that are compatible with phenotypic drug screening and functional characterization. Using this model system, we could demonstrate interactions between in vitro–generated TAMs and (i) cancer cells that led to increased cancer cell growth and motility, (ii) endothelial cells leading to the promotion of microvessel formation, and (iii) T cells causing inhibition of T-cell migration. Collectively, we present a physiologically relevant model system to study TAMs.

To document the signaling cascades that regulate complex TAM function, we performed molecular characterization of our in vitro–derived TAMs. We observed alterations of both gene expression and phosphorylation of proteins previously known to be involved in TAM function, including VEGFA, FN1, CXCR4, IL1B (8), CCL20 (41), phosphorylated STAT3 (42), AKT (43), and NFκB (44). The in vitro systems traced the TAM polarization process and allowed us to study time-dependent changes in multiple signaling pathways during the polarization (Supplementary Fig. S4G). Proinflammatory transcription factor NF-κB was phosphorylated on day 1, which was then downregulated at later time points, suggesting that tumor CM first induces inflammatory responses in TAMs then shift toward anti-inflammatory phenotypes. In contrast, STAT3 was unchanged in earlier time points but phosphorylated at later time points. These results describe serially programmed signaling cascades during TAM polarization and suggest that reprogramming this process may offer an opportunity for inhibiting the protumor activity of TAMs.

A combination of rich and uniform sources of in vitro–generated TAMs and robust and quantitative cellular elongation phenotype provides a reliable assay for high-throughput screening. To identify compounds that could compromise TAM function, we carried out a chemical inhibitor screen using a set of 85 kinase inhibitors covering most of the kinome. We identified BMS-794833 as a potent suppressor of cellular elongation in multiple cancer models, and suppresses protumoral activities of TAMs in vitro, underscoring the potential utility of our model system for phenotypic-based screening in drug discovery. The mechanism of action of BMS-794833 involves inhibition of phosphorylation status of multiple proteins, including FAK, SRC, p38 MAPK, and STAT3 (Figs. 5 and 6C). When administered to breast tumor–bearing mice, BMS-794833 could suppress tumor growth and metastasis (Fig. 4B–D). Although BMS-794833 programs TAMs toward more antitumoral phenotypes in vivo (Figs. 4E and F; Supplementary Fig. S3E), the tumor-suppressive effect of BMS-794833 seems primarily derived from tumoricidal activity and effects on nonimmune stromal cells, including endothelial cells (Fig. 4B and D; Supplementary Fig. S2B-S2D). Thus, a single agent, like BMS-794833, that targets multiple cell types in the tumor microenvironment could be desirable in clinical settings.

Several TAM-targeting therapies are currently in clinical studies, including CSF1–CSF1R, either as monotherapy or in combination with conventional treatment to augment therapeutic effects (45, 46). However, clinical studies have shown that monotherapies exhibit moderate effects (47), indicating the limited efficacy of targeted therapies in a complex environment. Consistently, our data showed that targeting single kinases was ineffective in inhibiting cellular elongation–based TAM polarization (Fig. 6A) while BMS-794833, which exhibits broad polypharmacology, effectively inhibits TAM polarization (Fig. 3D). Collectively, these data suggest simultaneous targeting of multiple signaling pathways in TAM polarization yields more effective treatment options than the use of more selective agents.

Overall, our study highlights the complex interplay of macrophage polarization with components of the TME. Broadly, our study provides the cancer community with a platform that allows analyses of TAMs under near-physiologic conditions. As we enter a new era of data-rich cancer biology, one of the primary challenges is integrating the knowledge of cellular and molecular information into a holistic understanding of cancer as a biological system. Quantitative assays and technologies that enable pharmacologic assessment with cellular and molecular phenotypes in the physiologically relevant environment, like the one described in this study, will form an essential component of system-based investigations.

T.S. Gujral reports grants from American Cancer Society, Concern Foundation, and grants from Breast Cancer Research Foundation during the conduct of the study. No disclosures were reported by the other authors.

N. Nishida-Aoki: Data curation, formal analysis, investigation, visualization, methodology, writing–original draft, writing-review and editing. T.S. Gujral: Conceptualization, resources, supervision, funding acquisition, visualization, methodology, writing–original draft, writing–review and editing.

This work was supported by grants from the Breast Cancer Research Foundation (BCRF 17-035) and the American Cancer Society (133870-RSG-19-197-01-CDD). N. Nishida-Aoki is a recipient of the Fred Hutch Interdisciplinary Training Grant in Cancer Research and the Japan Society for the Promotion of Science Overseas Research Fellowship. This research was supported by the Comparative Medicine, Scientific Imaging, and Flow Cytometry, and Experimental Histopathology Shared Resources of the Fred Hutch/University of Washington Cancer Consortium (P30 CA015704). The authors thank Drs. Milka Kostic, Thomas Bello, and Aleena Arakaki, and members of the Gujral lab for helpful comments and suggestions for the manuscript.

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