Low-dose photodynamic therapy promotes angiogenic potential and increases immunogenicity of human mesenchymal stromal cells

Journal of Photochemistry & Photobiology, B: Biology 199 (2019) 111596

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Low-dose photodynamic therapy promotes angiogenic potential and increases immunogenicity of human mesenchymal stromal cells

T

Olga O. Udartsevaa,c, , Olga V. Zhidkovac, Maria I. Ezdakovac, Irina V. Ognevac,d, Elena R. Andreevac, Ludmila B. Buravkovac, Sandra O. Gollnicka,b ⁎

a

Department of Cell Stress Biology, Roswell Park Comprehensive Cancer Center, Elm & Carlton Str, Buffalo, NY 14263, USA Department of Immunology, Roswell Park Comprehensive Cancer Center, Buffalo, NY 14263, USA c Institute of Biomedical Problems, Russian Academy of Science, Moscow, Russia d I.M. Sechenov First Moscow State Medical University, Moscow, Russia b

ABSTRACT

Photodynamic therapy (PDT) is a non-invasive FDA and EMA-approved anticancer treatment modality. Initially developed for elimination of malignant cells, PDT affects all cells in the tumor bed including stromal cells. Stroma represents not only an important component of tumor microenvironment, but has a significant impact on tumor susceptibility to PDT and other anticancer therapies. However, the effects of PDT on stromal cells are poorly investigated. During PDT the tumor stroma can receive low-dose irradiation as a result of chosen regimen or limited depth of light penetration. Here, we characterized response of human mesenchymal stromal cells (MSCs) to low-dose PDT. In an in vitro model we demonstrated that low-dose PDT resulted in activation of Erk1/2 and inhibition of GSK-3 signaling in MSCs. PDTmediated induction of intracellular reactive oxygen species (ROS) resulted in reorganization of MSC cytoskeleton and decreased cell motility. More importantly, lowdose PDT dramatically upregulated secretion of various proangiogenic factors (VEGF-A, IL-8, PAI-1, MMP-9, etc.) by MSCs and improved MSC ability to promote angiogenesis suggesting an increase in the pro-tumorigenic potential of MSCs. In contrast, co-cultivation of PDT-treated MSCs with lymphocytes resulted in significant decrease of MSC viability and potential increase in MSC immunogenicity, which may lead to increased anti-tumor immunity. Low-dose PDT in MSCs significantly inhibited secretion of CCL2 (MCP-1) potentially limiting infiltration of pro-tumorigenic macrophages. Altogether, our findings demonstrate that lowdose PDT significantly modifies functional properties of MSCs improving their pro-tumorigenic potential while simultaneously increasing potential immune stimulation suggesting possible mechanisms of stromal cell contribution to PDT efficacy.

1. Introduction Mesenchymal stromal cells (MSCs) are defined as adult non-hematopoietic fibroblast-like multipotent cells expressing CD73, CD90, CD105 and lacking CD45 expression [1,2]. While originally discovered in the bone marrow [3], studies of the last two decades show that MSCs have perivascular origin [4,5] and can be expanded from different tissues (bone marrow, adipose tissue, umbilical cord blood, etc.) [6,7]. Adipose tissue is currently considered the most abundant and accessible source of adult MSCs. Two key functional activities of MSCs, which define their physiological role, have been described. MSCs produce components of extracellular matrix (and enzymes for its remodeling) and secrete a variety of growth factors and cytokines to promote cell proliferation, angiogenesis and wound healing [8,9]. In addition, these cells demonstrate broad immunomodulatory abilities: MSCs have been shown to regulate immune responses by interacting with components of innate and adaptive immune systems and display both anti-inflammatory and pro-inflammatory effects [10–12]. These features make MSCs an attractive

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candidate for regenerative and cell therapies [8–10,13]. The same characteristics determine MSC pro-tumorigenic activity and make these cells a key player in cancer progression. MSCs that mobilize to a tumor site represent the main source of cancer-associated fibroblasts [14–16]. Dynamic and bidirectional interplay between cancer cells and stroma create tumor-supportive microenvironment and promote tumor growth. MSCs have been shown to interact with cancer cells directly through gap junctions and indirectly through soluble factors (cytokines, chemokines, growth factors, metabolites, and proteolytic enzymes), extracellular vesicles and exosomes. This cross-talk induces metabolic reprogramming of tumor cells, promotes their proliferation and epithelialmesenchymal transition, which implements an invasive phenotype and metastatic capacity of malignant cells [16,17]. MSC-derived pro-angiogenic factors (angiopoietins, VEGF-A, IL-8, IGF-1, FGFs) as well as MSCmediated remodeling of extracellular matrix stimulate tumor neovascularization [15,16,18]. At the same time MSCs orchestrate tumor-promoting inflammation regulating chemotaxis, activation, function, survival of immune cells and inducing immunosuppression [10–12,18]. Growing evidence supports the idea that MSCs not only promote

Corresponding author at: Department of Cell Stress Biology, Roswell Park Comprehensive Cancer Center, Elm & Carlton Str, Buffalo, NY 14263, USA E-mail address: [email protected] (O.O. Udartseva).

https://doi.org/10.1016/j.jphotobiol.2019.111596 Received 13 June 2019; Received in revised form 23 July 2019; Accepted 14 August 2019 Available online 16 August 2019 1011-1344/ © 2019 Published by Elsevier B.V.

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tumorigenesis, but also mediate resistance to anti-cancer therapies [18]. Tumor-associated MSCs can limit access of therapeutic agents to their target tissues, promote survival of chemotherapeutically-treated tumor cell, protect malignant cells from radiation and induce “pathway switch” during targeted therapies [19–21]. Photodynamic therapy (PDT) is a FDA and EMA-approved non-invasive therapeutic modality used for treatment of malignant and nonmalignant diseases [22,23]. PDT is based on the reaction of photosensitization – activation of photosensitive drug (photosensitizer) with the light of an appropriate wavelength [22,24]. This reaction requires the presence of oxygen, and results in generation of cytotoxic reactive oxygen species (ROS). The efficacy of PDT depends on photosensitizer uptake by target cells, penetration of light and chosen regimen [22,25]. Originally this technique was developed to ablate malignant or certain undesirable tissues by photodamage followed by acute inflammation [25,26]. Recently it has been demonstrated that low-dose PDT effectively stimulates anti-tumor immunity [27,28], and therefore can be used for combination therapies. PDT affects not only cancer cells, but all cells in the tumor microenvironment [23]. With both high- or low-dose PDT parts of the tumor including stromal cells receives low-dose irradiation, which induces low level of intracellular ROS as a result of chosen regimen or limited depth of light penetration [25]. ROS in non-toxic concentrations represent an important regulator of intracellular signaling [29,30]. In this study we for the first time characterized the effects of low-dose PDT on human MSCs and explore possible mechanisms involved in PDT-mediated regulation of MSC functional activity.

maintained under 20% O2 in α-MEM supplemented as indicated above. For some experiments MSCs were cultured under normobaric acute hypoxic conditions using hypoxic chambers (Stem Cell Technology, USA) where O2 concentration was controlled by an O2 sensor and maintained at 1%. MSCs from the 2nd to 4th passages and at 80–90% confluence were used for experiments. Human peripheral blood mononuclear cells (PBMCs) were isolated from blood samples collected from healthy volunteers who had given their written informed consent. Cells were separated using density gradient centrifugation with Histopaque-1077 (Sigma, USA) and maintained in RPMI-1640 (Gibco, Life Technologies, USA) supplemented with 5% heat-inactivated FBS, 50 U/ml penicillin, and 50 μg/ml streptomycin. For some experiments mononuclear cells were stimulated with 10 μg/ml phytohemagglutinin PHA-P (Sigma-Aldrich) for 24 h. 2.3. Co-cultivation of MSCs With PBMCs PBMCs were isolated from peripheral blood as described above. MSCs were plated in 6-well plates at a density of 105 cells/well, photosensitizer was added 24 h after plating followed by PDT treatment. 5*105 ml−1 PHA-activated or naϊve PBMCs were added to control MSC or PDT-treated MSCs (30 min after irradiation) for 24 h. 2.4. In vitro PDT Treatment Cells at 80–90% confluency were incubated with 10 μg/ml Photosens (Al-phthalocyanine, NIOPIK, Russia) for 24 h protected from light. Chosen concentration (10 μg/ml) corresponds to Photosens level in patient's serum before PDT according to Photosens pharmacokinetics provided by manufacturer. Use of lower Photosens concentrations resulted in significant decrease of its intracellular accumulation (Fig. S1a) and significantly impaired PDT efficacy (Fig. S1b). After incubation culture medium was replaced with fresh media without photosensitizer, and cells were irradiated using the diode laser light (λ = 675 nm) with a fluence of 500 mW/cm2 at a total dose of 0.25 J/cm2. Non-irradiated cells incubated with Photosens were used as a control. Effects of PDT were analyzed 0.5–24 h after irradiation. Conditioned media was collected 24 h after PDT. Dose-dependent analysis of PDT efficacy was performed using single concentration of Photosens (10 μg/ml) and variable irradiation doses (0.25–50 J/cm2). In the experiments on MEK1 inhibition, PD098059 (25 μM; Cayman, USA) was added to the MSC monolayers 1 h before and right after PDT in the fresh culture media.

2. Methods and Materials 2.1. Patients and Clinical Samples Adipose tissue samples were obtained from the subcutaneous abdominal depots of patients undergoing dermolipectomy at the Souz Multidisciplinary Clinic (Moscow, Russia), as part of a scientific agreement. Written informed consent was obtained from all participants, and all procedures were approved by the Biomedicine Ethics Committee of the Institute of Biomedical Problems, Russian Academy of Sciences (Physiology Section of the Russian Bioethics Committee, Russian Federation National Commission for UNESCO, Permit #314/ МCK/09/03/13). 2.2. Cell Isolation and Culture The adipose tissue samples were rinsed with sterile phosphate-buffered saline (PBS; Gibco, Life Technologies, USA), minced mechanically, and isolation of stromal vascular fraction cells was performed. Briefly, minced tissue pieces were enzymatically digested with 0.075% (equivalently 40.7 U/ml) collagenase type I (Sigma-Aldrich, USA) for 30 min at 37 °C with agitation. Subsequently, the enzyme was inactivated with an equal volume of α-MEM (Gibco, Life Technologies, USA) containing 10% fetal bovine serum (FBS; BI Biological Industries, Israel). Then the tissue was disintegrated by pipetting up and down and filtering through a 100 μm nylon mesh (Corning, USA). The stromal vascular fraction, containing the MSCs, was obtained by centrifuging the sample at 500 ×g for 10 min. Cells were plated in α-MEM supplemented with 10% FBS, 50 U/ml penicillin (Sigma, USA), and 50 μg/ml streptomycin (Sigma, USA) at 37 °C, 20% O2 and 5% CO2. After 24 h cells were carefully washed with PBS to remove non-adherent cells and debris. According to a joint statement of the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society for Cellular Therapy (ISCT), adherent cells are MSCs [1,2]; these cells were expanded until they reached 90% confluency. Then, cells were harvested and analyzed for the expression of MSC surface antigens and multilineage differentiation potential [31]. After characterization cells were split at a density of 3000 cells/cm2 and

2.5. Detection of Reactive Oxygen Species (ROS) and Mitochondrial Activity Mitochondrial activity was analyzed using 500 nM MitoTracker Red CMXRos (Thermofisher, USA). Dye was added to MSCs immediately after irradiation and incubated for 30 min. Cells were then harvested and analyzed by flow cytometry. The spectra overlay of Photosens and Mitotracker Red CMXRos is presented on Fig. S1c. In the experiments on MSCs and PBMCs co-cultivation, 5 μM JC-1 (Thermofisher, USA) was used for analysis of mitochondrial transmembrane potential according to manufacturer's instructions. Briefly, dye was added to control or PDT-treated MSCs for 1 h. Then cell were washed with PBS, co-cultivated with naïve or PHA-activated PBMCs overnight and analyzed by flow cytometry. Total of ROS and mitochondrial superoxide were detected using CM-H2DCFDA and MitoSOX Red (Thermofisher, USA) respectively according to manufacturer's instructions. Briefly, 10 μM CM-H2DCFDA or 5 μM MitoSOX Red in serum-free medium or PBS were added to MSCs pre-incubated with Photosens 5 min before irradiation. After irradiation cells were incubated for 15 min in a CO2-incubator protected from light, then harvested with trypsin and analyzed by flow cytometry. The spectra overlay of Photosens and MitoSOX Red is presented on Fig. S1d. 2

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2.6. Analysis of Cell Viability

phalloidin staining, cells were fixed with 4% formaldehyde in PBS for 15 min and permeabilized with 0.1% Triton X-100 for 10 min followed by staining with the primary anti-vimentin mouse monoclonal antibody (Chemicon, Millipore, USA) or anti-tubulin mouse monoclonal antibody (Santa Cruz Biotechnology, USA) for 1 h at 37 °C. Subsequently, rhodamine phalloidin (Molecular Probes, Life Technologies, USA) and Alexa Fluor 488-conjugated anti-mouse IgG secondary antibodies (Molecular Probes, Life Technologies, USA) were added for 1 h. Then, the samples were washed and mounted with Fluoroshield with DAPI (Sigma-Aldrich). Images were acquired using an LSM 780 (Carl Zeiss, Germany) confocal microscope.

Cell viability after PDT was analyzed by MTT test to detect the most appropriate irradiation dose. LD50 (median lethal dose or lethal dose, 50%) was defined as PDT dose which causes the death of 50% cells. Cell viability after co-cultivation with lymphocytes was analyzed using an Annexin V-FITC/PI Kit (Immunotech, France). Briefly, MSCs were incubated with Annexin V-fluorescein isothiocyanate (FITC) and propidium iodide (PI) for 15 min at 4 °C protected from light. Cells were then analyzed by flow cytometry. Cells negative for AnnexinV-FITC and PI staining were defined as viable, cells positive for Annexin V-FITC were defined as apoptotic, and cells negative for AnnexinV-FITC and positive for PI staining were defined as necrotic.

2.11. ELISA

2.7. Atomic Force Microscopy

Conditioned media was collected 24 h after PDT, centrifuged at 2500 g to remove cell debris, frozen and stored at −70 °C. Concentrations of IL-6, IL-8, MCP-1 and TGFβ were measured by Human IL-6/ IL-8/ MCP-1/ TGFβ ELISA kits (BD Bioscience, USA) respectively. VEGF-A was detected using Human VEGF-A ELISA kit (Peprotech, USA).

Cell transversal stiffness was measured using a Solver P47-Pro instrument (NT-MDT, Moscow, Russia). MSCs were grown on glass coverslips for cell culture (Corning, USA). When cells reached ~80–90% confluency, PDT was performed as described above. Twenty four hours after PDT coverslips with MSC monolayers were mounted onto the liquid cell of the atomic force microscope adjusting table. Force-distance curves were obtained in contact mode using a soft silicon cantilever. The cantilever spring constant (Microlever, Park Scientific Instruments, USA) and radius of the tip curvature were 0.01 N/m and 10 nm, respectively. The actual indentation depth and force applied were calculated using the following formula: hs = x − y · a, Fs = y · a · kc, where h s is the actual indentation depth (m), Fs is the actual force applied to a cell (N), and kc is the cantilever stiffness coefficient. At an indentation depth of 150 nm, the change in applied force was determined and cell stiffness was estimated using the following formula: ks = Fs/hs. The results were processed using MATLAB 6.5 software developed especially for this research.

2.12. Chorioallantoic Membrane Assay in Ovo MSC angiogenic activity after PDT was evaluated by CAM assay in ovo using Japanese quail embryos. The fertilized eggs were placed into the incubator and kept at 37 °C for 6 days followed by application of 30 μl MSC conditioned media collected before or 24 h after PDT to the surface of CAMs for 24 h. The embryos were fixed with 4% paraformaldehyde/2% glutaraldehyde solution in PBS and stained with Carracci's hematoxylin. Morphometric analysis was performed using AngioQuant software (www.cs.tut.fi). The vascularization of chorioallantoic membrane was evaluated by calculation of numbers of tubule complexes and junctions.

2.8. In Vitro Wound-Healing Assay

2.13. Protein Extraction and Western Blotting

MSC migration was evaluated in cell monolayers at a cell density of 104 cells per cm2 using the in vitro “scratch” assay. Twenty four hours after irradiation confluent monolayers were scratched with a sterile pipette tip to create a “wound” approximately 0.8–1.0 mm wide. Culture medium was replaced with α-MEM supplemented with 10% FBS to remove cell debris. All scratch assays were performed in six replicates. To estimate the wound closure, serial digital images were captured with a Nikon Eclipse Ti-U microscope (Nikon, Germany) immediately after and at specific time intervals (3, 6, 9, and 24 h) after the scratch. The images were analyzed using NIS-Elements software (Nikon, Germany) which measured the width of the scratch at previously marked points (five per Petri dish) along its length. The migration area was calculated as the difference between the initial and final wound squares.

Cells were lysed with a lysis buffer containing 63 mM Tris-HCl, 10% glycerol, 5% β-mercaptoethanol, 2% SDS (pH 6.8), and Halt Protease Inhibitor Cocktail (ThermoFisher Scientific, USA) on ice. After electrophoresis in SDS-polyacrylamide gel (SDS-PAGE) gel proteins were transferred onto nitrocellulose membranes followed by staining with specific primary, secondary antibodies and HRP-conjugated streptavidin-peroxidase. Membranes were developed using ECL substrate (BioRad, USA). Signal was detected using ChemiDoc XRS+ imaging system (Bio-Rad, USA) and analyzed using Image Lab Software (Bio-Rad, USA). 2.14. Protein Arrays Proteome Profiler Human Phospho-MAPK and Proteome Profiler Human Angiogenesis Array kits (R&D, USA) were used to profile phosphorylation of MAPKs or expression of angiogenesis-related proteins respectively. All procedures were performed according to manufacturer's instructions. Cells were lysed 30 min after PDT using Cell Lysis buffer included into Phospho-MAPK Array kit. Conditioned media collected 24 h after PDT was used for the Angiogenesis Array.

2.9. Transwell Migration Assay In these experiments PHA-activated peripheral blood mononuclear cells (PBMCs) secreting chemokines served as a chemoattractant for MSCs. Human PBMCs were stimulated with PHA-P for 24 h. PBMCs were washed with PBS, resuspended in culture media (5*105 cells/ml) and added to the lower chamber of Transwell according to the manufacturer's instructions. MSCs before or 24 h after low-dose PDT were harvested and plated into the upper chamber of an insert well (3000 cells/cm2). The experimental design of the experiment is schematically presented on Fig. 3c.

2.15. Quantitative RT-PCR Analysis To evaluate gene expression, total RNA was extracted using QIAzol Reagent (Qiagen, USA) and purified by the phenol/chloroform technique. Reverse transcription was performed using a QuantiTect Reverse Transcription Kit (Qiagen, USA). qPCR was performed using the Mx300P system (Stratagene, USA) using SYBR green MasterMix (Qiagen, USA). The expression of target genes was normalized to HPRT and quantified by the 2−ΔΔCT method.

2.10. Fluorescent Staining MSCs were grown on coverslips. For immunofluorescent and 3

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2.16. Statistical Analysis

effects of low-dose PDT on MSCs permanently cultured under 20% O2 (“normoxia”) and 1% O2 (“acute hypoxia”). Reduced mitochondrial activity and increased total intracellular ROS level were demonstrated for MSCs cultured under low oxygen tension (Fig. S2a,c) while level of mitochondrial superoxide wasn't affected by acute hypoxia (Fig. S2b). A range of light doses were tested to establish optimal conditions for low-dose PDT on MSCs. As shown in Fig. 1c and S2a, despite difference in the basal intracellular ROS level modulation of light dose at a constant photosensitizer concentration of 10 μg/ml resulted in similar dosedependent and controlled induction of ROS in MSCs cultivated under 20% and 1% O2. At the same time, induction of intracellular ROS by low-dose PDT was more effective for hypoxic MSCs, and the dose of 0.125 J/cm2 resulted in prominent increase of intracellular ROS (Fig. S2a). In MSCs cultured under normoxia, a dose of 0.5 J/cm2 induced a moderate level of total intracellular ROS (Fig. 1b) and high level of mitochondrial superoxide (Fig. 1c) which were measured with fluorescent dyes CM-H2DCFDA and MitoSOX, respectively. PDT-mediated production of mitochondrial superoxide was impaired under acute hypoxia (Fig. S2b), which can be explained by down-regulation of mitochondrial activity and shift of cellular metabolism towards glycolysis. In addition, the PDT dose of 0.5 J/cm2 decreased mitochondrial activity (Fig. 1d, S2c) and impaired MSC viability by 20% (Fig. 1e, S2d) under both normoxic and acute hypoxic conditions. A light dose of 0.25 J/cm2 induced a low level of intracellular ROS and mitochondrial superoxide, and didn't affect MSC viability. Therefore, dose of 0.25 J/cm2 was selected to examine the effect of low-dose PDT on MSCs.

Statistical analysis was performed using GraphPad Prism 7.03 software. Each experiment was performed at least thrice with consistent results. The Mann Whitney U test was used to assess difference between groups. Data were presented as median ± range. A two-tailed value of p < .05 was considered statistically significant. 3. Results 3.1. The Effect of PDT on Cell Viability and Intracellular ROS Production To evaluate susceptibility of tumor microenvironment to PDT, we compared cell viability of primary isolated human MSCs, PB-Mφ (macrophages from peripheral blood) and HUVECs (human umbilical vein endothelial cells) after PDT. Photosens which was used in the study as a photosensitizer, represents a water-soluble mixture of sulfonated Al-phthalocyanines. PDT using Photosens induced cell death in a dosedependent manner (Fig. 1a). Interestingly, endothelial cells were highly susceptible to PDT while MSCs and Mφ were more resistant: the LD50 was 0.5 and 2 J/cm2 for HUVECs and MSCs/Mφ respectively. In the present study we focused on the effect of non-toxic low-dose PDT on MSCs. MSCs were isolated from stromal vascular fraction of human adipose tissue samples and characterized as previously described [33]. Tumors are known to be low oxygenated while hypoxia significantly modulates cell functional activity and promotes tumor resistance to anti-cancer therapies. Therefore, we evaluated some

Fig. 1. PDT allows regulated intracellular ROS induction in stromal cells. a Susceptibility of endothelial cells (HUVECs), MSCs and PB-macrophages to PDT of different intensity (N = 5). Cells were incubated with Photosens (10 μg/ml) for 24 h prior to laser diode exposure (1 W/cm2; λ = 675 nm). b PDT-mediated dosedependent induction of intracellular ROS in MSCs (N = 5). CM-H2DCFDA was added to cells immediately after PDT for 30 min. Intracellular ROS level was analyzed by flow cytometry. c Flow cytometric analysis of mitochondrial superoxide detected by MitoSox Red staining in MSCs after PDT (N = 4). d Effect of 0.25 and 0.5 J/ cm2 PDT on mitochondrial activity (N = 3). Cell were stained with Mitotracker Red CMXRos and analyzed by flow cytometry. e Comparison of MSCs viability after 0.25 and 0.5 J/cm2 PDT (N = 5). Data are represented as median ± range. *P < .05, ***P < .001. 4

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Fig. 2. Low-dose PDT induces cytoskeleton reorganization. a Representative images of MSC morphology before and after 0.25 J/cm2 PDT (phase contrast microscopy; scale bar 100 um). b Quantification of MSC cell spread area before and after 0.25 J/cm2 PDT. Measurement of cell area was performed using ImageJ software. c Microtubule (left panel) and intermediate filament (right panel) organization in MSCs before and after PDT. Cells were stained with antibodies for β-tubulin and vimentin and analyzed by fluorescent confocal microscopy (scale bar 50 um). d Low-dose PDT induces microfilament reorganization in MSCs (left panel; scale bar 50 um). Confocal microscopy sections of the actin filament network at the basal surface (i) and apical surface (ii) of MSCs after 0.25 J/cm2 PDT (right panel; scale bar 20 um). Cells were stained for F-actin with phalloidin and analyzed by fluorescent confocal microscopy. e Representative force-distance curves obtained by atomic force microscopy during measurement of cell stiffness. The curves demonstrate a typical dependency of the applied force on the depth of indentation. Dots show experimental points; curves represent the least-squares fitting experimental equation. f Comparison of MSCs stiffness (pN/nm) before and after 0.25 J/cm2 PDT. Data represent median ± range. *P < .05.

3.2. Low-Dose PDT Leads to Cytoskeleton Reorganization and Increases Cell Stiffness

organization; therefore, we compared these parameters in MSCs before and 24 h after low-dose PDT. In untreated cells tubulin staining demonstrated a perinuclear distribution, with filaments extending towards cell periphery (Fig. 2c, upper left panel). PDT had no effect on microtubule organization, and tubulin structures remained stable after irradiation in spite of changes in cell shape (Fig. 2c, bottom left panel). Vimentin formed an extended network in control MSCs, from the

We found that low-dose PDT altered MSC morphology: cells became less flattened and more spindle-shaped, cell spread area was decreased, and cell-cell interactions were reduced (Fig. 2a–b, S3a). Cell morphology is usually determined by cell/matrix stiffness and cytoskeleton 5

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perinuclear region to the cell periphery (Fig. 2c, upper right panel). Upon low-dose PDT condensation of perinuclear vimentin and disassembly of peripheral intermediate filaments was observed (Fig. 2c, bottom right panel). The post-PDT reorganization of actin microfilaments was even more pronounced. In control cells confocal sectional images of actin staining revealed thick actin filament bundles organized into two typical types of structures: conventional basal stress fibers (ventral and dorsal) and perinuclear actin cap (Fig. 2d). Actin cap was represented with parallel microfilaments organized into a curved shell above the nucleus. It was aligned with the overall cell orientation and remained unchanged upon PDT-mediated intracellular ROS induction (Fig. 2d (ii)). At the same time, low-dose PDT induced disassembly of ventral stress fibers, formation of perinuclear basal star-like superstructures and stabilization of dorsal stress fibers and cortical actin (Fig. 2d (i)). Similar PDT-mediated disorganization of actin microfilaments was found in MSCs permanently cultivated under acute hypoxia (Fig. S3b). Next, we evaluated effects of low-dose PDT on cell stiffness (Fig. 2e). Stabilization of cortical actin has been associated with increased cell stiffness, whereas a reduction of cell spreading correlates with lower cell stiffness [32]. Accordingly, atomic force microscopy (AFM)-based indentation measurements revealed that MSCs 24 h after low-dose photodynamic treatment were significantly stiffer compared to non-irradiated ones (Fig. 2f). We conclude that low-dose PDT doesn't affect microtubules in MSCs, but induces reorganization of actin and intermediate filaments which is associated with cell stiffening.

low-dose PDT inhibited anti-angiogenic IGFBP-3 production, and stimulated secretion of pro-angiogenic IL-8, VEGF-A and serpin E1 as well as matrix metalloproteinases MMP-8 and MMP-9 (Fig. 4a–b, S4a). To validate these findings, we measured the levels of IL-8 and VEGF by ELISA in conditioned media of MSCs cultured under 20% and 1% O2 (Fig. 4c, S4b). In addition, expression of TGFβ and IL-6 – multifunctional cytokines with known pro-angiogenic activity – was analyzed by ELISA (Fig. 4c, S4b). TGFβ wasn't detected by the array (analyte #9, Fig. 4a, S4a) due to low expression and method sensitivity, and IL-6 wasn't included into the array as a target. We demonstrated that low-dose PDT stimulated secretion of IL-6, IL-8 and VEGF-A (consistent with the array observation) by MSCs cultured under 20% and 1% O2, but had no effect on TGFβ production. One of the approaches developed to evaluate effects of pro- and anti-angiogenic factors is the avian chorioallantoic membrane (CAM) assay which has been shown to be cost-effective and efficient tool for in vivo (in ovo) analysis of angiogenesis [33]. In the present study we evaluated effects of conditioned media from control and low-PDTtreated MSCs on CAM vascular architecture. CAMs treated with αMEM served as an internal control. Application of MSC conditioned media (Fig. 4d) didn't affect development of quail embryo itself (data not shown). A few blood vessels were detectable in CAMs treated with αMEM (Fig. 4e, f). Condition media from control MSCs significantly stimulated CAM vascularization (Fig. 4f) which was characterized by increased number of tubule complexes and junctions. Analysis of CAM vascular architecture after application of PDT-MSC conditioned media revealed remarkable increase of vessel density (Fig. 4f). In addition, the number and length of tubule complexes and total vessels, but not junctions, was significantly higher in CAM stimulated with PDT-MSCs compared with control ones (Fig. 4f, g). Altogether we demonstrated that low-dose PDT significantly improves angiogenic potential of MSCs.

3.3. Low-Dose PDT Reduces Cell Motility To analyze effects of low-dose PDT on MSC motility, in vitro “wound-healing” (scratch) and chemotaxis assays were performed with control MSCs or MSCs 24 h after low-dose PDT. MSCs permanently cultured under 1% O2 demonstrated reduced motility in scratch assay (Fig. S3c,d). Low-dose PDT under both normoxic and acute hypoxic conditions caused significant defect in MSC migration along the wound edge (Fig. 3a–b, S3c–d). The gap closure rates calculated 24 h after scratching were ≥4-fold lower for PDT-treated MSCs when compared to control MSCs (Fig. 3b, S3d). Chemotaxis was analyzed using the Transwell® migration assay. MSCs before or 24 h after low-dose PDT were plated into the upper chamber of an insert well, and human PHA-activated peripheral blood mononuclear cells (PBMCs) secreting chemokines were plated into the lower chamber (Fig. 3c). In the absence of chemoattractant (PHA-activated PBMCs) few MSCs passed through membrane pores, and there was no difference between control and PDT-treated cells (data not presented). Cell migration was significantly increased with the addition of PHA-activated PBMCs to the lower chamber. Low-dose PDT attenuated MSC chemotaxis: the number of PDT-MSCs migrating through Transwell® membrane was much lower compared with intact MSCs (Fig. 3d,e). Taken together, we demonstrated that low dose PDT prominently inhibits MSC migration and chemotaxis.

3.5. Low-Dose PDT Changes MSC-PBMC Interaction Interactions of MSCs with immune cells play an important role in the regulation of their functional activity. Here we analyzed effects of low-dose PDT on different aspects of MSC-immune cell interactions. We evaluated effects of low-dose PDT on monocyte chemoattractant protein-1 (MCP-1, CCL2) level in supernatants of MSCs cultured under 20% and 1% O2. In both cases PDT-treated MSCs secreted significantly less MCP-1 compared with control MSCs (Fig. 5a, S5a). However, the effect was less pronounced under acute hypoxia (Fig. S5a). Co-cultivation with non-activated PBMCs dramatically increased MCP-1 production, and PHA-mediated pre-activation of PBMCs further improved stimulation of MCP-1 secretion (Fig. 5b). At the same time low-dose PDT attenuated this effect, and MCP-1 levels in conditioned media from cocultures of PBMCs and PDT-treated MSCs were significantly lower compared with co-cultures of PBMCs with control MSCs (Fig. 5b). MSCs are considered to be hypo-immunogenic or immune evasive which allows allogenic transplantation to be safe and makes MSCs an attractive target for regeneration therapies. We evaluated MSC immunogenicity by assessing their viability after direct in vitro co-cultivation with allogenic PBMCs. We showed that control MSC were hypoimmunogenic as expected, and co-cultivation of MSCs with allogenic PBMCs had no impact on their viability (Fig. 5c–e, S5b). While low-dose PDT didn't affect MSC viability, it dramatically impaired their evasiveness and improved their recognition by PBMNs: co-cultivation with allogenic PBMCs resulted in dramatic decrease of MSC mitochondrial potential, induction of MSC cell death and cell lysis (Fig. 5c–e, S5b). We further analyzed effects of low-dose PDT on MSC-mediated activation of PBMCs. The activation was assessed by expression of CD25 and CD69 on CD3+ T-lymphocytes. Co-cultivation of PBMCs with either control or PDT-treated MSCs didn't affect T-lymphocyte activation (Fig. 5f, g). The number of Th (CD4+) cells was slightly increased after co-cultivation with control or PDT-treated MSCs (Fig. 5h) while no

3.4. Low-Dose PDT Promotes MSC Angiogenic Activity Stromal cells represent one of the main sources of proangiogenic factors in the tumor. Here we evaluated effects of low-dose PDT on MSC angiogenic activity. To identify key MSC-secreted proangiogenic factors affected by low-dose PDT, we utilized a human cytokine antibody array that allowed analyzing protein expression of up to 55 regulators of angiogenesis in conditioned media. It was demonstrated that MSCs predominantly secreted 3 proangiogenic cytokines (thrombospondin-1, VEGF-A and IL-8) and 1 anti-angiogenic cytokine (IGFBP-3), 2 matrix metalloproteinases (MMP-8, MMP-9) and 2 inhibitors of matrix metalloproteinases (serpin E1 and TIMP-1) (Fig. 4a–b, S4a). We found that 6

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Fig. 3. Low-dose PDT impairs MSC motility. a Analysis of MSCs migration before and after PDT by in vitro scratch assay. Representative images demonstrate reduced ability of low-dose PDT-treated MSCs to close the experimental wound in vitro (phase contrast microscopy; scale bar 100 um). b Quantification of gap closure (a) rates (N = 4). Scratch areas were measured right after scraping and 24 h after scraping using ImageJ software. Gap closure rates were calculated as difference of scratch areas between two measurements (0 h and 24 h). c Schematic design of Transwell chemotaxis assay. d Representative images demonstrating impaired ability of MSC to migrate towards PHA-activated PBMCs (chemoattractive stimuli). The membranes were fixed with methanol and stained with propidium iodide (fluorescent microscopy; scale bar 100 um). e Analysis of PDT effects on MSC chemotaxis by Transwell migration assay. Cells before or after 0.25 J/cm2 PDT were seed to the upper chamber as represented (c), and numbers of MSCs migrated through membrane were counted 24 h after. Data represent median ± range. *P < .05.

difference was found in percentage of B-cells (CD19+), CTL (CD8+), NK (CD3-CD56+) and NKT (CD3 + CD56+) (Fig. 5h, S5c). Increased recognition of MSCs subjected to low-dose PDT and their lysis by allogenic PBMCs can be related to PDT-mediated induction of MHC Class I expression in MSCs followed by activation of cytotoxic T-lymphocytes and natural killer (NK) cells in MSC-PBMC co-cultures. Therefore, we evaluated expression of MHC Class I-related molecules in MSCs and expression of stimulating and activating receptors in cytotoxic T-lymphocytes and NK cells. The analysis of HLA-ABC, MICA/B (NKG2D ligand) and Nectin-2 (DNAM-1 ligand) expression didn't reveal any difference between PDT-treated and non-treated MSCs (Fig. S5d). Cocultivation with MSCs subjected to low-dose PDT didn't affect expression of CD69, NKG2D and DNAM-1 in CD56/CD16+ cells (Fig. S5e). Thus, we demonstrated that low-dose PDT significantly modifies MSC interactions with leukocytes, which can affect PDT outcome.

demonstrated that low-dose PDT dramatically increased HSP27, Act2, JNK2, Erk1/2 and GSK3α/β phosphorylation (Fig. 6a, b). Activation of Erk1/2 and GSK3α/β signal transduction plays an important role in regulation of cell functional activity and fate; thus we chose these two kinases for further analysis: PDT-mediated Erk1/2 and GSK3α/β phosphorylation was analyzed in time-dependent manner. We showed that low-dose PDT rapidly induced phosphorylation of both kinases (30 min after irradiation). Erk1/2 phosphorylation dropped down by 6 h after PDT and returned to base line level by 24 h (Fig. 6c, d). The level of GSK3α/β phosphorylation remained elevated in PDT-treated MSCs compared to non-treated ones for at least 24 h following irradiation (Fig. 6e, f). However, the slow decrease of GSK3α/β phosphorylation was observed by 6 h after low-dose PDT, and it was further reducing for the next 18 h. Blocking of Erk1/2 phosphorylation by MEK1 inhibition with PD098059 prior PDT didn't affect MSC migration (Fig. S6a), but partially abrogated PDT-mediated enhancement of MSC proangiogenic activity (Fig. S6b). Recently the role of Erk1/2 activation and GSK3α/β inhibition in the induction of HIF1α expression and its stabilization under normoxic conditions was identified [34]. To evaluate whether low-dose PDT can induce HIF1α, HIF1α gene expression in MSCs was analyzed 15′, 1 h, 3 h, 6 h and 24 h after irradiation. We found that HIF1α expression remained stable during the first 3 h after PDT, prominently increased by

3.6. Low-Dose PDT Modifies MSC Functional Activity Through GSK-3/ ERK/HIF-1α Pathway Here we attempted to find mechanisms underlying PDT-mediated modulation of MSC functional activity. To identify key pathways involved in regulation of MSC functions upon low-dose PDT, we performed protein phospho-MAPK array with MSC lysates. It was 7

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Fig. 4. Low-dose PDT stimulates proangiogenic activity of MSCs. a Angiogenesis-related proteins in MSC supernatants before and after PDT were profiled using Human Angiogenesis Antibody Array. b Densitometric analysis of selected analytes in antibody arrays were quantified using ImageLab software (N = 4). c Level of VEGF-A, IL-8, IL-6 and TGFβ was measured in MSC conditioned media (CM) before and after 0.25 J/cm2 PDT. d Demonstration of sample loading to the quail CAM. e Schematic representation of CAM morphometric analysis using AngioQuant software (www.cs.tut.fi). Example of tubule complexes (1) and junctions (2) are shown with arrows. f Representative images for Chorioallantoic membrane (CAM) assay for angiogenesis in quail embryos treated with CM from control and PDT-MSCs. g Morphometric analysis of MSC-CM mediated induction of CAM vascularization. To evaluate vascularization rate, number of tubule complex and junction numbers was calculated using AngioQuant software. Data represent median ± range. *P < .05, **P < .01, ***P < .001, ***P < .0001.

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Fig. 5. Low-dose PDT modifies MSC-PBMC interactions. a Low-dose PDT treatment decreases MCP-1 (CCL2) secretion by MSCs. b Contact interaction of MSCs and PBMCs stimulates MCP-1 production. PDT of MSCs significantly reduces MCP-1 secretion in MSC-PBMC co-cultures. MSCs were treated with low-dose PDT before cocultivation with PBMCs. MCP-1 level was analyzed by ELISA (a,b) in CM of MSCs before and 24 h after PDT (N = 6). c Representative phase-contrast images showing induction of PDT-MSC, but not control MSC cell death upon interaction with PHA-activated allogenic PBMCs. d Mitochondrial transmembrane potential of MSCs cocultured with PBMCs was evaluated by flow cytometry after JC-1 staining (N = 3). e Co-cultivation with allogenic naïve/activated PBMCs significantly reduce viability of PDT-MSCs, but not control ones. f,g Co-cultivation with control or PDT-treated MSCs doesn't affect expression of activation markers CD25 (f) and CD69 (g) in allogenic lymphocytes. h Flow cytometric analysis of PBMCs subpopulation values after co-cultivation with control and PDT-MSCs. Data represent median ± range. *P < .05, **P < .01, ***P < .001.

Fig. 6. Effects of low-dose PDT on MSCs functional activity are mediated by Erk1/2 and GSK-3β phosphorylation. a Phosphorylation of different MAPK-related proteins in MSC lysates was profiled using Human Phospho-MAPK Antibody Array. b Densitometric analysis of selected analytes in antibody arrays were quantified using ImageLab software (N = 4). c,d Erk1/2 phosphorylation was analyzed by WB after 0.25 J/cm2 PDT at indicated time points. e,f Low dose PDT induces GSK-3β phosphorylation in MSCs. Data of WB in cell lysates. d,f Semi-quantative densitometric analysis of ERk1/2 and GSK-3β phosphorylation was performed using ImageLab software (N = 3). g Time course of HIF-1α expression in MSCs after low-dose PDT. Data of qPCR. Data represent median ± range. *P < .05, **P < .01, ***P < .001, ***P < .0001.

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Fig. 7. The proposed hypothetical model of low-dose PDT effects on MSC functional activity. Low-dose PDT results in generation of “regulatory” intracellular ROS followed by phosphorylation of Erk1/2 and GSK-3β and induction of HIF-1α expression. It inhibits MSC motility and promotes angiogenesis and extracellular matrix degradation.

6 h and returned back to the base line by 24 h (Fig. 6g, h). We conclude that activation of Erk1/2 and inactivation of GSK-3α/β signaling are involved in PDT-induced changes of MSC functional activity, and it can represent a possible mechanism underlying effects of low-dose PDT on stromal cells.

PDT, effects of low-dose PDT on MSCs are mediated by rapid phosphorylation of Erk1/2, JNK, GSK-3β and Akt2. One of Erk1/2 and Akt2 downstream targets is a small heat-shock protein 27 (HSP27) which has been associated with protection from programmed cell death and development of adaptive response to oxidative stress [38,39]. Our data reveal dramatic increase of HSP27 phosphorylation after low-dose PDT. Thus, we hypothesize that in our model PDT-mediated Erk1/2, Akt2 and JNK activation followed by phosphorylation of HSP27 could represent an important adaptive and pro-survival mechanism activated by MSC in response to ROS induction by low-dose PDT. Phosphorylation of GSK-3 by Akt has been shown to be the major mechanism of Aktmediated inhibition of GSK-3β activity [40,41]. Here, we find that PDT induced rapid and sustained GSK-3β phosphorylation at Ser9. Concomitant increase of Akt2 phosphorylation and its rapid kinetics makes it likely that accumulation of p-GSK-3β is the result of elevated Akt2 activity. It has been demonstrated that acute oxidative stress can activate GSK-3 followed by induction of cell death [41]. However, the role of GSK-3 in redox signaling remains mostly unknown. To our knowledge, this is the first report indicating inhibition of GSK-3 activity by intracellular ROS which can play an important role in regulation of cell survival and functional activity. Apart from survival pathways, Erk1/2 and GSK-3 have been shown to regulate HIF-1α signaling: Erk1/2 specifically upregulates the transactivation activity of HIF-1α while GSK-3-mediated phosphorylation results in HIF-1α destabilization in oxygen-independent manner [42,43]. These kinases also mediate ROStriggered HIF-1α stabilization and transcriptional activation in cancer cells, and it has been proposed as one of the mechanisms underlying tumor cell resistance to PDT [37,44]. Here, we show that low-dose PDT induces HIF-1α gene expression in MSCs. Erk1/2, GSK-3 and HIF-1α signaling has been implicated in regulation of metabolism, differentiation, migration, proliferation and survival. Thus, we propose that modulation of Erk1/2, GSK-3 and HIF-1α activity represents the key mechanism involved in low-dose PDT-mediated regulation of MSC functional activity. Both oxidative stress and redox signaling have been implicated in tumorigenesis: ROS have been shown to affect cancer cell phenotype driving proliferation, mutagenesis and chemoresistance [45]. The effects of ROS on MSC functional activity in the context of tumor growth are largely unknown: for a long time MSC biology was related exclusively to regenerative medicine. Now it is clear that MSCs represent a double-edged sword due to their role in regeneration and pathogenesis of different diseases (cancer, fibrosis, atherosclerosis, etc) [14,46]. However, most studies dedicated to redox signaling in MSCs are still focused on oxidative stress-mediated inhibition of MSC proliferation, differentiation and aging [47,48]. Here we demonstrate that ROS

4. Discussion PDT was originally developed as an ablation therapy utilizing high light doses to destroy tumors and eliminate cancer cells. Efficacy of PDT-mediated tumor eradication is determined by the direct induction of cell death and indirect effects mediated through tumor stroma represent by endothelial, stromal and immune cells [22]. While PDT effects on immune cells have been characterized [26,27], and tumor vasculature is a common target of PDT [22,35], effects of PDT on stromal cells remain poorly understood. Stromal cells are found throughout the tumor and thus are treated simultaneously with tumor during PDT. Light delivery during PDT is affected by numerous factors including tumor size and tissue optics. Thus, not all areas of the tumor receive equal light doses: some tumor areas will receive lower light doses due to limited depth of light penetration [25]. The present study was focused on the effects of low-dose PDT on MSCs, which represent key component of stroma and play a pivotal role in tumorigenesis. Here, we report for the first time that low-dose PDT dramatically modifies MSC phenotype (Fig. 7). These PDT-induced changes in MSC functional activity can be implicated into protection of cancer cells from PDT and can result into reduction of PDT efficacy. However, recent studies have been shown that low-dose PDT enhances anti-tumor immunity [27,28]. Importantly, low-dose PDT appears to increase immunogenicity of MSCs, potentially stimulating immune cells. PDT effects rely on generation of ROS as a result of photochemical reaction [25]. Our data show that modulation of light regimen during PDT allows regulated dose-dependent induction of intracellular ROS in MSCs. High dose PDT is known to induce oxidative damage and cell death via apoptosis, necrosis or necroptosis [22]. The regimen of low dose PDT chosen for the study doesn't affect cell viability. However, sublethal intracellular ROS induced by low-dose PDT can modulate cell functional activity as ROS in low concentrations represent an important mediator of intracellular signaling [30]. In tumor cells high-dose PDT has been shown to activate JNK- and p38-signaling which can be implicated in promotion of both survival and cell death pathways depending on cancer type, photosensitizer and level of oxidative damage [22,36]. Recently it has been shown that PDT-induced intracellular ROS activate pro-survival Erk1/2 signaling in SW480 colon carcinoma cells [37]. In line with that study, here we demonstrate that unlike high-dose 11

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mediate low-dose PDT-induced inhibition of MSC motility, promotion of proangiogenic potential and alteration of MSC immunomodulatory properties. One of the important factors defining cell functional activity is cytoskeleton organization. It coordinates migration, regulates cell proliferation and differentiation, and mediates intracellular signaling and micro-environmental sensing [49]. Disorganization of microfilaments (assembly of actin microspikes and disassembly of stress fibers) has been previously found in ALA-PDT resistant sublines of mammary adenocarcinoma LM3 [50]. These changes have been implicated in tumor cell resistance to PDT. Our data show that low-dose PDT induces similar reorganization of actin and vimentin filaments in MSCs. There are several mechanisms implemented in the regulation of cytoskeleton dynamic s by PDT: direct (oxidation of actin and actin-binding proteins) and indirect (activation of Erk1/2, Hsp27, etc.). ROS can alter structure and polymerization of microfilaments direct oxidation of actin and actin-binding proteins [51]. Erk1/2 is known to be responsible for the disruption of stress fibers via inhibition of Rho-ROCK-LIM kinase pathway [52]. HSP27 interacts with both actin and vimentin, modulating actin cytoskeleton dynamics [53] and promoting formation of vimentin aggregates [54]. Our data reveal that reorganization of MSC cytoskeleton by low-dose PDT is accompanied by increased MSC stiffness. This observation is in line with other studies implicating intermediate filaments in cell stiffness [55]. Recent studies have highlighted the role of mechanical properties of tumor microenvironment in tumorigenesis [56]. It has been shown that increase of ECM stiffness promotes cell proliferation and invasiveness (“durotaxis”), and modifies drug response through modulation of intracellular signaling [56,57]. Thus, our data suggest that low-dose PDT may promote tumor invasion through modulation of ECM-remodeling and stiffness of tumor microenvironment. Cytoskeletal dynamics regulates cell migration. Our findings show that PDT-induced cytoskeleton reorganization impairs MSC migration and chemotaxis. MSC motility plays an important role in their functional activity as well as it defines MSC ability to be recruited to the “target” sites of injury and inflammation where they can exert their functions [58,59]. Loss of MSC chemotaxis following PDT can result with high probability in tight MSC embedding into the tumor bed. Tumor vasculature represents a key component of tumor microenvironment and plays an important role in tumor progression, metastasis and recurrence. Formation of tumor blood vessels involves (1) pathological angiogenesis co-opting pre-existing local vessels [60] and (2) neovascularization recruiting bone marrow-derived and circulating endothelial precursors [61]. The regulation of tumor angiogenesis depends on the balance of pro- and anti-angiogenic factors. High-dose PDT usually induces immediate vascular shutdown followed by activation of IL-6 which has been proposed to potentiate tumor resistance to PDT [22,26,62]. Here we demonstrate that low-dose PDT can improve MSC angiogenic potential stimulating secretion of MMPs (MMP8, -9), proangiogenic (VEGF-A, IL-8) and proinflammatory (IL-6, -8) cytokines. At the same time, primary MMP function is a remodeling of ECM, which can be further implicated in the release of ECM-bound growth factors, promotion of angiogenesis and regulation of immune and cancer cell migration [63]. Induction of the factors described above by low-dose PDT correlates with modulation of Erk1/2 and GSK-3β signaling: GSK-3β as well as IGFBP-3 (insulin-like growth factor binding protein 3) has been implicated in the inhibition of angiogenesis [64] while Erk1/2 has the opposite effect stimulating MMP, VEGF-A, IL-6 and IL-8 production [65]. Thus, we hypothesize that promotion of MSC proangiogenic potential by low-dose PDT taken together with our observation on decreased MSC motility, may impair PDT outcome, especially in stroma-enriched tumors. The impact and detailed mechanisms of modulation of MSC immunoregulatory properties by PDT/oxidative stress are yet to be investigated. Here we uncover that low-dose PDT significantly increases MSC immunogenicity resulting in rapid MSC lysis by PBMCs. We and

others have previously demonstrated that low-dose PDT can induce specific anti-tumor immune response [66,67]. While precise mechanisms remain understudied, several studies have persuasively demonstrated that PDT-mediated stimulation of anti-tumor immunity depends on natural killer (NK) and CD8+ cells and involves release of cytokines and damage-associated molecular patterns (DAMPs) activating innate and adaptive immune response [68,69]. Here, we haven't found any difference in T-cell activation between PMBCs co-cultivated with naïve and low-dose PDT-treated MSCs. Previously it has been demonstrated that PDT treatment of cancer cells significantly increase macrophagemediated killing of these cells [70]. Moreover, activation of macrophages by adjuvant immunotherapy significantly improves anti-tumor efficacy of PDT [71]. Taken together, our results suggest that induction of PBMCs cytotoxicity against low-dose PDT-treated MSCs is related to the activation of innate, but not adaptive immune response. In present study we show that low-dose PDT doesn't affect expression of MHC Class I-related molecules in MSCs, and expression of stimulating and activating receptors in CD56/CD16+ cells remain unchanged after direct co-cultivation with PDT-treated or non-treated MSCs. Thus, we hypothesize that the NKG2D-DNAM-1 axis is not involved in the regulation of PDT-induced MSC immunogenicity. Cytokine release represents one of the mechanisms mediating immune effects of PDT [28,67]. Here, we demonstrate that low-dose PDT negatively regulates MCP-1 production and stimulates IL-6 secretion in MSCs. IL-6 is known to be frequently induced by PDT. Despite its pro-inflammatory activity, IL-6 has been shown to promote survival of tumor cells and negatively regulate antitumor immune memory [62]. MCP-1 is a pleotropic inflammatory cytokine that targets monocytes, NK cells, activated and memory T-lymphocytes, promotes angiogenesis and mediates both tumor promotion and immunosurveillance [72,73]. In addition, MCP-1 blockade has been associated with the induction of PD-L1 expression by activated CD4+ T-cells [74]. Active GSK-3 is required for MCP-1 expression [75], and oxidative stress is usually associated with MCP-1 induction [76]. Here we hypothesize that inhibition of MCP-1 by lowdose PDT is GSK-3-dependent. MCP-1 downregulation by low-dose PDT can potentially limit tumor infiltration with pro-tumorigenic macrophages and/or impair resolution of PDT-induced inflammation creating tumor-supportive microenvironment. Thus, PDT effects on MSC immunoregulatory properties are contradictory, and further studies are required to evaluate the impact of MSCs into PDT-mediated immune response. In summary, we have identified some of the possible mechanisms underlying stromal cell contribution to PDT efficacy and highlighted several pathways which could be implicated in PDT-mediated regulation of MSC functional activity. Our study demonstrates that role of stromal cells shouldn't be underestimated during development and optimization of PDT. As PDT efficacy relies on intracellular ROS induction, we anticipate that our findings can be useful for those who study ROS-related diseases with known MSC implication. Funding This study was supported by Russian Foundation for Basic Research (grant RFBR 16-04-01377 “A”) and Russian Academic Excellence Project 5-100. Declaration of Competing Interest None. Acknowledgements Dr. Sergey Buravkov assisted in confocal imaging. 12

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Appendix A. Supplementary data

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