Analytical Biochemistry 470 (2015) 22–24
Contents lists available at ScienceDirect
Analytical Biochemistry journal homepage: www.elsevier.com/locate/yabio
Notes & Tips
Apoptotic primary normal human dermal fibroblasts for in vitro models of fibrosis Elena García-Gareta ⇑, Nivedita Ravindran, Julian F. Dye RAFT Institute of Plastic Surgery, Mount Vernon Hospital, Northwood HA6 2RN, UK
a r t i c l e
i n f o
Article history: Received 27 August 2014 Received in revised form 13 October 2014 Accepted 15 October 2014 Available online 22 October 2014 Keywords: Apoptosis Cell viability Primary normal human dermal fibroblasts Fibrosis Ethanol
a b s t r a c t Recent studies show that apoptosis affects surrounding tissue, playing a role in diseases such as fibrosis, a significant global disease burden. Elucidating the mechanisms by which the different apoptotic cells present during fibrotic wound healing affect their environment would enable development of new therapies. We describe here a simple, rapid, and cost-effective method for inducing apoptosis of primary normal human dermal fibroblasts without affecting the overall cell viability of the population. Such population could be used for in vitro models of fibrotic wound healing in co-culture with other cells involved in this process to study events such as apoptosis-induced proliferation. Ó 2014 Elsevier Inc. All rights reserved.
Apoptosis, or programmed cell death, plays a crucial role throughout development as well as in repair and regeneration of damaged tissues. Moreover, it takes place during the necessary removal of cells that are not functional or needed any longer. The physical changes that cells undergo during apoptosis—cell membrane blebbing, rearrangement of cytoskeleton, DNA fragmentation, and cell disintegration, among others—are well-characterized. Important progress has been made in understanding the key biochemical elements of apoptosis. However, very little is still known about how apoptotic cells affect their surrounding environment [1,2]. Recent studies suggest that apoptosis is not a process that occurs without consequence to surrounding tissue; apoptotic cells have the ability to secrete mitogenic factors that induce proliferation of neighboring cells. This apoptosis-induced proliferation is caused by stress [1]. Questions such as how apoptotic cells communicate with their surroundings and what signals they send to other cells remain open [1]. More important, how apoptosis affects the environment plays a role in diseases such as fibrosis [3], a significant global disease burden [4]. Therefore, understanding this mechanism would be key to developing new therapies for treating these conditions. In the context of wound healing, apoptosis plays an important role in the homeostasis of the wound environment by keeping the balance between cell proliferation and elimination. In fibrotic outcomes of wound healing such as hypertrophic scars, this ⇑ Corresponding author. E-mail address: [email protected] (E. García-Gareta). http://dx.doi.org/10.1016/j.ab.2014.10.009 0003-2697/Ó 2014 Elsevier Inc. All rights reserved.
balance is altered toward cell proliferation [3]. Moreover, it has been shown that apoptotic endothelial cells may increase the presence and persistence of myofibroblasts, leading to fibrosis [5]. Elucidating the mechanisms by which the different cells present during wound healing affect their environment would enable the development of new therapies. To such means, the generation of viable apoptotic human dermal fibroblasts (HDFs),1 the main cell type present in the dermis, would be necessary to establish in vitro models of fibrosis in co-culture with other cells (myofibroblasts, endothelial cells, keratinocytes, or adult stem cells such as mesenchymal stem cells that are recruited into the wound area) involved in this pathological process. Events such as apoptosis-induced proliferation, cytokine/growth factor expression, and cell migration could be studied. In this article, we report a simple, rapid, and cost-effective method for inducing apoptosis of primary normal HDFs without affecting the overall cell viability of the population that could be used for in vitro models of fibrotic wound healing. This method uses ethanol to induce apoptosis by stress. Ethanol-induced apoptosis is well studied and affects a wide range of cell types, including fibroblasts [6–9]. Early apoptosis detection was done with Annexin V staining because it binds to the exposed phosphatidylserine on the cell surface, an early event in apoptosis. In addition, propidium iodide (PI), a fluorescent DNA dye that stains the nuclei of necrotic 1 Abbreviations used: HDF, human dermal fibroblast; PI, propidium iodide; DMEM, Dulbecco’s modified Eagle’s medium; PBS, phosphate-buffered saline; FITC, fluorescence isothiocyanate; UV, ultraviolet.
Notes & Tips / Anal. Biochem. 470 (2015) 22–24
cells, was used. Cell viability was assessed with alamarBlue, a colorimetric redox assay of metabolic activity and, therefore, of cell viability [10]. Primary normal HDF cultures were established from routine surgical excisions of normal skin obtained with informed consent and local ethics committee approval, as reported previously [11]. HDFs were cultured in Dulbecco’s modified Eagle’s medium (DMEM, Gibco) supplemented with 10% fetal calf serum (Gibco), 100 U/ml penicillin and streptomycin (Gibco), and 100 mM L-glutamine (Gibco) at 37 °C with 5% CO2. Medium was changed twice per week. Adherent HDF cultures were typically established within 3 weeks of isolation. Once cultures are established, cells can be either stored in liquid nitrogen to create a stock or used immediately. A first experiment was performed to investigate the threshold between apoptosis and necrosis of primary HDFs induced by ethanol treatment. Borosilicate glass coverslips (13 mm diameter, VWR International) were placed at the bottom of 12-well plates, and 1 104 HDFs were seeded per coverslip. After 1 h of incubation at 37 °C with 5% CO2, 2 to 3 ml of medium was added per well and plates were incubated overnight at 37 °C with 5% CO2. Medium was removed, and HDFs were incubated with 1 ml of different concentrations of ethanol (0, 0.1, 0.5, 1, 5, 10, 20, 30, and 40%) in both distilled water and phosphate-buffered saline (PBS) for 1 min (n = 3 per condition), after which the ethanol solution was removed and cells were washed twice in cold PBS. A TACS Annexin V–Biotin Kit (Trevigen), which contains both Annexin V and PI, was used. Streptavidin–FITC (fluorescence isothiocyanate) conjugate was used to detect Annexin V. Coverslips were placed on microscope slides and viewed under a fluorescence microscope (AXIOSKOP, Zeiss). Extensive cell death was seen on samples treated with more than 1% ethanol (5–40%), whereas different levels of apoptosis were observed on samples treated with 0.1 to 1% ethanol. A second experiment was then performed to find out which concentration(s) of ethanol induced apoptosis of HDFs without affecting the overall cell viability of the culture. As in the first experiment, coverslips were placed at the bottom of 12-well plates and 1 104 cells were seeded per coverslip. HDFs were incubated with 1 ml of 0, 0.1, 0.5, or 1% ethanol in PBS (n = 6 per condition) for 1 min, after which 3 samples per condition were washed twice in cold PBS (4 °C) and stained with the TACS Annexin V–Biotin Kit.
23
As described before, samples were viewed by fluorescence microscopy. The percentage of apoptotic cells was calculated by counting the total cell number as well as the number of apoptotic cells in 7 views across the middle section of the coverslip at 20 magnification. After the ethanol treatment, the remaining 3 samples per condition were used for an alamarBlue assay; after washing twice with cold PBS, 1 ml of 1% alamarBlue in phenol-free supplemented DMEM was added per well and incubated at 37 °C with 5% CO2 for 4 h. For each sample, 1 ml was transferred to a cuvette and, following the manufacturer’s instructions, absorbance was measured at 570 nm against air using an M550 double beam ultraviolet (UV)/visible spectrophotometer (Spectronic Camspec). Absorbance at 600 nm of phenol-free DMEM was subtracted from samples’ values. All of the results were statistically analyzed using one-way analysis of variance with SigmaStat 3.5 software (P value < 0.05 was considered a significant result). Fig. 1 shows the results from the second experiment. As can be seen from the fluorescence microscopy photos, control cells, which present a typical flattened morphology, appeared to be homogeneously stained for both green and red fluorescence. Nuclei of control cells were not stained, and only the cytoplasm appeared to be stained. Ethanol-treated cells showed a few cells with increased green and red fluorescence, which in some cases presented a disrupted morphology (cell shrinkage), and very rarely did the nucleus appear to be stained (indicating necrosis). Control cells showed some degree of apoptosis because apoptotic cells are always found in cultures [12]. The percentage of apoptosis increased with the concentration of ethanol, as expected, and it was significant only for 1% ethanol-treated cells compared with control cells (P = 0.041). Cell number decreased with increasing ethanol concentration, and it was significantly lower for 1% ethanol-treated cells compared with control cells (P = 0.006) and 0.1% ethanol-treated cells (P = 0.031). Cell viability represents the percentage of cells that died by necrosis [13]. As can be seen from the graph displayed at the bottom of Fig. 1, the cell viability of the 0.1 and 0.5% ethanol-treated cultures was almost identical to that of the control cultures. On the other hand, a reduction in cell viability was measured for the 1% ethanol-treated cultures. However, this reduction was not significant (P = 0.093 compared with control cells, P = 0.132 compared with 0.1% ethanoltreated cells, P = 0.075 compared with 0.5% ethanol-treated cells). Therefore, these results suggest that the generation of viable
Fig.1. Top: Fluorescence microscopy images of Annexin V–FITC (green)/PI (red) stained primary normal HDFs. Apoptotic cells (white arrows) present enhanced green/red fluorescence over background stained cells. Scale bar = 50 lm. Bottom: Percentage of apoptosis (⁄P = 0.041 compared with control cells), cell number as percentage of control (#P = 0.006 compared with control cells, P = 0.031 compared with 0.1% ethanol-treated cells), and cell viability of ethanol-treated primary normal HDFs. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)
24
Notes & Tips / Anal. Biochem. 470 (2015) 22–24
Fig.2. Flow diagram of the method proposed in this article for generating viable apoptotic primary normal HDFs for in vitro models of fibrosis.
apoptotic HDFs can be achieved by treatment with 1% ethanol for 1 min. Such cell population could be used for in vitro models of fibrosis in co-culture with other cells involved in this pathological process for the study of apoptosis-induced proliferation or cytokine/growth factor expression. The advantages of the method proposed in this article (Fig. 2) are that (i) ethanol is an inexpensive ubiquitous reagent in every laboratory, and its effects on apoptosis and cell viability are well studied; (ii) Annexin V/PI staining for dual detection of apoptosis and necrosis followed by fluorescence microscopy analysis is a fast procedure that uses common and easy-to-use laboratory equipment, in contrast to flow cytometry, another technique for measuring apoptosis/necrosis, which is expensive, time-consuming, and not easily accessible for every laboratory and requires highly trained personnel and high cell numbers; and (iii) alamarBlue is a well-studied and characterized colorimetric metabolic assay that is not toxic to the cells (and so cells can be used after the assay, unlike the tetrazolium salts MTT and XTT), does not generate toxic residues, and uses common laboratory equipment (UV/visible spectrophotometer or plate reader). Furthermore, we believe that the method we present here can be extended to other cell types. In conclusion, we have described a simple, rapid, and costeffective method for inducing apoptosis of primary normal HDFs without affecting the overall cell viability of the population. Apoptotic primary normal HDFs could be used for in vitro models of fibrotic wound healing in co-culture with other cells (myofibroblasts, endothelial cells, keratinocytes, or adult stem cells such as mesenchymal stem cells that are recruited into the wound area) involved in fibrosis. Thus, events such as apoptosis-induced proliferation and cytokine/growth factor expression could be studied. Acknowledgment This work was supported by the Restoration of Appearance and Function Trust (UK, registered charity 299811) charitable funds.
References [1] M. Suzanne, H. Steller, Shaping organisms with apoptosis, Cell Death Differ. 20 (2013) 669–675. [2] S. Vriz, S. Reiter, B. Galliot, Cell death: a program to regenerate, Curr. Top. Dev. Biol. 108 (2014) 121–151. [3] A. Johnson, L.A. DiPietro, Apoptosis and angiogenesis: an evolving mechanism for fibrosis, FASEB J. 27 (2013) 3893–3901. [4] World Health Organization, The Global Burden of Disease: 2004 Update, WHO, Geneva, Switzerland, 2004. [5] P. Laplante, I. Sirois, M.A. Raymond, V. Kokta, A. Beliveau, A. Prat, A.V. Pshezhetsky, M.J. Hebert, Caspase-3-mediated secretion of connective tissue growth factor by apoptotic endothelial cells promotes fibrosis, Cell Death Differ. 17 (2010) 291–303. [6] T. Kim, H. Tchah, E.H. Cho, M.S. Kook, Evaluation for safety of cultured corneal fibroblasts with cotreatment of alcohol and mitomycin C, Invest. Ophthalmol. Vis. Sci. 45 (2004) 86–92. [7] C.P. Chen, P. Kuhn, K. Chaturvedi, N. Boyadjieva, D.K. Sarkar, Ethanol induces apoptotic death of developing b-endorphin neurons via suppression of cyclic adenosine monophosphate production and activation of transforming growth factor-b1-linked apoptotic signalling, Mol. Pharmacol. 69 (2006) 706–717. [8] F. Castaneda, S. Rosin-Steiner, Low concentration of ethanol induces apoptosis in HepG2 cells: role of various signal transduction pathways, Int. J. Med. Sci. 3 (2006) 160–167. [9] L. Wang, J. Yang, W. Cui, Y.K. Shin, C. Wu, Involvement of promyelocytic leukemia protein in the ethanol-induced apoptosis n mouse embryo fibroblasts, Yakugaku Zasshi 128 (2008) 1067–1071. [10] M.M. Nociari, A. Shalev, P. Benias, C. Russo, A novel one-step, highly sensitive fluorometric assay to evaluate cell-mediated cytotoxicity, J. Immunol. Methods 213 (1998) 157–167. [11] E. García-Gareta, N. Ravindran, V. Sharma, S. Samizadeh, J.F. Dye, A novel multiparameter in vitro model of three-dimensional cell ingress into scaffolds for dermal reconstruction to predict in vivo outcome, BioRes. Open Access 2 (2013) 412–420. [12] T. Mammone, D. Gan, R. Foyouzi-Youssefi, Apoptotic cell death increases with senescence in normal human dermal fibroblast cultures, Cell Biol. Int. 30 (2006) 903–909. [13] M.G. Neuman, R.M. Nanau, L. Oruña, G. Coto, In vitro anti-inflammatory effects of hyaluronic acid in ethanol-induced damage in skin cells, J. Pharm. Pharm. Sci. 14 (2011) 425–437.
Contents lists available at ScienceDirect
Analytical Biochemistry journal homepage: www.elsevier.com/locate/yabio
Notes & Tips
Apoptotic primary normal human dermal fibroblasts for in vitro models of fibrosis Elena García-Gareta ⇑, Nivedita Ravindran, Julian F. Dye RAFT Institute of Plastic Surgery, Mount Vernon Hospital, Northwood HA6 2RN, UK
a r t i c l e
i n f o
Article history: Received 27 August 2014 Received in revised form 13 October 2014 Accepted 15 October 2014 Available online 22 October 2014 Keywords: Apoptosis Cell viability Primary normal human dermal fibroblasts Fibrosis Ethanol
a b s t r a c t Recent studies show that apoptosis affects surrounding tissue, playing a role in diseases such as fibrosis, a significant global disease burden. Elucidating the mechanisms by which the different apoptotic cells present during fibrotic wound healing affect their environment would enable development of new therapies. We describe here a simple, rapid, and cost-effective method for inducing apoptosis of primary normal human dermal fibroblasts without affecting the overall cell viability of the population. Such population could be used for in vitro models of fibrotic wound healing in co-culture with other cells involved in this process to study events such as apoptosis-induced proliferation. Ó 2014 Elsevier Inc. All rights reserved.
Apoptosis, or programmed cell death, plays a crucial role throughout development as well as in repair and regeneration of damaged tissues. Moreover, it takes place during the necessary removal of cells that are not functional or needed any longer. The physical changes that cells undergo during apoptosis—cell membrane blebbing, rearrangement of cytoskeleton, DNA fragmentation, and cell disintegration, among others—are well-characterized. Important progress has been made in understanding the key biochemical elements of apoptosis. However, very little is still known about how apoptotic cells affect their surrounding environment [1,2]. Recent studies suggest that apoptosis is not a process that occurs without consequence to surrounding tissue; apoptotic cells have the ability to secrete mitogenic factors that induce proliferation of neighboring cells. This apoptosis-induced proliferation is caused by stress [1]. Questions such as how apoptotic cells communicate with their surroundings and what signals they send to other cells remain open [1]. More important, how apoptosis affects the environment plays a role in diseases such as fibrosis [3], a significant global disease burden [4]. Therefore, understanding this mechanism would be key to developing new therapies for treating these conditions. In the context of wound healing, apoptosis plays an important role in the homeostasis of the wound environment by keeping the balance between cell proliferation and elimination. In fibrotic outcomes of wound healing such as hypertrophic scars, this ⇑ Corresponding author. E-mail address: [email protected] (E. García-Gareta). http://dx.doi.org/10.1016/j.ab.2014.10.009 0003-2697/Ó 2014 Elsevier Inc. All rights reserved.
balance is altered toward cell proliferation [3]. Moreover, it has been shown that apoptotic endothelial cells may increase the presence and persistence of myofibroblasts, leading to fibrosis [5]. Elucidating the mechanisms by which the different cells present during wound healing affect their environment would enable the development of new therapies. To such means, the generation of viable apoptotic human dermal fibroblasts (HDFs),1 the main cell type present in the dermis, would be necessary to establish in vitro models of fibrosis in co-culture with other cells (myofibroblasts, endothelial cells, keratinocytes, or adult stem cells such as mesenchymal stem cells that are recruited into the wound area) involved in this pathological process. Events such as apoptosis-induced proliferation, cytokine/growth factor expression, and cell migration could be studied. In this article, we report a simple, rapid, and cost-effective method for inducing apoptosis of primary normal HDFs without affecting the overall cell viability of the population that could be used for in vitro models of fibrotic wound healing. This method uses ethanol to induce apoptosis by stress. Ethanol-induced apoptosis is well studied and affects a wide range of cell types, including fibroblasts [6–9]. Early apoptosis detection was done with Annexin V staining because it binds to the exposed phosphatidylserine on the cell surface, an early event in apoptosis. In addition, propidium iodide (PI), a fluorescent DNA dye that stains the nuclei of necrotic 1 Abbreviations used: HDF, human dermal fibroblast; PI, propidium iodide; DMEM, Dulbecco’s modified Eagle’s medium; PBS, phosphate-buffered saline; FITC, fluorescence isothiocyanate; UV, ultraviolet.
Notes & Tips / Anal. Biochem. 470 (2015) 22–24
cells, was used. Cell viability was assessed with alamarBlue, a colorimetric redox assay of metabolic activity and, therefore, of cell viability [10]. Primary normal HDF cultures were established from routine surgical excisions of normal skin obtained with informed consent and local ethics committee approval, as reported previously [11]. HDFs were cultured in Dulbecco’s modified Eagle’s medium (DMEM, Gibco) supplemented with 10% fetal calf serum (Gibco), 100 U/ml penicillin and streptomycin (Gibco), and 100 mM L-glutamine (Gibco) at 37 °C with 5% CO2. Medium was changed twice per week. Adherent HDF cultures were typically established within 3 weeks of isolation. Once cultures are established, cells can be either stored in liquid nitrogen to create a stock or used immediately. A first experiment was performed to investigate the threshold between apoptosis and necrosis of primary HDFs induced by ethanol treatment. Borosilicate glass coverslips (13 mm diameter, VWR International) were placed at the bottom of 12-well plates, and 1 104 HDFs were seeded per coverslip. After 1 h of incubation at 37 °C with 5% CO2, 2 to 3 ml of medium was added per well and plates were incubated overnight at 37 °C with 5% CO2. Medium was removed, and HDFs were incubated with 1 ml of different concentrations of ethanol (0, 0.1, 0.5, 1, 5, 10, 20, 30, and 40%) in both distilled water and phosphate-buffered saline (PBS) for 1 min (n = 3 per condition), after which the ethanol solution was removed and cells were washed twice in cold PBS. A TACS Annexin V–Biotin Kit (Trevigen), which contains both Annexin V and PI, was used. Streptavidin–FITC (fluorescence isothiocyanate) conjugate was used to detect Annexin V. Coverslips were placed on microscope slides and viewed under a fluorescence microscope (AXIOSKOP, Zeiss). Extensive cell death was seen on samples treated with more than 1% ethanol (5–40%), whereas different levels of apoptosis were observed on samples treated with 0.1 to 1% ethanol. A second experiment was then performed to find out which concentration(s) of ethanol induced apoptosis of HDFs without affecting the overall cell viability of the culture. As in the first experiment, coverslips were placed at the bottom of 12-well plates and 1 104 cells were seeded per coverslip. HDFs were incubated with 1 ml of 0, 0.1, 0.5, or 1% ethanol in PBS (n = 6 per condition) for 1 min, after which 3 samples per condition were washed twice in cold PBS (4 °C) and stained with the TACS Annexin V–Biotin Kit.
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As described before, samples were viewed by fluorescence microscopy. The percentage of apoptotic cells was calculated by counting the total cell number as well as the number of apoptotic cells in 7 views across the middle section of the coverslip at 20 magnification. After the ethanol treatment, the remaining 3 samples per condition were used for an alamarBlue assay; after washing twice with cold PBS, 1 ml of 1% alamarBlue in phenol-free supplemented DMEM was added per well and incubated at 37 °C with 5% CO2 for 4 h. For each sample, 1 ml was transferred to a cuvette and, following the manufacturer’s instructions, absorbance was measured at 570 nm against air using an M550 double beam ultraviolet (UV)/visible spectrophotometer (Spectronic Camspec). Absorbance at 600 nm of phenol-free DMEM was subtracted from samples’ values. All of the results were statistically analyzed using one-way analysis of variance with SigmaStat 3.5 software (P value < 0.05 was considered a significant result). Fig. 1 shows the results from the second experiment. As can be seen from the fluorescence microscopy photos, control cells, which present a typical flattened morphology, appeared to be homogeneously stained for both green and red fluorescence. Nuclei of control cells were not stained, and only the cytoplasm appeared to be stained. Ethanol-treated cells showed a few cells with increased green and red fluorescence, which in some cases presented a disrupted morphology (cell shrinkage), and very rarely did the nucleus appear to be stained (indicating necrosis). Control cells showed some degree of apoptosis because apoptotic cells are always found in cultures [12]. The percentage of apoptosis increased with the concentration of ethanol, as expected, and it was significant only for 1% ethanol-treated cells compared with control cells (P = 0.041). Cell number decreased with increasing ethanol concentration, and it was significantly lower for 1% ethanol-treated cells compared with control cells (P = 0.006) and 0.1% ethanol-treated cells (P = 0.031). Cell viability represents the percentage of cells that died by necrosis [13]. As can be seen from the graph displayed at the bottom of Fig. 1, the cell viability of the 0.1 and 0.5% ethanol-treated cultures was almost identical to that of the control cultures. On the other hand, a reduction in cell viability was measured for the 1% ethanol-treated cultures. However, this reduction was not significant (P = 0.093 compared with control cells, P = 0.132 compared with 0.1% ethanoltreated cells, P = 0.075 compared with 0.5% ethanol-treated cells). Therefore, these results suggest that the generation of viable
Fig.1. Top: Fluorescence microscopy images of Annexin V–FITC (green)/PI (red) stained primary normal HDFs. Apoptotic cells (white arrows) present enhanced green/red fluorescence over background stained cells. Scale bar = 50 lm. Bottom: Percentage of apoptosis (⁄P = 0.041 compared with control cells), cell number as percentage of control (#P = 0.006 compared with control cells, P = 0.031 compared with 0.1% ethanol-treated cells), and cell viability of ethanol-treated primary normal HDFs. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)
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Notes & Tips / Anal. Biochem. 470 (2015) 22–24
Fig.2. Flow diagram of the method proposed in this article for generating viable apoptotic primary normal HDFs for in vitro models of fibrosis.
apoptotic HDFs can be achieved by treatment with 1% ethanol for 1 min. Such cell population could be used for in vitro models of fibrosis in co-culture with other cells involved in this pathological process for the study of apoptosis-induced proliferation or cytokine/growth factor expression. The advantages of the method proposed in this article (Fig. 2) are that (i) ethanol is an inexpensive ubiquitous reagent in every laboratory, and its effects on apoptosis and cell viability are well studied; (ii) Annexin V/PI staining for dual detection of apoptosis and necrosis followed by fluorescence microscopy analysis is a fast procedure that uses common and easy-to-use laboratory equipment, in contrast to flow cytometry, another technique for measuring apoptosis/necrosis, which is expensive, time-consuming, and not easily accessible for every laboratory and requires highly trained personnel and high cell numbers; and (iii) alamarBlue is a well-studied and characterized colorimetric metabolic assay that is not toxic to the cells (and so cells can be used after the assay, unlike the tetrazolium salts MTT and XTT), does not generate toxic residues, and uses common laboratory equipment (UV/visible spectrophotometer or plate reader). Furthermore, we believe that the method we present here can be extended to other cell types. In conclusion, we have described a simple, rapid, and costeffective method for inducing apoptosis of primary normal HDFs without affecting the overall cell viability of the population. Apoptotic primary normal HDFs could be used for in vitro models of fibrotic wound healing in co-culture with other cells (myofibroblasts, endothelial cells, keratinocytes, or adult stem cells such as mesenchymal stem cells that are recruited into the wound area) involved in fibrosis. Thus, events such as apoptosis-induced proliferation and cytokine/growth factor expression could be studied. Acknowledgment This work was supported by the Restoration of Appearance and Function Trust (UK, registered charity 299811) charitable funds.
References [1] M. Suzanne, H. Steller, Shaping organisms with apoptosis, Cell Death Differ. 20 (2013) 669–675. [2] S. Vriz, S. Reiter, B. Galliot, Cell death: a program to regenerate, Curr. Top. Dev. Biol. 108 (2014) 121–151. [3] A. Johnson, L.A. DiPietro, Apoptosis and angiogenesis: an evolving mechanism for fibrosis, FASEB J. 27 (2013) 3893–3901. [4] World Health Organization, The Global Burden of Disease: 2004 Update, WHO, Geneva, Switzerland, 2004. [5] P. Laplante, I. Sirois, M.A. Raymond, V. Kokta, A. Beliveau, A. Prat, A.V. Pshezhetsky, M.J. Hebert, Caspase-3-mediated secretion of connective tissue growth factor by apoptotic endothelial cells promotes fibrosis, Cell Death Differ. 17 (2010) 291–303. [6] T. Kim, H. Tchah, E.H. Cho, M.S. Kook, Evaluation for safety of cultured corneal fibroblasts with cotreatment of alcohol and mitomycin C, Invest. Ophthalmol. Vis. Sci. 45 (2004) 86–92. [7] C.P. Chen, P. Kuhn, K. Chaturvedi, N. Boyadjieva, D.K. Sarkar, Ethanol induces apoptotic death of developing b-endorphin neurons via suppression of cyclic adenosine monophosphate production and activation of transforming growth factor-b1-linked apoptotic signalling, Mol. Pharmacol. 69 (2006) 706–717. [8] F. Castaneda, S. Rosin-Steiner, Low concentration of ethanol induces apoptosis in HepG2 cells: role of various signal transduction pathways, Int. J. Med. Sci. 3 (2006) 160–167. [9] L. Wang, J. Yang, W. Cui, Y.K. Shin, C. Wu, Involvement of promyelocytic leukemia protein in the ethanol-induced apoptosis n mouse embryo fibroblasts, Yakugaku Zasshi 128 (2008) 1067–1071. [10] M.M. Nociari, A. Shalev, P. Benias, C. Russo, A novel one-step, highly sensitive fluorometric assay to evaluate cell-mediated cytotoxicity, J. Immunol. Methods 213 (1998) 157–167. [11] E. García-Gareta, N. Ravindran, V. Sharma, S. Samizadeh, J.F. Dye, A novel multiparameter in vitro model of three-dimensional cell ingress into scaffolds for dermal reconstruction to predict in vivo outcome, BioRes. Open Access 2 (2013) 412–420. [12] T. Mammone, D. Gan, R. Foyouzi-Youssefi, Apoptotic cell death increases with senescence in normal human dermal fibroblast cultures, Cell Biol. Int. 30 (2006) 903–909. [13] M.G. Neuman, R.M. Nanau, L. Oruña, G. Coto, In vitro anti-inflammatory effects of hyaluronic acid in ethanol-induced damage in skin cells, J. Pharm. Pharm. Sci. 14 (2011) 425–437.
