Transthyretin Stimulates Tumor Growth through Regulation of Tumor, Immune, and Endothelial Cells

Lung cancer is a very aggressive malignant form of cancer and is one of the biggest public health challenges facing the United States and many other countries. Although incidence rates have been stabilized, an estimated 154,050 Americans are expected to die of lung cancer in 2018, accounting for ∼25.3% of all cancer deaths (https://www.cancer.org/cancer/non-small-cell-lung-cancer/about/key-statistics.html). According to the World Health Organization, around 1.37 million people die of lung cancer each year worldwide (http://www.who.int/mediacentre/factsheets/fs297/en/). Lung cancer is by far the leading cause of cancer death among both men and women. Each year, more people die of lung cancer than of colon, breast, and prostate cancers combined. Lung cancer is a difficult disease to detect in its early stages, with >50% of patients diagnosed with lung cancer presenting with metastatic disease (http://seer.cancer.gov/statfacts/html/lungb.html). Early detection of lung cancer is an important opportunity for decreasing mortality while it is still treatable and curable (1). The overall 5-y survival rate is ∼15%. Thus, it is essential to better understand the mechanisms that initiate lung carcinogenesis and find easy-use biomarkers for more accurate lung cancer detection. Due to heterogeneity of lung cancers, a panel of biomarkers should be used for more accurate lung cancer detection and classification.

Signal transducer and activator of transcription 3 (Stat3) is well known for its lung cancer–promoting activity (2–4). The Stat3 expression level was upregulated in human lung cancers (5). To assess the consequences of STAT3 persistent activation in the lung, a doxycycline-controlled CCSP-rtTA/(tetO)₇-Stat3C bitransgenic mouse model was generated that overexpresses STAT3C (a constitutively active form of STAT3) in alveolar type II (AT II) epithelial cells. In sequential steps, Stat3C overexpression upregulated proinflammatory molecules, increased inflammatory cell infiltration, and caused adenocarcinomas in the lung (2). The GeneChip microarray analysis of lung tumor from the CCSP-rtTA/(tetO)₇Stat3C mice revealed around 800 up- and downregulated genes as potential lung cancer biomarkers, with at least 2-fold expression changes (p < 0.05) (2). Because most of these genes are intracellular proteins, it is inconvenient to use them for the purpose of clinical diagnosis without going through biopsy.

In this study, we report identification of 13 soluble and secretory proteins, which were selected from the Stat3 downstream gene list with 2-fold increase (p < 0.05) in lung tumors as a panel of biomarkers for lung cancer detection in humans using the sera. This panel of biomarkers can potentially be used to differentiate different types of lung cancers. To elucidate tumorigenic functions of these biomarkers, one of 13 protein biomarkers, transthyretin (TTR), was selected for further analysis for its role in lung cancer promotion. TTR (also called prealbumin) is a homotetramer plasma protein of ∼55 kDa, which is known for the transportation of thyroxine and retinol through binding to retinol-binding protein (6). However, TTR null mice suggest that TTR is not essential to thyroid hormone metabolism (7) and may not be crucial on retinol metabolism (8). We demonstrated that recombinant TTR protein enhanced myeloid cell differentiation and altered angiogenesis and promoted lung tumor cell proliferation in vitro and tumor growth in vivo.

The human serum samples of normal subjects and lung cancer patients (adenocarcinomas patients, squamous cell carcinomas patients, and small cell lung cancer patients) were obtained from the Biosample Repository Core Facility of Fox Chase Cancer Center in Philadelphia.

All scientific protocols involving the use of animals have been approved by the Institutional Animal Care and Use Committee in Indiana University School of Medicine and followed guidelines established by the Panel on Euthanasia of the American Veterinary Medical Association. Animals were housed under Institutional Animal Care and Use Committee–approved conditions in a secure animal facility in the Indiana University School of Medicine. Protocols involving the use of recombinant DNA or biohazardous materials have been approved by the Biosafety Committee of Indiana University School of Medicine and followed guidelines established by the National Institutes of Health. The generation of doxycycline-controllable and AT II epithelial cell–specific CCSP-rtTA/(tetO)₇-Stat3C bitransgenic mouse model was previously described (2). Lewis lung carcinoma (LLC) cells or B16 melanoma cells were purchased from the American Type Culture Collection.

The purified full-length TTR protein was used as the Ag for rabbit immunization by a custom Ab production service (Rockland Immunochemicals, Limerick, PA). The quality and the titer of TTR Ab in the serum was determined by ELISA and Western blotting.

The RNAs were extracted from mouse myeloid HD1A cells (Applied Biological Materials, Canada) using RNeasy kit (QIAGEN) and reversely transcribed into cDNA by oligo(dT) primer using ThermoScript transcriptase (Invitrogen, Carlsbad, CA) according to the manufacturer’s instruction. The cDNA was used in the PCR to amplify TTR amplicon by Phusion DNA polymerase (New England Biolabs, Ipswich, MA) using primers Xma I–TTR-F (5′-AGC CCC GGG TGC CAC CAT GGC TTC CCT TCG ACT CTT C-3′) and Not I–FLAG–TTR-R (5′-ACA GCT CAG AGC GGC CGC TCA CTT GTC ATC GTC ATC CTT GTA ATC ATT CTG GGG GTT GCT GAC GA-3′). The TTR amplicon (502 bp) was gel-purified and digested with Xma I and Not I restriction enzymes (New England Biolabs) and cloned into the pGEX-4T-1 expression vector (GE Healthcare Life Sciences, Pittsburgh, PA). A thrombin-specific cleavage site was present between GST and inserted TTR-Flag. The reading frame and the inserted TTR-Flag cDNA was confirmed by sequencing and named as pGEX-4T–TTR-Flag. The expression of GST-TTR-Flag fusion protein (∼42 kDa) was induced by IPTG in BL21 Escherichia coli, and the protein samples collected from preinduction, pellet, and supernatant were analyzed by SDS-PAGE and visualized by Coomassie blue staining. The GST-TTR-Flag fusion protein in the soluble fraction was purified by GST column and digested by thrombin. The TTR-Flag band (∼17 kDa) was monitored by Coomassie staining and confirmed by Western blot analyses using anti-FLAG Ab. The purified recombinant TTR-Flag protein was subjected to LPS removal by endotoxin removal column.

Bronchoalveolar lavage fluid (BALF) and serum were collected from wild-type (WT), doxycycline-treated, or -untreated CCSP-rtTA/(TetO)₇-Stat3C bitransgenic mice. Serum (1 μl) or BALF (6 μl) was mixed with Laemmli sample buffer (Bio-Rad, Hercules, CA) and heated at 95°C for 5 min, fractioned in Novex 4–20% Tris-Glycine Mini Gels (Invitrogen), and transferred to polyvinylidene difluoride membranes (Bio-Rad). The membranes were incubated with rabbit anti-TTR Ab (Abbiotec, San Diego, CA) at 4°C overnight and followed with secondary Ab (HRP-conjugated goat anti-rabbit IgG, 1:2000). TTR was detected by incubation with SuperSignal West Pico Chemiluminescent Substrate (Thermo Fisher Scientific), and the images were taken by ChemiDoc MP Imaging System (Bio-Rad).

To determine activation of signaling pathways by TTR treatment, LLC cells were seeded in six-well plates until ∼75% confluent and treated with LPS-removed TTR (1 and 0.1 μM) for 2 h in the presence of exotoxin inhibitor polymyxin B (PMB) (100 μg/ml). The treated LLC cells were harvested using Cellytic M cell lysis buffer with phosphatase and protease inhibitors. The cell lysates were centrifuged to remove insoluble cell debris. Protein concentrations in the lysates were determined by the BCA method. Twenty micrograms of total protein was analyzed by SDS-PAGE and transferred to a PVDF membrane. The membranes were probed by various Abs (1:1000) against p-mTOR, p–NF-κB p65, p-S6, p-Stat3, p-ERK, p-Akt, and p-p38 (Cell Signaling Technology, Danvers, MA), followed by secondary anti-rabbit IgG, HRP-conjugated Ab (1:2000).

The lungs from doxycycline-treated or -untreated CCSP-rtTA/(TetO)₇-Stat3C mice were inflated with 4% paraformaldehyde, harvested, stored in 4% paraformaldehyde overnight, and paraffin embedded as previously described (9). Tissue sections (5 μM) were incubated with rabbit anti-TTR Ab (1:500) at 4°C overnight and followed by biotinylated goat anti-rabbit Ab (1:1000). The color signals were detected with a Vectastain Elite ABC Kit following manufacturer’s instructions (Vector Laboratory, Burlingame, CA). Rabbit IgG was used as negative control.

Mouse B16 melanoma cells and LLC cells were seeded in 48-well plates (5 × 10³ cells per well). After overnight incubation, cells were treated with 0, 1, or 5 μM LPS-removed TTR in the presence of 100 μg/ml PMB. After 2 or 3 d, tumor cells were detached by Accutase (Sigma, St. Louis, MO), and the cell numbers were counted by a hemocytometer.

LLC cells (5 × 10⁵) or B16 melanoma cells (2 × 10⁵) were incubated with 20 μM TTR/PMB protein for 1 h and s.c. flank-injected into syngeneic C57BL/6 mice. Furthermore, LLC cells (5 × 10⁵) or B16 melanoma cells (1 × 10⁶) incubated with 20 μM TTR/PMB were s.c. flank-injected into allogeneic FVB/N mice. The tumor growth was estimated by measuring the maximal length (L) and width (W), and the tumor volume was calculated using the formula of 0.5 × L × W² (cubic millimeter).

LLC cells grown in a 60-mm dish until ∼70% confluence were transfected with three small interfering RNAs (siRNAs; against three independent regions) targeting Akt, mTOR, Akt/mTOR, or NF-κB p65 using Dharmacon siRNA transfection reagent according to the manufacturer’s instructions (Dharmacon, Lafayette, CO). After 2-d incubation, LLC cells (5 × 10⁵ cells) were harvested and treated with 5 μM TTR/PMB and s.c. flank-injected into WT C57BL/6 mice for tumor growth analyses. The effectiveness of siRNA knockdown was assessed by Western blot of each target protein.

For in vivo immune cell profiling, TTR (320 μg/mouse) was i.v. injected into WT mice twice a week for 2 wk, and PBS was used as control. Single-cell suspensions from the bone marrow, blood, and spleen were prepared as previously described (10). Approximately 1–3 × 10⁶ cells from various organs were incubated with FcR blocking Abs in FACS buffer (BD Biosciences, San Jose, CA) followed by isotype control or surface-specific primary Abs. For in vitro differentiation, the bone marrow cells from WT mouse were cultured in 96-well plates (1 × 10⁶ cells per well) and treated with LPS-removed TTR/PMB at concentrations of 0, 0.2, 1, or 5 μM for 2 d. Cells were harvested for surface staining with fluorescence-conjugated anti-mouse Abs. Anti-mouse MHC class II (MHC II) FITC, anti-mouse Ly6C FITC, anti-mouse CD11c PE, anti-mouse F4/80 allophycocyanin, anti-mouse CD11b, anti-mouse Ly6G (RB6-8c5), anti-mouse CD4 FITC, anti-mouse CD8 PE, and anti-mouse B220 allophycocyanin were purchased from eBioscience (San Diego, CA).

For TTR expression in mice, cells from the lung, blood, and spleen were prepared and stained with surface markers (SP-C, Ly6G, and CD11b Abs). Fixed cells were permeabilized using BD Cytofix/Cytoperm Fixation/Permeabilization Kit according to the manufacturer’s instruction. Cells were incubated with the anti-TTR Ab (1:500) at 4°C overnight. The next day, cells were washed and labeled with the secondary Ab for flow cytometry analysis. For expression of signaling molecules in bone marrow cells or endothelial cells (ECs), cells were treated with 50 μg/ml PMB or 1 μM TTR/PMB for 2 h and then stained with various cell surface markers, followed by intracellular staining of anti-mouse pAKT, anti-mouse pmTOR, anti-mouse pS6, anti-mouse pp38, anti-mouse pERK, and anti-mouse pp65 Abs (Cell Signaling Technology, Beverly, MA). Flow cytometry was analyzed on an LSR II machine (BD Biosciences). Data were analyzed using the BD FACStation Software (BD Biosciences). Quadrants were assigned using isotype control mAb. Data were processed using the CellQuest software.

WT FVB/N mouse femur bones were harvested and cut into small pieces aseptically to release bone marrow cells. The whole bone marrow cells were treated with 10 μM TTR/PMB for 2 d. The harvested cells were stained with Abs against various cell surface markers, including CD11b, CD11c, F4/80, Ly6G, and Ly6C, followed by intracellular staining of p-Akt, p-mTOR, pS6, p–NF-κB p65, p-ERK, and p-p38. The stained cells were subjective to flow cytometry analysis.

Fresh bone marrow cells (1 × 10⁶) from WT mice were recovered in RPMI 1640 medium (10% FBS) at 37°C for 1 h, followed by treatment with or without TTR (1, 5 μM) in PMB (50 μg/ml) for 1 h. Treated cells were stained with myeloid lineage–specific surface markers and 2 μmol/L 2', 7'-dichlorofluorescein diacetate (Invitrogen). The reactive oxygen species (ROS) level was measured by flow cytometry in gated areas using an LSR II machine (Becton Dickinson).

Freshly isolated spleen CD4⁺ T cells from the WT mice were labeled with CFSE and cultured in 96-well plates (0.2 × 10⁶ cells per well), which were precoated with anti-CD3 (2 μg/ml in PBS) and anti-CD28 (5 μg/ml in PBS) Abs in RPMI 1640 medium (10% FBS) at 37°C. The next day, freshly isolated Ly6G⁺ cells from the bone marrow of WT FVB/N mice that were treated with or without TTR (5 μM) overnight were added to CFSE-labeled CD4⁺ T cells at a 5:1 ratio and continuously incubated with or without 5 μM TTR in PMB (50 μg/ml) for 4 d. Cells were harvested and stained with allophycocyanin-labeled anti-CD4 mAb (eBioscience). Proliferation of CD4⁺ T cells was evaluated as CFSE dilution by flow cytometry.

ECs were isolated from the WT lung as previously described (11) and treated with 50 μg/ml PMB or 0.1 or 1 μM TTR/PMB for 24 h. Cells (5 × 10⁴) were seeded in 48-well plates precoated with 150 μl/well growth factor–reduced Matrigel (BD Biosciences). After 6 h, tube formation was observed with an inverted microscope (Nikon, Melville, NY) as a tube-like structure exhibiting a length four times its width. Images of tube morphology were taken in five random microscopic fields per sample at ×40 magnifications, and the cumulative tube lengths were measured by Nikon NIS Elements imaging software.

WT lung ECs (1.5 × 10⁵) were seeded into a 24-well plate and incubated overnight to form a confluent monolayer. After creating a scratch on the cell monolayer, 50 μg/ml PMB or 0.1 or 1 μM TTR/PMB was added into the medium in the presence of mitomycin C to prevent cell proliferation. Images were taken at 0 and 15 h after creating the scratch. ECs migration was estimated by measuring the distance from one side of the scratch to the other side using Nikon NIS Elements imaging software.

WT lung ECs (5 × 10⁴) were seeded into 24-well plates. Two days later, 50 μg/ml PMB or 0.1 or 1 μM TTR/PMB was added into the medium. Twenty-four hours later, EC cell numbers were counted.

Isolated WT lung ECs (5 × 10⁴) were added to the upper chamber of 24-well, 8.0-μm pore Transwell plates (Corning, Tewksbury, MA). After incubating at 37°C, 5% CO₂ for 48 h to form a monolayer, ECs were treated with 50 μg/ml PMB or 0.1 or 1 μM TTR/PMB for 24 h. The supernatant was removed, and CellTracker Green 5-chloromethylfluorescein diacetate–labeled bone marrow cells (1 × 10⁴ cells in 200 μl of media) were added to the upper well. Four hours later, transendothelial migration of bone marrow cells was determined by counting their numbers in the lower chamber from five randomly selected microscopic fields.

The effect of TTR on WT EC immunosuppressive function was analyzed by T cell proliferation assay. To isolate T cells, WT mouse splenocytes were incubated with biotinylated anti-CD4 Ab, followed by positive selection on magnetic beads, and were eluted from magnetic separation columns according to the manufacturer’s instructions (Miltenyi Biotec). The purified CD4⁺ T cells were labeled with 1 μM CFSE at 37°C for 10 min. After washing, cells were resuspended with growth medium and incubated at 37°C for 20 min. CFSE-labeled WT CD4⁺ T cells were cocultured with ECs that were pretreated with 50 μg/ml PMB or 0.1 or 1 μM TTR/PMB in 96-well plates, which were precoated with anti-CD3 mAb and anti-CD28 mAb for 4 d at 10:1 ratio between CD4⁺ T cells: WT ECs. The proliferation of CD4⁺ T cells was analyzed by flow cytometry. PBS was used as a negative control.

Statistical analyses were carried out in Word Excel 2016 (for animal studies) and SAS version 9.4 and R version 3.1.0 (for human studies). For animal studies, the data were mean values of multiple independent experiments and expressed as the mean ± SD. ANOVA and Tukey method based on log-transformed concentration level were used to evaluate the significance of the differences. Statistical significance level was set at p < 0.05. For human serum analyses of biomarkers, Kruskal–Wallis tests with pairwise Wilcoxon rank sum tests (Bonferroni-adjusted) were used to compare distributions of biomarkers among controls and cancer types. The area under the curve (AUC) was determined by cross-validated sensitivity, specificity, receiver operating characteristic (ROC) curve analysis to evaluate the diagnostic ability of biomarkers. To further distinguish between both cancer and controls and between different types of lung cancers, the classification and regression tree (CART) method (specifically RPART and TREE in R) was used. RPART uses all available cases. TREE, which only uses cases with all biomarkers available, was used to see if the classification results depended on the type of analytic method used.

The concentrations of all 13 secretory proteins showed statistically significant differences in at least one type of lung cancer compared with normal human subjects by ELISA (Table I). Optimal cutpoint for each biomarker using minimum specificity method (specificity = 0.80) and AUCs are listed in Table I. When comparing central tendencies among the four groups, markers 1, 2, 3, 4, 5, 6, and 10 all showed significant differences between each tumor type and control. For the other biomarkers, nonsignificant differences were seen mostly in either squamous cells, small cells, or both. The areas under the ROCs display similar patterns. The lowest areas are for squamous cell and/or small cell in biomarkers 7–13. Statistical analyses using CART method (specifically RPART and TREE) revealed that different combinations in this panel of secretory protein biomarkers distinguished different types of lung cancers (Table II). For example, the misclassification rate for cancer versus control was 11.2% using the RPART method.

Ups and downs of protein biomarkers in human patients implicate their functional roles in lung cancer formation. They are potential therapeutic targets for personalized lung cancer treatment if their pathogenic roles are clear. Based on its highly increased concentration and AUC in all three human lung cancer categories, TTR was chosen for further analysis. In addition to human lung cancers (Fig. 1A, Table I), TTR (monomer ∼14 kDa) was detected in the sera of CCSP-rtTA/(TetO)₇-Stat3C bitransgenic mice. In doxycycline-treated bitransgenic mice with spontaneous lung tumors (doxycycline-treated with tumor [CA] group), all mice showed higher TTR protein levels (Fig. 1B). In BALF from doxycycline-treated bitransgenic mice with spontaneous lung tumors, the TTR levels were also higher than those in BALF from the other three groups (group treated with doxycycline [+DOX], group without doxycycline treatment [−DOX], and WT lung and blood were analyzed by flow cytometry analyses with anti-TTR Ab and cell surface markers. In the lung, the TTR expression levels were higher in whole lung cells, SPC⁺ AT II epithelial cells, CD11b⁺Ly6G⁺ cells, and Ly6G⁺ cells in doxycycline treatment (Fig. 1C, Supplemental Fig. 1B). In the blood, the TTR expression levels were also higher in whole WBCs with doxycycline treatment but not in CD11b⁺Ly6G⁺ cells, CD11b⁺ cells, and Ly6G⁺ cells (Supplemental Fig. 1C). The spleen showed no difference of TTR expression levels in doxycycline-treated and -untreated mice (Data not shown). By immunohistochemical (IHC) staining of lung tissues, the TTR protein was positive in AT II epithelial cells (arrows) and alveolar macrophages in both doxycycline-treated and -untreated bitransgenic mice, with stronger signals in the doxycycline-treated group (Fig. 1D).

TTR cDNA was subcloned and expressed in bacteria (Supplemental Fig. 2A–C). The purified recombinant TTR protein (1 or 5 μM) after LPS removal (Supplemental Fig. 2F) was incubated with LLC cells or B16 melanoma cells for cell proliferation assessment in in vitro cell culture experiment. To eliminate the effect of residual LPS in recombinant TTR, PMB was used to block LPS. Comparing with the PMB control group, TTR treatment significantly enhanced LLC or B16 melanoma cell proliferation (Fig. 2A). Recombinant TTR without LPS removal showed a similar result (Supplemental Fig. 2G, 2H). In all subsequent studies, LPS-removed recombinant TTR was used in combination with PMB to minimize the LPS interference. In tumor growth in vivo study, LLC cells or B16 melanoma cells were pretreated with recombinant TTR/PMB and flank-injected into syngeneic C57BL/6 mice. The flank-injected LLC tumors with TTR/PMB pretreatment grew much faster than those with PMB-treated control mice at 11, 14, and 18 d postinjection (Fig. 2B). In the allogeneic FVB/N mouse model, LLC tumors showed the same results, although the tumor sizes were smaller in general (Fig. 2C). When the B16 melanoma model was tested, similar results were observed (Supplemental Fig. 2I, 2J).

The levels of phosphorylated forms of mitogenic molecules Akt1, mTOR, S6, ERK, p38, NF-κB p65, and Stat3 proteins were investigated by TTR/PMB treatment. TTR/PMB treatment of LLC cells led to increased activation of p-Akt1, p-mTOR, and p–NF-κB p65, whereas the levels of p-ERK and p-p38 were relatively unchanged (Fig. 2D). Inhibition of Akt1, mTOR, or Akt1/mTOR by siRNAs knockdown (Supplemental Fig. 2K) impaired LLC’s proliferative ability in response to TTR/PMB treatment (Fig. 2E). When tested in vivo, LLC cells were knocked down with Akt1, mTOR, or Akt1/mTOR siRNAs before s.c. flank injection to the WT C57BL/6 recipient mice. mTOR knockdown greatly decreased tumor growth by TTR/PMB stimulation comparing with control siRNA knockdown, which was further decreased by Akt1/mTOR double knockdown (Fig. 2F). NF-κB p65 knockdown (Supplemental Fig. 2K) also decreased tumor growth by TTR/PMB stimulation comparing with control siRNA knockdown (p < 0.05) (Fig. 2G). These results suggest that the Akt1/mTOR pathway and the NF-κB pathway in cancer cells are responsible for TTR stimulation.

In addition to directly stimulating cancer cells, it is intriguing to determine if TTR is involved in differentiation of various immune cells that are known to play important roles in the tumor environment. Purified and LPS-removed recombinant TTR was injected into WT mice twice a week for 2 wk. Profiling of total bone marrow cells by flow cytometry showed that the majority of immune cells in the myeloid compartment (CD11b⁺Ly6G⁺, CD11b⁺, Ly6G⁺, Ly6C⁺, F4/80⁺ cells) showed increased differentiation, whereas CD8⁺ and B220⁺ cells were decreased comparing with the control group (Fig. 3A, Supplemental Fig. 3A). This concludes that TTR has a cytokine-like function specific for increasing myeloid cell differentiation in the bone marrow.

To confirm these observations of myeloid differentiation, whole bone marrow cells were isolated from WT mice and cultured in vitro with TTR/PMB for 2 d. TTR/PMB stimulated CD11b^high+, CD11C⁺, and F4/80⁺ myeloid lineage differentiation in a dose-dependent manner (Fig. 3C, 3D, 3F), while stimulated Ly6C⁺, MHC II⁺ myeloid lineage differentiation starting at a low dosage of TTR (0.2 μM) (Fig. 3B, 3G). The Ly6G⁺ population was the only exception, showing downregulation (Fig. 3E) that contradicts the in vivo observation (Fig. 3A). The double-positive CD11b^high+Ly6C^high+ cells, which are myeloid progenitor cells, were also increased in a dose-dependent manner after TTR/PMB treatment (Fig. 3H). To show the activation status of immune cells treated by TTR/PMB, activation of various cellular signaling molecules were investigated. As demonstrated in Fig. 4A–C and Supplemental Fig. 4A–C, the phosphorylation levels of Akt1, mTOR, and S6 were increased after TTR/PMB treatment by flow cytometry, suggesting that the metabolic reprogramming controlled by the Akt1/mTOR pathway is involved in TTR-mediated myeloid lineage differentiation. Activation of ERK, p38, and NF-κB p65 molecules were also concomitantly increased in myeloid lineage cells (Fig. 4D–F, Supplemental Fig. 4D–F).

Functionally, TTR/PMB treatment increased cellular ROS production in CD11b⁺, Ly6G⁺, CD11b⁺Ly6G⁺, CD11C⁺, Ly6C⁺, MHC II⁺, and F4/80⁺ myeloid lineage cells (Fig. 5A, Supplemental Fig. 3B). High-level expression of ROS is a hallmark for the CD11b⁺Ly6G⁺ myeloid population to be immunosuppressive. This may explain why CD8⁺ or B220⁺ cell differentiation was suppressed after TTR/PMB in vivo treatment (Fig. 3A). To confirm this assumption, in vitro CFSE-labeled T cell proliferation was performed by incubating with TTR/PMB–treated CD11b⁺Ly6G⁺ cells. Although TTR/PMB treatment alone showed no effect on T cell proliferation (Fig. 5B, middle panel), TTR/PMB treatment of CD11b⁺Ly6G⁺ cells from the bone marrow showed significant suppression on CFSE-labeled WT splenocyte T cell proliferation in coculture experiment with stimulation of anti-CD3 mAb plus anti-CD28 mAb (Fig. 5B, lower panel). It seems that TTR treatment is able to convert WT bone marrow CD11b⁺Ly6G⁺ cells to myeloid-derived suppressive cells. Therefore, increased TTR concentration creates an immunosuppressive environment to benefit tumor growth.

ECs are a major cellular component of the alveolar structure and pulmonary vasculature, which control infiltration of cancer cells and immune cells into the lung. As shown in Fig. 6A, the TTR expression level was significantly increased in lung CD31⁺ ECs after doxycycline treatment of CCSP-rtTA/(TetO)₇-Stat3C bitransgenic mice. To determine whether TTR influences the formation of capillary-like tubes by lung ECs (an important parameter of angiogenesis) the in vitro Matrigel tube formation assay was performed. TTR/PMB–pretreated WT ECs formed more tubes than those of PMB-pretreated control ECs, demonstrating that TTR had ability to enhance EC tube formation in vitro (Fig. 6B). To test whether TTR/PMB affects EC migrating ability, monolayers of WT ECs were treated with PMB or TTR/PMB in the presence of mitomycin C to eliminate the potential effects of EC proliferation. As shown in Fig. 6C, 15 h after creating the scratch, TTR/PMB–treated WT ECs demonstrated increased migration, comparing with that of PMB-treated ECs in wound-healing assay, evidenced by a significantly reduced wound area lacking cells. In a separate experiment, TTR/PMB stimulated EC proliferation (Fig. 6D). Transwell assay was performed to determine the effect of TTR/PMB on the EC permeability. Freshly isolated WT bone marrow cells were labeled with 5-chloromethylfluorescein diacetate and loaded on primary lung EC monolayers that were pretreated with PMB or TTR/PMB for 24 h. As shown in the Fig. 6E, TTR/PMB–treated WT lung ECs showed increased permeability, with more bone marrow cells migrating to the lower chamber than that of PMB-treated ECs. ECs are known for their influence on T cell activity (12). When WT ECs were pretreated with TTR/PMB and cocultured with CFSE-labeled WT splenocyte CD4⁺ T cells with stimulation of anti-CD3 mAb plus anti-CD28 mAb, TTR/PMB–treated ECs suppressed T cell proliferation, comparing with that of PMB-pretreated ECs (Fig. 6F).

Lung cancers are inflammation-associated diseases. Inflammatory molecules are valuable tools to reveal lung cancer occurrence. At the moment, the clinical solution for lung cancer detection using protein biomarkers is challenging and premature (13). This is because lung cancers are heterogeneous diseases and caused by multiple factors; a particular biomarker may or may not be upregulated in a given patient. Conversely, a given biomarker can be upregulated in different types of diseases. Therefore, it is difficult to use one or two biomarkers for accurate lung cancer detection or prediction. To solve this clinical challenge, multiple biomarkers should be developed and used for more accurate lung cancer detection and classification. Different inflammatory diseases (including cancers) produce distinctive sets of molecules, which can be used as a panel of signature biomarkers for disease distinguishing. Because the high expression level of STAT3 was associated with advanced tumor stage (14), secretory protein biomarkers identified in this study have multiple advantages, especially that they detect lung cancer in the patient blood without going through biopsy (Table I). Combination of these biomarkers significantly eliminated false positive rate in lung cancer diagnosis. Importantly, they distinguished different types of human lung cancers (Table II).

Upregulation of protein biomarkers contributes to the tumor microenvironment that is in favor of tumor growth and metastasis. Therefore, it is critical to elucidate their functional roles in tumor formation and immune reaction to better use them for lung cancer prediction and further, for personalized treatment and precision medicine. Previously, chitinase 3-like 1 (CHI3L1), one of 13 biomarkers, has been shown to stimulate proliferation and growth of lung cancer cells (9). TTR was originally identified as a transport protein in the serum that carries the thyroid hormone thyroxine and retinol-binding protein bound to retinol (6, 15). It has been extensively studied in the brain and associated with neurologic diseases (16). In this report, TTR was highly expressed in the blood of human patients with adenocarcinoma, squamous carcinoma, and small cell lung cancer (Fig. 1A). The same observation was confirmed in the CCSP-rtTA/(tetO)₇-Stat3C lung tumor mouse model (Fig. 1B), which matched the result of gene profile analysis (2). In addition to being produced by AT II epithelial cells where Stat3C was overexpressed, TTR was also produced by pulmonary myeloid lineage cells (Fig. 1C, 1D) and lung ECs (Fig. 6A) in the CCSP-rtTA/(tetO)₇-Stat3C mouse model, probably through secondary inducible effects.

Based on our characterization, TTR has pleiotropic functions in multiple aspects of tumorigenesis. First, TTR directly targeted and stimulated LLC and B16 melanoma cell proliferation in vitro and growth in vivo (Fig. 2A–C, Supplemental Fig. 2G–J). This was mediated through activation of the Akt/mTOR and NF-κB pathways (Fig. 2D). mTOR is a master regulator of the serine/threonine protein kinase family member that regulates cell growth, proliferation, migration, survival, protein synthesis, and transcription in response to growth factors and mitogens (17). Oncogenic activation of the mTOR pathway has been reported to induce several processes required for cancer cell growth, survival, and proliferation (18). Akt1 is the upstream regulator of mTOR. Ablation of Akt1 and mTOR by siRNA knockdown reduced TTR stimulation of LLC proliferation and growth (Fig. 2E–F). These observations showed that metabolic master regulator mTOR is required for TTR stimulation of tumor cells. NF-κB is another important oncogenic pathway important for tumor growth (19). Ablation of NF-κB by siRNA knockdown reduced TTR stimulation of LLC tumor growth as well (Fig. 2G).

Second, in addition to directly stimulating cancer cell proliferation and growth, TTR demonstrated features of cytokines to regulate immune cell differentiation. Tumor initiation, growth, and metastasis are largely facilitated and dependent on their interactions with surrounding immune cells, which are critical components in the tumor environment. The high levels of TTR in the blood of human lung cancer patients and the CCSP-rtTA/(tetO)₇-Stat3C lung tumor animal model inevitably influence immune cell development and functions. Indeed, TTR injection into WT mice led to increased differentiation of myeloid lineage cells (CD11b⁺Ly6G⁺, CD11b⁺, Ly6G⁺, Ly6C⁺, F4/80⁺) and decreased CD8⁺ and B220⁺ cells (Fig. 3A). This was confirmed by in vitro study in which TTR stimulated myeloid lineage differentiation of isolated WT bone marrow cells (Fig. 3B–H). Similar to cancer cells, TTR activated Akt1, mTOR, and S6 phosphorylation in myeloid lineage cells (Fig. 4A–C). This is consistent with previous observations that mTOR-mediated metabolic reprogramming plays a critical role in myeloid-mediated tumor immunity (20, 21). ERK, p38, and NF-κB p65 were also activated by TTR in myeloid cells (Fig. 4D–F). In a separate mechanism, overproduction of ROS in myeloid lineage cells plays a critical role in their protumor activity. TTR treatment not only stimulated ROS production in WT myeloid cells but also converted WT bone marrow CD11b⁺Ly6G⁺ cells to become myeloid-derived suppressive cells that suppressed splenocyte T cell proliferation (Fig. 5). Antitumor rejection largely relies on proper T cell proliferation and functions.

Third, the increased TTR concentration in the circulation system influences angiogenesis and EC functions. ECs are the major component of blood vessels and actively participate in regulation of inflammatory and tumorigenic processes through controlling circulating cell migration, vessel permeability, and cell infiltration into organs (22). TTR regulated ECs functions in several folds. It had ability to influence angiogenesis by enhancing EC tube formation (Fig. 6B) and by stimulating EC proliferation (Fig. 6D). Permeability of EC depends on angiogenesis. TTR showed ability to increase both EC migration (Fig. 6C) and permeability (Fig. 6E). Migration, penetration, and function of leukocytes and cancer cells in the blood are influenced by ECs. TTR-treated ECs showed suppression of T cell proliferation (Fig. 6F). It is noted that TTR showed modest effects on some cell types. However, combination of these effects contribute enormously to stimulation of tumor growth in vivo by surged TTR (Fig. 2B, 2C, Supplemental Fig. 2I, 2J) and created an immunosuppressive environment to benefit tumor growth.

Taken together, Stat3 and its downstream inducible protein biomarkers show promise as a valuable tool for lung cancer detection and classification. These blood soluble biomarkers actively participate in tumor proliferation, growth, and invasion by directly stimulating cancer cells as well as by regulating immune cells and ECs in the tumor environment. Therefore, these biomarkers have potential for use as clinical targets for lung cancer personalized treatment if their expression levels are increased in a given lung cancer patient in the blood.

We thank Haijiao Xu for statistical analysis.

The authors have no financial conflicts of interest.