Basic & Clinical Pharmacology & Toxicology, 2014, 114, 352–359

Doi: 10.1111/bcpt.12186

Cytotoxicity of Lidocaine to Human Corneal Endothelial Cells In Vitro Hao-Ze Yu1,a, Yi-Han Li2,a, Rui-Xin Wang1, Xin Zhou1, Miao-Miao Yu1, Yuan Ge1, Jun Zhao1 and Ting-Jun Fan1 1

Laboratory for Corneal Tissue Engineering, College of Marine Life Sciences, Ocean University of China, Qingdao, China and 2School of Medicine, Shanghai Jiaotong University, Shanghai, China (Received 11 July 2013; Accepted 22 October 2013) Abstract: Lidocaine has been reported to induce apoptosis on rabbit corneal endothelial cells. However, the apoptotic effect and exact mechanism involved in cytotoxicity of lidocaine are not well-established in human corneal endothelial (HCE) cells. In this study, we investigated the apoptosis-inducing effect of lidocaine on HCE cells in vitro. After HCE cells were treated with lidocaine at concentrations of 0.15625–10.0 g/l, the morphology and ultrastructure of the cells were observed by inverted light microscope and transmission electron microscope (TEM). Cell viability was measured by MTT assay, and the apoptotic ratio was evaluated with flow cytometry and fluorescent microscopic counting after FITC–Annexin V/PI and AO/EB staining. DNA fragmentation was detected by electrophoresis, and the activation of caspases was evaluated by ELISA. In addition, changes in mitochondrial membrane potential were determined by JC-1 staining. Results suggest that lidocaine above 1.25 g/l reduced cellular viability and triggered apoptosis in HCE cells in a time- and dose-dependent manner. Diminishment of DΨm and the activation of caspases indicate that lidocaine-induced apoptosis was caspase dependent and may be related to mitochondrial pathway.

Human corneal endothelium (HCE), a functional monolayer forming the demarcation between cornea and anterior chamber, plays pivotal roles in maintaining corneal transparency by regulating corneal stromal water content [1]. Notwithstanding the evidence of proliferation when deprived of their nature environment, HCE cells are trapped in the G1 phase of the cell cycle and have a limited, if not restricted, proliferative capacity in vivo [2,3], accompanied by an annual attrition rate of 0.3–0.6% in cell density during adulthood [4]. In addition to ageing, excessive HCE cell loss may result from accidental injuries, surgical trauma and diseases [5–7] which consequently impair the physiology of cornea. Due to the inability of HCE to be repaired through cell number increase, wound healing can only be achieved by enlargement and migration of the neighbouring cells. However, if the decline of cell density transcends a threshold, the integrity of the corneal endothelium will be compromised, resulting in painful corneal oedema and ultimately loss of visual acuity. Recently, increased evidence has revealed corneal endothelium damage to be associated with abused administration of topical anaesthetics [8–10]. Lidocaine is one of the most frequently used topical anaesthetic agents in ophthalmic surgeries for its inherent potency, rapid onset, tissue penetration and efficiency [11]. It can result in corneal thickening, opacification and significant corneal endothelial cell loss when applied intracamerally [12]. In addition, administration of lidocaine has been shown to cause Author for correspondence: Ting-Jun Fan, Laboratory for Corneal Tissue Engineering, College of Marine Life Sciences, Ocean University of China, Qingdao 266003, Shandong Province, China (fax +86 532 82031793, e-mail [email protected]). a The first two authors contributed equally to this article.

an increase in apoptotic cells in corneal endothelium in rabbit models [13]. However, the side effects of lidocaine on HCE cells and their mechanism remain unknown. In this study, we aimed to investigate the apoptosis-inducing effect of lidocaine on HCE cells and its mechanism in vitro using untransfected HCE cell line recently established in our own laboratory [14]. Materials and Methods Cell culture. Human corneal endothelial cells were maintained and cultured in DMEM/F12 medium (Invitrogen, Carlsbad, CA, USA) containing 10% fetal bovine serum (Invitrogen) at 37°C and 5% CO2. Morphological observation of HCE cells. Human corneal endothelial cells were seeded onto a 24-well culture plate and cultured in DMEM/ F12 medium (Invitrogen) containing 10% fetal bovine serum (Invitrogen) at 37°C and 5% CO2. Cells at logarithmic phase were treated with lidocaine hydrochloride (Sigma-Aldrich, St. Louis, MO, USA) at concentrations (w/v) from 10 to 0.15625 g/l in step dilutions. HCE cells without lidocaine treatment were used as controls. The morphology and growing status of the cells were monitored under an Eclipse TS100 inverted light microscope (Nikon, Tokyo, Japan) every 4 hr. MTT assay. Cells were seeded into a 96-well cell culture cluster at a density of 1 9 104 cells per well and cultured 48 hr prior to MTT assay. 4–28 hr after lidocaine treatment, the medium was replaced by an equal volume (200 lL) of fresh medium containing 1.1 mM MTT (Sigma-Aldrich) and incubated for 4 hr at 37°C in the dark. The medium was discarded, and 150 lL of dimethyl sulfoxide (SigmaAldrich) was added to dissolve the formazan produced. Cell viability was determined by a colorimetric comparison of the optical density values of the samples using a microplate reader (Multiskan GO; Thermo Scientific, Waltham, MA, USA) at an absorption wavelength of 590 nm.

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CYTOTOXICITY OF LIDOCAINE TO CORNEAL ENDOTHELIAL CELLS Annexin V/propidium iodide (PI) analysis. Human corneal endothelial cells at logarithmic phase in a 24-well culture plate were treated with lidocaine and cultured as described above. After 8- to 24-hr treatment, cells were rinsed and collected by centrifugation for Annexin V/PI staining using FITC Annexin V Apoptosis Detection Kit I (BD Biosciences, Waltham, MA, USA). Each pellet was resuspended in 100 lL binding buffer. In addition, 5 lL Annexin V-FITC and 5 lL propidium iodide were added to each well and incubated for 15 min. Apoptotic ratios were determined by flow cytometry. AO/EB double staining. Alternation in plasma membrane permeability was measured by acridine orange (AO)/ethidium bromide (EB) double-fluorescent staining. HCE cells at logarithmic phase in a 24well culture plate were treated with lidocaine and cultured as mentioned above. The cells were harvested at 1- or 4-hr intervals by trypsin digestion (1–2 min.) and centrifugation. After resuspending the cell pellet with 100 lL serum-free DMEM/F12 medium, 4 lL of AO/ EB (Sigma-Aldrich) solution (100 mg/l AO: 100 mg/l EB = 1:1) was added, mixed and stained for 1 min. at room temperature. 20 lL stained cell suspension from each group was added onto a glass slide, covered with a coverslip and then observed under a Ti-S fluorescent microscope (Nikon, Tokyo, Japan). At least 300 cells were counted in each group. DNA fragmentation assay. DNA fragmentation was examined by agarose gel electrophoresis. Briefly, HCE cells in 25 cm2 flasks were treated with lidocaine at a concentration from 0.3125 to 10 g/l for 1–24 hr. Then, the cells were harvested by scrapping and centrifugation. Genomic DNA was isolated with the Quick Tissue/ Culture Cells Genomic DNA Extraction Kit (Dongsheng Biotech, Beijing, China) following the manufacturer’s instructions. The DNA sample from each group was separated in 1% agarose gel electrophoresis. Transmission electron microscopic observation. After treated with 1.25 g/l lidocaine for 16 hr, HCE cells were collected and fixed with 40 g/l glutaraldehyde in 0.1 M sucrose with 0.2 M sodium cacodylate buffer (pH 7.4) overnight at 4°C. After being washed with sodium cacodylate buffer and post-fixing with 10 g/l osmium tetroxide for 1.5 hr, the fixed cells were dehydrated and embedded in epoxy resin. Ultrathin sections were stained with 20 g/l uranyl acetate–lead citrate and observed by an H700 transmission electron microscope (TEM; Hitachi, Tokyo, Japan). Mitochondrial membrane potentials assay. JC-1 (Sigma-Aldrich) was employed to measure mitochondrial depolarization in HCE cells. Briefly, after cells cultured in 24-well plates were treated as described for the annexin V/PI analysis, cells were incubated with 500 lL JC-1 staining solution (5 mg/l) at 37°C for 20 min. and rinsed twice with PBS. Mitochondrial membrane potentials were monitored by determining the relative amounts of dual emissions from mitochondrial JC-1 monomers or aggregates using flow cytometry. Detection of caspase activation. Human corneal endothelial cells at logarithmic phase in a 25 cm2 culture flask were added with 2.5 g/l lidocaine. Cells were collected every 2 hr and counted. Whole-cell protein extracts were obtained by lysing 2 9 106 cells in 500 lL RIPA (Biotime, Beijing, China) supplemented with protease inhibitors PMSF (Biotime). Samples were mixed, incubated on ice for 30 min. and centrifuged at 16,000 9 g for 15 min. at 4°C to obtain supernatant. High-binding 96-well microtitre plates (Nunc, Waltham, MA, USA) were coated with 100 lL supernatant overnight at 4°C. After three washes in PBS containing 0.05% Tween-20 (PBST), the plates were blocked with 5% non-fat milk (BD Bioscience) at 37°C for 2 hr. After

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three washes with PBST, 100 lL rabbit anti-human caspase 3, 8, 9, 10 (active form) antibodies (Biosynthesis Biotechnology, Beijing, China) was added per well and incubated at 37°C for 2 hr. After three washes with PBST, the primary antibodies were conjugated with 100 lL HRP-labelled goat anti-rabbit secondary antibody (cwBiotech, Nanjing, China) diluted in PBST (1:3000) at 37°C for 1 hr. The plates were washed as described above, and a colorimetric reaction was induced by the addition of 100 lL chromogenic substrate (0.1 mg/mL tetramethylbenzidine, 100 mM acetate buffer, pH 5.6 and 1 mM urea hydrogen peroxide) for 25 min. at room temperature. Colour development was stopped with 50 lL H2SO4 (0.5 M), and the optical density at the wavelength of 450 nm and 630 nm was recorded using a microplate reader (Multiskan GO; Thermo Scientific). Statistics. Mean and standard deviation of the mean were calculated. Error bars are represented as standard deviation of the mean. Comparisons between groups were made by one-way ANOVA or unpaired two-tails assuming equal variance t-test. Differences were considered statistically significant when p < 0.05.

Results Morphological changes of HCE cells. Human corneal endothelial cells treated with lidocaine from concentration of 1.25–10 g/L exhibited morphological changes highly reminiscent to those of apoptotic cells including cytoplasmic vacuolation, cellular shrinkage, detachment from culture matrix and eventually death, while those treated 24 hr below 0.625 g/l lidocaine showed no morphological features of apoptotic cells (fig. 1). Viability changes of HCE cells. Human corneal endothelial cells treated with lidocaine from concentration of 1.25–10 g/l exhibited obvious viability decrease (p < 0.05 or p < 0.01) detected by MTT assay, while those treated with lidocaine below the concentration of 0.625 g/l showed no obvious viability changes (fig. 2). Structural and functional alternation of plasma membrane in HCE cells after lidocaine treatment. We used FITC Annexin V/propidium iodide (PI) to detect the externalization of membrane phospholipid phosphatidylserine (PS) and the loss of membrane integrity (fig. 3A–D). After treatment with 2.5 g/l lidocaine, increase in FITC Annexin V staining (fig. 3A–C K4) preceded the loss of membrane integrity (fig. 3C–D K2). AO/EB staining showed that the membrane integrity of HCE cells treated with 1.25–10 g/l lidocaine was destroyed and membrane permeability was elevated (fig. 4). It was also noticeable that the EB-stained nuclei were all in well-proportioned condensed round or pyknotic morphologies, while the green nuclei were all unevenly stained. The apoptotic rate of these cells, increasing with time and dosage varying, is shown in fig. 5. DNA fragmentation of HCE cells. Genomic DNA extracted from HCE cells treated with 1.25–10 g/l lidocaine for 4–24 hr showed typical DNA ladder or completely DNA degradation in agarose gel electrophoresis

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Fig. 1. Lidocaine induces morphological changes and retarded growth in human corneal endothelial (HCE) cells in vitro. Cultured HCE cells were treated with or without lidocaine (0.15625–10 g/l) for 32 hr. The dosage and time of lidocaine are shown in the top left of each photograph. One representative photograph from three independent experiments is shown. Scale bar, 50 lm.

in a dose-dependent manner (fig. 6A–E). No DNA fragmentation was found in HCE cells treated with 0.625 g/l lidocaine for 24 hr (fig. 6F), the same as the blank control (fig. 6G). Ultrastructural changes of HCE cells. TEM observations showed that some of HCE cells treated with 1.25 g/l lidocaine after 16 hr exhibited early apoptoticlike ultra structural characteristics, that is, cytoplasmic vacuolation, loss of microvilli and cytoplasmic disorganization (fig. 7A). Some cells exhibited middle-stage apoptotic-like ultrastructural characteristics, that is, structural disorganization, chromatin condensation and fragmentation (fig. 7B). And some exhibited ultrastructural characteristics of late apoptotic cells such as disaggregation of cell and nucleus, and shedding apoptotic bodies (fig. 7C). Contrarily, HCE cells in blank

control group preserved normal ultrastructure after cultured for 16 hr (fig. 7D). Apoptotic mechanisms of HCE cells induced by lidocaine. We further evaluated mitochondrial modifications in HCE cells at the early stage of apoptosis. Cells exposed to 2.5 g/l lidocaine exhibited a reduced JC-1 aggregation (altered from red fluorescence to green fluorescence) after 4 hr of treatment, an observation that reflected a drop in DΨm (fig. 8C–D), compared with untreated HCE cells (fig. 8A). We also observed, interestingly, a significant increase in the intake of the monomeric forms of JC-1 (green) into drug-treated HCE cells despite the fact that the JC-1 aggregates (red) remained unchanged (fig. 8B). To determine whether the activation of caspases was involved in lidocaine-induced apoptosis, HCE

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investigated are up-regulated during the early-phase of apoptosis (fig. 9A–D, 0-6 hr). Discussion

Fig. 2. Lidocaine affects the cell viability of human corneal endothelial (HCE) cells in vitro. Cultured HCE cells were treated with or without lidocaine (0.15625–10 g/l) for 28 hr and their viabilities were evaluated by MTT assay. * p < 0.05, ** p < 0.01 versus blank control.

cells were incubated in the presence of 2.5 g/l lidocaine for whole cellular protein extraction and ELISA. Active forms of one executor caspase (caspase 3), one initiator caspase (caspase 9) for intrinsic pathway and two initiator caspase (caspase 8 and caspase 10) for extrinsic pathway were detected by antibodies. Results showed that all the caspases

Lidocaine has been reported to induce apoptosis on rabbit corneal endothelial cells by intracameral injections [12,13]. However, the apoptotic effect and exact mechanism involved in cytotoxicity of lidocaine are not well-established in HCE cells. Inverted light microscopic and TEM observation showed that HCE cells, treated with lidocaine at a concentration above 1.25 g/l, underwent dramatic morphological changes including cytoplasmic vacuolation, cellular shrinkage, detachment and death with time and concentration, which was similar to those of apoptotic cells [15,16]. It was also found that plasma membrane permeability of HCE cells, treated with lidocaine at a concentration above 1.25 g/l, increased with time and concentration followed the externalization of membrane phospholipid phosphatidylserine (PS), one of the earliest features of apoptosis. DNA extracted from lidocaine-treated HCE cells showed typical DNA ladder in agarose gel electrophoresis in a dosedependent manner. In a word, the present study showed that lidocaine significantly induced apoptosis on HCE cells. Based on these observations, a series of experiments were performed to further determine apoptotic mechanisms.

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Fig. 3. Annexin V/PI staining pattern of human corneal endothelial cells treated with lidocaine. A, blank control; B, 2.5 g/l lidocaine, 8 hr; C, 2.5 g/l lidocaine, 16 hr; D, 2.5 g/l lidocaine, 24 hr. (K1 necrosis, K2 late apoptosis, K3 normal cell, K4 early apoptosis).

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Fig. 4. Lidocaine induces membrane integrity collapse in human corneal endothelial (HCE) cells. Cultured HCE cells were treated with or without lidocaine (0.3125–10 g/l) for 24 hr and harvested every 4 hr, respectively. Live and apoptotic populations were observed and analysed by fluorescent microscopy with acridine orange/ethidium bromide double staining. One representative photograph from three independent experiments is shown. Scale bar, 50 lm.

Fig. 5. Apoptotic rate of human corneal endothelial cells treated with lidocaine at different concentrations for different time. * p < 0.05, ** p < 0.01 versus blank control.

Conventional apoptosis can be triggered via two signalling pathways, the intrinsic or mitochondria-mediated pathway and the extrinsic or death receptor-mediated pathway, both of which result in the activation of caspases [17–19]. The present study demonstrated that caspases typical for both the extrinsic and intrinsic pathways were found to be involved in apoptosis induced by lidocaine (fig. 9A–D). Moreover, JC-1 staining showed a drop in DΨm compared with untreated cells. Depolarization of the mitochondria and loss of mitochondrial membrane potential (DΨm) is a sensitive indicator of mitochondrial damage caused by several toxic triggers [20], which is also regarded as one of the first events during apoptosis and may even be a prerequisite for cytochrome c release

Fig. 6. DNA Electrophoresis of human corneal endothelial cells treated with different concentrations of lidocaine after different time intervals. M, D2000 DNA marker, their bp numbers are shown on the left; A, 10 g/l, 4 hr; B, 10 g/l, 2 hr; C, 5 g/l, 4 hr; D, 2.5 g/l,16 hr; E, 1.25 g/l, 16 hr; F, 0.625 g/l, 24 hr; G, blank control, 24 hr. 1% agarose gel was used.

and the activation of caspase 9. Mitochondria with normal DΨm incorporate JC-1 into aggregates (red fluorescence), while in depolarized mitochondria, JC-1 forms monomers (green fluorescence; fig. 8A–D). Together with the rapid activation of caspases, our results indicated that lidocaine-induced apoptosis on HCE cells in a caspase-dependent manner, and it might initiate the process of apoptosis through mitochondrial pathway.

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Fig. 7. Ultrastructural changes of human corneal endothelial cells after treated with lidocaine. A-C, 1.25 g/l lidocaine, 16 hr; D, normal control, 16 hr. V, vacuole; N, nucleus; m, mitochondrion; sc, secreting vesicle; mv, microvillus; *, apoptotic body. Scale bar, 1 lm.

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Fig. 8. Decrease in mitochondrial membrane potential in human corneal endothelial (HCE) cells after treated with lidocaine. HCE cells were cultured in the presence of lidocaine were stained with JC-1 followed by flow cytometry analysis. Mitochondria with normal DΨm incorporate JC-1 into aggregates (red fluorescence), while in depolarized mitochondria, JC-1 forms monomers (green fluorescence). (A) control; (B) 2.5 g/l lidocaine 2 hr; (C) 2.5 g/l lidocaine 4 hr; (D) 2.5 g/l lidocaine 8 hr.

A noteworthy finding was a significant increase in the uptake of the monomeric forms of JC-1 into drug-treated HCE cells, while JC-1 aggregates remained unchanged, which was

thought to be associated with the increase in mitochondrial mass [21–23]. Mitochondrion is a strictly regulated cellular organelle. The mechanisms to increase the mitochondrial mass

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Fig. 9. Detection of caspase activation in 2.5 g/l lidocaine-treated human corneal endothelial cells by ELISA. Rabbit anti-human (A) caspase 9, (B) caspase 3, (C) caspase 8 and (D) caspase 10 antibodies were recognized by secondary antibody conjugated with HRP. (* p < 0.05; ** p < 0.01 versus blank control).

are highly complicated and require orchestration with nuclear genomes as in approximately 1100 various proteins found in mitochondria, only 13 subunits of respiratory chain complexes are encoded by the mitochondrial DNA (mtDNA) [24,25]. Interestingly, there are mounting reports of similar increases in mitochondrial mass in a variety of cell types during apoptotic cell death [26,27]. It is plausible that it constitutes an early response to stress by which the cells adapt to new energy demands and repair injuries, thus increasing their chances for survival. In our case, however, the mitochondrial membrane potential dropped drastically after the relegation of caspase cascade. Therefore, further studies are needed to demonstrate the role of mitochondria mass increase in lidocaine-induced apoptosis on HCE cells. Acknowledgements This work was supported by the National High Technology Research and Development Program (“863” Program) of China (No. 2006AA02A132). References 1 Peh GS, Beuerman RW, Colman A, Tan DT, Mehta JS. Human corneal endothelial cell expansion for corneal endothelium transplantation: an overview. Transplantation 2011;91:811–9.

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CYTOTOXICITY OF LIDOCAINE TO CORNEAL ENDOTHELIAL CELLS 11 Li Z, Chai Y, Gong C, Guizhi D, Liu J, Yang J. Evaluation of the antinociceptive effects of lidocaine and bupivacaine on the tail nerves of healthy rats. Basic Clin Pharmacol 2013;113:31–6. 12 Borazan M, Karalezli A, Oto S, Aydin Akova Y, Karabay G, Kocbiyik A et al. Induction of apoptosis of rabbit corneal endothelial cells by preservative-free lidocaine hydrochloride 2%, ropivacaine 1%, or levobupivacaine 0.75%. J Cataract Refract Surg 2009;35:753–8. 13 Chang Y-S, Tseng S-Y, Tseng S-H, Wu C-L. Cytotoxicity of lidocaine or bupivacaine on corneal endothelial cells in a rabbit. Cornea 2006;25:590–6. 14 Fan T, Zhao J, Ma X, Xu X, Zhao W, Xu B. Establishment of a continuous untransfected human corneal endothelial cell line and its biocompatibility to denuded amniotic membrane. Mol Vis 2011;17:469–80. 15 Yagami T, Yamamoto Y, Kohma H, Nakamura T, Takasu N, Okamura N. L-type voltage-dependent calcium channel is involved in the snake venom group IA secretory phospholipase A2-induced neuronal apoptosis. Neurotoxicology 2013;35:146–53. 16 Periasamy VS, Alshatwi AA. Tea polyphenols modulate antioxidant redox system on cisplatin-induced reactive oxygen species generation in a human breast cancer cell. Basic Clin Pharmacol 2013;112:374–84. 17 Matthews GM, Newbold A, Johnstone RW. Intrinsic and extrinsic apoptotic pathway signaling as determinants of histone deacetylase inhibitor antitumor activity. Adv Cancer Res 2012;116: 165–97. 18 Inoue S, Browne G, Melino G, Cohen GM. Ordering of caspases in cells undergoing apoptosis by the intrinsic pathway. Cell Death Differ 2009;16:1053–61. 19 Ehrhardt H, Wachter F, Maurer M, Stahnke K, Jeremias I. Important role of caspase-8 for chemosensitivity of all cells. Clin Cancer Res 2011;17:7605.

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Cytotoxicity of lidocaine to human corneal endothelial cells in vitro.

Lidocaine has been reported to induce apoptosis on rabbit corneal endothelial cells. However, the apoptotic effect and exact mechanism involved in cyt...
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