Affiliations of authors: Departments of Human Cancer Genetics (GBL, SVK, WEC), Epidemiology and Biometrics (LS), Medical Oncology (KK), and Surgery (MW, WEC), Arthur G. James Cancer Hospital and Richard J. Solove Research Institute, The Ohio State University, Columbus, OH; Primetrics, Hilliard, OH (TC)
Correspondence to: William E. Carson III, MD, Division of Surgical Oncology, The Ohio State University, N924 Doan Hall, 410 W. 10th Ave., Columbus, OH 43210 (e-mail: carson-1{at}medctr.osu.edu)
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ABSTRACT |
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INTRODUCTION |
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The binding of IFN to its receptor results in activation of the Jak-STAT signal transduction pathway. The heterodimeric receptor for IFN
consists of two subunits, IFN
receptor 1 (IFNAR1) and IFN
receptor 2 (IFNAR2), and is widely expressed on the surface of tumor cells and immune effector cells (6,7). Binding of IFN
to its receptor activates Janus kinase 1 (Jak1) and tyrosine kinase 2 (Tyk2), which phosphorylate tyrosine residues within the cytoplasmic region of IFNAR1. The phosphotyrosine residues provide docking sites for cytoplasmic transcription factors that belong to the signal transducer and activation of transcription (STAT) family of proteins, which are phosphorylated (activated) by the Janus kinases (8). The prototypical IFN
signaling reaction facilitates the formation of interferon-stimulated gene factor 3 (ISGF3), a DNA-binding complex that consists of STAT1
(or STAT1
), STAT2, and interferon regulatory factor 9 (IRF9) (9). ISGF3 rapidly translocates to the cell nucleus and binds to interferon-stimulated response elements located in the promoter regions of IFN-responsive genes (10). This binding induces the expression of a variety of immunoregulatory genes and largely determines the pattern of immune cell activation following IFN
administration (1113).
The ability to study signal transduction in distinct subsets of immune cells following IFN treatment has been limited by the efficiency and qualitative nature of the available techniques. The use of phosphorylation statespecific antibodies for intracellular flow cytometry has unique potential for the evaluation of signaling events in immune effectors following the administration of immunomodulatory cytokines. We have developed a novel flow cytometric technique for the analysis of STAT1 phosphorylation among immune effector cell subsets that is rapid, highly quantitative, and extremely sensitive. We have used this method to examine Jak-STAT signal transduction in peripheral blood mononuclear cells (PBMCs) isolated from healthy blood donors in vitro and from melanoma patients who were undergoing IFN
immunotherapy.
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MATERIALS AND METHODS |
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Recombinant human IFN -2b (specific activity = 2 x 108 IU/mg) was obtained from Schering-Plough (Kenilworth, NJ) and resuspended in phosphate-buffered saline (PBS) supplemented with 0.1% human albumin (Sigma, St. Louis, MO). Anti-Phospho-STAT1 (Tyr701) monoclonal antibody was obtained from Cell Signaling Technology (Beverly, MA). RosetteSep NK and T cell enrichment cocktails were obtained from Stem Cell Technologies (Vancouver, British Columbia, Canada). Recombinant human granulocytemacrophage colony-stimulating factor (GM-CSF), interleukin 4 (IL-4), and tumor necrosis factor-alpha (TNF-
) were obtained from R&D Systems (Minneapolis, MN).
Blood Samples
This study was approved by the Institutional Review Board of The Ohio State University (OSU 99H0348). Peripheral blood was obtained from melanoma patients who were evaluated in the Melanoma Multi-Disciplinary Clinic of The Ohio State University Comprehensive Cancer Center and who provided written informed consent. Patients were considered eligible for this study if they had histologic or cytologic documentation of cutaneous melanoma, clinical evidence of metastatic disease, had not previously received cytokine treatment for metastatic disease (i.e., IL-2 or IFN ), and had not received chemotherapy, radiotherapy, or antihormonal therapy within the 3 weeks before peripheral blood was drawn and IFN
treatment was initiated. PBMCs from healthy adult blood donors were obtained from source leukocytes (American Red Cross, Columbus, OH). Flow cytometric analysis of STAT1 phosphorylation was conducted on PBMCs from five melanoma patients who were undergoing treatment with IFN
-2b. PBMCs were separated from peripheral blood or source leukocytes by density gradient centrifugation with Ficoll-Paque (Amersham Pharmacia Biotech, Uppsala, Sweden) and immediately used for flow cytometric analysis (14).
Intracellular Staining for STAT1, STAT2, and Phosphorylated STAT1
PBMCs (5 x 105 cells per condition) were cultured in RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS) or human AB serum (Pel-Freez Clinical Systems, Brown Deer, WI) and IFN -2b or PBS for different times, incubated with antibodies specific for STAT1, STAT2 or phosphorylated STAT1 (P-STAT1), and subjected to intracellular flow cytometry as described by Fleisher et al. (15), with modifications. Briefly, cells were harvested from culture by centrifugation at 290g, resuspended in 100 µL of RPMI-1640 medium supplemented with 10% FBS, treated with either PBS or IFN
-2b, fixed by incubating in 100 µL of Fix & Perm Reagent A (Caltag Laboratories, Burlingame, CA) for 23 minutes at room temperature, and incubated for 10 minutes in 3 mL of cold methanol. Cells were then washed in flow buffer (PBS supplemented with 5% FBS) and permeabilized with 100 µL of Fix & Perm Reagent B (Caltag Laboratories), according to the manufacturers specifications. Cells were incubated for a total of 30 minutes at room temperature in Fix & Perm Reagent B containing 1 µg of a mouse anti-human STAT1 antibody (BD Transduction Laboratories, San Diego, CA), 1.256 µg of a rabbit anti-human STAT2 antibody (Biosource International, Camarillo, CA), 7.5 ng of a rabbit anti-human P-STAT1 (Tyr701) antibody (Cell Signaling Technologies), or an appropriate isotype control antibody. The cells were washed with flow buffer, incubated with a fluorescein isothiocyanateconjugated goat anti-mouse secondary antibody (STAT1) or an Alexafluor 488conjugated goat anti-rabbit secondary antibody (P-STAT1, STAT2) (Molecular Probes, Eugene, OR) for 30 minutes at room temperature, washed with flow buffer, fixed in 1% formalin, and stored at 4 °C. Flow cytometric analysis of STAT1 expression was conducted on PBMCs from normal donors (n = 15) and melanoma patients (n = 17). Flow cytometric analysis of STAT2 expression was conducted on PBMCs from normal donors (n = 17) and melanoma patients (n = 19). Analysis of P-STAT1 activation following ex vivo stimulation with IFN
-2b was conducted on PBMCs from normal donors (n = 18) and melanoma patients (n = 19).
Multiparametric Staining
Briefly, PBMCs were stained for P-STAT1 or STAT1 as described above except that a primary antibody specific for a subset of immune cells was added to the cells along with the Alexafluor 488conjugated secondary antibody. For this particular technique, the detection of intracellular P-STAT1 and STAT1 was best achieved using Alexafluor 488conjugated secondary antibodies. The following antibodies were used to detect immune cell subsets: anti-CD56 ([NK1-RD1] for NK cells; Beckman Coulter, Miami, FL); anti-CD14-APC (for monocytes; Beckman Coulter); anti-CD3-APC (for T lymphocytes; BD Pharmingen, San Diego, CA); and anti-CD21-APC (for B lymphocytes; BD Pharmingen). Nonspecific intracellular and extracellular antibody binding were blocked by incubating the cells with normal goat serum and normal mouse serum, respectively, for 10 minutes before the fluorochrome-conjugated antibodies were added. Separate aliquots of PBMCs were stained with appropriate intracellular and extracellular isotype control antibodies (Beckman Coulter and BD Pharmingen).
Flow Cytometric Analysis
Flow cytometry was performed with the use of a Becton-Dickinson FACScalibur cytometer (BD Immunocytometry Systems, San Jose, CA) equipped with a 488-nm air-cooled argon laser and a 633-nm heliumneon laser. Each analysis was performed using at least 10 000 cells that were gated in the region of the lymphocyte population, as determined by light scatter properties (forward scatter versus side scatter). To analyze monocyte (i.e., CD14-positive) cell populations, cells were gated in both the lymphocyte and monocyte regions. For multiparametric analysis, percent positive values were determined from quadrants set with isotype control antibodies. Data files were processed with the use of WinMDI software (created by Joseph Trotter; available at: http://pingu.salk.edu/software.html). Amplified fluorescence signals were displayed on four-decade log scales and expressed as specific fluorescence (Fsp = Ft Fb), where Ft represents the median value of total staining, and Fb represents the median value of background staining (obtained by staining with the isotype antibody control). The specific activation of STAT1 following IFN treatment of PBMCs was calculated as FspIFN-treated FspPBS-treated, where FspIFN-treated and FspPBS-treated represent the median levels of specific fluorescence for P-STAT1 in PBMCs treated with IFN
and PBS, respectively. We collected a minimum of three data files for each condition analyzed to control for inter-assay variability.
Immunoblot Analysis and Electrophoretic Mobility Shift Assay
PBMCs (5 x 106 cells) isolated from source leukocytes obtained from healthy adult donors were cultured in RPMI-1640 medium supplemented with 10% FBS and various concentrations of IFN -2b or PBS for 15 minutes. We prepared cell lysates from these cultures and subjected equal amounts of protein per lane to immunoblot analysis, as previously described (16), using the same rabbit anti-human P-STAT1 antibody used for flow cytometry or a
-actin antibody (Sigma), followed by quantitative densitometry using Optimas 6.51 image analysis software (Media Cybernetics, Carlsbad, CA). Lysates from A431 cells (Cell Signaling Technology) were used as a positive control for measuring STAT1 expression by immunoblot analysis. We simultaneously prepared whole-cell extracts from these cultures (5 x 106 cells), as previously described (17), and used them in an electrophoretic mobility shift assay (EMSA) with a double-stranded serum-inducible element of the c-fos promoter oligonucleotide that has affinity for activated human STAT proteins (5'-GATCCGATTCCGGGAATCA-3') (18).
Generation of Dendritic Cells
Mature dendritic cells were generated as previously described (19). Briefly, PBMCs were isolated from source leukocytes of a healthy blood donor by density gradient centrifugation and aliquots were placed in each well of 6-well plastic culture dishes. After 3 days in culture, monocytes were separated from total PBMCs on the basis of their adherence to the plastic. Fresh complete RPMI-1640 medium containing 10% FBS, 800 IU/mL GM-CSF, and 500 IU/mL IL-4 was then added to the remaining adherent cells and they were cultured for 57 days. Dendritic cells were generated from these cultures under endotoxin-free conditions, and fresh medium with cytokines was added on day 3. Mature dendritic cells were obtained following the addition of 200 U/mL TNF- from day 5 to day 7. To confirm that mature dendritic cells had been generated, we performed direct cell-surface staining with anti-human CD11c, anti-human CD14, and anti-human CD83 antibodies and the appropriate isotype control antibodies (BD Pharmingen).
Real-Time Reverse TranscriptionPolymerase Chain Reaction Analysis of IFN Stimulated Gene Expression
Real-time reverse transcriptionpolymerase chain reaction (RT-PCR) was used to quantitate levels of mRNAs expressed by known IFN stimulated genes present within PBMCs. Briefly, PBMCs isolated from source leukocytes of healthy donors or from the peripheral blood of melanoma patients were cultured (5 x 106 cells) in RPMI-1640 medium supplemented with 10% FBS and various concentrations of IFN
-2b or PBS for 4 hours. Total RNA was isolated from the cultured PBMCs with the use of an RNeasy RNA Isolation Kit (Qiagen, Valencia, CA) and quantitated by using a RiboGreen RNA Quantitation Kit (Molecular Probes). Reverse transcription was performed using 2 µg of total RNA and random hexamers (PerkinElmer, Norwalk, CT) as primers for first-strand synthesis of cDNA and the following conditions: 70 °C for 2 minutes, 42 °C for 60 minutes, and 94 °C for 5 minutes. We used 2 µL of the resulting cDNA as template to measure the levels of mRNA for ISG-15, ISG-54, and 2',5'-oligoadenylate synthetase 1 (OAS-1) by real-time RT-PCR with pre-designed primer/probe sets (Assays On Demand; Applied Biosystems, Foster City, CA) and 2x Taqman Universal PCR Master Mix (Applied Biosystems). Pre-designed primer/probe sets for human
-actin (Applied Biosystems) were used as an internal control in each reaction well. Real-time RT-PCR reactions were performed in triplicate in capped 96-well optical plates. The following amplification scheme was used: 50 °C for 2 minutes, 95 °C for 10 minutes, 40 cycles of 95 °C for 15 seconds, and 60 °C for 1 minute. Real-time RT-PCR data were analyzed using Sequence Detector software, version 1.6 (PE Applied Biosystems, Foster City, CA).
Statistical Analysis
The specific fluorescence (Fsp) and specific activation of STAT1 within NK cells, T cells, and total PBMCs were of primary interest. Primary analyses focused on differences in mean values and variances between PBMCs from healthy donors and melanoma patients. We used Levenes test for equality of variances (20) to determine whether healthy donors and patients had statistically significantly different within-group variances. Students t test for equality of means was then used to determine whether the group means were statistically significantly different. A variation of the t test that does not assume equality of variances was used for assays where statistically significant inequality of variance was found. Analyses were also repeated using the MannWhitney U test (a nonparametric test equivalent to the t test) to confirm that the same pattern of results emerged (data not shown). We used the Friedman test for k-dependent samples to test whether P-STAT1 levels were similar between different dose levels of IFN . All statistical tests were two-sided; a P value of less than .05 was considered statistically significant.
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RESULTS |
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PBMCs freshly isolated from healthy adult donors (n = 3) were treated with different doses of IFN -2b or PBS for 15 minutes to evaluate the utility of our flow cytometry assay for detecting P-STAT1. We observed a rapid and dose-dependent increase in P-STAT1 within PBMCs treated with IFN
-2b (Fig. 1). P-STAT1 was detected even after PBMCs were treated with relatively low concentrations of IFN
-2b (i.e., 1100 IU/mL). The Friedman test for k-dependent samples revealed that equivalence between the dose levels was not rejected (df = 3; P = .241), suggesting that different dose levels (102105 IU/mL) can produce similar responses. Examination of P-STAT1 levels in PBMCs treated with increasing doses of IFN
-2b (200-IU/mL increments) revealed a smooth doseresponse curve (data not shown). Treatment of PBMCs with 103 IU/mL IFN
-2b routinely led to the highest levels of P-STAT1, and higher doses of IFN
-2b did not markedly increase the level of P-STAT1 (data not shown). In addition, we routinely detected P-STAT1 in PBS-treated PBMCs, further demonstrating the sensitivity of this technique for detecting the low levels of P-STAT1 in cells with unactivated Jak-STAT signaling pathways. Although this assay required only 5 x 105 cells per condition, we could obtain meaningful data using as few as 1 x 105 cells (data not shown). Further experiments revealed that cryopreservation (i.e., overnight freezing in a Nalgene Cryo 1 °C Freezing Container (Nalgene, Rochester, NY) at 80 °C in a 10% dimethyl sulfoxide/90% FBS solution followed by storage at 130 °C in liquid nitrogen for 24 hours) of PBMCs from healthy adult donors before ex vivo stimulation with IFN
-2b decreased the level of STAT1 phosphorylation in the T and B lymphocyte, monocyte, and NK cell populations when compared with freshly isolated lymphocytes from the same donor (data not shown).
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To confirm our flow cytometry results, we treated PBMCs freshly isolated from a single healthy adult donor with different concentrations of IFN -2b and subjected aliquots of the treated cells to flow cytometry, immunoblot analysis, and an EMSA. Flow cytometry and immunoblot analyses, both of which used the same anti-P-STAT1 antibody, revealed that PBMCs treated with IFN
-2b displayed a dose-dependent increase in the level of P-STAT1 (Fig. 2, A and B). However, unlike the flow cytometry assay, the immunoblot assay and EMSA could not detect basal P-STAT1 levels in PBS-treated cells, and the response to 1 IU/mL IFN
-2b was almost undetectable with these assays (Fig. 2, B and C). Whereas our flow cytometry data indicated that maximal activation of STAT1 occurred when cells were treated with 103 IU/mL IFN
-2b, densitometric analysis of the immunoblot data suggested that maximal activation of STAT1 occurred at a dose level of 104105 IU/mL IFN
-2b (Fig. 2, B; data not shown). EMSAs using lysates made from these cells to detect the generation of DNA binding activity revealed signals of similar intensities for cells treated with IFN
-2b doses ranging from 102 to 105 IU/mL (Fig. 2, C). Thus, our flow cytometry assay appeared to be substantially better at detecting low levels of activated STAT1 and differentiating between the levels of activated STAT1 induced by different doses of IFN
-2b than the immunoblot assay and EMSA (Fig. 2, A). It is important to note that these latter two assays required 10-fold more cells per condition (i.e., 5 x 106 cells) than did flow cytometry. Results of time-course studies using flow cytometry (which were also confirmed by immunoblot and EMSA analysis) revealed that maximal induction of P-STAT1 in PBMCs occurred at 15 minutes after treatment with 104 IU/mL IFN
-2b, after which P-STAT1 levels slowly returned to basal levels (i.e., the levels of P-STAT1 in untreated or PBS-treated cells) over a 4-hour period (data not shown).
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We next examined the relationship between IFN -2b induction of P-STAT1 and the expression of known IFN
stimulated genes. PBMCs from a healthy donor were treated with various doses of IFN
-2b and aliquots were processed for flow cytometric analysis of P-STAT1 (after 15 minutes of treatment) and real-time RT-PCR analysis of ISG-15, ISG-54, and OAS-1 gene expression (after 4 hours of treatment). We observed dose-dependent increases in the levels of P-STAT1 and ISG-54 and OAS-1 mRNAs following stimulation with IFN
-2b (Fig. 3, AC). We also observed an attenuated dose-dependent increase in the level of ISG-15 mRNA in PBMCs following stimulation with IFN
-2b: ISG-15 mRNA levels were comparable following stimulation with 103 or 104 IU/mL of IFN
-2b (Fig. 3, D). These observations suggest that dose-specific and variable patterns of gene expression occur in immune effector cells following exogenous administration of IFN
-2b.
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A systematic analysis of IFN -induced signal transduction in immune cell subsets has not previously been reported. We therefore developed a dual-parameter flow cytometric technique that uses antibodies that stain intracellular and extracellular proteins to assay the activation of STAT1 within T lymphocytes, NK cells, B lymphocytes, and monocytes. PBMCs treated with IFN
-2b (104 IU/mL for 15 minutes) displayed a rapid induction of P-STAT1 in the T lymphocyte, NK cell, B lymphocyte, and monocyte compartments when compared with PBMCs treated with PBS (P<.05 for each comparison) (Fig. 4, A). Although every cell type tested displayed an increase in the level of P-STAT1 after IFN
treatment, the most robust response to this cytokine appeared to occur within T lymphocytes and monocytes. Multiparametric staining and immunoblot analysis indicated that the increased levels of activated STAT1 observed in IFN
-treated T lymphocytes and monocytes probably reflect the greater overall levels of STAT1 protein in T lymphocytes and monocytes than in NK cells and B lymphocytes (Fig. 5, A and B; data not shown). We also examined Jak-STAT signal transduction in cultured dendritic cells because recent studies (21,22) have indicated that IFN
may play an important role in modulating the maturation and function of this antigen-presenting cell type. Mature dendritic cells (i.e., cells that were >98% CD11-positive, 100% CD83-positive, and <1% CD14-positive) treated with IFN
-2b displayed an increase in the level of P-STAT1 compared with mature dendritic cells treated with PBS (Fig. 4, B; data not shown). These data suggest that the major subsets of immune cells exhibit different responses to IFN
in terms of their levels of Jak-STAT signal transduction. These data also suggest that the therapeutic effects of IFN
therapy may involve the actions of multiple cellular compartments.
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Other studies have documented that patients with advanced malignancies have reduced levels of critical signaling intermediates in their T cells compared with that in T cells from healthy subjects (2325). We therefore used flow cytometry to examine the intracellular levels of STAT1 and STAT2, another protein involved in Jak-STAT signal transduction, in PBMCs from patients with malignant melanoma and from healthy donors (Fig. 6). The mean levels of unphosphorylated STAT1 in PBMCs obtained from melanoma patients and healthy donors were not statistically significantly different (mean Fsp: 57.1 in healthy donors versus 48.8 in patients, difference = 8.3, 95% CI = 5.9 to 22.6, P = .232). The mean levels of STAT2 in PBMCs obtained from melanoma patients and healthy donors were also not statistically significantly different (mean Fsp: 54.3 in healthy donors versus 60.6 in patients, difference = 6.3, 95% CI = 25.7 to 13.2, P = .529). However, PBMCs from melanoma patients displayed statistically significantly more variability in median levels of STAT2 than PBMCs from healthy donors (P = .045).
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We used multiparametric staining to examine STAT1 activation within immune effector cells following treatment with IFN -2b ex vivo. PBMCs freshly procured from healthy donors and from patients with metastatic malignant melanoma were treated with PBS or IFN
-2b (104 IU/mL) for 15 minutes and analyzed for their levels of P-STAT1. Healthy donors had statistically significantly higher basal levels of P-STAT1 (FspPBS; i.e., the level of P-STAT1 in PBS-treated cells) in total PBMCs, NK cells, and T cells than melanoma patients (mean FspPBS in total PBMCs: 5.5 in healthy donors versus 1.6 in patients, difference = 3.9, 95% CI = 1.4 to 6.5, P = .004; mean FspPBS in NK cells: 4.6 in healthy donors versus 0.9 in patients, difference = 3.7, 95% CI = 1.7 to 5.7, P = .001; mean FspPBS in T cells: 6.8 in healthy donors versus 0.9 in patients, difference = 5.9, 95% CI = 2.5 to 9.3, P = .002) (Fig. 7). However, after ex vivo stimulation of PBMCs with IFN
-2b, P-STAT1 levels (FspIFN
) in total PBMCs, NK cells, and T cells from melanoma patients were not statistically significantly different from those in PBMCs, NK cells, and T cells, respectively, of healthy donors (mean FspIFN
in total PBMCs: 28.3 in healthy donors versus 32.5 in patients, difference = 4.2, 95% CI = 16.0 to 7.6, P = .472; mean FspIFN
in NK cells: 14.8 in healthy donors versus 15.6 in patients, difference = 0.8, 95% CI = 6.3 to 4.8, P = .79; mean FspIFN
in T cells: 46.0 in healthy donors versus 51.0 in patients, difference = 5.0, 95% CI = 23.7 to 13.7, P = .587) (Fig. 7). To determine the amount of STAT1 phosphorylation induced specifically in response to IFN
treatment, we used these values to calculate the specific activation of STAT1 (FspIFN
FspPBS). The specific activation of STAT1 in total PBMCs was not statistically significantly different between patients and healthy donors (mean specific activation in total PBMCs was 22.8 in healthy donors versus 30.9 in patients, difference = 8.1, 95% CI = 19.1 to 2.8, P = .139). Similar results were obtained for the specific activation of STAT1 in CD3-positive T lymphocytes (mean specific activation in CD3+ cells was 39.2 in healthy donors versus 50.1 in patients, difference = 10.9, 95% CI = 26.1 to 4.8, P = .263) and in CD56-positive NK cells (mean specific activation in CD56+ cells was 10.3 in healthy donors versus 14.7 in patients, difference = 4.4, 95% CI = 10.0 to 1.1, P = .111).
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We used our flow cytometry method to analyze STAT1 phosphorylation in immune effector cells obtained from five melanoma patients who were undergoing immunotherapy with IFN -2b. We examined the specific activation of STAT1 following administration of IFN
-2b as well as the relative efficacy of different doses of interferon with respect to activation of STAT1. For these experiments, we used PBMCs isolated from peripheral venous blood obtained immediately before and 1 hour after the patients had received various doses of IFN
-2b. Levels of P-STAT1 were strongly induced in the PBMCs of patient A, who received a high intravenous dose of IFN
-2b (20 MU/m2 [MU = million units]); most (>98%) of the PBMCs analyzed from this patient exhibited STAT1 activation (P-STAT1 Fsppretreatment = 0.35; P-STAT1 Fspposttreatment = 10.24; data not shown). Analysis of PBMCs from patient B, who received 1 MU/m2 IFN
-2b by subcutaneous injection, revealed that P-STAT1 levels also increased in response to a very low dose of IFN
-2b (P-STAT1 Fsppretreatment = 0.48; P-STAT1 Fspposttreatment = 1.34; data not shown). We also analyzed PBMCs from patient C, who received subcutaneous injections of 5 MU/m2 IFN
-2b and 10 MU/m2 IFN
-2b (the 10 MU/m2 sample was obtained 2 months after IFN
immunotherapy with 5 MU/m2 was initiated on a thrice-weekly injection schedule). P-STAT1 levels in the PBMCs of this patient also increased in response to treatments with IFN
-2b but did not increase further with the increased dosage of IFN
-2b (Fig. 8, A). Additional venous blood procured from this patient at each time point was used to isolate RNA from total PBMCs to measure activation of known IFN
-responsive genes relative to pretreatment values. Real-time RT-PCR analysis indicated that dose escalation of IFN
-2b was associated with an increase in expression of OAS-1 mRNA, but with only modest increases in the expression of ISG-54 and ISG-15 mRNAs (Fig. 8, B).
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DISCUSSION |
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Analysis of signaling within NK cells, T lymphocytes, B lymphocytes, monocytes, and dendritic cells revealed that each of these subsets of immune cells responded to IFN with an increase in P-STAT1. However, we observed that T lymphocytes and monocytes displayed more P-STAT1 after IFN
treatment than the other immune cell subsets; further analysis revealed that these differences reflected the higher basal levels of STAT1 protein in these two subsets than in the other subsets. We also found that flow cytometry was useful for measuring the basal expression of STAT1 and for monitoring variation in STAT1 activation within subsets of immune effector cells of patients who were undergoing immunotherapy. P-STAT1 was detectable in circulating PBMCs of patients who received subcutaneous administration of the lowest dose of IFN
(1 x 106 U/m2) that is used clinically. Although high-dose IFN
(i.e., 20 MU/m2 administered intravenously) routinely stimulated the phosphorylation of STAT1 in more than 98% of PBMCs, there appeared to be some inter-patient variability in the overall levels of P-STAT1 within both the NK cell and T cell compartments. In addition, analysis of a patient who received escalating doses of IFN
revealed that low levels of IFN
were just as effective in the induction of STAT signal transduction and gene expression as were higher doses that were considerably more toxic.
We have previously shown that STAT1 in melanoma cells is rapidly and reproducibly activated in response to clinically relevant doses of IFN (18). However, our interest in analyzing IFN
signal transduction in immune cell subsets was predicated on our recent discoveries that exogenous IFN
did not prolong the survival of STAT1-deficient mice challenged with STAT1-competent melanoma cells and that restoring STAT1 expression to a STAT1-deficient murine melanoma cell line did not prolong the survival of tumor-bearing normal mice receiving IFN
(4). Theoretically, the antitumor activity of IFN
in patients with malignant melanoma might involve effects on both tumor cells and immune cells. In this case, an analysis of IFN
-induced STAT1 activation in subsets of PBMCs would have to be interpreted with caution because the results might reflect the signaling events that took place within the immune cells rather than those that took place within the melanoma cells. We were therefore intrigued by recent reports that flow cytometry could be used to measure the activation state of MAP kinase, STAT1, and STAT4 on a per-cell basis (15,27,28). For example, an initial report by Fleisher et al. (15) demonstrated that flow cytometric analysis of P-STAT1 was feasible; however, this study was limited to the detection of P-STAT1 in PBMCs from healthy individuals following ex vivo stimulation with IFN-gamma and did not explore the application of this assay to the evaluation of patient PBMCs following immunotherapy with IFN
-2b (15). An elegant study by Perez and Nolan (29) extended this technology to simultaneously detect activated members of the mitogen-activated protein kinase family (i.e., p38 MAPK, p44/42 MAPK, and JNK/SAPK) and activated members of cell survival pathways (e.g., AKT/PKB) in PBMCs. The technique we describe for analyzing signal transduction within specific subsets of immune cells does not require isolation of the individual cell types from total PBMCs. Moreover, it is rapid, requires very few cells for the generation of meaningful data, and can be applied to cell preparations that have been subjected to minimal manipulation. Indeed, our comprehensive analysis of IFN
responsiveness within the NK cells, T lymphocytes, monocytes, and B lymphocytes from a single individual is the first of its kind and is not subject to the technical limitations of EMSA and immunoblot analysis.
We observed that maximal STAT1 activation and expression of IFN -responsive genes in PBMCs occurred concomitantly after in vitro stimulation of isolated PBMCs with low doses of IFN
(102103 U/mL) and that higher doses of IFN
did not lead to increased signal transduction. These findings suggested that maximal signal transduction might occur at different doses of IFN
in different patients and that this flow cytometric technique might be useful for determining the optimal doses of IFN
for patients. We found that PBMCs from a patient treated with escalating subcutaneous doses of IFN
(i.e., 5 MU/m2 and 10 MU/m2) displayed only a minimal increase in the levels of P-STAT1 and IFN
stimulated gene expression at the higher dose level than at the lower dose level. Importantly, this patient experienced a clinically significant increase in toxicity following the dose escalation to 10 MU/m2. The fact that escalated dosage of IFN
was accompanied by clinical toxicity but not increased signaling or gene expression in PBMCs challenges the assumption that IFN
must be administered at the highest tolerated dose for all patients to receive clinical benefits.
Our data also suggest that individual responsiveness to IFN at the level of Jak-STAT signal transduction may contribute to the low response rates and variable results that have been achieved with IFN
immunotherapy among patients with malignant melanoma. In a recent report by Whitney et al. (30), microarray technology was used to explore the extent of interindividual variation in gene expression within the unstimulated PBMCs of 75 healthy volunteers. They found that the greatest degree of interindividual variation occurred within a cluster of 15 genes known to be responsive to IFN. In this study, we observed some variability in the levels of P-STAT1 among melanoma patients who received high-dose IFN
. However, additional patients must be analyzed to determine whether the inter-patient variability is statistically significant. Choosing an appropriate dosage of IFN
on a patient-by-patient basis is a serious challenge for oncologists. To date, dosage determination has been accomplished either through the use of a maximally tolerated dose followed by reductions for clinically significant toxicities or by using dose-escalation schemas that also have toxicity as their primary endpoint. Our results suggest that further analysis of patient responsiveness to IFN
using flow cytometry for P-STAT1 levels and microarray techniques for gene expression is warranted.
In this study, we used flow cytometry to examine differences in components of the Jak-STAT pathway between healthy adult donors and malignant melanoma patients. We found that unstimulated PBMCs from melanoma patients had statistically significantly less P-STAT1 than unstimulated PBMCs from healthy donors, despite the fact that unstimulated PBMCs from patients and healthy donors had similar levels of the unphosphorylated STAT1 protein. The presence of reduced basal P-STAT1 levels in PBMCs from patients suggests that the Jak-STAT signal transduction pathway might be altered in patients with advanced malignancy or that immune effector cells of patients were exposed to an altered cytokine milieu. Although patient NK cells, T cells, and PBMCs have statistically significantly lower basal levels of P-STAT1 than do cells from healthy donors, the fact that this difference is lost following in vitro activation of immune cells with IFN suggests that the lower level of basal activation in patient cells is readily reversible. General defects in host immunity are common to cancer patients, and altered immune function has been routinely reported for tumor-bearing animals and cancer patients. Some of the observed defects include diminished levels of lymphocyte cytotoxic activity and proliferation (31,32), impaired production of Th1 cytokines, and reduced NK cell activity (33,34). Alterations in the levels of specific signal transduction molecules have also been reported in both tumor-infiltrating lymphocytes and peripheral lymphocytes (23). For example, defects in the T-cell receptor zeta chain, p56 (lck) kinase, ZAP-70 expression, and nuclear factor
B activation are associated with stage of disease and prognosis in patients with melanoma and renal cell carcinoma (24,25).
Despite numerous reports of abnormal cytokine signal transduction in patients with advanced malignancies, the mechanisms responsible for these signaling abnormalities remain unclear. However, because melanoma cells can secrete cytokines such as IL-6 and IL-10 (3537), it is conceivable that these factors may lead to the induction of negative regulators of IFN signal transduction and reduced levels of activated STAT proteins. Thus, an altered cytokine profile could lead to inhibitory effects on immune cells of patients with advanced malignancies (3840). Further studies involving microarray analysis of gene expression in patients treated with IFN
and continued clinical observation will be necessary to determine whether the patterns of Jak-STAT signaling that we report here are associated with a patients response to IFN
immunotherapy. However, the flow cytometric technique we used has also been used to demonstrate the ability of IL-12 pretreatments to enhance IFN
-induced signal transduction (41) and is currently being used to assess STAT1 activation in PBMCs of malignant melanoma patients who are enrolled in a nationwide Cancer and Leukemia Group B (CALGB)sponsored phase II clinical trial of IL-12 and IFN
(CALGB 500001).
We have used a sensitive and efficient flow cytometric assay to demonstrate that maximal activation of STAT1 in immune effector cells occurs in response to relatively low doses of IFN and that the host response to immunotherapy with IFN
-2b is highly variable among patients and among immune cell subsets. Until the precise molecular determinants of IFN
responsiveness are identified, it seems reasonable to use signal transduction as a surrogate marker of IFN
action in patients undergoing immunotherapy. We anticipate that this flow cytometric method could be used to rapidly determine IFN
sensitivity by using patient PBMCs ex vivo. This method could also be used as a means of identifying the dose of IFN
that produces optimal Jak-STAT signal transduction and gene regulation on a patient-by-patient basis.
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We thank Dr. James W. Jacobberger for helpful suggestions with data analysis and Mark Kotur in the Dorothy Davis Heart and Lung Research Institute Flow Cytometry and Cell Analysis Core (The Ohio State University) for technical assistance.
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Manuscript received November 3, 2003; revised June 28, 2004; accepted July 20, 2004.
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