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Int. J. Mol. Sci. 2013, 14(7), 13542-13558; doi:10.3390/ijms140713542

Review
Regulation of miRNA Expression by Low-Level Laser Therapy (LLLT) and Photodynamic Therapy (PDT)
Toshihiro Kushibiki *, Takeshi Hirasawa , Shinpei Okawa and Miya Ishihara
Department of Medical Engineering, National Defense Medical College 3-2 Namiki, Tokorozawa, Saitama 359-8513, Japan
*
Author to whom correspondence should be addressed; E-Mail: toshi@ndmc.ac.jp; Tel.: +81-4-2995-1211 (ext. 2241); Fax: +81-4-2996-5199.
Received: 31 May 2013; in revised form: 19 June 2013 / Accepted: 20 June 2013 /
Published: 27 June 2013

Abstract

: Applications of laser therapy, including low-level laser therapy (LLLT), phototherapy and photodynamic therapy (PDT), have been proven to be beneficial and relatively less invasive therapeutic modalities for numerous diseases and disease conditions. Using specific types of laser irradiation, specific cellular activities can be induced. Because multiple cellular signaling cascades are simultaneously activated in cells exposed to lasers, understanding the molecular responses within cells will aid in the development of laser therapies. In order to understand in detail the molecular mechanisms of LLLT and PDT-related responses, it will be useful to characterize the specific expression of miRNAs and proteins. Such analyses will provide an important source for new applications of laser therapy, as well as for the development of individualized treatments. Although several miRNAs should be up- or down-regulated upon stimulation by LLLT, phototherapy and PDT, very few published studies address the effect of laser therapy on miRNA expression. In this review, we focus on LLLT, phototherapy and PDT as representative laser therapies and discuss the effects of these therapies on miRNA expression.
Keywords:
low-level laser therapy (LLLT); phototherapy; photodynamic therapy (PDT); miRNA

1. Low-Level Laser Therapy (LLLT) and Its Effects on miRNA Expression

A laser (light amplification by stimulated emission of radiation) is a device that generates electromagnetic radiation that is relatively uniform in wavelength, phase and polarization. This technology was originally described by Maiman in 1960 in the form of a ruby laser [1]. The properties of lasers have allowed for numerous medical applications, including their use in surgery, activation of photodynamic agents and various ablative therapies in cosmetics, all of which are based on heat generated by the laser beam, in some cases, leading to tissue destruction [29]. These applications of lasers are considered “high-energy”, because of their intensities, which range from about 1–100 watt (W)/cm2.

This paper will address another type of laser application, low-level laser therapy (LLLT), which elicits its effects through non-thermal means. This field was initiated by the work of Mester et al., who in 1967 reported non-thermal effects of lasers on mouse hair growth [10]. In a subsequent study, the same group reported acceleration of wound healing and improvement in the post-wounding regeneration ability of muscle fibers using a 1 J/cm2 ruby laser [11]. Since those early days, numerous in vitro and in vivo studies of LLLT in the context of regenerative medicine have demonstrated a wide variety of therapeutic effects, including reduction of pain, anti-inflammatory effects and wound healing. According to da Silva et al. [12], the types of laser most frequently used for wound healing and tissue repair are helium neon (He-Ne) lasers and diode lasers, including gallium-aluminum-arsenic (Ga-Al-As), arsenic-gallium (As-Ga) and indium-gallium-aluminum-phosphide (In-Ga-Al-P) lasers.

One of the most distinctive features of LLLT relative to other modalities is that the effects are mediated not through induction of thermal effects, but rather, through a process, still not clearly defined, called “photobiostimulation”. Because this effect of LLLT apparently does not depend on coherence, it is therefore possible to achieve photobiostimulation using non-laser light-generating devices, such as inexpensive light-emitting diode (LED) technology [1317]. To date, several mechanisms of biological action have been proposed, although none have been clearly established. These include augmentation of cellular ATP levels [1820], manipulation of inducible nitric oxide synthase (iNOS) activity [2125], suppression of inflammatory cytokines, such as TNF-alpha [19,2628], IL-1beta [2830], IL-6 [28,3134] and IL-8 [28,31,32,35], upregulation of growth factors, such as PDGF, IGF-1, NGF and FGF-2 [30,3638], alteration of mitochondrial membrane potential [3942], due to chromophores found in the mitochondrial respiratory chain [4345], stimulation of protein kinase C (PKC) activation [46], manipulation of NF-kappaB activation [47], induction of reactive oxygen species (ROS) [48,49], modification of extracellular matrix components [50], inhibition of apoptosis [39], stimulation of mast cell degranulation [51] and upregulation of heat shock proteins [52]. We have also proposed that LLLT influences cell differentiation following laser stimulation [5355].

Unfortunately, these effects have been demonstrated using a variety of laser devices in non-comparable models. To add to the confusion, dose-dependency seems to be confined to a very narrow range, and in numerous systems, the therapeutic effects disappear with increased dose. Consequently, only two studies of miRNA expression dynamics following LLLT have been reported to date, by Wang et al. [56] and Gu et al. [57]. With the exception of those studies, no data are currently available regarding the overall changes in the global expression of many hundreds of miRNAs following LLLT. Wang et al. [56] showed that LLLT increases the migration, proliferation and viability of rat mesenchymal stem cells (MSCs) and, also, activates the expression of various miRNAs. Using a diode laser (wavelength: 635 nm, 0.5 J/cm2), they found that the proliferation rate and expression of cell cycle-associated genes increased in a time-dependent manner following LLLT treatment of MSCs. Microarray assays revealed subsets of miRNAs that were regulated by LLLT: 19 miRNAs were upregulated and 15 miRNAs were downregulated (Table 1); these dynamic changes were confirmed by quantitative real-time PCR.

The most highly upregulated miRNA was miR-193. Gain- and loss-of-function experiments demonstrated that miR-193 levels regulate the proliferation of MSCs of both humans and rats; in particular, blockade of miR-193 repressed the MSCs proliferation induced by LLLT. However, this miRNA apparently does not affect apoptosis or differentiation. In addition, Wang et al. found that miR-193 regulated expression of cyclin-dependent kinase 2 (CDK2). Bioinformatic analyses and luciferase reporter assays revealed that inhibitor of growth family, member 5 (ING5), was the most likely target of miR-193 to functionally regulate proliferation and CDK2 expression; indeed, the mRNA and protein levels of ING5 are regulated by miR-193. Furthermore, inhibition of ING5 by small interfering RNA (siRNA) upregulated both MSC proliferation and the expression of CDK2. Another miRNA, miR-335, has been shown by others to regulate the proliferation and migration of MSCs [58], so it is likely to play an important role in MSC proliferation after LLLT. Moreover, several studies have shown that LLLT also stimulates cell differentiation [5355,5975], and future work should reveal miRNAs specifically involved in mediating this effect.

Although some literature reported that tumor or apoptosis related miRNAs were induced by UV irradiation to cells [7681], Gu et al. reported UV-phototherapy and its effect on miRNA expression [57]. They showed the effect of narrow-band ultraviolet B (NB-UVB) irradiation on miR-21 and -125b expression in psoriatic epidermis. Psoriasis is an inflammatory skin disease in which dysregulation of p63, a member of the p53 family that is crucial for skin development and maintenance, has been demonstrated [8284]. Involvement of miR-203, miR-21 and miR-125b were implicated in the regulation of p63 or p53 in the pathogenesis of psoriasis. Skin biopsies from 12 psoriasis patients were collected before, during and after NB-UVB therapy. The p63 expression was not significantly affected, whereas NB-UVB phototherapy significantly decreased expression of miR-21and increased miR-125b levels. Since NB-UVB phototherapy is commonly used in the treatment of psoriasis [8587], those results indicate a complex mechanism of p63 regulation, which merits further investigation in order to achieve better long-term clinical improvement.

2. Photodynamic Therapy and Its Effects on miRNA Expression

Photodynamic therapy (PDT), a class of laser therapy, is a photochemical modality approved for the treatment of various cancers and diseases in which neovascularization occurs [88,89]. The PDT process consists of injecting a photosensitizer, which selectively accumulates at the lesion site, followed by local irradiation of the tumor with light of an appropriate wavelength to activate a specific drug [90]. Irradiation leads to the generation of singlet oxygen and other reactive oxygen species (ROS) [91]. PDT is being considered not only as palliative therapy, but also as a treatment option for early-stage skin, lung, cervical and esophageal cancers, as well as basal-cell carcinomas. Currently, PDT has been approved for localized diseases and precancerous lesions, such as bladder cancers, pituitary tumors and glioblastomas [92,93]. Furthermore, numerous ongoing clinical studies have been designed to optimize the conditions for PDT; subsequently, PDT has been approved in several countries.

Upon absorption of one or more photons, the excited photosensitizer undergoes one of two possible reactions (type I or/and II) with a neighboring oxygen molecule, yielding ROS [94]. These ROS oxidize various cellular substrates, affecting cellular functions and resulting in cell death. The ROS that are produced during PDT destroy tumors by multiple mechanisms: in contrast to most conventional cytotoxic agents, which usually only trigger apoptotic cell death, PDT can cause cell death by necrosis and/or apoptosis.

The direct destruction of cancer cells (necrosis) by PDT is caused by irreversible damage to the plasma membrane and intracellular organelles, including the mitochondria, lysosomes, Golgi apparatus and endoplasmic reticulum (ER). The mechanisms of PDT-induced apoptosis have been described by many studies. Apoptosis, or programmed cell death, is one mechanism that mediates toxicity in the target tissue following PDT [95]. Apoptosis involves a cascade of molecular events leading to orderly cellular death without an inflammatory response [9698]. The initiation of apoptosis involves a complex network of signaling pathways, both intrinsic and extrinsic to the individual cell, which are regulated, in part, by pro- and anti-apoptotic factors [96]. The initial damage can involve different molecules, ultimately leading to activation of specific death pathways. Mitochondria-localized photosensitizers can cause immediate and light-dependent photodamage to mitochondrial components, such as the anti-apoptotic Bcl-2, Bcl-xL and the other apoptosis-related proteins, prompting the release of caspase-activating molecules [99]. Photosensitizers that accumulate in the lysosomes or mitochondria and which were excited by laser light can induce Bax-mediated caspase activation (Figure 1).

Another important cellular factor induced by PDT and released from necrotic tumor cells is heat-shock protein 70 (Hsp70) [100]. Hsp70 is significantly induced after stress; when it remains within the cell, it chaperones unfolded proteins and prevents cell death by inhibiting the aggregation of cellular proteins. Hsp70 directly binds to the caspase-recruitment domain of apoptotic-protease activating factor 1 (Apaf-1), thereby preventing the recruitment of Apaf-1 oligomerization and association of Apaf-1 with procaspase 9. These properties not only enable intracellular Hsp70 to inhibit cancer-cell death by apoptosis, but also promote the formation of stable complexes with cytoplasmic tumor antigens. These antigens can then either be expressed at the cell surface or escape intact from dying necrotic cells to interact with antigen-presenting cells, thereby stimulating an anti-tumor immune response.

The mechanisms of cell death following PDT have been thoroughly summarized in the literature [95,101104]. A better understanding of the molecular differences between apoptosis and necrosis and identification of the crosstalk between these programs will certainly be crucial to the development of new PDT modalities aimed at increasing the efficiency of cancer-cell killing.

Another inherent consequence of PDT is local hypoxia, which can arise either directly, from oxygen consumption during treatment [105107], or indirectly, from the destruction of tumor vasculature as a result of effective treatment [108,109]. Hypoxia is a major stimulus for angiogenesis, via its stabilization of the hypoxia-inducible factor-1α (HIF-1α) transcription factor [110,111]. HIF-1 is a heterodimeric complex of two helix-loop-helix proteins, HIF-1α and HIF-1β (ARNT). ARNT is constitutively expressed, whereas HIF-1α is rapidly degraded under normoxic conditions. Hypoxia induces the stabilization of the HIF-1α subunit, which, in turn, allows formation of the transcriptionally active protein complex. A number of HIF-1–responsive genes have been identified, including those encoding vascular endothelial growth factor (VEGF), erythropoietin and glucose transporter-1 [112,113]. Following PDT, increases in VEGF secretion and angiogenic responses stimulated via HIF-1 pathways have been documented in vivo [114117]. VEGF induction can contribute to tumor survival and regrowth and, therefore, may represent one of the factors that prevent PDT from achieving its full tumoricidal potential. PDT has been considered for both palliative therapy and as an early treatment option for cancer. Numerous ongoing clinical studies have been designed to optimize PDT conditions. However, no standardized biological markers of cell death and PDT efficacy, other than cell viability itself, have been reported.

Human cancer is associated with changes in miRNA expression. The pattern of miRNA expression varies dramatically across tumor types, and miRNA profiles reflect the developmental lineage and differentiation state of a tumor [118]. miRNA is also likely to play critical roles in various aspects of hematopoiesis, including the differentiation of hematopoietic stem/progenitor cells, as well as in events that lead to hematological disorders. Nonetheless, very few miRNA expression patterns of specific diseases are available. Moreover, no profiles of miRNA expression after PDT have been reported. Cheng et al. found that inhibition of miR-95, -124, -125, -133, -134, -144, -150, -152, -187, -190, -191, -192, -193, -204, -211, -218, -220, -296 and -299 resulted in a decrease in cell growth, whereas inhibition of miR-21 and miR-24 profoundly increased cell growth in HeLa cells [119]. In addition, they identified miRNAs, whose expression increased levels of apoptosis (miR-7, -148, -204, -210, -216 and -296). Those data suggest that specific miRNAs are involved in the cell-death response. We have shown that a miRNA specific to apoptosis is expressed at increased levels in HeLa cells in response to PDT using talaporfin sodium as a photosensitizer [120]. Our study was the first to characterize miRNA expression levels following PDT. In our experiments, miR-210 and miR-296 expression levels increased significantly 1 h after PDT in cells treated with 50 μg/mL talaporfin sodium, relative to the control group (i.e., 0 μg/mL talaporfin sodium), as shown in Figure 2. However, the expression levels of other miRNAs, e.g., miR-7, -148a, -204 and -216, were indistinguishable from those of the control group after PDT.

miR-210 is the miRNA most consistently stimulated under hypoxic conditions [121]. Because hypoxia and stabilization of intracellular HIF are inherent consequences of PDT [92], Giannakakis et al. investigated miR-210 expression in the context of its hypoxic effect, and they reported evidence for the involvement of the HIF signaling pathway in miR-210 regulation. To study the biological impacts of a partial or complete loss of miR-210 functions, they also identified the putative mRNA targets of miR-210. According to their report, miR-210 targets important regulators of transcription, cell metabolism, differentiation and development, i.e., processes that are critically affected by hypoxia [121]. The identification of key regulators of important cellular processes among miR-210 target mRNAs, as well as the high frequency of gene copy-number aberrations in tumors, underscore the involvement of miR-210 in oncogenesis and highlight miR-210 as a potential link between hypoxia and cell-cycle control in cancer cells.

Würdinger et al. reported a role for miR-296 in promoting angiogenesis in tumors [122], and in particular, they showed that VEGF alone is capable of increasing miR-296 expression levels. Their results revealed a feedback loop, wherein VEGF induces miR-296 expression, which targets the hepatocyte growth factor-regulated tyrosine kinase substrate (HGS), which, in turn, results in increased levels of VEGF receptor 2 and platelet-derived growth factor (PDGF) receptor β protein and, ultimately, in an increased response to VEGF. Because increased VEGF sensitivity of cancer cells is one of the inherent consequences of PDT [115], our results suggest that inhibition of miR-296 expression should improve PDT efficacy [120]. Our study also suggested that hypoxia induced by PDT induces miR-210 expression, followed by an increased expression of both VEGF and miR-296 [120]. Hence, we reported that miR-210 and miR-296 expression levels represent markers for the efficacy of talaporfin sodium-mediated PDT in cancer cells.

Furthermore, a recently published paper by Bach et al. described a comprehensive analysis of changes in miRNA levels following PDT, using polyvinylpyrrolidone hypericin (PVPH) as a photosensitizer, against A431 human epidermoid carcinoma cells [123]. That study was the first comprehensive analysis of changes in miRNA induced by PDT. Using microarray analysis, Bach et al. identified eight miRNAs that were significantly differentially expressed 5 hr after treatment, compared with baseline levels, and three miRNAs with more than two-fold differential expression that could be detected in one or two biological replicates. The verification of these results by quantitative real-time PCR, including a detailed time course, revealed an up to 15-fold transient upregulation of miR-634, -1246 and -1290 relative to their basal levels (Table 2).

In silico prediction of the targets of these miRNAs yielded numerous mRNAs encoding proteins, including the apoptotic protease activating factor-1 interacting protein and the BMI1 polycomb ring finger oncogene in the apoptosis/cell death category, cyclin-dependent kinase 20 and the cell division cycle 25 homolog C in the proliferation/cell cycle category, frizzled family receptor 3 and bone morphogenetic protein 4 in the cell signaling/adhesion category and the DNA excision repair protein ERCC-8 and peroxiredoxin-6 in the cell stress category. Although several studies have investigated the PDT-induced changes in the transcriptome and proteome, no comprehensive data are currently available regarding the effect of PDT on the miRNA transcriptome. Using a comprehensive microarray platform covering 1223 mature human miRNAs, Bach et al. did not observe up- or down-regulation by PDT of the miRNAs reported in our study (miR-210 and -296 [120]). This difference is likely attributable to the PDT conditions, such as cell type, photosensitizer and laser dose. Furthermore, the significant increase in the apoptosis-related miRNAs (3–4-fold increase) observed in our study was measured in a mixed population of cells, consisting predominantly of surviving cells [124]. Given these discrepancies, there is a need for additional experiments that might uncover additional miRNAs that are transiently regulated following photodynamic damage. It will be also of paramount interest to study miRNA-related cellular responses under explicitly non-lethal PDT conditions, as this approach could identify possible miRNA targets, whose manipulation might increase cells’ sensitivity towards PDT.

Interestingly, Bach et al. also found that the incubation with the photosensitizer induced a slight to moderate increase in the expression of several miRNAs (i.e., miR-1260b, -1260, -1280, -3182, -1290 and -1246), particularly at later time points [123]. Conversely, several miRNAs were transiently up-regulated by light-only treatment, especially at earlier time points (miR-1260b, -1260, -1280, -3182 and -1290). They concluded that the detailed functions of the increased expression of these miRNAs following apoptosis induced by PDT remain to be elucidated [123].

3. Conclusions

In this review, we focused on miRNA expression after LLLT and PDT. As mentioned above, only a few papers have been published regarding miRNA expression in this context, and those few reports discuss only a small number of laser therapy conditions. The ability of LLLT to induce growth-factor production, inhibition of inflammation, stimulation of angiogenesis, pain reduction and direct effects on stem cells suggests that there is an urgent need to combine this modality with regenerative medicine. PDT has been employed in the treatment of many tumor types, and its effectiveness as a curative and palliative treatment is well documented, especially in the context of skin cancer. A detailed understanding of LLLT-, phototherapy- and PDT-related molecular mechanisms, including the specific effects on miRNA and protein expression, will provide an important source for new applications of laser therapy and for the development of individualized treatments.

Acknowledgments

This work was supported by Japan Society for the Promotion of Science (JSPS) KAKENHI, Grant Number 25713009.

Conflict of Interest

The authors declare no conflict of interest.

References

  1. Maiman, T.H. Stimulated optical radiation in ruby. Nature 1960, 187, 493–494. [Google Scholar]
  2. Khatri, K.A.; Mahoney, D.L.; McCartney, M.J. Laser scar revision: A review. J. Cosmet. Laser Ther 2011, 13, 54–62. [Google Scholar]
  3. Chung, S.H.; Mazur, E. Surgical applications of femtosecond lasers. J. Biophotonics 2009, 2, 557–572. [Google Scholar]
  4. Zhao, Z.; Wu, F. Minimally-invasive thermal ablation of early-stage breast cancer: A systemic review. Eur. J. Surg. Oncol 2010, 36, 1149–1155. [Google Scholar]
  5. Siribumrungwong, B.; Noorit, P.; Wilasrusmee, C.; Attia, J.; Thakkinstian, A. A systematic review and meta-analysis of randomised controlled trials comparing endovenous ablation and surgical intervention in patients with varicose vein. Eur. J. Vasc. Endovasc. Surg 2012, 44, 214–223. [Google Scholar]
  6. Vuylsteke, M.E.; Mordon, S.R. Endovenous laser ablation: A review of mechanisms of action. Ann. Vasc. Surg 2012, 26, 424–433. [Google Scholar]
  7. Vogel, A.; Venugopalan, V. Mechanisms of pulsed laser ablation of biological tissues. Chem. Rev 2003, 103, 577–644. [Google Scholar]
  8. Casas, A.; di Venosa, G.; Hasan, T.; Al, B. Mechanisms of resistance to photodynamic therapy. Curr. Med. Chem 2011, 18, 2486–2515. [Google Scholar]
  9. Anand, S.; Ortel, B.J.; Pereira, S.P.; Hasan, T.; Maytin, E.V. Biomodulatory approaches to photodynamic therapy for solid tumors. Cancer Lett 2012, 326, 8–16. [Google Scholar]
  10. Mester, E.; Szende, B.; Gartner, P. The effect of laser beams on the growth of hair in mice. Radiobiol. Radiother 1968, 9, 621–626. [Google Scholar]
  11. Mester, E.; Spiry, T.; Szende, B.; Tota, J.G. Effect of laser rays on wound healing. Am. J. Surg 1971, 122, 532–535. [Google Scholar]
  12. Da Silva, J.P.; da Silva, M.A.; Almeida, A.P.; Lombardi, I., Junior; Matos, A.P. Laser therapy in the tissue repair process: A literature review. Photomed. Laser Surg 2010, 28, 17–21. [Google Scholar]
  13. Buravlev, E.A.; Zhidkova, T.V.; Vladimirov, Y.A.; Osipov, A.N. Effects of laser and led radiation on mitochondrial respiration in experimental endotoxic shock. Lasers Med. Sci 2012, 28, 785–790. [Google Scholar]
  14. De Sousa, A.P.; Santos, J.N.; dos Reis, J.A., Jr; Ramos, T.A.; de Souza, J.; Cangussu, M.C.; Pinheiro, A.L. Effect of led phototherapy of three distinct wavelengths on fibroblasts on wound healing: A histological study in a rodent model. Photomed. Laser Surg. 2010, 28, 547–552. [Google Scholar]
  15. Lev-Tov, H.; Brody, N.; Siegel, D.; Jagdeo, J. Inhibition of fibroblast proliferation in vitro using low-level infrared light-emitting diodes. Dermatol. Surg 2012, 39, 422–425. [Google Scholar]
  16. Nishioka, M.A.; Pinfildi, C.E.; Sheliga, T.R.; Arias, V.E.; Gomes, H.C.; Ferreira, L.M. Led (660 nm) and laser (670 nm) use on skin flap viability: Angiogenesis and mast cells on transition line. Lasers Med. Sci 2012, 27, 1045–1050. [Google Scholar]
  17. Pinheiro, A.L.; Soares, L.G.; Cangussu, M.C.; Santos, N.R.; Barbosa, A.F.; Silveira, L., Junior. Effects of led phototherapy on bone defects grafted with mta, bone morphogenetic proteins and guided bone regeneration: A raman spectroscopic study. Lasers Med. Sci 2012, 27, 903–916. [Google Scholar]
  18. AlGhamdi, K.M.; Kumar, A.; Moussa, N.A. Low-level laser therapy: A useful technique for enhancing the proliferation of various cultured cells. Lasers Med. Sci 2012, 27, 237–249. [Google Scholar]
  19. Gao, X.; Xing, D. Molecular mechanisms of cell proliferation induced by low power laser irradiation. J. Biomed. Sci 2009, 16, 4. [Google Scholar]
  20. Tafur, J.; van Wijk, E.P.; van Wijk, R.; Mills, P.J. Biophoton detection and low-intensity light therapy: A potential clinical partnership. Photomed. Laser Surg 2010, 28, 23–30. [Google Scholar]
  21. Gavish, L.; Perez, L.S.; Reissman, P.; Gertz, S.D. Irradiation with 780 nm diode laser attenuates inflammatory cytokines but upregulates nitric oxide in lipopolysaccharide-stimulated macrophages: Implications for the prevention of aneurysm progression. Lasers Surg. Med 2008, 40, 371–378. [Google Scholar]
  22. Lindgard, A.; Hulten, L.M.; Svensson, L.; Soussi, B. Irradiation at 634 nm releases nitric oxide from human monocytes. Lasers Med. Sci 2007, 22, 30–36. [Google Scholar]
  23. Moriyama, Y.; Moriyama, E.H.; Blackmore, K.; Akens, M.K.; Lilge, L. In vivo study of the inflammatory modulating effects of low-level laser therapy on inos expression using bioluminescence imaging. Photochem. Photobiol 2005, 81, 1351–1355. [Google Scholar]
  24. Moriyama, Y.; Nguyen, J.; Akens, M.; Moriyama, E.H.; Lilge, L. In vivo effects of low level laser therapy on inducible nitric oxide synthase. Lasers Surg. Med 2009, 41, 227–231. [Google Scholar]
  25. Tuby, H.; Maltz, L.; Oron, U. Modulations of vegf and inos in the rat heart by low level laser therapy are associated with cardioprotection and enhanced angiogenesis. Lasers Surg. Med 2006, 38, 682–688. [Google Scholar]
  26. Fukuda, T.Y.; Tanji, M.M.; Silva, S.R.; Sato, M.N.; Plapler, H. Infrared low-level diode laser on inflammatory process modulation in mice: Pro- and anti-inflammatory cytokines. Lasers Med. Sci. 2012. [Google Scholar] [CrossRef]
  27. Oliveira, R.G.; Ferreira, A.P.; Cortes, A.J.; Aarestrup, B.J.; Andrade, L.C.; Aarestrup, F.M. Low-level laser reduces the production of tnf-alpha, ifn-gamma, and il-10 induced by ova. Lasers Med. Sci. 2013. [Google Scholar] [CrossRef]
  28. Yamaura, M.; Yao, M.; Yaroslavsky, I.; Cohen, R.; Smotrich, M.; Kochevar, I.E. Low level light effects on inflammatory cytokine production by rheumatoid arthritis synoviocytes. Lasers Surg. Med 2009, 41, 282–290. [Google Scholar]
  29. Aimbire, F.; Ligeiro de Oliveira, A.P.; Albertini, R.; Correa, J.C.; Ladeira de Campos, C.B.; Lyon, J.P.; Silva, J.A., Jr; Costa, M.S. Low level laser therapy (lllt) decreases pulmonary microvascular leakage, neutrophil influx and il-1beta levels in airway and lung from rat subjected to lps-induced inflammation. Inflammation 2008, 31, 189–197. [Google Scholar]
  30. Safavi, S.M.; Kazemi, B.; Esmaeili, M.; Fallah, A.; Modarresi, A.; Mir, M. Effects of low-level he-ne laser irradiation on the gene expression of il-1beta, tnf-alpha, ifn-gamma, tgf-beta, bfgf, and pdgf in rat’s gingiva. Lasers Med. Sci 2008, 23, 331–335. [Google Scholar]
  31. Boschi, E.S.; Leite, C.E.; Saciura, V.C.; Caberlon, E.; Lunardelli, A.; Bitencourt, S.; Melo, D.A.; Oliveira, J.R. Anti-inflammatory effects of low-level laser therapy (660 nm) in the early phase in carrageenan-induced pleurisy in rat. Lasers Surg. Med 2008, 40, 500–508. [Google Scholar]
  32. Shiba, H.; Tsuda, H.; Kajiya, M.; Fujita, T.; Takeda, K.; Hino, T.; Kawaguchi, H.; Kurihara, H. Neodymium-doped yttrium-aluminium-garnet laser irradiation abolishes the increase in interleukin-6 levels caused by peptidoglycan through the p38 mitogen-activated protein kinase pathway in human pulp cells. J. Endod 2009, 35, 373–376. [Google Scholar]
  33. Houreld, N.N.; Sekhejane, P.R.; Abrahamse, H. Irradiation at 830 nm stimulates nitric oxide production and inhibits pro-inflammatory cytokines in diabetic wounded fibroblast cells. Lasers Surg. Med 2010, 42, 494–502. [Google Scholar]
  34. Simunovic-Soskic, M.; Pezelj-Ribaric, S.; Brumini, G.; Glazar, I.; Grzic, R.; Miletic, I. Salivary levels of tnf-alpha and il-6 in patients with denture stomatitis before and after laser phototherapy. Photomed. Laser Surg 2010, 28, 189–193. [Google Scholar]
  35. Fushimi, T.; Inui, S.; Nakajima, T.; Ogasawara, M.; Hosokawa, K.; Itami, S. Green light emitting diodes accelerate wound healing: Characterization of the effect and its molecular basis in vitro and in vivo. Wound Repair Regen 2012, 20, 226–235. [Google Scholar]
  36. Saygun, I.; Karacay, S.; Serdar, M.; Ural, A.U.; Sencimen, M.; Kurtis, B. Effects of laser irradiation on the release of basic fibroblast growth factor (bfgf), insulin like growth factor-1 (igf-1), and receptor of igf-1 (igfbp3) from gingival fibroblasts. Lasers Med. Sci 2008, 23, 211–215. [Google Scholar]
  37. Schwartz, F.; Brodie, C.; Appel, E.; Kazimirsky, G.; Shainberg, A. Effect of helium/neon laser irradiation on nerve growth factor synthesis and secretion in skeletal muscle cultures. J. Photochem. Photobiol. B 2002, 66, 195–200. [Google Scholar]
  38. Yu, W.; Naim, J.O.; Lanzafame, R.J. The effect of laser irradiation on the release of bfgf from 3t3 fibroblasts. Photochem. Photobiol 1994, 59, 167–170. [Google Scholar]
  39. Hu, W.P.; Wang, J.J.; Yu, C.L.; Lan, C.C.; Chen, G.S.; Yu, H.S. Helium-neon laser irradiation stimulates cell proliferation through photostimulatory effects in mitochondria. J. Invest. Dermatol 2007, 127, 2048–2057. [Google Scholar]
  40. Lan, C.C.; Wu, C.S.; Chiou, M.H.; Chiang, T.Y.; Yu, H.S. Low-energy helium-neon laser induces melanocyte proliferation via interaction with type iv collagen: Visible light as a therapeutic option for vitiligo. Br. J. Dermatol 2009, 161, 273–280. [Google Scholar]
  41. Wu, S.; Xing, D.; Gao, X.; Chen, W.R. High fluence low-power laser irradiation induces mitochondrial permeability transition mediated by reactive oxygen species. J. Cell. Physiol 2009, 218, 603–611. [Google Scholar]
  42. Zungu, I.L.; Hawkins Evans, D.; Abrahamse, H. Mitochondrial responses of normal and injured human skin fibroblasts following low level laser irradiation—An in vitro study. Photochem. Photobiol 2009, 85, 987–996. [Google Scholar]
  43. Karu, T. Photobiology of low-power laser effects. Health Phys 1989, 56, 691–704. [Google Scholar]
  44. Karu, T.I. Mitochondrial signaling in mammalian cells activated by red and near-ir radiation. Photochem. Photobiol 2008, 84, 1091–1099. [Google Scholar]
  45. Tiphlova, O.; Karu, T. Role of primary photoacceptors in low-power laser effects: Action of he-ne laser radiation on bacteriophage t4-escherichia coli interaction. Lasers Surg. Med 1989, 9, 67–69. [Google Scholar]
  46. Zhang, L.; Xing, D.; Zhu, D.; Chen, Q. Low-power laser irradiation inhibiting abeta25–35-induced pc12 cell apoptosis via pkc activation. Cell. Physiol. Biochem 2008, 22, 215–222. [Google Scholar]
  47. Aimbire, F.; Santos, F.V.; Albertini, R.; Castro-Faria-Neto, H.C.; Mittmann, J.; Pacheco-Soares, C. Low-level laser therapy decreases levels of lung neutrophils anti-apoptotic factors by a nf-κ dependent mechanism. Int. Immunopharmacol 2008, 8, 603–605. [Google Scholar]
  48. Kushibiki, T.; Hirasawa, T.; Okawa, S.; Ishihara, M. Blue laser irradiation generates intracellular reactive oxygen species in various types of cells. Photomed. Laser Surg 2013, 31, 95–104. [Google Scholar]
  49. Lipovsky, A.; Nitzan, Y.; Lubart, R. A possible mechanism for visible light-induced wound healing. Lasers Surg. Med 2008, 40, 509–514. [Google Scholar]
  50. Ignatieva, N.; Zakharkina, O.; Andreeva, I.; Sobol, E.; Kamensky, V.; Lunin, V. Effects of laser irradiation on collagen organization in chemically induced degenerative annulus fibrosus of lumbar intervertebral disc. Lasers Surg. Med 2008, 40, 422–432. [Google Scholar]
  51. Silveira, L.B.; Prates, R.A.; Novelli, M.D.; Marigo, H.A.; Garrocho, A.A.; Amorim, J.C.; Sousa, G.R.; Pinotti, M.; Ribeiro, M.S. Investigation of mast cells in human gingiva following low-intensity laser irradiation. Photomed. Laser Surg 2008, 26, 315–321. [Google Scholar]
  52. Coombe, A.R.; Ho, C.T.; Darendeliler, M.A.; Hunter, N.; Philips, J.R.; Chapple, C.C.; Yum, L.W. The effects of low level laser irradiation on osteoblastic cells. Clin. Orthod. Res 2001, 4, 3–14. [Google Scholar]
  53. Kushibiki, T.; Awazu, K. Controlling osteogenesis and adipogenesis of mesenchymal stromal cells by regulating a circadian clock protein with laser irradiation. Inter. J. Med. Sci 2008, 5, 319–326. [Google Scholar]
  54. Kushibiki, T.; Awazu, K. Blue laser irradiation enhances extracellular calcification of primary mesenchymal stem cells. Photomed. Laser Surg 2009, 27, 493–498. [Google Scholar]
  55. Kushibiki, T.; Tajiri, T.; Ninomiya, Y.; Awazu, K. Chondrogenic mrna expression in prechondrogenic cells after blue laser irradiation. J. Photochem. Photobiol. B 2010, 98, 211–215. [Google Scholar]
  56. Wang, J.; Huang, W.; Wu, Y.; Hou, J.; Nie, Y.; Gu, H.; Li, J.; Hu, S.; Zhang, H. Microrna-193 pro-proliferation effects for bone mesenchymal stem cells after low-level laser irradiation treatment through inhibitor of growth family, member 5. Stem Cells Dev 2012, 21, 2508–2519. [Google Scholar]
  57. Gu, X.; Nylander, E.; Coates, P.J.; Nylander, K. Effect of narrow-band ultraviolet b phototherapy on p63 and microrna (mir-21 and mir-125b) expression in psoriatic epidermis. Acta Derm. Venereol 2011, 91, 392–397. [Google Scholar]
  58. Tome, M.; Lopez-Romero, P.; Albo, C.; Sepulveda, J.C.; Fernandez-Gutierrez, B.; Dopazo, A.; Bernad, A.; Gonzalez, M.A. Mir-335 orchestrates cell proliferation, migration and differentiation in human mesenchymal stem cells. Cell Death Differ 2011, 18, 985–995. [Google Scholar]
  59. Bouvet-Gerbettaz, S.; Merigo, E.; Rocca, J.P.; Carle, G.F.; Rochet, N. Effects of low-level laser therapy on proliferation and differentiation of murine bone marrow cells into osteoblasts and osteoclasts. Lasers Surg. Med 2009, 41, 291–297. [Google Scholar]
  60. Da Silva, A.P.; Petri, A.D.; Crippa, G.E.; Stuani, A.S.; Stuani, A.S.; Rosa, A.L.; Stuani, M.B. Effect of low-level laser therapy after rapid maxillary expansion on proliferation and differentiation of osteoblastic cells. Lasers Med. Sci 2012, 27, 777–783. [Google Scholar]
  61. Ebrahimi, T.; Moslemi, N.; Rokn, A.; Heidari, M.; Nokhbatolfoghahaie, H.; Fekrazad, R. The influence of low-intensity laser therapy on bone healing. J. Dent 2012, 9, 238–248. [Google Scholar]
  62. Fujimoto, K.; Kiyosaki, T.; Mitsui, N.; Mayahara, K.; Omasa, S.; Suzuki, N.; Shimizu, N. Low-intensity laser irradiation stimulates mineralization via increased bmps in mc3t3-e1 cells. Lasers Surg. Med 2010, 42, 519–526. [Google Scholar]
  63. Hou, J.F.; Zhang, H.; Yuan, X.; Li, J.; Wei, Y.J.; Hu, S.S. In vitro effects of low-level laser irradiation for bone marrow mesenchymal stem cells: Proliferation, growth factors secretion and myogenic differentiation. Lasers Surg. Med 2008, 40, 726–733. [Google Scholar]
  64. Kim, H.; Choi, K.; Kweon, O.K.; Kim, W.H. Enhanced wound healing effect of canine adipose-derived mesenchymal stem cells with low-level laser therapy in athymic mice. J. Dermatol. Sci 2012, 68, 149–156. [Google Scholar]
  65. Lin, F.; Josephs, S.F.; Alexandrescu, D.T.; Ramos, F.; Bogin, V.; Gammill, V.; Dasanu, C.A.; de Necochea-Campion, R.; Patel, A.N.; Carrier, E.; et al. Lasers, stem cells, and copd. J. Transl. Med 2010, 8, 16. [Google Scholar]
  66. Luo, L.; Sun, Z.; Zhang, L.; Li, X.; Dong, Y.; Liu, T.C. Effects of low-level laser therapy on ros homeostasis and expression of igf-1 and tgf-beta1 in skeletal muscle during the repair process. Lasers Med. Sci 2013, 28, 725–734. [Google Scholar]
  67. Medrado, A.P.; Soares, A.P.; Santos, E.T.; Reis, S.R.; Andrade, Z.A. Influence of laser photobiomodulation upon connective tissue remodeling during wound healing. J. Photochem. Photobiol. B 2008, 92, 144–152. [Google Scholar]
  68. Nogueira, G.T.; Mesquita-Ferrari, R.A.; Souza, N.H.; Artilheiro, P.P.; Albertini, R.; Bussadori, S.K.; Fernandes, K.P. Effect of low-level laser therapy on proliferation, differentiation, and adhesion of steroid-treated osteoblasts. Lasers Med. Sci 2012, 27, 1189–1193. [Google Scholar]
  69. Renno, A.C.; McDonnell, P.A.; Parizotto, N.A.; Laakso, E.L. The effects of laser irradiation on osteoblast and osteosarcoma cell proliferation and differentiation in vitro. Photomed. Laser Surg 2007, 25, 275–280. [Google Scholar]
  70. Rosa, A.P.; de Sousa, L.G.; Regalo, S.C.; Issa, J.P.; Barbosa, A.P.; Pitol, D.L.; de Oliveira, R.H.; de Vasconcelos, P.B.; Dias, F.J.; Chimello, D.T.; et al. Effects of the combination of low-level laser irradiation and recombinant human bone morphogenetic protein-2 in bone repair. Lasers Med. Sci 2012, 27, 971–977. [Google Scholar]
  71. Saito, K.; Hashimoto, S.; Jung, H.S.; Shimono, M.; Nakagawa, K. Effect of diode laser on proliferation and differentiation of pc12 cells. Bull. Tokyo Dent. Coll 2011, 52, 95–102. [Google Scholar]
  72. Soleimani, M.; Abbasnia, E.; Fathi, M.; Sahraei, H.; Fathi, Y.; Kaka, G. The effects of low-level laser irradiation on differentiation and proliferation of human bone marrow mesenchymal stem cells into neurons and osteoblasts—An in vitro study. Lasers Med. Sci 2012, 27, 423–430. [Google Scholar]
  73. Song, S.; Zhou, F.; Chen, W.R. Low-level laser therapy regulates microglial function through src-mediated signaling pathways: Implications for neurodegenerative diseases. J. Neuroinflammation 2012, 9, 219. [Google Scholar]
  74. Stein, A.; Benayahu, D.; Maltz, L.; Oron, U. Low-level laser irradiation promotes proliferation and differentiation of human osteoblasts in vitro. Photomed. Laser Surg 2005, 23, 161–166. [Google Scholar]
  75. Stein, E.; Koehn, J.; Sutter, W.; Wendtlandt, G.; Wanschitz, F.; Thurnher, D.; Baghestanian, M.; Turhani, D. Initial effects of low-level laser therapy on growth and differentiation of human osteoblast-like cells. Wien. Klin. Wochenschr 2008, 120, 112–117. [Google Scholar]
  76. Dziunycz, P.; Iotzova-Weiss, G.; Eloranta, J.J.; Lauchli, S.; Hafner, J.; French, L.E.; Hofbauer, G.F. Squamous cell carcinoma of the skin shows a distinct microrna profile modulated by uv radiation. J. Invest. Dermatol 2010, 130, 2686–2689. [Google Scholar]
  77. Glorian, V.; Maillot, G.; Poles, S.; Iacovoni, J.S.; Favre, G.; Vagner, S. Hur-dependent loading of mirna risc to the mrna encoding the ras-related small gtpase rhob controls its translation during uv-induced apoptosis. Cell Death Differ 2011, 18, 1692–1701. [Google Scholar]
  78. Guo, L.; Huang, Z.X.; Chen, X.W.; Deng, Q.K.; Yan, W.; Zhou, M.J.; Ou, C.S.; Ding, Z.H. Differential expression profiles of micrornas in nih3t3 cells in response to UVB irradiation. Photochem. Photobiol 2009, 85, 765–773. [Google Scholar]
  79. Pothof, J.; Verkaik, N.S.; van IJcken, W.; Wiemer, E.A.; Ta, V.T.; van der Horst, G.T.; Jaspers, N.G.; van Gent, D.C.; Hoeijmakers, J.H.; Persengiev, S.P. Microrna-mediated gene silencing modulates the uv-induced DNA-damage response. EMBO J 2009, 28, 2090–2099. [Google Scholar]
  80. Tan, G.; Niu, J.; Shi, Y.; Ouyang, H.; Wu, Z.H. Nf-kappab-dependent microrna-125b up-regulation promotes cell survival by targeting p38alpha upon ultraviolet radiation. J. Biol. Chem 2012, 287, 33036–33047. [Google Scholar]
  81. Tan, G.; Shi, Y.; Wu, Z.H. Microrna-22 promotes cell survival upon uv radiation by repressing pten. Biochem. Biophys. Res. Commun 2012, 417, 546–551. [Google Scholar]
  82. Gu, X.; Lundqvist, E.N.; Coates, P.J.; Thurfjell, N.; Wettersand, E.; Nylander, K. Dysregulation of tap63 mrna and protein levels in psoriasis. J. Invest. Dermatol 2006, 126, 137–141. [Google Scholar]
  83. Okuyama, R.; Ogawa, E.; Nagoshi, H.; Yabuki, M.; Kurihara, A.; Terui, T.; Aiba, S.; Obinata, M.; Tagami, H.; Ikawa, S. P53 homologue, p51/p63, maintains the immaturity of keratinocyte stem cells by inhibiting notch1 activity. Oncogene 2007, 26, 4478–4488. [Google Scholar]
  84. Shen, C.S.; Tsuda, T.; Fushiki, S.; Mizutani, H.; Yamanishi, K. The expression of p63 during epidermal remodeling in psoriasis. J. Dermatol 2005, 32, 236–242. [Google Scholar]
  85. Menter, A.; Korman, N.J.; Elmets, C.A.; Feldman, S.R.; Gelfand, J.M.; Gordon, K.B.; Gottlieb, A.; Koo, J.Y.; Lebwohl, M.; Lim, H.W.; et al. Guidelines of care for the management of psoriasis and psoriatic arthritis: Section 5. Guidelines of care for the treatment of psoriasis with phototherapy and photochemotherapy. J. Am. Acad. Dermatol 2010, 62, 114–135. [Google Scholar]
  86. Ozawa, M.; Ferenczi, K.; Kikuchi, T.; Cardinale, I.; Austin, L.M.; Coven, T.R.; Burack, L.H.; Krueger, J.G. 312-nanometer ultraviolet b light (narrow-band UVB) induces apoptosis of t cells within psoriatic lesions. J. Exp. Med 1999, 189, 711–718. [Google Scholar]
  87. Schneider, L.A.; Hinrichs, R.; Scharffetter-Kochanek, K. Phototherapy and photochemotherapy. Clin. Dermatol 2008, 26, 464–476. [Google Scholar]
  88. Celli, J.P.; Spring, B.Q.; Rizvi, I.; Evans, C.L.; Samkoe, K.S.; Verma, S.; Pogue, B.W.; Hasan, T. Imaging and photodynamic therapy: Mechanisms, monitoring, and optimization. Chem. Rev 2010, 110, 2795–2838. [Google Scholar]
  89. Dolmans, D.E.; Fukumura, D.; Jain, R.K. Photodynamic therapy for cancer. Nat. Rev. Cancer 2003, 3, 380–387. [Google Scholar]
  90. Verma, S.; Watt, G.M.; Mai, Z.; Hasan, T. Strategies for enhanced photodynamic therapy effects. Photochem. Photobiol 2007, 83, 996–1005. [Google Scholar]
  91. Buytaert, E.; Dewaele, M.; Agostinis, P. Molecular effectors of multiple cell death pathways initiated by photodynamic therapy. Biochim. Biophys. Acta 2007, 1776, 86–107. [Google Scholar]
  92. Brown, S.B.; Brown, E.A.; Walker, I. The present and future role of photodynamic therapy in cancer treatment. Lancet Oncol 2004, 5, 497–508. [Google Scholar]
  93. Dougherty, T.J. An update on photodynamic therapy applications. J. Clin. Laser Med. Surg 2002, 20, 3–7. [Google Scholar]
  94. Tomioka, Y.; Kushibiki, T.; Awazu, K. Evaluation of oxygen consumption of culture medium and in vitro photodynamic effect of talaporfin sodium in lung tumor cells. Photomed. Laser Surg 2010, 28, 385–390. [Google Scholar]
  95. Oleinick, N.L.; Morris, R.L.; Belichenko, I. The role of apoptosis in response to photodynamic therapy: What, where, why, and how. Photochem. Photochem. Photobiol. Sci 2002, 1, 1–21. [Google Scholar]
  96. Danial, N.N.; Korsmeyer, S.J. Cell death: Critical control points. Cell 2004, 116, 205–219. [Google Scholar]
  97. Ferri, K.F.; Kroemer, G. Organelle-specific initiation of cell death pathways. Nat. Cell Biol 2001, 3, E255–E263. [Google Scholar]
  98. Hengartner, M.O. The biochemistry of apoptosis. Nature 2000, 407, 770–776. [Google Scholar]
  99. Piette, J.; Volanti, C.; Vantieghem, A.; Matroule, J.Y.; Habraken, Y.; Agostinis, P. Cell death and growth arrest in response to photodynamic therapy with membrane-bound photosensitizers. Biochem. Pharmacol 2003, 66, 1651–1659. [Google Scholar]
  100. Helbig, D.; Simon, J.C.; Paasch, U. Photodynamic therapy and the role of heat shock protein 70. Int. J. Hyperthermia 2011, 27, 802–810. [Google Scholar]
  101. Matroule, J.Y.; Volanti, C.; Piette, J. Nf-kappab in photodynamic therapy: Discrepancies of a master regulator. Photochem. Photobiol 2006, 82, 1241–1246. [Google Scholar]
  102. Agostinis, P.; Buytaert, E.; Breyssens, H.; Hendrickx, N. Regulatory pathways in photodynamic therapy induced apoptosis. Photochem. Photobiol. Sci 2004, 3, 721–729. [Google Scholar]
  103. Dewaele, M.; Verfaillie, T.; Martinet, W.; Agostinis, P. Death and survival signals in photodynamic therapy. Methods Mol. Biol 2010, 635, 7–33. [Google Scholar]
  104. Kessel, D.; Oleinick, N.L. Photodynamic therapy and cell death pathways. Methods Mol. Biol 2010, 635, 35–46. [Google Scholar]
  105. Chen, Q.; Huang, Z.; Chen, H.; Shapiro, H.; Beckers, J.; Hetzel, F.W. Improvement of tumor response by manipulation of tumor oxygenation during photodynamic therapy. Photochem. Photobiol 2002, 76, 197–203. [Google Scholar]
  106. Henderson, B.W.; Busch, T.M.; Vaughan, L.A.; Frawley, N.P.; Babich, D.; Sosa, T.A.; Zollo, J.D.; Dee, A.S.; Cooper, M.T.; Bellnier, D.A.; et al. Photofrin photodynamic therapy can significantly deplete or preserve oxygenation in human basal cell carcinomas during treatment, depending on fluence rate. Cancer Res 2000, 60, 525–529. [Google Scholar]
  107. Sitnik, T.M.; Hampton, J.A.; Henderson, B.W. Reduction of tumour oxygenation during and after photodynamic therapy in vivo: Effects of fluence rate. Br. J. Cancer 1998, 77, 1386–1394. [Google Scholar]
  108. Engbrecht, B.W.; Menon, C.; Kachur, A.V.; Hahn, S.M.; Fraker, D.L. Photofrin-mediated photodynamic therapy induces vascular occlusion and apoptosis in a human sarcoma xenograft model. Cancer Res 1999, 59, 4334–4342. [Google Scholar]
  109. Fingar, V.H.; Kik, P.K.; Haydon, P.S.; Cerrito, P.B.; Tseng, M.; Abang, E.; Wieman, T.J. Analysis of acute vascular damage after photodynamic therapy using benzoporphyrin derivative (bpd). Br. J. Cancer 1999, 79, 1702–1708. [Google Scholar]
  110. Keith, B.; Johnson, R.S.; Simon, M.C. Hif1alpha and hif2alpha: Sibling rivalry in hypoxic tumour growth and progression. Nat. Rev. Cancer 2012, 12, 9–22. [Google Scholar]
  111. Semenza, G.L. Hypoxia-inducible factors in physiology and medicine. Cell 2012, 148, 399–408. [Google Scholar]
  112. Forsythe, J.A.; Jiang, B.H.; Iyer, N.V.; Agani, F.; Leung, S.W.; Koos, R.D.; Semenza, G.L. Activation of vascular endothelial growth factor gene transcription by hypoxia-inducible factor 1. Mol. Cell. Biol 1996, 16, 4604–4613. [Google Scholar]
  113. Takenaga, K. Angiogenic signaling aberrantly induced by tumor hypoxia. Front. Biosci 2011, 16, 31–48. [Google Scholar]
  114. Deininger, M.H.; Weinschenk, T.; Morgalla, M.H.; Meyermann, R.; Schluesener, H.J. Release of regulators of angiogenesis following hypocrellin-a and -b photodynamic therapy of human brain tumor cells. Biochem. Biophys. Res. Commun 2002, 298, 520–530. [Google Scholar]
  115. Ferrario, A.; von Tiehl, K.F.; Rucker, N.; Schwarz, M.A.; Gill, P.S.; Gomer, C.J. Antiangiogenic treatment enhances photodynamic therapy responsiveness in a mouse mammary carcinoma. Cancer Res 2000, 60, 4066–4069. [Google Scholar]
  116. Jiang, F.; Zhang, Z.G.; Katakowski, M.; Robin, A.M.; Faber, M.; Zhang, F.; Chopp, M. Angiogenesis induced by photodynamic therapy in normal rat brains. Photochem. Photobiol 2004, 79, 494–498. [Google Scholar]
  117. Schmidt-Erfurth, U.; Schlotzer-Schrehard, U.; Cursiefen, C.; Michels, S.; Beckendorf, A.; Naumann, G.O. Influence of photodynamic therapy on expression of vascular endothelial growth factor (vegf), vegf receptor 3, and pigment epithelium-derived factor. Invest. Ophthalmol. Vis. Sci 2003, 44, 4473–4480. [Google Scholar]
  118. Lu, J.; Getz, G.; Miska, E.A.; Alvarez-Saavedra, E.; Lamb, J.; Peck, D.; Sweet-Cordero, A.; Ebert, B.L.; Mak, R.H.; Ferrando, A.A.; et al. Microrna expression profiles classify human cancers. Nature 2005, 435, 834–838. [Google Scholar]
  119. Cheng, A.M.; Byrom, M.W.; Shelton, J.; Ford, L.P. Antisense inhibition of human mirnas and indications for an involvement of mirna in cell growth and apoptosis. Nucleic Acids Res 2005, 33, 1290–1297. [Google Scholar]
  120. Kushibiki, T. Photodynamic therapy induces microrna-210 and -296 expression in HeLa cells. J. Biophotonics 2010, 3, 368–372. [Google Scholar]
  121. Giannakakis, A.; Sandaltzopoulos, R.; Greshock, J.; Liang, S.; Huang, J.; Hasegawa, K.; Li, C.; O’Brien-Jenkins, A.; Katsaros, D.; Weber, B.L.; et al. Mir-210 links hypoxia with cell cycle regulation and is deleted in human epithelial ovarian cancer. Cancer Biol. Ther 2008, 7, 255–264. [Google Scholar]
  122. Wurdinger, T.; Tannous, B.A.; Saydam, O.; Skog, J.; Grau, S.; Soutschek, J.; Weissleder, R.; Breakefield, X.O.; Krichevsky, A.M. Mir-296 regulates growth factor receptor overexpression in angiogenic endothelial cells. Cancer Cell 2008, 14, 382–393. [Google Scholar]
  123. Bach, D.; Fuereder, J.; Karbiener, M.; Scheideler, M.; Ress, A.L.; Neureiter, D.; Kemmerling, R.; Dietze, O.; Wiederstein, M.; Berr, F.; et al. Comprehensive analysis of alterations in the mirnome in response to photodynamic treatment. J. Photochem. Photobiol. B 2013, 120, 74–81. [Google Scholar]
  124. Sato, M.; Kubota, N.; Inada, E.; Saitoh, I.; Ohtsuka, M.; Nakamura, S.; Sakurai, T.; Watanabe, S. Hela cells consist of two cell types, as evidenced by cytochemical staining for alkaline phosphatase activity: A possible model for cancer stem cell study. Adv. Stem. Cell 2013. [Google Scholar] [CrossRef]
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Figure 1. Representative signaling pathways of apoptosis induced by photodynamic therapy (PDT). Depending on the nature of the photosensitizer and its intracellular localization, the initial photodamage can involve different molecules, with the consequent activation of specific death pathways that converge on mitochondria. Mitochondria-localized photosensitizer can cause immediate and light-dependent photodamage to the anti-apoptotic Bcl-2 and Bcl-xL proteins, prompting the release of caspase-activating molecules. Lysosomal hydrolases and ER stress also induce Bax-mediated caspase activation.

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Figure 1. Representative signaling pathways of apoptosis induced by photodynamic therapy (PDT). Depending on the nature of the photosensitizer and its intracellular localization, the initial photodamage can involve different molecules, with the consequent activation of specific death pathways that converge on mitochondria. Mitochondria-localized photosensitizer can cause immediate and light-dependent photodamage to the anti-apoptotic Bcl-2 and Bcl-xL proteins, prompting the release of caspase-activating molecules. Lysosomal hydrolases and ER stress also induce Bax-mediated caspase activation.
Ijms 14 13542f1 1024
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Figure 2. Expression of miR-210 and miR-296 after PDT in HeLa cells. miR-210 and miR-296 expression levels were significantly increased 1 h after PDT (60 mW/cm2, 90 s) in cells treated with 50 μg/mL talaporfin sodium relative to levels in the control group (i.e., talaporfin sodium concentration of 0 μg/mL) (1 × 104 cells/well). The asterisk, * indicates p < 0.05, a significant difference between the relative expression levels of PDT-treated cells and non-PDT-treated cells. All experiments were performed four times independently. All data are expressed as the means ± SD of four replicates from four experiments (Adapted from [120]).

Click here to enlarge figure

Figure 2. Expression of miR-210 and miR-296 after PDT in HeLa cells. miR-210 and miR-296 expression levels were significantly increased 1 h after PDT (60 mW/cm2, 90 s) in cells treated with 50 μg/mL talaporfin sodium relative to levels in the control group (i.e., talaporfin sodium concentration of 0 μg/mL) (1 × 104 cells/well). The asterisk, * indicates p < 0.05, a significant difference between the relative expression levels of PDT-treated cells and non-PDT-treated cells. All experiments were performed four times independently. All data are expressed as the means ± SD of four replicates from four experiments (Adapted from [120]).
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Table Table 1. Aberrations in miRNA expression after low-level laser therapy (LLLT) to mesenchymal stem cells by using a diode laser (wavelength: 635 nm, 0.5 J/cm2) [56].

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Table 1. Aberrations in miRNA expression after low-level laser therapy (LLLT) to mesenchymal stem cells by using a diode laser (wavelength: 635 nm, 0.5 J/cm2) [56].
UpregulationDownregulation
miR-30e *
miR-15b
miR-30b-5pmiR-204 *
miR-322miR-7a
miR-215miR-423
miR-449amiR-678
miR-126miR-25 *
miR-133bmiR-327
miR-21 *miR-351
miR-455miR-23a
miR-759miR-667
miR-872 *miR-770
miR-29bmiR-324-3p
miR-192miR-30c-2 *
miR-219-1-3pmiR-758
miR-301amiR-320
miR-551bmiR-466c
miR-224
miR-193

miRNAs expression confirmed by quantitative real-time PCR are indicated by underlining.The asterisk, * indicates the star-form of miRNA.

Table Table 2. Aberrations in miRNA expression after PDT to human epidermoid carcinoma cells (A431) by using polyvinylpyrrolidone hypericin (PVPH) [123].

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Table 2. Aberrations in miRNA expression after PDT to human epidermoid carcinoma cells (A431) by using polyvinylpyrrolidone hypericin (PVPH) [123].
UpregulationDownregulation
miR-1290miR-1260b
miR-634miR-720
miR-1246miR-1260
miR-1280
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