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. 2015 Jul 21;182(1):101–107. doi: 10.1111/cei.12671

Langerhans cell-like dendritic cells stimulated with an adjuvant direct the development of Th1 and Th2 cells in vivo

K Matsui 1,✉, A Mori 1, R Ikeda 1
PMCID: PMC4578513  PMID: 26084192

Abstract

It is well known that Langerhans cells (LCs) work as the primary orchestrators in the polarization of immune responses towards a T helper type 1 (Th1) or Th2 milieu. In this study, we attempted to generate LCs from murine bone marrow cells and elicit a Th1- or Th2-prone immune response through the LCs after stimulation with Th1 or Th2 adjuvant. LCs were generated from murine bone marrow cells using granulocyte–macrophage colony-stimulating factor (GM-CSF), interleukin (IL)-4 and transforming growth factor (TGF)-β, and were obtained as I-Ad positive cells. Mice were primed with Th1/Th2 adjuvant- and ovalbumin (OVA)-pulsed LCs and then given a booster injection of OVA 2 days later via the hind footpad. Five days after the OVA injection, the cytokine response in the draining popliteal lymph nodes was investigated by reverse transcription–polymerase chain reaction (RT–PCR) flow cytometry and enzyme-linked immunosorbent assay (ELISA). The generated LCs expressed typical LC surface markers, E-cadherin and Langerin, and were classified accordingly as LC-like dendritic cells (LDCs). Administration of Th1 adjuvant, cytosine–phosphate–guanosine (CpG)-DNA- and OVA-pulsed LDCs into the hind footpads of mice induced a Th1-prone immune response, as represented by up-regulation of IFN-γ production and down-regulation of IL-4 production in the lymph node cells. Conversely, Th2 adjuvant, histamine-pulsed LDCs induced a Th2-prone immune response, as represented by up-regulation of IL-4 production and down-regulation of IFN-γ production. These results suggest that LDCs may be used as a substitute for LCs and have the ability to induce the development of Th1 and Th2 cells in vivo. Our experimental system would therefore be useful for screening of inhibitors of Th1/Th2 differentiation in order to control allergic disease.

Keywords: IFN-γ, IL-4, Langerhans cells, Th1, Th2

Introduction

Langerhans cells (LCs) are a subpopulation of bone marrow-derived dendritic cells (DCs). They are antigen-presenting cells (APCs) capable of internalizing and processing antigen 1. Because they reside in the skin and mucosal membrane epithelia, they may be the primary target cells for antigens entering the skin, oral mucosa and airways 2–5. After antigen uptake they migrate to regional lymph nodes where peptides, in the context of major histocompatibility complex (MHC) class II molecules, are presented to naive T helper (Th) cells with appropriate T helper (Th) cell receptors. This first signal delivered to naive Th cells, together with a second signal, delivered in part by interaction between the CD80 and CD86 molecules on LCs and CD28 on Th cells, results in activation of the Th cells 6,7. Furthermore, LCs work as the primary orchestrators in the polarization of immune responses towards Th type 1 (Th1) or Th2 immune responses. It is likely that the development of Th17 cells or regulatory T cells (Treg) cells would also be regulated by LCs. The nature of the polarization is influenced by a number of factors and, in particular, the development of Th2 cells, producing type II cytokines such as interleukin (IL)-4, IL-5 and IL-13, plays pivotal roles in inducing allergic inflammation 8. Therefore, LCs could become an appropriate target for inhibition of Th2 differentiation, and allergic inflammation might be controllable through down-regulation of LC function. In our previous study, although we used murine LCs purified from epidermis as APCs 9, problems such as the complicated nature of the experimental procedure and low cell recoveries remained. To solve these problems, in the present study we tried to generate LCs from murine bone marrow and to develop a Th1- or Th2-prone immune response through stimulation of the LCs with Th1 or Th2 adjuvants to confirm whether the cells would work as APCs.

Materials and methods

Mice

Female specific pathogen-free BALB/c mice were obtained from Japan SLC (Hamamatsu, Japan) and used at the age of 6–8 weeks. They were housed in plastic cages with sterilized paper bedding in a clean, air-conditioned room at 24°C and allowed free access to a standard laboratory diet and water. All procedures performed on the mice were in accordance with the guidelines of the Animal Care and Use Committee of Meiji Pharmaceutical University, Tokyo.

Generation of murine Langerhans cells (LCs)

Bone marrow cells were cultured in RPMI-10 [RPMI-1640 medium with L-glutamine (Sigma-Aldrich, St Louis, MO, USA) containing 10% fetal bovine serum (Sigma-Aldrich), 25 mM HEPES (Sigma-Aldrich), 100 U/mL penicillin and 100 μg/ml streptomycin (Gibco RBL, Grand Island, NY, USA)] supplemented with recombinant murine granulocyte–macrophage colony-stimulating factor (GM-CSF) (20 ng/ml; PeproTech, Rocky Hill, NJ, USA), recombinant murine IL-4 (100 ng/ml; PeproTech) and recombinant human transforming growth factor (TGF)-β1 (10 ng/ml; PeproTech) at 37°C in a humidified atmosphere with 5% CO2. In a previous study 10, the above three cytokines were used to generate Langerin-positive DCs derived from blood monocytes, and in the present study the concentration of each cytokine was newly optimized to generate LCs from bone marrow cells. Half the total volume of the culture medium was changed every 48 h, and 7 days after the start of culture the grown cells were treated with mouse anti-mouse I-Ad monoclonal antibody (clone 34-5-3s, mouse IgG2a) (1 : 200; Cedarlane, Ontario, Canada) in RPMI-10 for 1 h on ice. The cells reactive with anti-I-Ad antibody were then purified using a CELLectionTM pan mouse immunoglobulin (Ig)G kit (Invitrogen Dynal AS, Oslo, Norway), and used as LCs. LCs, i.e. the I-Ad positive cells, were purified to approximately 95% as determined by flow cytometry (FCM). In addition, expression of E-cadherin and Langerin on the LCs was measured by FCM using phycoerythrin-conjugated anti-mouse E-cadherin monoclonal antibody (clone 114420; rat IgG2a) (R&D Systems, Minneapolis, MN, USA) and phycoerythrin-conjugated anti-mouse CD207 (Langerin) monoclonal antibody (clone eBioL31; rat IgG2a) (eBioscience, San Diego, CA, USA), respectively.

Th1 and Th2 cell development by Langerhans cell-like dendritic cells (LDCs)

LCs obtained were placed as LDCs. LDCs were adjusted to 2 × 105 cells/mL in RPMI-10 and then incubated with 30 μg/ml ovalbumin (OVA) in the absence or presence of 1 μM cytosine–phosphate–guanosine (CpG) DNA or 100 μM histamine at 37°C in a humidified atmosphere with 5% CO2. Phosphorothioate-stabilized CpG DNA (5′TCCATGACGTTCCTGATGCT-3′) was obtained from Eurofins (Tokyo, Japan). Histamine dihydrochloride was purchased from Nacalai (Kyoto, Japan). The cells were collected after incubation for 18 h, washed in RPMI-1640, and administered at a dose of 5 × 104 cells into both hind footpads of mice. After 2 days, 30 μg OVA was injected into both hind footpads of the mice as a booster, and draining popliteal lymph nodes were harvested 5 days after the OVA injection. Th1/Th2 cytokine expression in the lymph node cells was confirmed by reverse transcription–polymerase chain reaction (RT–PCR), FCM and enzyme-linked immunosorbent assay (ELISA).

Detection of mRNA for Th1/Th2 cytokine and chemokine receptor by RT–PCR

In order to determine the levels of mRNA expression for Th1/Th2 cytokine and chemokine receptor, mRNA was extracted from lymph node cells using a Quick Prep Micro mRNA purification kit (GE Healthcare, Little Chalfont, UK). Then, the cDNA was synthesized from 160 ng of the mRNA using a first-strand cDNA synthesis kit (GE Healthcare). PCR was performed using the following primers: β-actin [540 base pairs (bp)] 5′ primer, 5′-GTGGGCCGCTCTAGGCACCAA-3′ and 3′ primer, 5′-CTCTTTGATGTCACGCACGATTTC-3′; interferon (IFN)-γ (405 bp) 5′ primer, 5′-GCTACACACTGCATCTTGGCTTTG-3′ and 3′ primer, 5′-CACTCGGATGAGCTCATTGAATGC-3′; IL-4 (400 bp) 5′ primer, 5′-AGTTGTCATCCTGCTCTTCTTTCTC-3′ and 3′ primer, 5′-CGAGTAATCCATTTGCATGATGCTC-3′; IL-10 (274 bp) 5′ primer, 5′-GTGAAGACTTTCTTTCAAACAAAG-3′ and 3′ primer, 5′-CTGCTCCACTGCCTTGCTCTTATT-3′; TGF-β1 (406 bp) 5′ primer, 5′-CGGGGCGACCTGGGCACCATCCATGAC-3′ and 3′ primer, 5′-CTGCTCCACCTTGGGCTTGCGACCCAC-3′; IL-17A (300 bp) 5′ primer, 5′-TCTCATCCAGCAAGAGATCC-3′ and 3′ primer, 5′-AGTTTGGGACCCCTTTACAC-3′; chemokine (C-X-C motif) receptor 3 (CXCR3) (301 bp) 5′ primer, 5′-ATCAGCGCCTCAATGCCAC-3′ and 3′ primer, 5′-TGGCTTTCTCGACCACAGTT-3′; chemokine (C-C motif) receptor 4 (CCR4) (199 bp) 5′ primer, 5′-TCTACAGCGGCATCTTCTTCAT-3′ and 3′ primer, 5′-CAGTACGTGTGGTTGTGCTCTG-3′ and CCR6 (933 bp) 5′ primer, 5′-ATGAATTCCACAGAGTCCTA-3′ and 3′ primer, 5′-AATAAACGCATACAACACGG-3′. Each PCR was performed using a GeneAmp PCR System 9700 (Perkin-Elmer, Norwalk, CT, USA) in 25 μl of reaction mixture comprising 1·5 μl cDNA (corresponding to 16 ng of mRNA starting material), 200 μM deoxynucleotide triphosphate mixture, 400 nM each PCR primer and 25 U/mL ExTaq DNA polymerase (Takara, Shiga, Japan). The reaction conditions were as follows: one 4-min cycle at 94°C, 35 cycles comprising 45 s at 94°C, 45 s at 61°C and 2 min at 72°C, followed by one 7-min cycle at 72°C, and the PCR products were separated on a 2% agarose gel containing ethidium bromide.

Measurement of intracellular Th1/Th2 cytokine by FCM

Lymph node cells were stained with allophycocyanin-conjugated anti-mouse CD4 monoclonal antibody (clone H129·19; rat IgG2a) (Santa Cruz Biotechnology, Santa Cruz, CA, USA), and then subsequently fixed and permeabilized with a BD Cytofix/CytopermTM Fixation/Permeabilization Kit (BD Biosciences, San Diego, CA, USA). Intracellular cytokines of CD4+ cells were detected by flow cytometry using fluorescein isothiocyanate-conjugated anti-mouse IFN-γ monoclonal antibody (clone XMG1.2; rat IgG1) (Santa Cruz Biotechnology) and phycoerythrin-conjugated anti-mouse IL-4 monoclonal antibody (clone 11B11; rat IgG1) (Santa Cruz Biotechnology).

Quantification of Th1/Th2 cytokine by ELISA

Lymph node cells were adjusted to 1 × 106 cells/ml in RPMI-10. The cultures (0·2 ml/well) were incubated in 96-well culture plates (Nunc, Roskilde, Denmark) in the presence of Dynabeads® mouse T-Activator CD3/CD28 (Invitrogen Dynal AS, Oslo, Norway) at 37°C in a humidified atmosphere with 5% CO2. The culture supernatants were collected after incubation for 48 h, and the IFN-γ and IL-4 concentrations were measured using ELISA kits for quantification of murine IFN-γ and IL-4, respectively (R&D Systems, Minneapolis, MN, USA).

Statistical analysis

The data were expressed as means [± standard deviation (s.d.)], and differences between means were analysed using Student’s t-test with a two-tailed test of significance. Differences at P < 0·05 were considered to be statistically significant.

Results

Expression of E-cadherin and Langerin on LCs generated from bone marrow cells

LCs are immature DCs of the epidermis and mucosal tissues. It is well known that E-cadherin and Langerin are typical LC surface markers 11,12. LCs were generated from mouse bone marrow cells by using three cytokines, GM-CSF, IL-4 and TGF-β1, and subsequently separated as MHC class II-positive cells. FCM analysis of the purified LCs showed that E-cadherin was expressed largely on their cell surface (Fig. 1). Although expression of Langerin was also detected on LCs, the level was weaker than that of E-cadherin. However, the levels of E-cadherin and Langerin expression on these LCs resembled those on primary fresh LCs obtained from trypsinized murine epidermis 13,14. Therefore, these LCs were placed as Langerhans cell-like dendritic cells (LDCs) and used in this study.

Figure 1.

Figure 1

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Expression of E-cadherin and Langerin on Langerhans cell-like dendritic cells (LDCs). LDCs were generated from mouse bone marrow cells, stained with an isotype-matched control monoclonal antibody (solid line), an anti-E-cadherin monoclonal antibody (shaded area) and an anti-Langerin monoclonal antibody (shaded area), and analysed by flow cytometry (FCM). The data shown are representative results of four independent experiments.

Induction of Th1- and Th2-related mRNA expression through LDCs

LDCs were pulsed with OVA for 18 h in the absence or presence of CpG-DNA or histamine. The OVA-pulsed LDCs were injected into the hind footpads of mice, and the mice were boosted with an OVA injection into the hind footpads 2 days later. Popliteal lymph node cells were harvested 5 days after the OVA injection, and the expression of IFN-γ and IL-4 mRNA was confirmed by RT–PCR. Each time-point selected in this experiment was set to detect a maximum immune response. The data in Fig. 2 indicate that LDCs treated with the Th1 adjuvant, CpG-DNA, induced a Th1-prone immune response, as shown by enhanced expression of IFN-γ mRNA and suppressed expression of IL-4 mRNA. Conversely, LDCs treated with the Th2 adjuvant, histamine, induced a Th2-prone immune response, as shown by enhanced expression of IL-4 mRNA and suppressed expression of IFN-γ mRNA. Furthermore, injection of LDCs treated with CpG-DNA enhanced the mRNA expression of the Th1 cell marker, CXCR3, and suppressed the mRNA expression of the Th2 cell and Treg cell marker, CCR4, in popliteal lymph node cells. In contrast, injection of LDCs treated with histamine suppressed the expression of CXCR3 mRNA and enhanced the expression of CCR4 mRNA in popliteal lymph node cells. However, expression of mRNAs for the Treg cytokines IL-10 and TGF-β1, the Th17 cytokine IL-17A and the Th17 cell marker CCR6 was not influenced by CpG-DNA or histamine. These findings indicate that LDCs treated with CpG-DNA and histamine may induce the development of Th1- and Th2-type cells, respectively.

Figure 2.

Figure 2

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T helper type 1 (Th1)- and Th2-related mRNA expression in lymph node cells. Langerhans cell-like dendritic cells (LDCs) were pulsed with ovalbumin (OVA) for 18 h in the absence or presence of cytosine–phosphate–guanosine (CpG)-DNA (a) or histamine (b). The OVA-pulsed LDCs were injected into the hind footpads of mice, and the mice were boosted with an OVA injection into the hind footpads 2 days later. Lymph node cells were harvested 5 days after the OVA injection, and cytoplasmic mRNA was extracted, reverse-transcribed and amplified by polymerase chain reaction (PCR) using primer sets for β-actin, interferon (IFN)-γ, interleukin (IL)-4, IL-10, transforming growth factor (TGF)-β1, IL-17A, CXCR3, CCR4 and CCR6. The data shown are the representative results of four independent experiments.

Development of Th1 cells and Th2 cells through LDCs

LDCs were pulsed with OVA for 18 h in the absence or presence of CpG-DNA or histamine. The OVA-pulsed LDCs were injected into the hind footpads of mice, and the mice were boosted with an OVA injection into the hind footpads 2 days later. Popliteal lymph node cells were harvested 5 days after the OVA injection, and intracellular IFN-γ and IL-4 of CD4+ cells were analysed by FCM. Each time-point selected in this experiment was set to detect a maximum immune response. As shown in Fig. 3, in comparison with control mice that had been injected with non-stimulated LDCs, an increase in the number of IFN-γ−positive cells and a decrease of the number of IL-4-positive cells among CD4+ cells were observed after injection of CpG-DNA-stimulated LDCs. However, the injection of histamine-stimulated LDCs induced an increase in the number of IL-4-positive cells and a decrease in the number of IFN-γ positive cells among CD4+ cells.

Figure 3.

Figure 3

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Detection of intracellular T helper type 1 (Th1)- and Th2 cytokines of CD4+ lymph node cells. Langerhans cell-like dendritic cells (LDCs) were pulsed with ovalbumin (OVA) for 18 h in the absence or presence of cytosine–phosphate–guanosine (CpG)-DNA or histamine. The OVA-pulsed LDCs were injected into the hind footpads of mice, and the mice were boosted with an OVA injection into the hind footpads 2 days later. Lymph node cells were harvested 5 days after the OVA injection, and intracellular IFN-γ and IL-4 in CD4+ cells were analysed by flow cytometry (FCM). The data shown are representative results of four independent experiments.

Furthermore, T lymphocytes among popliteal lymph node cells obtained from the LDC-injected mice were stimulated through surface CD3/CD28 molecules, and the IFN-γ and IL-4 concentrations in the culture supernatants were determined by ELISA. As shown in Fig. 4, the CpG-DNA-pulsed LDCs induced a Th1-prone immune response, as represented by enhanced IFN-γ production and suppressed IL-4 production. Conversely, the histamine-pulsed LDCs induced a Th2-prone immune response, as represented by suppressed IFN-γ production and enhanced IL-4 production.

Figure 4.

Figure 4

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Quantification of T helper type 1 (Th1)- and Th2 cytokine production from T lymphocytes in lymph node. Langerhans cell-like dendritic cells (LDCs) were pulsed with ovalbumin (OVA) for 18 h in the absence or presence of cytosine–phosphate–guanosine (CpG)-DNA or histamine. The OVA-pulsed LDCs were injected into the hind footpads of mice, and the mice were boosted with an OVA injection into the hind footpads 2 days later. Lymph node cells were harvested 5 days after the OVA injection and stimulated through surface CD3/CD28 molecules, and the interferon (IFN)-γ and interleukin (IL)−4 concentrations in the culture supernatants were determined by enzyme-linked immunosorbent assay (ELISA). Each culture was prepared in triplicate, and the mean value was obtained as a representative result for one experiment. The same experiment was repeated six times, and the results are expressed as means ± standard deviation (s.d.) (n = 6). *P < 0·01 versus non-treatment.

Discussion

The Th1/Th2 immune balance plays an integral role in various immunological diseases, including allergy. Many investigators have demonstrated that Th2 immunity is responsible for allergic immune responses and the subsequent pathogenesis of allergic inflammatory diseases 15,16. As a result of modern lifestyles, the prevalence of allergies has been increasing steadily due to reduced exposure to microbial components. The proposed allergy-preventing potential of these factors is no longer present in sufficient quality and/or quantity, leading to an imbalance of the immune system with a predisposition to development of allergic disorders 17. Therefore, pharmacotherapy is required to correct this disparity of the immune system in allergic patients.

LCs are bone marrow-derived MHC class II-positive APCs localized in the epidermis and mucosa. They belong to a DC lineage and are crucial for primary and secondary T cell-dependent immune responses 2. DCs are crucial in determining the outcome of antigen encounter, integrating signals derived from the antigen, its inflammatory context and the host environment into a form that can be read by naive T cells in lymphoid tissues and by effector T cells in peripheral tissues. LDCs used in this study closely resembled LCs in their expression of LC-specific cell surface markers such as E-cadherin and Langerin.

In the present work, we observed an increase in the expression of mRNA for the Th1 cytokine, IFN-γ, and a decrease in the expression of mRNA for the Th2 cytokine, IL-4, in lymph node cells of mice injected with CpG-DNA-stimulated LDCs, whereas in lymph node cells of mice injected with histamine-stimulated LDCs, we observed a decrease in the expression of IFN-γ mRNA and an increase in the expression of IL-4 mRNA. However, the expression of mRNAs for Treg cytokines such as IL-10 and TGF-β1, and Th17 cytokine such as IL-17, were not influenced by injection of either CpG-DNA-stimulated LDCs or histamine-stimulated LDCs. CpG-DNA, which is a powerful activator of Th1 immune responses, has shown promise as an immunomodulator for prevention of Th2-dominant diseases such as allergic rhinitis, pollen allergy, atopic dermatitis and bronchial asthma 18–20. Furthermore, it is well known that histamine is an activator of Th2 immune responses, mediating allergic inflammation 21–24. We further observed an increase in the expression of mRNA for the Th1 cell marker, CXCR3, and a decrease in the expression of mRNA for the Th2 cell marker, CCR4, in lymph node cells of mice injected with CpG-DNA-stimulated LDCs whereas, in contrast, we found that lymph node cells of mice injected with histamine-stimulated LDCs showed decreased expression of CXCR3 mRNA and increased expression of CCR4 mRNA. However, expression of CCR6 mRNA was not influenced by injection of either CpG-DNA-stimulated LDCs or histamine-stimulated LDCs. It has been reported that CXCR3 and CCR6 are expressed on the surface of Th1 and Th17 cells, respectively 25–27. Also, it is known that CCR4 is expressed on the surface of both of Th2 and Treg cells 25,28–31. Therefore, comparison of cytokine patterns and detection of the expression of these chemokine receptors in lymph nodes would predict the development of each T cell type. The available data suggest that CPG-DNA and histamine regulate the development of Th1 and Th2 cells but not that of Treg and Th17 cells, and also that the level of CCR4 mRNA expression controlled by CPG-DNA and histamine reflects the development of Th2 cells but not that of Treg cells. In addition, the results of FCM also demonstrated that the level of expression of mRNAs for Th1 and Th2 cytokines in lymph node cells was correlated with the number of Th1 and Th2 cells that differentiated in the lymph nodes. Furthermore, when the lymph node cells obtained from mice injected with CpG-DNA-treated LDCs or histamine-treated LDCs were stimulated in vitro through the CD3 and CD28 molecules, the culture supernatants showed cytokine production that reflected the Th1/Th2 balance in the lymph nodes. These results demonstrated that LDCs treated with CpG-DNA and histamine induced a Th1 and a Th2 immune response, respectively.

DCs generated from bone marrow cells by the use of GM-CSF alone, which are used widely as APCs, comprise more than three subpopulations, including the progenitor cells of LCs 32. Furthermore, as these GM-CSF DCs do not induce T cell polarization towards a Th2 cell milieu 33, they cannot become substitute cells for LCs. Our present findings suggest that LDCs work as the primary orchestrators in the polarization of immune responses towards a Th1 or Th2 immune milieu as well as primary fresh LCs 9, and that regulation of Th1/Th2 immune balance through LDCs would be beneficial for screening of anti-allergic drugs. To our knowledge, no LC cell line is currently available, making it difficult to investigate potential medication that would target LCs. The LDCs employed in the present study, which were generated from bone marrow cells using GM-CSF, IL-4 and TGF-β1, expressed both E-cadherin and Langerin. Among the DC populations present in murine and human skin, only LCs express both E-cadherin and Langerin, and other DC populations expressing E-cadherin are not detectable 34. These facts suggest that LDCs could be used as substitute cells for LCs. Although the level of Langerin expression on LDCs was weak, it approximated that on primary fresh LCs from trypsinized murine epidermis, as was also the case for E-cadherin expression 13,14. The weak expression of Langerin on LDCs would be attributable to both LDC immaturity and endocytosis of Langerin by LDCs during the process of their purification 14,32. Therefore, when LDCs were administered into the hind footpads of mice, Langerin expression by LDCs might have been augmented upon maturation of the cells. Furthermore, with regard to DC function, it is known that Langerin expression on DCs is induced by peripheral factors such as TGF-β, in addition to cell maturation. Therefore, when mouse bone marrow cells are cultured in the presence of GM-CSF and IL-4, without TGF-β, Langerin-negative DCs are induced 32. As these GM-CSF + IL-4 DCs induce both a Th1 and a Th2 immune response 33, Langerin-negative DCs would also be able to work as appropriate APCs. However, because GM-CSF + IL-4 DCs comprise two DC populations, MHC class IIhigh DC and MHC class IIlow DC 32, the addition of TGF-β is necessary in order to change the properties of GM-CSF + IL-4 DCs so that they resemble more closely those of immature LCs, which are detected as MHC class IIlow + Langerinlow DCs and express MHC class II and Langerin strongly upon maturation 10,32. Therefore, our experimental system using LDCs would be useful for studies of allergy control through correction of the Th1/Th2 balance by targeting LCs.

Acknowledgments

This work was supported by JSPS KAKENHI Grant number 26460238.

Disclosure

The authors declare no conflicts of interest.

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