Title Molecular physiological characterization of TRP channels asmediators of cellular redox status( Dissertation_全文 )

Author(s) Heba, Abdallah Elsaid Badr

Citation Kyoto University (京都大学)

Issue Date 2016-11-24

URL https://doi.org/10.14989/doctor.k20064

Right

Type Thesis or Dissertation

Textversion ETD

Kyoto University

Molecular physiological characterization

of TRP channels as mediators of cellular

redox status

Heba Abdallah Elsaid Badr

2016

1

Preface

This is a thesis submitted by the author to Kyoto University for the degree of Doctor of

Engineering. The studies presented in this thesis have been carried out under the direction of

Professor Yasuo Mori at the Laboratory of Molecular Biological Chemistry, Department of Synthetic

Chemistry and Biological Chemistry, Graduate School of engineering, Kyoto University during

September 2012 to 2016.

It is my great pleasure to express my sincere gratitude to Professor Yasuo Mori, who taught

me to the milestone of this research work, for suggesting the topics of this work, and for his kind

guidance, valuable suggestion and advice.

I am greatly indebted to Dr. Tomohiro Numata, for his kind supervision, moral support,

and valuable guidance all through the research work. The author is also deeply grateful to Associate

professor Masayuki Mori and Assistant professor Tatsuki Kurokawa, Department of Synthetic

Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University. I would

like to express my gratitude Dr Daisuke Kozai and Assistant Professor Reiko Sakaguchi, Department

of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto

University.

I would like to express my thankfulness to everybody who has helped me with this thesis. It

should be emphasized that the contribution of Ms. Emiko Mori and all laboratory members is greatly

appreciated.

I extend my thanks to my father, mother, husband and all members of my family, for their

encouragement and understanding throughout the entire period of my study.

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Lastly but not the least, I am grateful to all my Senior Staff and Colleagues for their sincere

feeling and help in bringing out this presentation to light.

Heba Badr

Laboratory of Molecular Biological Chemistry

Department of Synthetic Chemistry and Biological Chemistry

Graduate School of Engineering

Kyoto University

2016

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4

Contents

General Introduction………………..……………………….………………5

Chapter 1

Different contribution of redox-sensitive transient receptor potential channels

to acetaminophen-induced death of human hepatoma cell line……….….....15

Chapter 2

TRPM2 regulates the development of rheumatoid arthritis in the anti-

collagen-induced mouse model…………….……………….……………….64

Chapter 3

Functional and structural divergence in human trpv1 channel subunits by

oxidative cysteine modification……………………….……………………..86

General Conclusion…………………………………………………….....118

List of Publications…………………………………………..…….……...120

5

General Introduction

Alteration of intracellular calcium and oxidative stress

Calcium ion (Ca2+) functions as a second messenger that can regulate many physiological

function, including gene transcription, cell cycle regulation, neurotransmitter release, muscle

contraction, fertilization, gene expression, cell proliferation, differentiation to apoptosis and

irreversible cell damage (Tsien and Tsien, 1990). Calcium signaling occurs through a rise in the

intracellular Ca2+ concentration ([Ca2+]i), which is usually maintained around 100 nM. Elevated

[Ca2+]i is due to enhanced Ca2+ entry across the plasma membrane and/or the release of Ca2+ from

the ER endoplasmic/sarcoplasmic reticulum, Golgi apparatus, and mitochondria. Ca2+ entry is

mediated by store-operated (SOC), receptor-activated, stretch-activated, and ligand-gated Ca2+-

permeable channels (Barritt et al., 2008; El Boustany et al., 2008). Cells usually control [Ca2+]i using

a variety of different channels, and transporters to regulate the movement of Ca2+ across plasma

membrane or between the aforementioned intracellular organelles. The dysregulation of intracellular

Ca2+ ([Ca2+]i) homeostasis leads to many pathological disease including cerebral ataxia,

atherosclerosis, hepatocellular damage, ischemic cell death, neural degeneration, polycystic kidney

disease, irreversible cell damage, and death (Dröge, 2000; Brieger et al., 2012; Tabima et al., 2012).

Recently, there is an intimate interplay between cytosolic Ca2+ signaling and oxygen-free

radicals to induce oxidative stress (Berliocchi et al. 2005; Chvanov et al., 2005; Godfraind, 2005;

Hidalgo 2005; Peers et al., 2005). Where it is known that oxidative stress is a result of the disruption

of the balance between the production of oxidative compounds and natural cellular antioxidant

activity.

Cellular oxidative compounds can be divided in to two entities; reactive oxygen species (ROS)

and reactive nitrogen species (RNS). These species (ROS and RNS) are produced by the metabolism

of normal cells. The ROS are involved in redox biology or oxidative stress. The ROS comprise

superoxide anion (O2-); a highly reactive compound generated during energy production and can

generate hydrogen peroxide (H2O2, less toxic than O2-.), hydroxyl radicals (OH.) (Michael and

Navdeep, 2014), or react with nitric oxide (NO) to produce RNS, especially peroxynitrite (Squadrito

and Pryor, 1998). ROS are generated endogenously as in the process of mitochondrial oxidative

phosphorylation, or interactions with xenobiotic compound (Inoue et al., 2003) and are mainly

detoxified by cellular antioxidants, for example, glutathione and superoxide dismutase (Rahman,

2007).

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ROS can be beneficial or harmful to cells and tissues. It has been recently reported that low

levels of ROS act as a redox messengers in intracellular signaling to regulate physiological process

(Finkel, 2011). Whereas oxidative stress due to high levels of ROS and RNS contribute to oxidative

damage of cellular macromolecules such as DNA, lipid, and protein, and lead to increased levels of

8-hydroxideoxoguanosine as a marker of DNA damage, lipid peroxidation, ultimately inhibiting

protein function, and promoting cell death (Circu and Aw, 2012). The mechanisms involved in

oxidative stress-mediated cellular damage are the same in both intrinsic and extrinsic pathways,

where reactive species accumulate in cells, react with all component of the cell, and subsequently

disrupt their function (Hoye et al., 2008). In addition, ROS can directly mediate cysteine modification

of protein by oxidation of cysteine sulfhydryl within protein in vitro and in vivo (Berlett and Stadtman,

1997). Moreover, alterations in the intracellular thiol/disulfide redox state have been shown to trigger

redox-responsive signaling proteins and pathways (NF-kB transcription factor in human T cells, JNK,

and p38 MAPK signaling pathways) were subsequently shown to be activated under oxidative

conditions including H2O2 (Galter et al., 1994; Hehener et al., 2000). These MAPK pathways activate

pro-apoptotic enzymes, such as the Bcl2 family, inducing cell death. There are also antioxidants

present in the cells that neutralize ROS and RNS, including GSH and ascorbic acid (Valko et al.,

2006).

Oxidative stress can damage the cells through a variety of mechanisms, including activation of

protein kinases, mitochondrial dysfunction, and a [Ca2+]i overload (Hoye et al., 2008). Therefore,

elevation in [Ca2+]i is one of the crucial steps occurring during oxidative stress. In oxidative stress,

[Ca2+]i is increased, promoting a rise of in mitochondrial [Ca2+]i, disrupting mitochondrial membrane

integrity, activating caspases, and destructive enzymes leading to irreversible cell death. It is well

established that high [Ca2+]i levels can be deleterious to normal cell function and has a pivotal role

in the cellular damage mediated by ROS (Yan et al., 2006).

Recently, a group of calcium ion channels have been considered among the redox-responsive

signaling pathways. Transient receptor potential (TRP) channels have been found to play a crucial

role in ROS-induced increases [Ca2+]i (Miller, 2006). Therefore, maintaining ROS homeostasis is

crucial for normal cell growth and survival. For this reason, cells finely control the balancing of ROS

generation and scavenging. An excessive and/or sustained increase in ROS, is associated with many

pathological states, such as abnormal cell growth, diabetes mellitus, rheumatoid arthritis,

ischemia/reperfusion injury, atherosclerosis, programmed cell death, cancer, aging,

neurodegenerative, and liver diseases (Dröge, 2002).

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Redox sensitive transient receptor potential (TRP) channels

Transient receptor potential (TRP) proteins, were first identified in Drosophlia melanogaster

(Montella and Rubin, 1989). There are currently 28 known members of mammalian TRP channel

superfamily (Montella, 2005). The TRP protein has six putative transmembrane domains (TM) and

a pore region between the fifth and sixth transmembrane domains, and it assembles into homo- or

hetero-tetramers to form non-selective Ca2+ permeable cation channels, maintain Ca2+ levels (Figure

1) (Clapham, 2003; Voets et al., 2005), are activated by a myriad of different stimuli. Mammalian

TRP homologs are classified into six subfamilies based on their amino acid sequence homology:

canonical (C), vanilloid (V), melastatin (M), polycystic kidney disease (P), mucolipin (ML), and

ankyrin (A) (Figure 2). TRP channels are expressed and function in a great variety of multicellular

organisms, including yeasts, worms, fruit flies, mice, and humans (Venkatachalam and Montell,

2007). TRP channels play a crucial role in many mammalian senses, including touch, taste, and smell.

Figure 1. Transmembrane topology of TRP channels (Kozai et al., 2014).

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Figure 2. Phylogenetic tree of mammalian TRP channels based on their homology (Takahashi et al.,

2012).

Some of these channels are known to be involved in the rise of [Ca2+]i during ROS

overproduction (Miller and Zhang, 2011). Among the 28 known TRP channels, some are activated

during oxidative stress, these include some members of TRPM, TRPC, and TRPV subfamilies.

TRP-melastatin subfamily (TRPM), named for the tumour suppressor melastatin and being

activated by [Ca2+]i and H2O2 or through the generation of intracellular adenosine diphosphate

ribose (ADPR) (Fleig and Penner, 2004 a,b). The TRPM2 and TRPM7 channels are activated in

oxidative stress. TRPM2 is activated by ROS through various mechanisms, contributing to the rise

in [Ca2+]i, leading to cellular death (Hara et al., 2002; Miller and Zhang 2011). TRPM7 is another

member of the TRPM channels that is believed to be involved in the damage mediated by ROS.

TRPM7 is expressed in most tissues and is non-selectively permeable to Na+ and Ca2+ and is

activated by either ROS or RNS and mediated anoxic neuronal death (Aarts et al., 2003).

TRPC channel is non-selective Ca2+ permeable channel, and its expression is broad (Montell et

al., 2002; Montell et al., 2005) and can be activated by metabotropic changes after receptor

stimulation. TRPC1 activates Ca2+ entry after agonist, growth factor, and phosphokinase C (PKC)

stimulation in endothelial cells, platelets, smooth muscle cells, and B-lymphocytes (Mori et al.,

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2002; Dietrich et al., 2005; Tiruppathi et al. 2006; Authi, 2007). Of the TRPC channel subfamily,

the TRPC3 and TRPC4 channels have been shown to have a role in oxidative stress (Miller 2006;

Miller and Zhang 2011). We have demonstrated that H2O2 and NO activate TRPC5 through

modification of cysteine residues (Yoshida et al., 2006).

The TRPV responds to noxious stimuli including capsaicin, heat, anandamide, and extracellular

acidification (Nilius et al., 2003; Nilius et al., 2004; Vennekens et al., 2008; Vriens et al., 2009).

TRPV1 is the only channel of the TRPV subfamily that is known to be activated during oxidative

stress. However, the mechanism of activation and the role of TRPV1 in oxidative stress-mediated

damage are not known (Miller and Zhang, 2011).

TRPA1 is the only member of TRPA channels, activated by noxious cold and different chemicals

including allyl isothiocyanate, allicin, cinnamaldehyde, mentol, tetra-hydrocannabinoid, nicotine,

and bradykinin (Jordt et al., 2004; Hinman et al., 2006; Macpherson et al., 2007; Andersson et al.,

2009; Hu et al., 2010).

Recently, it has been revealed that a class of TRP channels is sensitive to ROS or RNS mediate

cellular redox status and is notably susceptible to modifications of cysteine residues, such as

oxidation, electrophilic reaction, and S-nitrosylation of sulfhydryls. In literature, H2O2 activates

redox sensitive TRPM2 mediating Ca2+ entry, oxidative stress and cell death (Hara et al., 2002), and

TRPM7 stimulates Ca2+ influx and anoxic cell death (Aarts et al., 2003). Meanwhile, certain families

of TRPC and TRPV as TRPC1, TRPC4, TRPC5, and TRPV1-V4 are activated by ROS and RNS

through cysteine residues oxidation (Yoshida et al., 2006; Takahashi et al., 2011). Moreover, TRPA1

is also activated by exogenous and endogenous electrophilic products of oxidative stress mediating

cellular redox state (Anderson et al., 2008; Bessac et al., 2008).

Survey of this thesis.

This thesis consists of three chapters illustrating the physiological, molecular, and functional role

of redox-sensitive TRP channels in various ROS-related disease. The first chapter describes an

essential role of TRPV1, TRPC1, TRPM2, and TRPM7 in acetaminophen (APAP)-induced

hepatocellular death. The second chapter shows that TRPM2 could be a novel potential therapeutic

strategy for treating rheumatoid arthritis. The last chapter describes that cysteine residues involved

in inter-subunit disulfide bond formation of TRPV1 are implicated in sensing oxidation.

10

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Chapter 1

Different contribution of redox-sensitive transient receptor potential

channels to acetaminophen-induced death of human hepatoma cell line.

ABSTRACT

Acetaminophen (APAP) is a safe analgesic antipyretic drug at prescribed doses. Its overdose,

however, can cause life-threatening liver damage. Though involvement of oxidative stress is

widely acknowledged in APAP-induced hepatocellular death, the mechanism of this increased

oxidative stress and the associated alterations in Ca2+ homeostasis are still unclear. Among

members of transient receptor potential (TRP) channels activated in response to oxidative

stress, we here identify that redox-sensitive TRPV1, TRPC1, TRPM2, and TRPM7 channels

underlie Ca2+ entry and downstream cellular damages induced by APAP in human hepatoma

(HepG2) cells. Our data indicate that APAP treatment of HepG2 cells resulted in increased

reactive oxygen species (ROS) production, glutathione (GSH) depletion, and Ca2+ entry leading

to increased apoptotic cell death. These responses were significantly suppressed by

pretreatment with the ROS scavengers N-acetyl-L-cysteine (NAC) and 4,5-dihydroxy-1,3-

benzene disulfonic acid disodium salt monohydrate (Tiron), and also by preincubation of cells

with the glutathione inducer Dimethylfumarate (DMF). TRP subtype-targeted

pharmacological blockers and siRNAs strategy revealed that suppression of either TRPV1,

TRPC1, TRPM2, or TRPM7 reduced APAP-induced ROS formation, Ca2+ influx, and cell

death; the effects of suppression of TRPV1 or TRPC1, known to be activated by oxidative

cysteine modifications, were stronger than those of TRPM2 or TRPM7. Interestingly, TRPV1

and TRPC1 were labeled by the cysteine-selective modification reagent, 5,5’-dithiobis (2-

nitrobenzoic acid)-2biotin (DTNB-2Bio), and this was attenuated by pretreatment with APAP,

suggesting that APAP and/or its oxidized metabolites act directly on the modification target

cysteine residues of TRPV1 and TRPC1 proteins. In human liver tissue, TRPV1, TRPC1,

TRPM2, and TRPM7 channels transcripts were localized mainly to hepatocytes and Kupffer

cells. Our findings strongly suggest that APAP-induced Ca2+ entry and subsequent

hepatocellular death are regulated by multiple redox-activated cation channels, among which

TRPV1 and TRPC1 play a prominent role.

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INTRODUCTION

Acetaminophen (N-acetyl-para-aminophenol, APAP) is a widely used and safe over-the-counter

analgesic antipyretic drug (Rumack, 2004). However, an accidental APAP overdose can result in

potentially lethal hepatotoxicity in both humans and experimental animals (Thomas, 1993; Hinson

et al., 2010). Adverse reactions of APAP account for a substantial number of acute liver failure cases

necessitating liver transplantation (Larson et al., 2005; Myers et al., 2007; Chun et al., 2009).

Determining the exact pathways underlying APAP-induced hepatocellular death should provide

important cues for preventing its lethal side effects.

It has been established that APAP overdose causes a constellation of co-related cellular events

(Hinson et al., 2010). APAP is largely converted by hepatic cytochrome P450-dependent oxidases to

a reactive intermediate metabolite, which depletes glutathione (GSH) and covalently binds to cellular

proteins and lipids. Research on its mechanisms of toxicity has recently focused on oxidative stress

exerted through accumulation of reactive oxygen species (ROS) and reactive nitrogen species (RNS),

due to the depletion of GSH. APAP also causes mitochondrial dysfunction, deregulation of Ca2+

homeostasis, and DNA fragmentation (Muriel, 2009; Hinson et al., 2010). Furthermore, increased

intracellular Ca2+ ([Ca2+]i) induces the mitochondrial membrane permeability transition (MPT),

which promotes ROS production, loss of mitochondrial potential, cessation of ATP synthesis and

eventually hepatocellular apoptosis or necrosis (Bessems and Vermeulen, 2001). Though increased

[Ca2+]i in hepatocytes is a consequence of APAP overdose, how GSH depletion/ROS accumulation,

[Ca2+]i increases and mitochondrial dysfunction interact to induce APAP overdose-induced

hepatocellular death remains elusive.

Transient receptor potential (TRP) proteins and their homologs have six putative transmembrane

segments that assemble in tetramers to form Ca2+-permeable non-selective cation channels that

regulate [Ca2+]i levels (Clapham, 2003; Voets and Nilius, 2003). Mammalian TRP channels are

classified into six subfamilies: TRPV, TRPC, TRPM, TRPA, TRPP, and TRPML and are activated

by myriad extra- or intracellular physical and chemical stimuli (Numata et al., 2011). Members of

the TRPV, TRPC, TRPM, and TRPA subclasses were reported to act as potential sensors of changes

in cellular redox status and to contribute to ROS-induced [Ca2+]i increases (Numata et al., 2011;

Takahashi et al., 2011; Kozai et al., 2014). Several TRP family channels have been detected in liver

cells (Fonfria et al., 2006; Rychkov and Barritt, 2011). TRPC1 and TRPM7 were reported to be

expressed in H4-IIE rat liver hepatoma cell lines (Brereton et al., 2001; Barritt et al., 2008), while

TRPM2 channels were recently linked to APAP-induced oxidative stress and Ca2+ overload in mouse

hepatocytes (Kheradpezhouh et al., 2014). However, expression and localization of such redox-

sensitive TRP channels in human liver tissues have not been thoroughly investigated. It remains an

17

open question how respective redox-sensitive TRP channels expressed in human hepatocytes

contribute to Ca2+ entry and cell death induced by APAP overdose.

The goal of the present study was to profile redox-sensitive TRP channels expressed in human

hepatoma cell line (HepG2), serving as a model of human liver hepatocytes, and to investigate the

roles of these channels in APAP-induced oxidative stress, Ca2+ influx and consequent cell death. Our

study included assessment of the ameliorative effects of ROS scavengers or GSH inducers on Ca2+

influx during APAP overdose, relevant to expression of these channels. To profile cell-specific

expression of redox-sensitive TRP channels, in situ hybridization was used to map cellular

distribution of TRP mRNAs in normal human liver tissue sections. Our results identified, for the first

time, the redox-activated TRPV1, TRPC1, TRPM2, and TRPM7 channels as being critical in the

mechanism of APAP-induced Ca2+ entry and subsequent HepG2 cell death. These channels were

confirmed to be localized to human liver hepatocytes. Among these channels, functional inhibition

by pharmacological agents and expression suppression by siRNA strategy revealed that the

contributions of TRPV1 and TRPC1 to APAP-induced responses of HepG2 cells were bigger than

those of the other TRP channels. These TRP channels might represent new therapeutic targets for

reducing hepatocellular damage caused by APAP overdoses.

18

MATERIALS AND METHODS

Reagents. N-acetyl-para-aminophenol (APAP), capsazepine (CPZ), 2-aminoethyl diphenylborinate

(2-APB), clotrimazole (CTZ), 2-(12-hydroxydodeca-5,10-diynyl)-3,5,6-trimethyl-p-benzoquinone

(AA861), N-acetyl-L-cysteine (NAC), dimethylfumarate (DMF), metaphosphoric acid,

triethanolamine, and cyclosporine A (CsA) were from Sigma-Aldrich (St. Louis, MO, USA).

Hydrogen peroxide (H2O2) was from Wako Pure Chemical Industries (Osaka, Japan). 4,5-

Dihydroxy-1,3-benzene disulfonic acid disodium salt monohydrate (tiron) was from Tokyo Kasei

Kogyo chemical Co. Ltd (Tokyo, Japan). Mitogen activated protein kinase (MAPK) inhibitors

including extracellular signal-regulated kinase (ERK) inhibitor, (U0126), c-jun N-terminal kinase

(JNK) inhibitor, (SP600125), and p38 kinase inhibitor, (SB203580) were from Calbiochem (La Jolla,

CA, USA). N-(6-Aminohexyl)-5-chloro-2-naphthalenesulfonamide (W-7) was from Santa Cruz

Biotechnology (Santa Cruz, CA, USA). Allyl isothiocyanate (AITC) was from Nacalai Tesque Inc

(Kyoto, Japan).

cDNA Cloning and Recombinant Plasmid Construction. The plasmids of pCI-neo vector carrying

human TRPV1, human TRPV2, human TRPV3, human TRPV4, mouse TRPC1, mouse TRPC4β,

mouse TRPC5, human TRPM2, human TRPM7, and human TRPA1 were used as previously

described (Yoshida et al., 2006; Takahashi et al., 2011). Plasmids of the pCI-neo vector carrying

human TRPC1 were used as previously described (Mori et al., 2002).

Cell Culture and cDNA Expression. Human embryonic kidney cell lines (HEK293, HEK293T)

and HepG2 were cultured in Dulbecco’s modified Eagle’s medium (DMEM) (Sigma) containing

10% fetal bovine serum (FBS), 30 U/ml penicillin, and 30 µg/ml streptomycin (Meiji Seika pharma

Co., Ltd, Tokyo, Japan). Human lung fibroblast (WI-38) cells were cultured in modified Eagle’s

medium (MEM) containing 10% FBS, 30 U/ml penicillin, and 30 µg/ml streptomycin. All cells were

grown at 37°C in a humidified atmosphere of 95% air, 5% CO2. HepG2 (RCB1886) and WI-38

(RCB0702) cells were purchased from RIKEN BRC (Tsukuba, Japan).

HEK293 cells were co-transfected with the recombinant plasmids and pEGFP-F (Clontech

Laboratories, Palo Alto, CA, USA) as a transfection marker using SuperFect Transfection Reagent

(QIAGEN, Valencia, CA, USA) according to the manufacturer’s instructions. Transfected cells were

grown for 36-40 h prior to performing [Ca2+]i measurements. HEK293T cells were transfected with

the recombinant plasmids using Lipofectamine 2000 transfection reagent (Invitrogen, Life

Technologies Corporation, Grand Island, NY, USA) according to the manufacturer’s instructions

and the transfected HEK293T cells were grown for 36 h prior to performing in situ hybridization.

19

siRNA Construction. Small interfering RNA (siRNA) sequences targeting the coding regions of

human TRPV1 mRNA (5′-AACCTATGTAATTCTCACCTACATCCT-3′), human TRPC1 mRNA

(5′-AAGCTTTTCTTGCTGGCGTGC-3′), human TRPM2 mRNA (5′-

AAAGCCTCAGTTCGTGGATTCTT-3′), and human TRPM7 mRNA (5′-

AAGAACAAGCTATGCTTGATGCT-3′) were used. The oligonucleotide sequence used for

synthesis of non-targeting siRNA is 5′-GGGTATACTAGTGAATTAG-3′ (forward) and 5′-

CTAATTCACTAGTATACCC-3′ (reverse). To construct siRNA oligomers, the Silencer siRNA

Construction Kit (Ambion, Life Technologies Corporation, Carlsbad, CA, USA) was used according

to the manufacturer’s protocol. Transfection of siRNAs at 100 nM for human TRPV1, human

TRPC1, and human TRPM2 or 300 nM for human TRPM7 to HepG2 cells were carried out using

Lipofectamine 2000. Cells were treated with siRNAs of human TRPV1 or human TRPC1 for 24 h

and siRNAs of human TRPM2 or human TRPM7 for 48 h. They were then subjected to RT-PCR,

western blotting, [Ca2+]i measurements, intracellular ROS measurements, trypan blue exclusion

assays, Hoechst33342/propidium iodide (PI) assays, caspase 3/7 activity assays, or assessment of

intracellular cytochrome c levels, as indicated in the individual experiments.

[Ca2+]i Measurements. Transfected HEK293, HepG2, and siRNA-transfected HepG2 cells were

subjected to [Ca2+]i measurements 3-16 h after plating onto poly-L-lysine-coated glass coverslips.

Fura-2-AM (Dojindo, Kumamoto, Japan) fluorescence was measured in HEPES-buffered saline

(HBS) containing the following: 107 mM NaCl, 6 mM KCl, 1.2 mM MgSO4, 2 mM CaCl2, 11.5 mM

glucose, and 20 mM HEPES (pH was adjusted to 7.4 with NaOH). Fluorescence images of cells were

recorded and analyzed with the video image analysis system AQUACOSMOS (Hamamatsu

Photonics, Shizuoka, Japan) according to the manufacturer’s instructions. The 340:380 nm ratio

images were obtained on a pixel-by-pixel basis. Fura-2 measurements were performed at 37°C in

HEPES-buffered saline. The 340:380 nm ratio images were converted to Ca2+ concentrations by in

vivo calibration using 10 μM ionomycin (Calbiochem/EMD Chemicals, San Diego, CA, USA) as

described previously (Takahashi et al., 2008).

Reverse Transcriptase Polymerase Chain Reaction (RT-PCR) and PCR. Total RNA from

HepG2, WI-38, and siRNA-transfected HepG2 cells was extracted using ISOGEN (Wako Pure

Chemical Industries) according to the manufacturer’s instructions. The concentration and purity of

RNA were determined using a NanoVue Plus spectrophotometer (GE Healthcare Life Science,

Chalfont, Buckinghamshire, UK). Total RNA samples (0.2 µg) were reverse-transcribed at 42°C for

30 min with Avian Myeloblastosis virus reverse transcriptase using the RNA LA PCR kit (TaKaRa-

Bio, Shiga, Japan). Expression levels of TRPV1-4, TRPC1, TRPC4, TRPC5, TRPM2, TRPM7, and

TRPA1 in the cDNA from HepG2 and the cDNA library of human liver (TaKaRa-Bio, Code No.

20

9505, LotNo. A602) were determined by PCR. TRPA1 expression in the cDNA of WI-38 cells was

also determined by PCR. As a positive control, we amplified the glyceraldehyde-3-phosphate-

dehydrogenase (GAPDH) sequence. Suppression of RNA expression was confirmed by RT-PCR

analysis. PCR was performed with LA Taq polymerase (TaKaRa) and conducted in a thermal cycler

(Gene Amp PCR System 9600, Perkin Elmer Life Sciences, Boston, MA, USA) under the following

conditions: initial heating at 94°C for 2 min, followed by 32-35 cycles of denaturation at 94°C for 2

min, annealing at 55-63°C for 1 min and final extension at 72°C for 1 min. Sequences of gene-

specific primers (synthesized by Sigma-Aldrich), predicated lengths of PCR products and

experimental conditions are listed in Table 1.

Table 1. Primer sequences used in RT-PCR experiments.

Western Blot Analysis. HepG2 and siRNA-transfected HepG2 cells were lysed in ice-cold lysis

RIPA buffer (50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 1% NP-40, 0.1% sodium dodecyl sulfate

(SDS), 0.5% sodium deoxycholate, 0.1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl

fluoride, 10 μg/ml leupeptin, and 5 μg/ml aprotinin) at 4°C for 30 min. The supernatant, containing

protein, was collected by centrifuging at 15,000 rpm for 20 min at 4°C. Protein concentrations of

samples were determined using a Pierce BCA Protein Assay kit (Thermo Fisher Scientific, Rockford,

IL, USA) and samples were fractionated by electrophoresis through 7.5% SDS-polyacrylamide gel

electrophoresis (SDS-PAGE). Protein bands were then transferred onto a polyvinylidene difluoride

(PVDF) membrane (Millipore Corp, Bedford, MA, USA). TRPV1, TRPC1, TRPM2, and TRPM7

were detected by western blotting using anti-TRPV1 (1:1000, Santa Cruz Biotechnology, sc-12498),

anti-TRPC1 (1:1000, Alomone Laboratory, Jerusalem, Israel, ACC-010), anti-TRPM2 (1:1000)

(Lang et al., 2009), and anti-TRPM7 antibodies (1:1000) (Hanano et al., 2004), respectively, with

Genes Sense / Antisense sequence Annealing temperatures (°C) / number of the thermal cycle Expected size 1 HepG2 cDNA of human liver (bp) 2 3

TRPV1 5′-ACGCTGATTGAAGACGGGAAGA-3′

5′-TGCTCTCCTGTGCGATCTTGTT-3′ 58/32 58/32 295

55/35 58/32 476

55/35 58/32 328

55/32 58/32 286

63/32 58/32 371

58/32 58/32 415

58/32 58/32 501

58/32 58/32 262

58/32 58/32 216

58/32 58/32 540

55/32 58/32 451

TRPV2 5′-AGCAGTGGGATGTGGTAAGCTA-3′

5′-TTTGTTCAGGGGCTCCAAAACG-3′

TRPV3 5′-CGAGGATGATTTCCGACTGT-3′

5′-GGGTGCACTCTGCTTCTAGG-3′

TRPV4 5-TGGGGTCTTTCAGCACATCATC-3′

5-GAGACCACGTTGATGTAGAAGG-3′

TRPC1 5′-CAAGATTTTGGAAAATTTCTTG-3′

5′-TTTGTCTTCATGATTTGCTAT-3′

TRPC4 5′-TCTGCAAATATCTCTGGGAAGAATGC-3′

5′-AAGCTTTGTTCGTGCAAATTTCCATTC-3′

TRPC5 5′-GTGGAGTGTGTGTCTAGTTCAG-3′

5′-AGACAGCATGGGAAACAGGAAC-3′

TRPM2 5′-CTGGGAGACGGAGTTCCTGA-3′

5′-TGGGGTACAGCGTGTGGTTG-3′

TRPM7 5′-GTCAGGTGTTTTTGTGGTCGCT-3′

5′-AGCCTCACATACCTTAGCTCTG-3′

TRPA1 5′-GACCACAATGGCTGGACAGCTT-3′

5′-GTACCATTGCGTTGAGGGCTGT-3′

GAPDH 5′-ACCACAGTCCATGCCATCAC-3′

5′-TCCACCACCACCCTGTTGCTGTA-3′

4

21

detection by the ECL system (Amersham Pharmacia Biotech, Piscataway, NJ). For loading

normalization, α-tubulin was detected with an anti-α-tubulin antibody (1:3000, Sigma-Aldrich,

T6074). Chemiluminescence was detected by the luminescent image analyzer LAS-3000 (Fuji film,

Tokyo, Japan). The intensity of the protein bands observed for TRPV1, TRPC1, TRPM2, and

TRPM7 was quantified and normalized to the intensity of α-tubulin bands using the ImageJ software.

DTNB-2Bio Labeling Assay. The 5,5’-dithiobis (2-nitrobenzoic acid)-2biotin (DTNB-2Bio)

labeling assay was performed as previously described [Yoshida et al., 2006]. HEK293T cells

transfected with human TRPV1-GFP, human TRPV4-GFP, human TRPC1-Flag, or vector (5 106

cells) were washed with phosphate-buffered saline (PBS). Cell surface membranes were

permeabilized with PBS containing 0.001% digitonin (Sigma) for 5 min. Cells were then collected

and incubated in HBS solution with or without 20 mM APAP for 3 h at 37°C followed by 100 µM

DTNB-2Bio for 40 min at room temperature. Cells were washed with HBS and lysed in RIPA buffer

(pH 8.0). Cell lysates were incubated batchwise with NeutrAvidin-Plus beads (Thermo-Scientific)

overnight at 4°C with constant shaking. Beads were rinsed three times with RIPA buffer by

centrifugation at 15,000 rpm for 1 min. Proteins were eluted in RIPA buffer containing 50 mM

dithiothreitol (DTT) for 60 min and denatured in SDS sample buffer containing 50 mM DTT for 30

min at room temperature. Proteins were analyzed by 7.5% SDS-PAGE, transferred to a PVDF

membrane, and detected by western blotting with an anti-GFP (Clontech) or anti-Flag (Sigma)

antibody.

Intracellular Reactive Oxygen Species (ROS) Measurements. Intracellular ROS levels in HepG2

cells were measured using 2,7-dichlorofluorescein diacetate (DCF-DA, Sigma-Aldrich) following

the manufacture’s protocol. Cells were incubated with 20 µM DCF-DA dissolved in Hanks’ balanced

salt solution (HBSS) in the dark for 45 min at 37°C, then washed once with HBSS. After loading in

this manner, cells were stimulated with APAP or H2O2, with or without inhibitors, for 3 h.

Fluorescence intensity was measured using a Tecan Infinite M200 microplate reader (Tecan Group

Ltd., Mannedorf, Switzerland) with excitation and emission wavelengths of 485 nm and 530 nm,

respectively.

Reduced Glutathione (GSH) Measurements. GSH content of HepG2 cells was determined using

a GSH assay kit (Cayman Chemical Co, Ann Arbor, MI, USA), according to the manufacturer’s

instructions. This assay was performed on cells that had been treated with APAP or H2O2, with or

without ROS scavengers (NAC, and tiron) or GSH inducer (DMF). Absorbance was measured at

410 nm on a 96-well plate using a Tecan Infinite M200 microplate reader. Samples and standards

were assayed three times in triplicate. The concentrations of GSH were calculated from a standard

curve produced using a range of known GSH concentrations.

22

Trypan Blue Exclusion Assay. Cell viability was assessed by trypan blue exclusion by counting

cells after a 5 min incubation with 0.4% trypan blue (Maeno et al., 2000). Data are represented as the

mean values of triplicates from each separate experiment.

Hoechst33342 and Propidium iodide (PI) Cell Death Assays. Cell death was evaluated by staining

cells with Hoechst33342 (2'-[4-ethoxyphenyl]-5-[4-methyl-1-piperazinyl]-2,5'-bi-1H-

benzimidazole trihydrochloride trihydrate solution) (Dojindo) and propidium iodide (PI) (Setareh

Biotech, Eugene, OR, USA). After 24 h stimulation with various test reagents, as indicated for the

individual experiments, cells were incubated with Hoechst33342 (1 µg/ml) and PI (0.2 µg/ml) for 10

min at 37°C in 95% air, 5% CO2 to stain the nuclei and dead cells, respectively. Staining was

visualized under a fluorescence microscope (Olympus IX81, Olympus, Tokyo, Japan) with

MetaMorph software (Molecular Devices). The proportion of dead cells was calculated by dividing

the number of PI-positive cells by the number of nuclei and this was expressed as percent PI-positive

cells.

Caspase 3/7 Enzymatic Activity Assay. To detect caspase 3/7 activity, HepG2 cells (2×105 cells

per well) were cultured in 96-well plates. After overnight adherence, cells were stimulated with

APAP or H2O2, with or without inhibitors, and caspase 3/7 activity analyzed using ApoONE

Homogeneous Caspase 3/7 Assay kit (Promega, Madison, WI, USA) according to the manufacturer’s

instructions. Fluorescence was read at an excitation wavelength of 485 nm and emission wavelength

of 527 nm using a Tecan Infinite M200 microplate reader.

Measurements of Cytochrome c Release. To detect levels of cytochrome c released, HepG2 cells

(1×106 cells) were plated in a 6-cm cell culture dish. After overnight adherence, cells were stimulated

with APAP or H2O2 with or without inhibitors, then harvested by scraping, washed with cold PBS

and re-suspended in ice-cold cytosol extraction buffer containing 10 mM Tris pH 7.4, 100 mM NaCl,

1 mM EDTA, 1 mM EGTA, 1 mM NaF, 20 mM Na4P2O7, 2 mM Na3VO4, 1% Triton X-100, 10%

glycerol, 0.1% SDS, 0.5% deoxycholate, and protease inhibitor (0.1 mM sodium orthovanadate, 1

mM phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, and 5 μg/ml aprotinin). The cell lysate was

centrifuged at 10,000 rpm for 30 min at 4°C. The supernatant (cytosolic fraction) was collected and

stored at 80°C. A cytochrome c ELISA kit (Invitrogen, Carlsbad, CA, USA) was used to estimate

cytochrome c protein content in the HepG2 cell extracts according to the manufacturer’s instructions.

Measurements were performed in triplicate and the absorbance at 450 nm was determined with a

Tecan Infinite M200 microplate reader. Cytochrome c levels were calculated from a standard curve

produced with a range of known cytochrome c concentrations.

23

DNA Fragmentation Assay. To detect DNA fragmentation, HepG2 cells (1×106 cells) were plated

on 6-cm culture plates and allowed to adhere overnight. Cells were then treated with various doses

of APAP or H2O2 for 24 h, collected by scraping and centrifuged at 1,000 × g for 3 min at 4°C. DNA

was then isolated using the Apoptosis Ladder Detection Kit (Wako Pure Chemical Industries Inc.)

according to the manufacturer’s protocol. DNA fragments were electrophoretically separated on a

1.5% agarose gel in Tris-acetate EDTA buffer containing 40 mM Tris acetate and 1 mM EDTA. The

bands were stained with SYBR® Green I (Molecular probes Inc., Eugene, Oregon, USA).

In Situ Hybridization. hTRPV1 (69-428), hTRPC1 (1419-1923), hTRPM2 (3889-4151), hTRPM7

(371-1020), and hTRPA1 (474-960) cDNA fragments were amplified from the plasmids of pCI-neo

vector carrying human TRPV1, human TRPC1, human TRPM2, human TRPM7, and human TRPA1

using two sets of primers, summarized in Table 2, and cloned into pGEM-T Easy vector (Promega).

In vitro transcription was performed using the digoxigenin (DIG) RNA Labeling Mix (Roche

Applied Science, Roche Diagnostics Deutschland GmbH, Mannheim, Germany) for synthesis of the

sense or antisense DIG-labelled RNA probes according to the manufacturer’s protocol. In situ

hybridization of these transcripts was performed in HEK293T expressing TRP of interest or HepG2

cells according to the manufacturer’s protocol. Human liver paraffin sections obtained from

BioChain Institute, Inc. (San Leandro, CA, USA) were digested for 30 min at 37°C with a freshly

prepared solution of proteinase K (Sigma-Aldrich) at a final concentration 5 µg/ml in 100 mM Tris-

HCl/50 mM EDTA (pH 8.0). After deparaffinization and rehydration, sections were acetylated with

0.25% acetic anhydride in 0.1 M triethanolamine (pH 8.0). Hybridization was performed for 14-16

h at 50°C with 400 ng/ml antisense or sense probe in 5 standard sodium citrate (SSC, 150 mM NaCl

and 15 mM sodium citrate, pH 7.4), 0.5 M EDTA (pH 8.0), 1 Denhardt’s solution (0.02% Ficoll,

0.02% polyvinylpyrrolidone, 0.2 mg/ml RNasefree bovine serum albumin), 50 mg/ml Heparin, 10

mg/ml yeast tRNA, 10% Tween 20, 10% CHAPS, 10% dextran sulphate, and 50% formamide.

Sections were washed twice in 2 SSC with 50% formamide at 50°C for 15 min. They were then

washed further at increasingly high stringencies up to a final condition of 0.2 SSC at 50°C for 20

min. Washed slides were then incubated in a blocking reagent containing 100 mM Tris-HCl (pH

7.5), 150 mM NaCl, 1% normal goat serum (NGS), and 0.1% Triton X-100 at room temperature for

1 h. Immunohistochemical detection of the hybridized probes was performed using alkaline

phosphatase-conjugated anti-DIG antibody (Roche Applied Science, Cat. No. 11093274910) diluted

to 1:500 in blocking buffer for 16 h. Alkaline phosphatase activity was visualized by incubating for

16-24 h with nitroblue tetrazolium (Roche Applied Science) and 5-bromo-4-chloro-3-indolyl

phosphate (Roche Applied Science) in buffer containing 100 mM Tris-HCl (pH 9.5), 50 mM MgCl2,

100 mM NaCl and 1 mM levamisole (Sigma-Aldrich), protected from light. The colorimetric reaction

was stopped by washing sections twice for 10 min in buffer containing 0.1 M Tris-HCl and 1 mM

24

EDTA, pH 8.0. Sections were then counterstained with 0.02% fast green FCF (Sigma-Aldrich) and

coverslips were mounted with Entellan® New (Merck Millipore, Darmstadt, Germany). All sections

were analyzed with an Olympus IX81 microscope.

Table 2. Primer sequences used in RT-PCR experiments.

Statistical Analysis. All data are expressed as means ± SEM. We collected data for each condition

from at least three independent experiments. Statistical analysis was performed with the Student’s t

test for comparing two groups. In experiments involving more than two conditions, statistical

analysis was performed with a one way analysis of variance (ANOVA) and Bonferroni post-hoc

analysis. A P value of < 0.05 was considered statistically significant. All statistical analyses were

performed using Prism 6.02 (GraphPad Software, Inc., San Diego, CA, USA).

Genes Sense / Antisense sequence 1 2 3

TRPV1 5′-CCCCCTGGATGGAGACCCTA-3′

5′-CTGCAGAAGAGCAAGAAGCA-3′

TRPC1 5′-TTCTGTGGATTATTGGGATGA-3′

5′-CAGAACAAAGCAAAGCAGGTG-3′

TRPM2 5′-CTGGGAGACGGAGTTCCTGA-3′

5′-TGGGGTACAGCGTGTGGTTG-3′

TRPM7 5-TCCAGGATGTCAAATTTG-3′

5-TATGAAATGGGAATGCAG-3′

TRPA1 5′-CCCCTCTGCATTGTGCTGTA-3′

5′-AAAATGTGCCTGGACAATGG-3′

4

25

RESULTS

[Ca2+]i Increases Are Evoked by Acetaminophen and H2O2 in HepG2 Cells

Intracellular signaling molecules such as Ca2+ and ROS normally regulate diverse cellular

functions (Yan et al., 2006). Nonetheless, ROS-induced oxidative stress and impaired Ca2+

homeostasis could potentially contribute to hepatocellular death induced by APAP overdose (Hinson

et al., 2010). We first tested whether APAP and H2O2 administration induced Ca2+ entry in HepG2

cells, using these cells as a model of human hepatocytes. Treatment of HepG2 cells with APAP at

various concentrations (5, 15 or 20 mM) for 10 min resulted in a gradual, dose-dependent increase

in [Ca2+]i (Figure 1A), whereas H2O2 administration, at concentrations > 100 µM, evoked abrupt

[Ca2+]i increases at 37°C (Figure 1B). Thus, Ca2+ entry positively correlates with the dose of APAP

and ROS used for stimulation.

Figure 1. [Ca2+]i responses induced by acetaminophen (APAP) or H2O2 in HepG2 cells. (A) [Ca2+]i rises

induced by APAP for 10 min at indicated concentrations in HepG2 cells. Average time courses (left) and

maximal rise of [Ca2+]i (Δ[Ca2+]i) (right) (n = 23-56). (B) [Ca2+]i rises induced by H2O2 for 10 min at indicated

concentrations in HepG2 cells. Average time courses (left) and Δ [Ca2+]i (right) (n = 21-45). Data points are

mean ± SEM. P ≥ 0.05, **P < 0.01, and ***P < 0.001 compared with control. Differences not statistically

significant are labelled as (ns). All data were analyzed by Student's t-test.

26

APAP-Evoked ROS Production Elicits Ca2+ Responses in HepG2 Cells

APAP induces ROS generation (DeVries, 1981). We confirmed that treatment with 20 mM

APAP or 1 mM H2O2 for 3 h significantly increased ROS levels in HepG2 cells, as compared with

in untreated controls. Pretreatment for 3 h with the ROS scavengers N-acetyl-L-cysteine (NAC) and

tiron, each at 1 mM, significantly reduced ROS levels produced after treatment with either 20 mM

APAP or 1 mM H2O2 for additional 3 h (Figure 2A,B). Administration of either of the ROS

scavengers alone did not significantly change ROS levels, as compared with those in untreated

control cells. In [Ca2+]i measurements, increases in [Ca2+]i in response to 20 mM of APAP or 1 mM

H2O2 were also significantly suppressed by pretreatment with either NAC or tiron, at 1 mM, for 6

min (Figure 2C,D). Together, these results indicate that ROS production evoked by APAP and H2O2

induces the Ca2+ responses in HepG2 cells.

27

Figure 2. Attenuation of APAP- or H2O2-induced ROS production and Ca2+ responses by ROS

scavengers in HepG2 cells. (A) Suppression of ROS levels induced by APAP (20 mM) treatment for 3 h by

either N-acetyl-L-cysteine (NAC) or tiron (1 mM). (B) Suppression of ROS levels induced by H2O2 treatment

(1 mM) for 3 h by either NAC or tiron (1mM). (C) Effects of either NAC or tiron (1 mM) on APAP (20 mM)

induced-[Ca2+]i responses in HepG2 cells. Average time courses (left) and Δ[Ca2+]i (right) (n = 24-62). (D)

Effects of either NAC or tiron (1 mM) on H2O2 (1 mM) induced-Ca2+ entry in HepG2 cells. Average time

courses (left) and Δ[Ca2+]i (right) (n = 19-33). ROS scavengers are applied for 3 h before and during APAP or

H2O2 stimulation. Data points are mean ± SEM. P ≥ 0.05, **P < 0.01, and ***P < 0.001. Differences not

statistically significant are labelled as (ns). All data of [Ca2+]i measurements were analyzed by Student's t-test,

while those of ROS measurements were analyzed by ANOVA and Bonferroni post-hoc.

Characterization of redox-Sensitive TRP Channels Found in HepG2 using HEK293 Cell

System

The data above suggested a role for ROS in mediating [Ca2+]i increases in response to APAP

overdose in HepG2 cells. TRPV1, TRPV3, TRPV4, TRPC1, TRPC4, TRPC5 (Yoshida et al., 2006),

TRPM2 (Hara et al., 2002), TRPM7 (Aarts et al., 2003), and TRPA1 channels (Takahashi et al.,

2008) were reportedly activated by H2O2. Therefore, we hypothesized that these channels might be

involved in the aforementioned APAP-induced Ca2+ responses in HepG2 cells. To investigate this,

we first examined expression of mRNAs for redox-sensitive TRPV1-4, TRPC1, TRPC4, TRPC5,

TRPM2, TRPM7, and TRPA1 channels by RT-PCR in HepG2 cells. As shown in Figure 3A, RNAs

encoding TRPV1-4, TRPC1, TRPM2, and TRPM7 were detected in HepG2 cells. Next, we tested

whether 20 mM APAP activates recombinant ROS-sensitive TRP channels expressed in HEK293

cells. APAP, at 20 mM, evoked [Ca2+]i responses in HEK293 cells expressing TRPV1, TRPC1,

TRPM2, TRPM7, or TRPA1 but not in cells expressing TRPV2, TRPV3, TRPV4, TRPC4, or TRPC5

(Figure 3B). In contrast, all TRP channels except TRPC4 responded to H2O2 (Figure 3C), as

previously reported (Hara et al., 2002; Aarts et al., 2003; Yoshida et al., 2006; Takahashi et al., 2008).

These results illustrate the sensitivity of HEK293 cells expressing TRPV1, TRPC1, TRPM2,

TRPM7, and TRPA1 to both APAP and H2O2. Though, TRPA1, among all the channels, showed the

highest Ca2+ responses to APAP, its molecular expression had not, as noted above, been detected in

HepG2 cells. This was confirmed by comparing HepG2 with a positive control, demonstrating

TRPA1 mRNA expression in WI-38 human lung fibroblast cells (Figure 3D), which express

functional TRPA1 (Jaquemar et al., 1999; Hu et al., 2010; Kozai et al., 2014). In addition, we found

that 100 µM AITC, a TRPA1 agonist (Jordt et al., 2004), failed to elicit a Ca2+ response in HepG2

cells (Figure 3E). Therefore, TRPV1 is the most responsive to APAP, among the recombinant TRPs

that are found being expressed in HepG2 cells. Overall, these results raise a possibility that redox-

sensitive TRPV1, TRPC1, TRPM2, and TRPM7 contribute to the APAP-induced Ca2+ responses in

HepG2 cells.

28

Figure 3. Expression of redox-sensitive TRP channels in HepG2 cells and their responses when expressed

in HEK293 cells. (A) Expression of redox-sensitive TRP channel mRNAs (TRPV1-4, TRPC1, TRPC4, TRPC5, TRPM2, TRPM7, TRPA1, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH)) detected by

RT-PCR in total RNA isolated from HepG2 cells. Specific PCR primers used are listed in Table 1. (B, C)

[Ca2+]i responses evoked by 20 mM APAP (B) or 1 mM H2O2 (C) in HEK293 cells expressing human TRPV1,

29

human TRPV2, human TRPV3, human TRPV4, mouse TRPC1, mouse TRPC4β, mouse TRPC5, human

TRPM2, human TRPM7, human TRPA1, or vector. Average time courses (left) and Δ[Ca2+]i (right) (n = 16-

64). Data points are mean ± SEM. P ≥ 0.05, *P < 0.05, **P < 0.01, and ***P < 0.001 compared to vector.

Differences not statistically significant are labelled as (ns). (D) RT-PCR analysis of TRPA1 mRNA expression

in HepG2 and WI38 cells as a positive control. (E) [Ca2+]i induced by 100 µM allyl isothiocyanate (AITC) in

HepG2 cells for 10 min. Average time courses (left) and Δ [Ca2+]i (right) (n = 72-92). Data points are mean ±

SEM. P ≥ 0.05 compared to control. Differences not statistically significant are labelled as (ns). All data were

analyzed by Student's t-test.

APAP-Evoked Ca2+ Entry and ROS Production in HepG2 Cells are mediated by TRPV1,

TRPC1, TRPM2, and TRPM7

To characterize involvement of endogenous redox-sensitive TRPV1, TRPC1, TRPM2, and

TRPM7 in the APAP-induced Ca2+ responses in HepG2 cells, we employed blockers for these TRP

channels. These were capsazepine (CPZ), 2-aminoethyl diphenylborinate (2-APB), clotrimazole

(CTZ), and 2-(12-hydroxydodeca-5,10-diynyl)-3,5,6-trimethyl-p-benzoquinone (AA861),

previously shown to block recombinant TRPV1 (McIntyre et al., 2001), TRPC1 (Lievremont et al.,

2005), TRPM2 (Kheradpezhouh et al., 2014), and TRPM7 (Chen et al., 2010), respectively.

For [Ca2+]i measurements, HepG2 cells were pretreated with 10 µM CPZ, 100 µM 2-APB, 50

µM CTZ, or 10 µM AA861 for 6 min, then with APAP (20 mM) or H2O2 (1 mM). [Ca2+]i increases

induced by APAP in HepG2 cells were significantly lower if cells were also pretreated with CPZ, 2-

APB, CTZ, or AA861 (Figure 4A). Strikingly, CPZ and 2-APB exerted relatively strong, however

insignificantly inhibition among the blockers employed. Similar inhibition was observed against

[Ca2+]i increases induced by 1 mM H2O2 (Figure 4B). For intracellular ROS measurements,

treatment of HepG2 cells with TRP channel blockers alone did not affect ROS levels (Figure 4C).

Levels of intracellular ROS at 3 h after incubation of HepG2 with either APAP (Figure 4D) or H2O2

(Figure 4E) were also significantly suppressed by pretreatment with the four TRP channel blockers.

Thus, native TRPV1, TRPC1, TRPM2, and TRPM7 are likely to be involved in the APAP-induced

oxidative stress and Ca2+ overload we observed in HepG2 cells.

To confirm this interpretation, knockdown of individual channels was accomplished by treating

HepG2 cells with a small interfering RNA (siRNA) duplex targeting either TRPV1, TRPC1, TRPM2,

or TRPM7. Reduced mRNA (Figure 5A) and protein levels (Figure 5B, Figure 6A) of the

corresponding TRPV1, TRPC1, TRPM2, and TRPM7 channels were observed with specific siRNA,

as compared with control siRNA (siScramble). Importantly, HepG2 cells transfected with siTRPV1

or siTRPC1 showed a significant reduction in APAP- or H2O2-induced Ca2+ influx (Figure 5C,D,

respectively) and intracellular ROS levels (Figure 5E,F, respectively). This effect was greater than

that in HepG2 cells transfected with siTRPM2 or siTRPM7. No significant changes in ROS levels

were found in unstimulated HepG2 cells transfected with each of these siRNAs, as compared with

30

siScramble (Figure 6B). These results suggest that TRPV1 and TRPC1 were more involved in

APAP-induced oxidative stress and Ca2+ overload in HepG2 cells than were TRPM2 and TRPM7.

Figure 4. Inhibition of APAP- or H2O2 -induced Ca2+ entry and ROS production by selective TRP

channel blockers in HepG2 cells. (A) Effects of selective blockers capsazepine (CPZ; 10 µM) for TRPV1, 2-

APB (100 µM) for TRPC1, clotrimazole (CTZ; 50 µM) for TRPM2, or AA861 (10 µM) for TRPM7 on [Ca2+]i

rises evoked by APAP (20 mM). Average time courses (left) and Δ[Ca2+]i (right) (n = 18-57). (B) Effects of

10 µM CPZ, 100 µM 2-APB, 50 µM CTZ, and 10 µM AA861 on [Ca2+]i rises evoked by H2O2 (1 mM).

Average time courses (left) and Δ[Ca2+]i (right) (n = 19-43). (C) Inconsiderable changes in the ROS levels in

31

HepG2 cells treated with TRP channels blockers (CPZ, 2-APB, CTZ, and AA861; 1 µM). (D, E) Effects of

selective blockers of TRP channels on ROS production evoked by APAP (20 mM) (D) or H2O2 (1 mM) (E).

Blockers are applied for 3 h before and during APAP or H2O2 stimulation. Data points are mean ± SEM. P ≥

0.05 and ***P < 0.001 compared to DMSO. Differences not statistically significant are labelled as (ns). All

data of [Ca2+]i measurements were analyzed by Student's t-test, while those of ROS measurements were

analyzed by ANOVA and Bonferroni post-hoc.

Figure 5. Inhibition of APAP- or H2O2-induced Ca2+ responses and ROS production by siRNA-mediated

knockdown of either TRPV1, TRPC1, TRPM2, or TRPM7 channel in HepG2 cells. (A) Effects of specific

siRNA (siTRP) on channels of TRPV1, TRPC1, TRPM2, and TRPM7 mRNA levels detected by RT-PCR.

GAPDH is used as a positive control. (B) Effects of siTRPV1, siTRPC1, siTRPM2, and siTRPM7 on levels of

corresponding proteins detected by western blot. An anti-α-tubulin is used as a loading control. (C, D) [Ca2+]i

changes induced by 20 mM APAP (C) or 1 mM H2O2 (D) in cells treated with siTRPM2, siTRPM7, siTRPV1,

32

siTRPC1, or siScramble. Average time courses (left) and Δ[Ca2+]i (right) (n = 50-125). (E, F) Inhibitory effects

of siTRPM2, siTRPM7, siTRPV1, siTRPC1, or siScramble on ROS produced by 20 mM APAP (E) or 1 mM

H2O2 for 3 h (F). Data points are mean ± SEM. *P < 0.05 and ***P < 0.001 compared to siScramble. $P <

0.05, $$P < 0.01, and $$$P < 0.001 compared to siTRPV1 or siTRPC1. All data of [Ca2+]i measurements were

analyzed by Student's t-test, while those of ROS measurements were analyzed by ANOVA and Bonferroni

post-hoc.

Figure 6. (A) Quantification of intensity of bands of TRPV1, TRPC1, TRPM2, and TRPM7 (for 2-3

independent measurements for each channel), normalized by α-tubulin is represented by percentage (%). Data

points are mean ± SEM. P ≥ 0.05, **P<0.01, and ***P < 0.001 compared to control or siScramble. (B)

Inconsiderable changes in the ROS levels in HepG2 treated by siRNA-mediated knockdown of TRPV1, TR

PC1, TRPM2, and TRPM7. Data points are mean ± SEM. P ≥ 0.05 compared to control, DMSO, or siScramble.

Differences not statistically significant are labelled as (ns). All data were analyzed by ANOVA and Bonferroni

post-hoc.

Apoptosis Is Involved in APAP or H2O2-Induced HepG2 Cell Death

Acetaminophen has been reported to induce cell death by apoptosis (Boulares et al., 2002).

HepG2 cells were cultured in serum-free DMEM and treated with different doses of APAP (5, 15

and 20 mM) (Figure 7A) or H2O2 (0.25, 0.5 and 1 mM) (Figure 7B) for 24 h. A dose dependent

decrease in HepG2 cell viability was shown by trypan blue exclusion. Similar dose dependent

decreases in cell viability were also observed with 6 and 12 h treatments with APAP (Figure 7C) or

H2O2 (Figure 7D). HepG2 cell death, assessed by Hoechst33342/PI staining, indicated a significant

increase in the number of PI-positive dead cells (red) after incubation with 20 mM APAP or 1 mM

H2O2 for 24 h (Figure 7E).

Internucleosomal DNA fragmentation and caspase 3/7 activation represent essential steps in the

apoptotic cell death (Denault and Salvesen, 2002). HepG2 cells treated with APAP or H2O2 for 24 h

showed a dose dependent increase in DNA fragmentation compared with control cells (Figure 7F,G,

respectively). To examine whether HepG2 cell death elicited by APAP or H2O2 occurs via apoptosis,

we measured the caspase 3/7 activity. There was an increase in caspase 3/7 activity in HepG2 cells

proportional to the duration of treatment with either APAP (20 mM) or H2O2 (1 mM) (Figure 7H,

Figure 7I). These results suggest that APAP and H2O2 induce apoptotic death of HepG2 cells.

33

Figure 7. APAP-induced HepG2 cell death. (A, B) Dose dependence of viabilities of HepG2 cell treated

with APAP (A) or H2O2 (B) for 24 h. (C, D) Dose dependence of viabilities of HepG2 cell treated with APAP

(C) or H2O2 (D) for 6 h and 12 h.Cell viability is presented as percentage of viable cells in trypan blue exclusion

assay. (E) HepG2 cell death upon exposure to 20 mM APAP or 1 mM H2O2 for 24 h. Hoechst33342- and

propidium iodide (PI)-staining were visualized by fluorescence microscopy. Scale bar, 100 µm. (F, G) DNA

fragmentation in HepG2 cells. Cells were exposed for 24 h to APAP (F) (5, 15 and 20 mM) or H2O2 (G) (250

µM, 500 µM and 1 mM). M is the DNA marker. (H) Caspase 3/7 activity in HepG2 cells stimulated with 20

mM APAP or 1 mM H2O2 for 24 h. (I) Caspase 3/7 activity in HepG2 cells stimulated with 20 mM APAP

(left) or 1 mM H2O2 (right) for 4, 6, and 12 h. (G) Effects of either Ca2+-calmodulin antagonist (W-7; 1 µM),

cyclosporine A (CsA; 4 µM), or different MAPK-specific inhibitors (U0126, SP600125 and SB203580; 20

µM) on HepG2 cell death induced by APAP for 24 h in Hoechst33342- and PI-staining assays. Data points are

mean ± SEM. P ≥ 0.05, *P <0.05, **P < 0.01, and ***P < 0.001 compared to DMSO or control. Differences

not statistically significant are labelled as (ns). All data were analyzed by ANOVA and Bonferroni post-hoc.

We next assessed involvement of ERK, JNK or P38 MAPK pathways in APAP-induced HepG2

cell death. These pathways regulate the cell cycle, differentiation and growth as well as apoptosis

(Tidyman and Rauen, 2009) and the mitochondrial permeability transition (MPT) (Qian et al., 1997).

For these experiments, we employed the specific inhibitors W-7 for Ca2+-calmodulin, cyclosporine

34

A (CsA) for MPT, U0126 for ERK, SP600125 for JNK, and SB203580 for P38 MAPK. Compared

with untreated cells, there was no significant cell death in HepG2 cells treated with 1 µM W-7, 4 µM

CsA, or 20 µM of either U0126, SP600125 or SB203580 (Figure 8A). HepG2 cells treated with

some of these inhibitors were partially protected from APAP compared to the cells that were treated

only with APAP. However, at 20 µM, U0126 or SB203580 did not significantly decrease APAP-

induced cell death (Figure 8B). This finding suggests that APAP induced apoptosis in HepG2 cells

was via activation of JNK and MPT, while the p38 and the ERK MAPK pathways did not contribute.

Figure 8. APAP-induced HepG2 cell death. (A) Representative images of HepG2 treated with Ca2+-

calmodulin antagonist (W-7; 1 µM), cyclosporine A (CsA; 4 µM) or different MAPK-specific inhibitors

(U0126, SP600125 and SB203580; 20 µM) for 24 h. Hoechst33342- and PI-staining were visualized by

fluorescence microscopy. Scale bar, 100 µm. (B) Representative images of HepG2 treated with 1 µM W-7, 4

µM CsA, or different MAPK-specific inhibitors (U0126, SP600125 and SB203580; 20 µM) on HepG2 cell

death induced by APAP for 24 h in Hoechst33342- and PI-staining assays. Hoechst33342- and PI-staining

were visualized by fluorescence microscopy. Scale bar, 100 µm. Data points are mean ± SEM. P ≥ 0.05, *P

<0.05 compared to DMSO. Differences not statistically significant are labelled as (ns). All data were analyzed

by ANOVA and Bonferroni post-hoc.

To investigate the correlations among ROS levels, [Ca2+]i elevation and cell death, we examined

effects of ROS scavengers on HepG2 cell viability in the presence of APAP or H2O2. Preincubation

of HepG2 cells with of either NAC or tiron (1 mM) for 3 h prior to treatment with APAP or H2O2

significantly improved cell viability (Figure 9A) and reduced the number of PI-positive cells (red)

(Figure 9B and Figure 10A). The ROS scavengers alone showed no significant effects on cell

viability or PI-staining in HepG2 cells (Figure 11A,B).

35

Strikingly, pretreatment of HepG2 cells with TRP channel blockers (1 µM of either CPZ, 2-APB,

CTZ, or AA861) for 3 h prior to treatment with APAP or H2O2 significantly improved cell viability

(Figure 9C) and reduced the numbers of PI-positive cells (red) (Figure 9D and Figure 10B). Similar

results were observed after siRNA-mediated knockdown of TRPV1, TRPC1, TRPM2, and TRPM7

(Figure 9E,F and Figure 10C). Being consistent with the contribution of TRPV1 and TRPC1 to

APAP-induced Ca2+ entry, the siRNAs for TRPV1 and TRPC1 worked more efficiently than those

of the others, a finding consistent with measurements of Ca2+ responses (Figure 5C,D) and ROS

levels (Figure 5E,F). The channel blockers or siRNAs alone did not significantly affect cell viability

and number of PI-positive cell compared to untreated HepG2 cells (Figure 11C-F).

Figure 9. TRPV1, TRPC1, TRPM2, and TRPM7 confer susceptibility to APAP-induced cell death via

ROS in HepG2 cells. (A and B) ROS scavengers, NAC and tiron (1 mM), suppressed APAP- or H2O2-induced

losses of cell viability (A) and increases of cell death (B) in HepG2 cells. (C, D) Selective TRP channels

36

blockers, 1 µM of either CPZ, 2-APB, CTZ, or AA861 suppressed APAP- or H2O2-induced losses of cell

viability (C) and increases of cell death (D) in HepG2 cells. (E, F) siRNA-mediated knockdown of TRPV1,

TRPC1, TRPM2, and TRPM7 suppressed APAP- or H2O2-induced losses of cell viability (E) and increases of

cell death (F) in HepG2 cells. Inhibitors are applied for 3 h before and during APAP or H2O2 stimulation. Data

points are mean ± SEM. *P <0.05, **P < 0.01, and ***P < 0.001 compared to the DMSO, siScramble or

control. All data were analyzed by ANOVA and Bonferroni post-hoc.

37

Figure 10. Hoechst33342- and PI-

assay of HepG2 cell death. (A) Representative images of HepG2 that

were treated with ROS scavengers,

NAC or tiron (1 mM), on 20 mM

APAP- or 1 mM H2O2-induced cell

death for 24 h. Hoechst33342- and PI-

staining were visualized by

fluorescence microscopy. Scale bar,

100 µm. (B) Representative images of

HepG2 treated with TRP channel

blockers (CPZ, 2-APB, CTZ, and

AA861; 1 µM) on 20 mM APAP- or 1

mM H2O2-induced cell death for 24 h.

Hoechst33342- and PI-staining were

visualized by fluorescence

microscopy. Scale bar, 100 µm. (C)

Representative images of HepG2

treated with siRNA for knockdown of

TRPV1, TRPC1, TRPM2, and

TRPM7 on 20 mM APAP- or 1 mM

H2O2-induced cell death for 24 h.

Hoechst33342- and PI-staining were

visualized by fluorescence

microscopy. Scale bar, 100 µm.

38

Figure 11. (A,B) Percentages of viable cells (A) and percentages of PI-positive cells (B) in HepG2 treated

with NAC or tiron (1mM), for 24 h. (C,D) Percentages of viable cells (C) and percentages of PI-positive cells

(D) in HepG2 cells treated with TRP channels blockers (CPZ, 2-APB, CTZ, and AA861; 1 µM). (E and F)

Percentages of viable cells (E) and percentages of PI-positive cells (F) in HepG2 treated with siRNA for

knockdown of TRPV1, TRPC1, TRPM2, and TRPM7. (G,H) Caspase 3/7 activity (G) and the level of

intracellular cytochrome c (H) in HepG2 cell treated with TRP blockers (CPZ, 2-APB, CTZ, and AA861; 1

µM). (I,J) Caspase 3/7 activity (I) and the level of intracellular cytochrome c (J) in HepG2 cell treated with

siRNA for knockdown of TRPV1, TRPC1, TRPM2, and TRPM7. Data points are mean ± SEM. P ≥ 0.05

compared to DMSO or siScramble. Differences not statistically significant are labelled as (ns). All data were

analyzed by ANOVA and Bonferroni post-hoc.

Increases in caspase 3/7 activity and intracellular cytochrome c release, induced by APAP or

H2O2, were also ameliorated by pretreatment with ROS scavengers (1 mM of either NAC or tiron;

Figure 12A,B), selective TRP channel blockers (1 µM of either CPZ, 2-APB, CTZ, or AA861;

Figure 12C,D) as well as by siRNA-mediated knockdown of TRPV1, TRPC1, TRPM2, or TRPM7

(Figure 12E,F). These results suggest that APAP-induced HepG2 cell apoptosis was more dependent

39

on TRPV1 and TRPC1 than on TRPM2 and TRPM7, when assessed by Ca2+ entry, ROS levels, and

mitochondrial membrane depolarization.

Figure 12. Effects of APAP on the activity of caspase 3/7 and the level of intracellular cytochrome c in

HepG2 cell. (A and B) ROS scavengers, NAC or tiron, (1 mM) suppressed APAP- or H2O2-induced increases

of the activity of caspase 3/7 (A) and the level of intracellular cytochrome c (B) in HepG2 cells. (C, D) Selective

Ca2+ channels blockers, 1 µM of either CPZ, 2-APB, CTZ, and AA861 suppressed APAP- or H2O2-induced

increases of the activity of caspase 3/7 (C) and the level of intracellular cytochrome c (D) of HepG2 cells. (E,

F) siRNA-mediated knockdown of TRPV1, TRPC1, TRPM2, and TRPM7 suppressed APAP- or H2O2-

induced increases of the activity of caspase 3/7 (E) and the level of intracellular cytochrome c (F) in HepG2

cells. Data points are mean ± SEM. P ≥ 0.05, *P < 0.05, **P < 0.01, and ***P < 0.001 compared to the DMSO,

siScramble or control. Differences not statistically significant are labelled as (ns). All data were analyzed by

ANOVA and Bonferroni post-hoc.

40

Involvement of Cysteine Residues in Redox and APAP Sensing of TRPV1 and TRPC1

An intriguing question is the mechanism that explains the relative importance of TRPV1 and

TRPC1 compared with other TRPs. It has been reported that free thiol groups of cysteine residues

are among the major targets of oxidation and electrophiles in proteins. This is also the case for TRP

channels; oxidants and electrophiles induce their activation (Yoshida et al., 2006; Takahashi et al.,

2008). To examine involvements of cysteine residues of TRPV1 and TRPC1 in their APAP

responsiveness, we incubated permeabilized HEK293T cells transfected with GFP-tagged TRPV1

or Flag-tagged TRPC1 with DTNB-2Bio, which reacts with free thiol groups and can be purified by

avidin-based affinity purification (Yoshida et al., 2006; Takahashi et al., 2008). DTNB-2Bio was

incorporated into TRPV1 (Figure 13B) and TRPC1 (Figure 13C), and this was blocked by

pretreatment with 20 mM APAP. In contrast, no blockade by APAP was detected for DTNB-2Bio

incorporation into TRPV4, which had shown only a minor contribution to APAP-induced responses,

as compared with TRPV1 and TRPC1 (Figure 13D). Thus, the action of APAP via the modification

target cysteine residues may at least in part account for the important roles of TRPV1 or TRPC1 in

APAP-induced responses in human hepatoma cells.

GSH Depletion Modulates Sensitivity of Ca2+ Influx and Cell Death to APAP in HepG2 Cells

In addition to those of ROS, glutathione (GSH) levels serve as pivotal regulators of cellular redox

status. Previous studies showed that intracellular GSH suppressed oxidative stress-induced TRPM2

activation and Ca2+ entry in DRG neurons (Nazıroğlu et al., 2011). It is, therefore, important to study

the role of GSH in suppressing APAP-induced ROS increases, Ca2+ overload via TRP channels, and

cell death in HepG2 cells. Moreover, serum deprivation induces excess ROS production linked to

depletion of intracellular GSH (Pandey et al., 2003). We found that serum deprivation enhanced the

Ca2+ responses and cell death induced by 20 mM APAP or 1 mM H2O2 (Figure 14A,B) in HepG2,

as compared with that in normally cultured cells. However, serum deprivation did not affect caspase

3/7 activity (Figure 14C), though the GSH content of serum-deprived HepG2 was significantly

lower than that of cells cultured normally (Figure 14D). Pretreatment of serum-deprived HepG2

with ROS scavengers (NAC or tiron at 1 mM) for 3 h significantly reversed APAP- or H2O2-induced

depletion of GSH content (Figure 14E).

Dimethylfumarate (DMF) was previously reported to upregulate antioxidant activities and

suppress cell death induced by H2O2 (Duffy et al., 1998). Treatment of serum-deprived HepG2 with

50, 100 or 200 µM DMF for 24 h significantly increased GSH content in a dose-dependent manner

(Figure 15A). There were no significant changes in cell viability, except for a significant decrease

with 200 µM DMF (Figure 15B). Intracellular ROS levels 3 h after incubation of HepG2 with either

41

Figure 13. The effects of APAP on the covalent incorporation of DTNB-2Bio in TRPV1, TRPC1, and

TRPV4 proteins. (A) Chemical structure of DTNB-2Bio. (B-D) Western blot using an anti-GFP or anti-Flag

42

antibody of avidin-bound fractions (top panel) and total cell lysates (bottom panel) prepared from HEK293T

cells which were transfected with EGFP-TRPV1 (B), TRPC1-Flag (C), EGFP-TRPV4 (D), and then incubated

with DTNB-2Bio in the presence or absence of 20 mM APAP.

Figure 14. Serum deprivation enhanced Ca2+ responses and death induced by APAP in HepG2 cells. (A) [Ca2+]i rises induced by APAP (20 mM) (left) or H2O2 (1 mM) (right) in HepG2 cells cultured in normal

condition and serum-deprived HepG2 cells for 10 min. Average time courses (left) and Δ[Ca2+]i (right) (n =

42-98). (B) Cell death in HepG2 cells cultured in normal condition and serum-deprived HepG2 cells upon

exposure to APAP (20 mM) or H2O2 (1 mM) for 24 h. Hoechst33342- and PI-staining were visualized by

fluorescence microscopy. Scale bar, 100 µm. (C,D) Caspase 3/7 activity (C) and GSH content (D) in HepG2

cells cultured in normal condition and serum-deprived HepG2 cells for 24 h. (E) ROS scavengers, NAC and

tiron, (1 mM) suppressed APAP- or H2O2-induced depletion of GSH content in HepG2 cells. Data points are

mean ± SEM. P ≥ 0.05, *P < 0.05, ***P < 0.001. Differences not statistically significant are labelled as (ns).

All data of [Ca2+]i measurements were analyzed by Student's t-test while other data were analyzed by ANOVA

and Bonferroni post-hoc.

43

20 mM APAP or 1 mM H2O2 were also significantly reduced by pretreatment with 100 µM DMF

(Figure 15C). Furthermore, GSH content in HepG2 cells after 24 h incubation with either APAP or

H2O2 was also significantly higher in cells that had been pretreated with 100 µM DMF (Figure 15D).

Preincubation of serum-deprived HepG2 cells with 100 µM DMF for 24 h, followed by

pretreatment with 100 µM DMF for 6 min, attenuated the [Ca2+]i increases induced by 20 mM APAP

or 1 mM H2O2 (Figure 15E). In addition, DMF attenuated the loss of cell viability (Figure 15F),

increase in caspase 3/7 activity (Figure 15G), intracellular cytochrome c levels (Figure 15H), and

cell death (Figure 15I) induced by APAP or H2O2. No significant cytotoxicity was observed when

DMF was administrated in combination with APAP as shown in Figure 15. These results support a

role for GSH depletion, reversible with GSH inducer DMF, in APAP-induced oxidative stress, Ca2+

influx, and HepG2 cell death.

Expression and Cellular Localization of Native Redox-Sensitive TRP Channels in the Human

Liver

To confirm the validity of using HepG2 cells as a model system for hepatotoxicity, we conducted

histological experiments to obtain definitive evidence that redox sensitive TRP channels are

expressed in the human liver. Expression profiles of TRPV1-4, TRPC1, TRPC4, TRPC5, TRPM2,

TRPM7, and TRPA1 were analyzed using PCR in cDNA of normal human liver. TRPV1, TRPC1,

TRPM2, TRPM7, and TRPA1 were expressed (Figure 16A).

In situ hybridization of normal human liver sections with antisense probes yielded strong staining

for TRPV1, TRPC1, TRPM2, and TRPM7, localized in the nuclei of hepatocytes and Kupffer cells

(Figure 16B). The sinusoidal endothelial lining and Kupffer cells were also positive for TRPA1

expression but at a lower level, while hepatocytes were negative for this channel (Figure 16B). No

hybridization signal was observed with sense probes applied as negative controls (Figure 16B).

Recombinant TRPV1, TRPC1, TRPM2, TRPM7, and TRPA1 transcripts were intensely labeled

mainly in the nucleus of HEK293T cells (Figure 17A). RNA transcripts of these TRPs, except for

TRPA1, were expressed in HepG2 cells (Figure 17A).

44

Figure 15. GSH inducer DMF attenuate APAP- or H2O2-induced Ca2+ influx and ROS-mediated death

in HepG2 cells. (A) The effects of 50, 100 and 200 µM DMF on GSH content in HepG2 cells. (B) The effects

of 50, 100 and 200 µM DMF on cell viabilities in HepG2 cells. (C, D) DMF attenuated APAP- or H2O2-

induced increases of ROS levels (C) and GSH depletion (D). (E) Averaged time courses and Δ [Ca2+]i of

APAP-(left) or H2O2 -induced Ca2+ response (right) in the presence and absence of 100 µM DMF. (F-I) Effects

of DMF on APAP- or H2O2-induced losses of cell viability (F) and increases of the activity of caspase 3/7 (G),

the level of intracellular cytochrome c (H), and the percentages of PI-positive cells (I) in HepG2 cells. P ≥

0.05, *P < 0.05, **P < 0.01, and ***P < 0.001 compared to DMSO or control. Differences not statistically

significant are labelled as (ns). Data points are mean ± SEM. All data of [Ca2+]i measurements were analyzed

by Student's t-test, while other data were analyzed by ANOVA and Bonferroni post-hoc.

45

Figure 16. Expression and localization of redox-sensitive TRP channel in the human liver tissue. (A) Expression of mRNAs for TRPV1-4, TRPC1, TRPC4, TRPC5, TRPM2, TRPM7, TRPA1, and GAPDH

46

detected by PCR in the cDNA library from normal human liver. Specific PCR primers used are listed in the

Table 1. (B) In situ hybridization analysis of TRPV1, TRPC1, TRPM2, TRPM7, and TRPA1 mRNAs in the

paraffin section of the human liver. mRNAs of TRPV1 (panel a, f), TRPC1 (panel b, g), TRPM2 (panel c, h),

TRPM7 (panel d, i) were localized in hepatocytes (*) and Kupffer cells (arrow). TRPA1 (panel e, j) was

localized in the lining of sinusoids and Kupffer cells of the human liver tissue. The images from the analysis

of both the antisense (left) and sense probes (right) are shown. Scale bar, 100 µm.

Figure 17. In situ hybridization analysis of redox-sensitive TRP channels expressed in HEK 293T and

HepG2 cells. Expression of mRNAs encoding TRPV1 (panel A), TRPC1 (panel B), TRPM2 (panel C),

TRPM7 (panel D), and TRPA1 (panel E) are observed except for TRPA1 in HepG2 cells. The images from

the analysis of both antisense (left) and sense probes (right) are shown. Scale bar, 100 µm.

47

Figure 18. Schematic representation of the proposed signaling mechanism underlying cell death

regulated by Ca2+ entry via redox-sensitive TRPV1, TRPC1, TRPM2, and TRPM7 channels activated

by APAP overdose in HepG2. TRPV1 and TRPC1 channels are activated by oxidative modification of free

sulfhydryl groups of cysteine residues, which APAP is able to reach to enhance activation. Initial Ca2+ influx

by APAP-induced ROS induces the mitochondrial and nuclear damages mediating the release of ADPR, which

is released into the cytosol to activate TRPM2. ROS activates TRPM7. Suppression of TRPV1, TRPC1,

TRPM2, and TRPM7 using blockers, siRNAs and ROS scavengers (NAC or tiron) alleviates APAP-induced

Ca2+ entry and HepG2 death. APAP, N-acetyl-para-aminophenol; [Ca2+]i, calcium ions; HepG2, human

hepatoma cell line; TRP, transient receptor potential channels; TRPV1, type 1 vanilloid receptor; TRPC1, type

1 canonical receptor; TRPM2, type 2 melastatin receptor; TRPM7, type 7 melastatin receptor; CaMs,

calmodulins; CPZ, capsazepine (TRPV1 antagonist); 2-APB, 2-aminoethyl diphenylborinate (TRPC1

antagonist); CTZ, clotrimazole (TRPM2 antagonist); AA861, 2-(12-hydroxydodeca-5,10-diynyl)-3,5,6-

trimethyl-p-benzoquinone (TRPM7 antagonist); CsA, cyclosporine A (inhibitor of mitochondrial

permeability transition, MPT); H2O2, hydrogen peroxide; O2-, superoxide anion; ROS, reactive oxygen species;

GSH, glutathione; ADPR, adenosine diphosphate ribose; JNK, c-jun NH2-terminal kinase; NAC, N-acetyl-L-

cysteine.

48

DISCUSSION

In this study, we found that APAP overdose elicited ROS production, [Ca2+]i increases, and GSH

depletion triggering HepG2 cell death. All responses were markedly reduced by pretreatment with

the ROS scavengers, NAC or tiron, or with the GSH inducer DMF. In HepG2 cells, expression of

redox-sensitive TRP channels including TRPV1, TRPC1, TRPM2, and TRPM7 was observed.

Suppression of TRP channels, particularly, TRPV1 and TRPC1, with channel blockers and siRNAs

in terms of function and expression, respectively, substantially reduced APAP-induced ROS

formation, Ca2+ influx, and cell death, as summarized in Figure 18. Additionally, in situ

hybridization analysis of normal human liver sections showed that transcripts for TRPV1, TRPC1,

TRPM2, and TRPM7 channels were primarily localized to hepatocytes and Kupffer cells, while

mRNA for TRPA1 was visible, to a lesser extent, in Kupffer cells and sinusoidal endothelium but

was absent in hepatocytes.

Earlier reports described the presence of TRPV1-4, TRPC1, and TRPM7 mRNAs but not TRPC4

and TRPC5 in HepG2 cells (Vriens et al., 2004; El Boustany et al., 2008). Other studies showed

expression of the TRPM7 channel in rat HSC-T6 hepatic stellate, RLC-18 and WIF-B hepatoma cells

(Liu et al., 2012; Lam et al., 2012) and of the TRPC1 channel in rat H4-IIE hepatoma cells (Chen

and Barritt, 2003). In addition, our study newly reveals, by RT-PCR and in situ hybridization, the

existence of TRPM2 mRNA in HepG2 cells. Furthermore, this is the first in situ hybridization

analysis to show localization and distribution of TRPV1, TRPC1, TRPM2, and TRPM7 transcripts

in the hepatocyte of human liver. Regarding functional expression, HEK293 cells heterologously

expressing TRPV1, TRPC1, TRPM2, or TRPM7 responded well to H2O2, in agreement with previous

reports (Hara et al., 2002; Aarts et al., 2003; Yoshida et al., 2006; Salazar et al., 2008), and as well

to APAP as indicated by increased Ca2+ entry. In contrast, HEK293 cells expressing TRPV2, TRPV3,

TRPV4, TRPC4, or TRPC5 failed to respond to APAP. Such differences are presumably due to

structural variations among various TRP channel subfamily members (Figure 3). Interestingly, it has

been reported that TRPV subunits can assemble into hetero-oligomeric channel complexes, for

example, between TRPV1 and TRPV2 (Hellwig et al., 2005) or TRPV1 and TRPV3 (Smith et al.,

2002). Therefore, we cannot exclude the possibility that TRPV1 could operate as hetero-multimeric

channel as well as homo-multimeric channel in response to APAP or H2O2.

ROS overproduction and the resulting cell damaging events are often accompanied by rises in

[Ca2+]i (Yan et al., 2006). A previous report proposed that H2O2 alters cellular redox status through

increased [Ca2+]i, decreases GSH level and induces lipid peroxidation, eventually causing cell death

in the human hepatoma cell line SMMC-7721 (Li et al., 2000). Other reports suggested that APAP

exposure leads to increased levels of superoxide anions, which upon dismutation generate H2O2 (Dai

49

and Cederbaum, 1995). To clarify whether such mechanisms were involved in our experiments, we

used NAC, a sulfhydryl drug, which contributes to repletion of GSH by diminishing ROS production

via removal of superoxide radicals, H2O2, and hydroxyl radicals (Sen and Packer, 2000). NAC is the

most widely used antidote for APAP-induced hepatotoxicity (Smilkstein et al., 1991). We also

employed tiron, an intracellularly active scavenger of superoxide anions and hydroxyl radicals

(Krishna et al., 1992; Taiwo, 2008). There had been no previous investigations addressing whether

NAC or tiron actually suppress ROS and [Ca2+]i in HepG2 cells. Importantly, we found that

pretreatments with either NAC or tiron strongly attenuated APAP-induced elevations of ROS and

[Ca2+]i in HepG2 cells. This supports the hypothesized link between APAP overdose and ROS

formation in HepG2 cells. Interestingly, it was reported that NAC reduced oxidative stress and

suppressed TRPV1 channel-mediated Ca2+ entry in neutrophils from patients with polycystic ovary

syndrome (Köse and Nazıroğlu, 2015). NAC may also play a protective role against Ca2+ influx

through regulation of TRPM2 channels in neurons (Özgül and Nazıroğlu, 2012). It was further

suggested that tiron reduced Ca2+ entry through TRPC1 channels in mouse muscle fibers (Gervásio

et al., 2008). Taken together, NAC or tiron might work through suppression of APAP-elicited

oxidative stress and ensuing activation of redox-sensitive TRP channels whose molecular expression

was detected in HepG2 cells.

Consistent with our results, it was reported that TRPV1, TRPC1, TRPM2, and TRPM7 channels

were modulated by ROS and induced deleterious responses such as cell death (Miller, 2006; Kozai

et al., 2014). For example, TRPV1 activation has previously been shown to increase [Ca2+]i, oxidative

stress and apoptotic cell death (Shin et al., 2003; Kim et al., 2006). TRPV1 and TRPC1 are activated

by oxidative modification of free sulfhydryl groups of cysteine residues (Yoshida et al., 2006). We

reported that TRPM2 induced cell death during conditions of oxidative stress (Hara et al., 2002).

Suppression of TRPM7 expression by RNA interference blocked TRPM7 currents, anoxic Ca2+

influx, and ROS production, protecting cortical neurons from anoxia. This suggested a role for

endogenous TRPM7 in anoxic neuronal death (Aarts et al., 2003). Thus, redox-sensitive TRP

channels are widely involved in oxidative stress linked to cellular disorders.

Ca2+ dysregulation was reported to be involved in APAP-induced hepatocellular damage (Shen

et al., 1991). It was suggested that [Ca2+]i increases are primarily caused by inhibition of Ca2+-Mg2+

ATPase and accompany, but do not cause, hepatocellular damage (Tsokos-Kuhn et al., 1988). In our

study, pretreatment with antagonists of TRPV1, TRPC1, TRPM2, and TRPM7 channels (CPZ, 2-

APB, CTZ, and AA861, respectively) protected the HepG2 cells from deleterious increases in Ca2+

entry and ROS production and the resulting cell death induced by APAP. Importantly, knockdown

of these channels by a siRNA strategy resulted in significantly suppressed Ca2+ overload and ROS

50

production in response to either APAP or H2O2. CTZ was previously reported to protect the liver

against normothermic ischemia-reperfusion injury in rats (Iannelli et al., 2009). Likewise, CPZ

treatment conferred significant protection against liver dysfunction, as indicated by serum alanine

transaminase (ALT) and aspartate transaminase (AST) activities, in sepsis (Ang et al., 2011).

Moreover, 2-APB reduced APAP-induced liver injury in mouse primary hepatocytes in vivo (Du et

al., 2013). In a recent study, AA861 pretreatment significantly attenuated acute liver failure in rats

by inhibiting macrophage activation (Li et al., 2014). Hence, abundant evidence indicates that

activation of these TRP channels contributes to APAP-induced hepatotoxicity, raising a possibility

that their blockers can ameliorate the APAP-induced damages of human hepatocytes.

Our data demonstrate that TRPC1 and TRPV1 play prominent roles in regulation of the APAP-

induced responses among TRP channels expressed in HepG2 cells. Previously, we have reported that

TRPV1 and TRPC1 are directly activated through oxidative modifications of cysteine residues by

ROS, RNS, and electrophiles (Yoshida et al. 2006; Takahashi et al., 2011; Kozai et al., 2014). In

addition, previous reports have shown that endogenous expression of cytochrome P450 enzymes

(CYPs), which convert APAP to its oxidized metabolite N-acetyl-p-benzoquinoneimine NAPQI,

were detected in HepG2 cells (Sumida et al., 2000; Maruyama et al., 2007; El Gendy and El-Kadi,

2009; Hart et al., 2010, Wang et al., 2011). It is therefore possible that TRPV1 and TRPC1 are

activated by NAPQI together with ROS/RNS generated upon APAP treatment to evoke pronounced

downstream responses, compared with TRPM2 and TRPM7, which is most likely to be only

activated by indirect action of ROS/RNS generated upon APAP treatment (Hara et al., 2002; Aarts

et al., 2003). Interestingly, TRPA1, which is the most sensitive to oxidants and electrophiles among

TRP channels (Takahashi et al., 2011), has been reported to respond to NAPQI to mediate spinal

nociception induced by APAP in sensory neurons (Andersson et al., 2011). Alternatively, APAP

itself may act directly but non-covalently to the DTNB-2Bio target cysteine residue(s), and activates

TRPV1 and TRPC1. This possibility is likely when we consider our finding that APAP efficiently

activates TRPV1 and TRPC1 expressed recombinant in HEK cells, in which expression of CYPs has

not been clarified yet. At this point, we cannot conclude whether the action of APAP is through

covalent binding of NAPQI or non-covalent binding of APAP itself, because both possibilities are

consistent with our finding that treatment of permeabilized cells with APAP suppresses incorporation

of DTNB-2Bio into TRPV1 and TRPC1 proteins (Figure 13). Thus, cysteine residues susceptible to

oxidative modifications are critical for exacerbation of APAP-mediated hepatotoxicity in human,

suggesting that TRP channels such as TRPC1 and TRPV1 susceptible to this type of modification

are potential targets for the treatment of APAP-induced liver toxicity.

51

In our study, decrease in HepG2 cell viability as well as increases in DNA fragmentation,

intracellular cytochrome c level, and caspase 3/7 activity were observed after either APAP overdose

or H2O2 treatment. These responses were significantly reversed by ROS scavengers or TRP channel

blockers. APAP overdose was previously reported to cause DNA fragmentation in primary cultured

mouse hepatocytes (Shen et al., 1991) and mitochondrial cytochrome c release in isolated mouse

liver cells (El-Hassan et al., 2003). Kim and colleagues suggested that oxidative stress and increased

Ca2+ levels are both stimuli of mitochondrial dysfunction in rat hepatocytes (Kim et al., 2003).

Downstream signaling of Ca2+ influx and Ca2+ overload in mitochondria can cause opening of the

MPT pore (Zoratti and Szabo, 1995), and eventually promote apoptosis (Liu et al., 1996; Polster and

Fiskum, 2004). The time required for this signaling cascade can possibly account for the time elapsed

between Ca2+ concentration changes that took place within minutes and the observation of maximal

HepG2 cell death at 24 h or more (Zhivotovsky and Orrenius, 2011). Collectively, these results

suggest that mitochondrial Ca2+ overload might have contributed to APAP-induced HepG2 cell death

in our study. Some reports claim that caspase activation and apoptosis were suggested to only occur

in HepG2 but not in primary hepatocytes (Shen et al., 1991). However, activation of caspases in

APAP-induced liver injury has been reported previously (Hu and Colletti, 2010; Sharma et al., 2011;

Li et al., 2013), and such activation of caspases due to APAP treatment was significantly reduced in

cells treated with anti-apoptotic caspases inhibitors (Ferret et al., 2001), which altogether would

support an apoptotic pathway for APAP toxicity in hepatocytes.

We demonstrated that serum-deprived HepG2 cells are more susceptible than normally cultured

cells to APAP- or H2O2-induced cell damage, depletion in GSH stores, increased Ca2+ influx, and

loss of cell viability. Serum, a mixture of hundreds of proteins, contains various factors needed for

proliferation of cells in culture (van der Valk et al., 2004). Serum deprivation may induce oxidative

stress, caused by excess production of ROS or decreased GSH, and this can be inhibited by

intracellular antioxidants (Pandey et al., 2003; Zhuge and Cederbaum, 2006).

Intracellular GSH is important for detoxification of a variety of chemicals, acting as a reductant

in the metabolism of peroxides and free radicals (Circu and Aw, 2012). In addition, GSH is a crucial

endogenous agent to restore the normal cellular redox state when it is compromised. Previous studies

suggested that depletion of GSH by H2O2 is important for apoptosis and that H2O2-mediated lipid

peroxidation might be a primary event leading to other biochemical changes (Li et al., 2000). It is

possible that a decline in GSH content during APAP treatment similarly altered the cellular redox

state in HepG2 cells. Our results showed that, in HepG2 cells, the GSH inducer DMF attenuated

APAP- or H2O2-induced decrease in GSH content, increased ROS levels, Ca2+ overload, and cell

death associated events such as intracellular cytochrome c release and caspase 3/7 activation. DMF

52

has been used successfully to treat multiple sclerosis (Fox et al., 2012), and psoriasis (Bovenschen

et al., 2010), to attenuate renal fibrosis (Oh et al., 2012), and for cardioprotection (Ashrafian et al.,

2012). It is believed to exert such benefits by restoring the balance between pro-oxidant and

antioxidant systems during oxidative stress. DMF may therefore represent an alternative therapy to

NAC, the current treatment for APAP hepatotoxicity.

In conclusion, this is the first study to systematically and comparatively analyze the contributions

of redox-sensitive TRP channels such as TRPV1, TRPC1, TRPM2, and TRPM7 to human hepatoma

cell death induced by APAP overdose. Our findings add considerably to current understanding of the

mechanism of APAP-induced liver toxicity. Further comparative studies using various TRP

knockout mice are important to establish individual roles in vivo of these TRP channels. Once our

observation is confirmed in other models in vitro and in vivo, these TRP channels could represent

potential drug targets for treating APAP overdose over a wide range of post-ingestion time.

ABBREVIATIONS: 2-APB, 2-aminoethyl diphenylborinate; AA861, 2-(12-hydroxydodeca-

5,10-diynyl)-3,5,6-trimethyl-p-benzoquinone; AITC, allyl isothiocyanate; APAP, N-acetyl-para-

aminophenol; bp, base pair; [Ca2+]i, intracellular Ca2+ concentration; CPZ, capsazepine; CsA,

cyclosporine A; CTZ, clotrimazole; DCF-DA, 2,7-dichlorofluorescein diacetate; DMF,

dimethylfumarate; H2O2, hydrogen peroxide; HepG2, human hepatoma cell line; MAPK, mitogen

activated protein kinase; MPT, mitochondrial permeability transition; NAC, N-acetyl-L-cysteine; PI,

propidium iodide; RT-PCR, reverse transcriptase polymerase chain reaction; siRNA, small

interfering RNA; tiron, 4,5-dihydroxy-1,3-benzene disulfonic acid disodium salt monohydrate; TRP,

transient receptor potential; W-7, N-(6-aminohexyl)-5-chloro-1-naphthalene sulfonamide; WI-38,

human lung fibroblast cell line.

53

ACKNOWLEDGEMENTS

Human hepatoma cell line (HepG2) (RCB1886) and human lung fibroblast cell line (WI-38)

(RCB0702) were provided by RIKEN Bio Resource Center with the support of the National Bio

Resource Project of the MEXT, Tsukuba, Japan. The author thank Dr Sawamura for experimental

advice.

54

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Chapter 2

TRPM2 regulates the development of rheumatoid arthritis in the anti-

collagen-induced mouse model

ABSTRACT

Rheumatoid arthritis (RA) is an autoimmune inflammatory disease of joints. In developing a

novel therapeutic approach, it is critical to understand the molecular mechanisms underlying

RA symptoms. Because, RA has recently been characterized by the augmentation of reactive

oxygen species (ROS), we investigate the role of TRPM2, a ROS-sensitive transient receptor

potential (TRP) cation channels, in mouse anti-type II collagen antibody-induced arthritis

(CAIA). Compared with wild-type mice with CAIA, paw swelling is prominently suppressed

in Trpm2-deficient mice, despite the binding of the collagen antibody to cartilage of which type

II collagen is a major component. Furthermore, the massive infiltration of immune cells into

the joint cavity, the damage of bones, and the production of inflammatory cytokines are

dramatically attenuated in Trpm2 deficient mice. These results suggest that TRPM2 plays an

essential role in the development of CAIA, and we propose that inhibition of TRPM2 activity

is a novel therapeutic strategy for treating RA.

65

INTRODUCTION

Rheumatoid arthritis (RA) is an autoimmune disease that is characterized by autoantibodies,

severe inflammation of the joints, and marked damage of bones (Lee and Weinblatt, 2001; Firestein,

2003; Scott et al., 2010). To reduce pain and progressive joint damage, different treatments have been

utilized, including non-steroidal anti-inflammatory drugs (NSAIDs), disease-modifying anti-

rheumatic drugs (DMARDs) such as methotrexate, and biological agents which are inhibitors of

tumor necrosis factor alpha (TNF-α), an inflammatory cytokine involved in the pathogenesis of RA

(Scott et al., 2010). However, these drugs also have serious negative impacts such as infections. Thus,

to develop novel therapeutics, it is critical to elucidate molecular mechanisms responsible for the

development of RA.

Well-validated autoantibodies in RA patients includes the antibody that targets various regions

of type II collagen, which is a major constituent of articular cartilage matrix proteins (Rowley et al.,

2008). Anti-type II collagen antibody-induced arthritis (CAIA) has been established as a simple

mouse model of RA, in which arthritis can be induced by the administration of a cocktail of

monoclonal anti-type II collagen antibodies, together with endotoxin lipopolysaccharide (LPS)

(Nandakumar et al., 2003; Khachigian, 2006). In CAIA, LPS administration induces the production

of some vasoactive mediators, leading to the collagen antibody gaining access to the joint, and the

subsequent activation of innate immune processes and inflammation [Wipke et al., 2004]. It has been

reported that complement such as C5a, signaling proteins such as Fcγ receptors for immunoglobulin

G (IgG) and spleen tyrosine kinase (Syk), and inflammatory cytokines such as TNF-α and interleukin

(IL)-1β are essential for CAIA development (Kagari et al., 2002; Kagari et al., 2003; Tanaka et al.,

2006; Daha et al., 2011; Ozaki et al., 2012). Furthermore, the infiltration of neutrophils into the joint

cavity greatly contributes to CAIA development (Nandakumar et al., 2003; Tanaka et al., 2006).

Although, CAIA is not a T cell-driven arthritis model and is independent of B and mast cells

(Nandakumar et al., 2004; Mitamura et al., 2007; Zhou et al., 2007), the pathogenic features of CAIA

have striking similarities with human RA (Khachigian, 2006).

Oxidative stress plays an important role in the pathogenesis of inflammation. ROS, signaling

molecules that regulate various physiological and pathological events (Dröge, 2002; Brieger et al.,

2012), have been suggested to contribute to RA (Gelderman et al., 2007; Filippin et al., 2008). ROS

can be a potential biomarker of the degree of inflammation in RA patients (Filippin et al., 2008;

Kundu et al., 2012). It has been reported that ROS production mediated by toll-like receptor (TLR)

2 activation and nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (NOX) 2 complex

leads to severe granulocyte-dependent inflammation in CAIA (Kelkka et al., 2012). Thus, the

66

development of inflammation in CAIA has been characterized by the augmentation of ROS. However,

molecular mechanisms that connect ROS and CAIA development remain to be elucidated.

Drosophila Melanogaster TRP protein and its homologs consist of putative six transmembrane

subunits that assemble into tetramers to form Ca2+-permeable cation channels (Clapham, 2003; Voets

et al., 2005). Mammalian TRP channels are divided into six subfamilies: canonical (C), vanilloid (V),

melastatin (M), polycystic kidney disease (P), mucolipin (KL), and ankyrin (A). TRP channels are

activated by diverse stimuli, including receptor stimulation, heat, osmotic pressure, mechanical stress,

and environmental irritants from the extracellular environment and from inside the cell. It has been

revealed that some members of the TRPM, TRPC, TRPV, and TRPA subfamilies act as cell sensors

for ROS (Hara et al., 2002; Aarts et al., 2003; Yoshida et al., 2006; Andersson et al., 2008; Takahashi

et al., 2008; Takahashi et al., 2011). Notably, TRPM2 is activated by hydrogen peroxide (H2O2) (Hara

et al., 2002), and is abundantly expressed in immune cells such as neutrophils, monocytes, and

macrophages (Yamamoto et al., 2008; Hiroi et al., 2013). Previously, our group have reported that

TRPM2 activation mediated by H2O2 contributes to the aggravation of ROS-related disease models,

including dextran sulfate sodium-induced ulcerative colitis, myocardial ischemia/reperfusion injury,

and irradiation-induced salivary gland dysfunction (Yamamoto et al., 2008; Hiroi et al., 2013; Liu et

al., 2013). Theses paper lead us to hypothesize that ROS-sensitive TRPM2 in immune cells is

important in the exacerbation of RA. In this work, we study the roles of TRPM2 in CAIA by using

TRPM2 knockout mice.

67

MATERIALS AND METHODS

Animals. The generation of Trpm2 knockout mice has been previously reported (Yamamoto et al.,

2008). Male Trpm2 knockout mice and littermate wild-type control (C57BL/6J background) were

used. The animals used in this study were handled in accordance with the National Institute of Health

Guide for the Care and Use of Laboratory Animals and the experimental protocol was approved by

the Experimental Animal Committee of Showa University.

Genotyping of TRPM2 transgenic mice. Genomic DNA was extracted from tissue samples

obtained by tail tips using Tris-EDTA Buffer and proteinase K. PCR was performed using Taq

polymerase with the primer sets of PTRPM2-13F, PTRPM2-10R, and Pneo-5'a was carried out using

genomic DNA from mice; sequences are PTRPM2-13F (5’-CTTGGGTTGCAGTCATATGCAGGC-

3’), PTRPM2-10R (5’-GCCCTCACCATCCGCTTCACGATG-3’), and Pneo5'a (5’-

GCCACACGCGTCACCTTAATATGCG-3’). The PCR conducted in a thermal cycler (Gene Amp

PCR System 9600, Perkin Elmer Life Sciences, Boston, MA, USA) under the following conditions:

initial heating at 94°C for 2 min, followed by 34 cycles of denaturation at 94°C for 40 Sec, annealing

at 56°C for 30 Sec, and then final extension at 72°C for 30 Sec. The PCR samples were analyzed by

electrophoresis using 2% agarose gels.

Reverse transcriptase polymerase chain reaction (RT-PCR), Subcloning and sequencing. Total

RNA was isolated from mouse brain wild type (WT) mice and Trpm2 -knockout (KO) using ISOGEN

(Wako Pure Chemical Industries, Osaka, Japan) according to the manufacturer's instructions. The

concentration and purity of RNA were determined using NanoVue plus spectrophotometer (GE

Healthcare Life Science, Chalfont, Buckinghasmshire, UK). Total RNA samples (0.2 µg) were

reverse-transcribed at 42°C for 30 min with Avian Myeloblastosis virus reverse transcriptase using

RNA LA PCR kit (TaKaRa-Bio, Shiga, Japan). Sequences of gene-specific primer pairs that were

synthesized by Sigma-Aldrich and used to amplify three fragments of mouse brain TRPM2 cDNA

(Genebank accession number: AJ878415) are listed in Table 1. PCR was performed with LA Taq

polymerase (TaKaRa) and conducted in a thermal cycler (Gene Amp PCR System 9600) under the

following conditions: initial heating at 94°C for 2 min, followed by 32-35 cycles of denaturation at

94°C for 2 min, annealing at 55-63°C for 1 min, and then final extension at 72°C for 3 min. cDNAs

of TRPM2 were subcloned via Eco RI into pGEM®-T easy vector (Promega, Madison, USA). The

sequences were analyzed using 3130 Genetic Analyzer (life technology, Tokyo, Japan). The mouse

brain nucleotide sequence and predicted amino acid sequence were retrieved and analyzed through

the Genome Net WWW server using BLASTN, BLASTX, and BLASTP searches of the

nonredundant database (Altschul et al., 1990). Prediction of transmembrane region was performed

through the GenomeNet WWW server using the SOSUI program

68

(http://www.tuat.ac.jp/;mitaku/sosui/).

Induction of Arthritis in Mice. Arthritis was induced by using a cocktail of mouse 5 monoclonal

IgG antibodies recognizing the conserved epitopes of type II collagen (anti-CII Ab) with LPS,

according to the manufacturer’s instruction (Chondrex, Redmond, WA). Mice were intraperitoneally

injected with 5 mg of anti-CII Ab, and 3 days later injected with 50 μg LPS. Changes in paw thickness

were measured by using a micrometer caliper (Mitsutoyo, Kawasaki, Japan).

Histology. For histopathological analyses, right front paws of mice were obtained by cutting. These

were fixed in 10% formalin solution (Wako Pure Chemical Industries, Osaka, Japan), decalcified,

and embedded in paraffin. Sections of joints were made by slicing, and metacarpophalangeal joints

were stained with hematoxylin and eosin (H&E).

Immunohistochemistry. Paraffin sections of right front paw joints were blocked with buffer (Dako,

Glostrup, Denmark) at room temperature for 10 min. Sections were then incubated with horseradish

peroxidase-conjugated anti-mouse IgG antibody (Sigma-Aldrich, St. Louis, MO) at room

temperature for 1.5 h. Sections were stained by using diaminobenzidine (DAB) methods (Dako).

Bone CT. For bone computed tomography (CT) analyses, front left paws of mice were obtained by

cutting between wrist and elbow. These were fixed in 10% formalin solution. Scanning was

performed using a microfocus X-ray CT system, according to the manufacturer’s protocol (inspeXio

SMX-90CT, Shimadzu, Kyoto, Japan). The 3D images were constructed by using the Tri3d BON

software (Ratoc, Tokyo, Japan).

Measurement of cytokine concentration in paws. Hind right or left paws of mice were obtained

by cutting above the ankle. The paws were homogenized in lysis buffer (100 mM Tris pH 7.8, 66

mM EDTA, 1% NP-40, 0.4% sodium deoxycholate, and supplemented with protease inhibitor

cocktail) using a Polytron homogenizer (KINEMATICA, Luzern, Switzerland) and a sonicater

(VibraCell, Newtown, CT). The volume of homogenization buffer was adjusted to 170 mg of

tissue/ml buffer. The homogenate was centrifuged for 5 min at 15,000 r.p.m., and the supernatants

were subjected to cytokine analyses using ELISA analysis (R&D Systems, Minneapolis, MN) and

mouse inflammation cytometric bead array measurement (BD Bioscience, San Jose, CA) according

to the manufactures’ instructions. The concentration of total protein in the supernatants was measured

using a protein assay dye reagent (Bio-Rad, Hercules, CA). The concentration of cytokines was

expressed as picograms per milligram of protein.

Statistical analysis. All data are expressed as means ± S.E.M. The statistical analysis were

performed using the Student’s t-test. A value of P < 0.05 was considered significant.

69

RESULTS

Sequence characteristics of TRPM2 channels in wild type and Trpm2 knockout mice.

TRPM2 is highly expressed in the brain (Nagamine et al., 1998). Currently, there is no TRPM2

specific inhibitor (Hecquet and Malik, 2009), so we used transgenic mice in which TRPM2

expression was knocked out (Figure 1A). The nucleic acid of sequence of the mouse brain TRPM2

gene was characterized by performing a homology search with public and proprietary sequence

databases using the mouse TRPM2 protein (AJ878415). We found disruption of transmembrane

structure of Trpm2 knockout mice brain (Figure 1C). Our data indicates that wild type TRPM2

sequence displays 1507 amino acid residue, containing a long cytosolic N-terminal region, six

transmembrane segments (S1-S6) domains with putative pore regions, and a C-terminal region. On

the other hand, Trpm2 knockout displays the predicated 834-amino-acid protein, containing cytosolic

N-terminal region, two transmembrane segments (S1-S2) domains (Figure 1C,D). Immediately after

the introduced stop codon, a new in frame initiation codon follows, which may lead to generation of

two separated TRPM2 proteins. Meanwhile, the entire nucleotide sequence of Trpm2 wild type is

4521 bases and for Trpm2 knockout is 4097 bases (Figure 2). One notable region of difference

between mouse wild-type TRPM2 brain and knockout type Trpm2 is the deletion of exons 17, 18,

and 19 that leads to a frame shift and forming stop codon.

Table. 1. Primer pairs sequence used for identification of the sequence of TRPM2 in WT and

KO brain tissue.

Primer name Primer sequence

1(+) 5-ATGGAGTCCTTGGACCGGAG -3

1500(-) 5-CCTCACGAACTCAGGCTTGT-3

1475(+) 5-TCTCCAACAAGCCTGAGTTCGTG -3

3033 (-) 5-CGTCCAGTCGCTCTCAGGACACTTAG-3

2851(+) 5-GCCATTCTCATACATAACGA-3

4521(-) 5-TCAGAAGTGAGCTCCAAACAG-3

70

Figure 1. Molecular expression, structure, and transmembrane topology of mouse brain Trpm2 channel. (A)

Genomic PCR analysis of DNAs from wild-type (WT) and Trpm2 knockout (KO) mice. (B) Expression of

TRPM2 channel mRNA were detected by RT-PCR from total RNA isolated from Trpm2 WT or KO mouse

brain tissue. (C) Transmembrane topology in WT brain (left) and Trpm2 KO TRPM2 brain (right). (D) Amino

acid residues corresponding to putative transmembrane segments (S1–S2) of WT and KO brain tissue.

71

WT ATGGAGTCCTTGGACCGGAGAAGAACTGGCTCTGAGCAGGAGGAGGGCTTTGGGGTGCAGTCAAGGAGGG 70

KO ATGGAGTCCTTGGACCGGAGAAGAACTGGCTCTGAGCAGGAGGAGGGCTTTGGGGTGCAGTCAAGGAGGG 70

WT CCACTGACCTGGGCATGGTCCCCAATCTCCGACGAAGCAATAGCAGCCTTTGCAAGAGCAGGAGATTTCT 140

KO CCACTGACCTGGGCATGGTCCCCAATCTCCGACGAAGCAATAGCAGCCTTTGCAAGAGCAGGAGATTTCT 140

WT GTGCTCTTTCAGCAGTGAGAAGCAAGAAAACCTTAGCTCATGGATTCCCGAGAATATCAAGAAGAAGGAG 210

KO GTGCTCTTTCAGCAGTGAGAAGCAAGAAAACCTTAGCTCATGGATTCCCGAGAATATCAAGAAGAAGGAG 210

WT TGTGTGTACTTCGTGGAAAGTTCCAAACTCTCGGACGCAGGGAAGGTAGTGTGTGCGTGTGGTTATACCC 280

KO TGTGTGTACTTCGTGGAAAGTTCCAAACTCTCGGACGCAGGGAAGGTAGTGTGTGCGTGTGGTTATACCC 280

WT ACGAGCAACACTTGGAGGTGGCCATCAAGCCACACACCTTCCAGGGCAAGGAGTGGGATCCAAAGAAACA 350

KO ACGAGCAACACTTGGAGGTGGCCATCAAGCCACACACCTTCCAGGGCAAGGAGTGGGATCCAAAGAAACA 350

WT CGTCCAAGAGATGCCCACAGATGCCTTTGGTGACATCGTTTTCACAGACCTGAGCCAGAAAGTGGGGAAG 420

KO CGTCCAAGAGATGCCCACAGATGCCTTTGGTGACATCGTTTTCACAGACCTGAGCCAGAAAGTGGGGAAG 420

WT TATGTCCGGGTCTCCCAGGACACGCCCTCCAGTGTCATCTACCAGCTCATGACCCAGCACTGGGGCCTAG 490

KO TATGTCCGGGTCTCCCAGGACACGCCCTCCAGTGTCATCTACCAGCTCATGACCCAGCACTGGGGCCTAG 490

WT ATGTCCCCAACCTCCTCATCTCCGTGACCGGTGGGGCCAAGAACTTCAACATGAAGCTGAGGCTGAAGAG 560

KO ATGTCCCCAACCTCCTCATCTCCGTGACCGGTGGGGCCAAGAACTTCAACATGAAGCTGAGGCTGAAGAG 560

WT CATCTTCCGGAGAGGCTTGGTCAAGGTGGCTCAAACCACGGGGGCCTGGATCATCACTGGAGGATCCCAC 630

KO CATCTTCCGGAGAGGCTTGGTCAAGGTGGCTCAAACCACGGGGGCCTGGATCATCACTGGAGGATCCCAC 630

WT ACAGGCGTGATGAAGCAGGTGGGCGAAGCGGTACGGGACTTCAGTCTGAGCAGCAGCTGCAAAGAAGGTG 700

KO ACAGGCGTGATGAAGCAGGTGGGCGAAGCGGTACGGGACTTCAGTCTGAGCAGCAGCTGCAAAGAAGGTG 700

WT AAGTCATCACTATTGGCGTAGCCACGTGGGGCACCATCCACAACCGCGAGGGACTGATCCATCCCATGGG 770

KO AAGTCATCACTATTGGCGTAGCCACGTGGGGCACCATCCACAACCGCGAGGGACTGATCCATCCCATGGG 770

WT AGGCTTCCCCGCCGAGTACATGCTGGATGAGGAAGGCCAAGGGAACCTGACCTGCTTGGACAGCAACCAT 840

KO AGGCTTCCCCGCCGAGTACATGCTGGATGAGGAAGGCCAAGGGAACCTGACCTGCTTGGACAGCAACCAT 840

WT TCCCACTTCATCTTGGTGGATGATGGGACCCACGGGCAATATGGTGTGGAGATTCCGCTGAGGACTAAGC 910

KO TCCCACTTCATCTTGGTGGATGATGGGACCCACGGGCAATATGGTGTGGAGATTCCGCTGAGGACTAAGC 910

WT TGGAAAAGTTCATCTCAGAGCAAACGAAGGAAAGGGGAGGTGTGGCCATCAAGATCCCCATTGTCTGCGT 980

KO TGGAAAAGTTCATCTCAGAGCAAACGAAGGAAAGGGGAGGTGTGGCCATCAAGATCCCCATTGTCTGCGT 980

WT GGTGTTGGAGGGTGGCCCTGGCACTCTGCATACAATCTACAATGCCATCAACAATGGCACACCCTGCGTG 1050

KO GGTGTTGGAGGGTGGCCCTGGCACTCTGCATACAATCTACAATGCCATCAACAATGGCACACCCTGCGTG 1050

WT ATAGTGGAGGGCTCTGGCCGAGTGGCTGACGTCATCGCTCAGGTAGCTACTCTGCCTGTCTCTGAGATCA 1120

KO ATAGTGGAGGGCTCTGGCCGAGTGGCTGACGTCATCGCTCAGGTAGCTACTCTGCCTGTCTCTGAGATCA 1120

WT CCATCTCCTTGATCCAGCAGAAGCTCAGCATATTCTTCCAGGAGATGTTTGAGACTTTCACCGAAAACCA 1190

KO CCATCTCCTTGATCCAGCAGAAGCTCAGCATATTCTTCCAGGAGATGTTTGAGACTTTCACCGAAAACCA 1190

WT GATTGTGGAATGGACCAAAAAGATCCAAGACATTGTCCGGAGGCGGCAGCTGCTGACGATCTTCCGGGAA 1260

KO GATTGTGGAATGGACCAAAAAGATCCAAGACATTGTCCGGAGGCGGCAGCTGCTGACGATCTTCCGGGAA 1260

WT GGCAAGGATGGTCAGCAGGATGTGGATGTGGCCATTCTGCAGGCGTTACTGAAAGCCTCTCGAAGCCAAG 1330

72

KO GGCAAGGATGGTCAGCAGGATGTGGATGTGGCCATTCTGCAGGCGTTACTGAAAGCCTCTCGAAGCCAAG 1330

WT ACCACTTTGGCCACGAGAACTGGGACCACCAACTGAAGTTGGCTGTGGCCTGGAACCGCGTGGACATCGC 1400

KO ACCACTTTGGCCACGAGAACTGGGACCACCAACTGAAGTTGGCTGTGGCCTGGAACCGCGTGGACATCGC 1400

WT TCGCAGTGAGATCTTCACCGATGAATGGCAGTGGAAGCCTGCAGATCTGCATCCCATGATGACGGCCGCT 1470

KO TCGCAGTGAGATCTTCACCGATGAATGGCAGTGGAAGCCTGCAGATCTGCATCCCATGATGACGGCCGCT 1470

WT CTCATCTCCAACAAGCCTGAGTTCGTGAGGCTCTTTCTGGAGAATGGGGTGCGGCTCAAGGAGTTTGTCA 1540

KO CTCATCTCCAACAAGCCTGAGTTCGTGAGGCTCTTTCTGGAGAATGGGGTGCGGCTCAAGGAGTTTGTCA 1540

WT CTTGGGATACTCTTCTCTGCCTGTACGAGAACCTGGAGCCGTCCTGTCTCTTCCACAGCAAGCTACAGAA 1610

KO CTTGGGATACTCTTCTCTGCCTGTACGAGAACCTGGAGCCGTCCTGTCTCTTCCACAGCAAGCTACAGAA 1610

WT GGTGCTGGCCGAAGAACAGCGCTTAGCCTATGCATCTGCAACACCCCGCCTGCACATGCACCATGTGGCC 1680

KO GGTGCTGGCCGAAGAACAGCGCTTAGCCTATGCATCTGCAACACCCCGCCTGCACATGCACCATGTGGCC 1680

WT CAGGTGCTTCGTGAACTCCTGGGGGACTCCACGCAGCTGCTGTACCCCCGGCCCCGGTACACTGACAGGC 1750

KO CAGGTGCTTCGTGAACTCCTGGGGGACTCCACGCAGCTGCTGTACCCCCGGCCCCGGTACACTGACAGGC 1750

WT CACGGCTCTCGATGACCGTGCCACACATCAAGCTTAACGTGCAGGGAGTGAGCCTCCGGTCCCTCTATAA 1820

KO CACGGCTCTCGATGACCGTGCCACACATCAAGCTTAACGTGCAGGGAGTGAGCCTCCGGTCCCTCTATAA 1820

WT GCGATCAACAGGCCACGTTACCTTCACCATTGACCCAGTCCGTGACCTTCTCATTTGGGCCGTTATCCAG 1890

KO GCGATCAACAGGCCACGTTACCTTCACCATTGACCCAGTCCGTGACCTTCTCATTTGGGCCGTTATCCAG 1890

WT AACCACAGGGAGCTGGCAGGCATCATCTGGGCTCAGAGTCAGGACTGTACTGCCGCAGCACTGGCCTGTA 1960

KO AACCACAGGGAGCTGGCAGGCATCATCTGGGCTCAGAGTCAGGACTGTACTGCCGCAGCACTGGCCTGTA 1960

WT GCAAGATCCTGAAGGAGCTGTCCAAGGAGGAGGAAGATACAGACAGCTCTGAGGAGATGCTGGCACTGGC 2030

KO GCAAGATCCTGAAGGAGCTGTCCAAGGAGGAGGAAGATACAGACAGCTCTGAGGAGATGCTGGCACTGGC 2030

WT AGACGAGTTTGAGCACAGAGCTATAGGCGTCTTCACTGAGTGCTACAGGAAGGATGAGGAAAGAGCCCAG 2100

KO AGACGAGTTTGAGCACAGAGCTATAGGCGTCTTCACTGAGTGCTACAGGAAGGATGAGGAAAGAGCCCAG 2100

WT AAGCTGCTTGTCCGTGTGTCTGAGGCCTGGGGGAAGACCACCTGCCTGCAGCTGGCCCTAGAGGCCAAGG 2170

KO AAGCTGCTTGTCCGTGTGTCTGAGGCCTGGGGGAAGACCACCTGCCTGCAGCTGGCCCTAGAGGCCAAGG 2170

WT ACATGAAATTCGTGTCTCATGGAGGCATCCAGGCTTTCCTAACCAAGGTGTGGTGGGGCCAGCTCTGTGT 2240

KO ACATGAAATTCGTGTCTCATGGAGGCATCCAGGCTTTCCTAACCAAGGTGTGGTGGGGCCAGCTCTGTGT 2240

WT GGACAATGGCCTGTGGAGGATCATCCTGTGCATGCTGGCCTTCCCGCTGCTCTTCACCGGCTTCATCTCC 2310

KO GGACAATGGCCTGTGGAGGATCATCCTGTGCATGCTGGCCTTCCCGCTGCTCTTCACCGGCTTCATCTCC 2310

WT TTCAGGGAAAAGAGGCTGCAGGCACTGTGTCGCCCAGCCCGCGTCCGCGCCTTCTTCAATGCGCCTGTGG 2380

KO TTCAGGGAAAAGAGGCTGCAGGCACTGTGTCGCCCAGCCCGCGTCCGCGCCTTCTTCAATGCGCCTGTGG 2380

WT TCATCTTCCACATGAATATCCTCTCCTACTTTGCCTTCCTCTGCCTGTTCGCCTACGTGCTCATGGTGGA 2450

KO TCATCTTCCACATGAATATCCTCTCCTACTTTGCCTTCCTCTGCCTGTTCGCCTACGTGCTCATGGTGGA 2450

WT CTTCCAGCCTTCTCCATCCTGGTGCGAGTACCTCATCTACCTGTGGCTCTTCTCCCTGGTGTGCGAAGAG 2520

KO CTTCCAGCCTTCTCCATCCTGGTGCGAGTACCTCATCTACCTGTGGCTCTTCTCCCTGGTGTGCGAAGAG 2520

WT ACTCGGCAGCTATTCTATGATCCTGATGGCTGTGGACTAATGAAGATGGCGTCCCTGTACTTCAGTGACT 2590

KO .ACTCGGCAG 2529

73

WT TCTGGAACAAACTGGACGTTGGGGCCATTCTGCTCTTCATAGTAGGACTGACCTGTCGGCTCATCCCAGC 2660

KO

WT GACGCTGTACCCTGGGCGCATCATCCTGTCTTTGGACTTCATCATGTTCTGTCTCCGTCTCATGCACATC 2730

KO

WT TTCACTATTAGCAAGACACTGGGGCCCAAGATAATCATCGTGAAGCGGATGATGAAGGACGTCTTCTTCT 2800

KO

WT TCCTCTTTCTCCTGGCGGTGTGGGTGGTGTCCTTCGGCGTAGCTAAGCAGGCCATTCTCATACATAACGA 2870

KO

WT GAGCCGCGTGGACTGGATCTTCCGTGGGGTTGTCTATCACTCTTACCTGACCATCTTTGGCCAGATCCCA 2940

KO

WT ACCTACATTGACGGTGTGAATTTCAGCATGGACCAGTGCAGCCCCAATGGCACGGACCCCTACAAGCCTA 3010

KO………. GTGTGAATTTCAGCATGGACCAGTGCAGCCCCAATGGCACGGACCCCTACAAGCCTA 2586

WT AGTGTCCTGAGAGCGACTGGACGGGACAGGCACCTGCCTTCCCCGAGTGGCTGACTGTCACCCTGCTCTG 3080

KO AGTGTCCTGAGAGCGACTGGACGGGACAGGCACCTGCCTTCCCCGAGTGGCTGACTGTCACCCTGCTCTG 2656

WT CCTCTACCTGCTCTTTGCCAACATCCTGCTGCTTAACCTGCTCATCGCCATGTTCAACTACACCTTCCAG 3150

KO CCTCTACCTGCTCTTTGCCAACATCCTGCTGCTTAACCTGCTCATCGCCATGTTCAACTACACCTTCCAG 2726

WT GAGGTGCAGGAACACACAGACCAGATCTGGAAATTCCAGCGCCACGACCTGATCGAGGAGTACCATGGCC 3220

KO GAGGTGCAGGAACACACAGACCAGATCTGGAAATTCCAGCGCCACGACCTGATCGAGGAGTACCATGGCC 2796

WT GTCCCCCGGCACCTCCCCCACTCATCCTCCTCAGCCACCTGCAGCTCCTGATCAAGAGGATTGTCCTGAA 3290

KO GTCCCCCGGCACCTCCCCCACTCATCCTCCTCAGCCACCTGCAGCTCCTGATCAAGAGGATTGTCCTGAA 2866

WT GATCCCTGCCAAGAGGCATAAGCAGCTCAAGAACAAGCTGGAGAAGAACGAGGAGACAGCGCTCCTGTCT 3360

KO GATCCCTGCCAAGAGGCATAAGCAGCTCAAGAACAAGCTGGAGAAGAACGAGGAGACAGCGCTCCTGTCT 2936

WT TGGGAACTGTACCTGAAGGAGAACTACCTGCAGAACCAGCAGTACCAGCAGAAACAGCGTCCAGAGCAGA 3430

KO TGGGAACTGTACCTGAAGGAGAACTACCTGCAGAACCAGCAGTACCAGCAGAAACAGCGTCCAGAGCAGA 3006

WT AAATCCAAGACATCAGTGAGAAAGTGGACACCATGGTGGATCTGCTGGACATGGACCAGGTGAAGAGGTC 3500

KO AAATCCAAGACATCAGTGAGAAAGTGGACACCATGGTGGATCTGCTGGACATGGACCAGGTGAAGAGGTC 3076

WT AGGCTCCACAGAGCAGAGACTGGCTTCCCTGGAGGAACAGGTGACTCAGGTGACCAGAGCTTTGCACTGG 3570

KO AGGCTCCACAGAGCAGAGACTGGCTTCCCTGGAGGAACAGGTGACTCAGGTGACCAGAGCTTTGCACTGG 3146

WT ATCGTGACAACCCTGAAGGACAGTGGCTTTGGCTCAGGAGCAGGTGCGCTGACCCTGGCACCCCAGAGGG 3640

KO ATCGTGACAACCCTGAAGGACAGTGGCTTTGGCTCAGGAGCAGGTGCGCTGACCCTGGCACCCCAGAGGG 3216

WT CCTTCGATGAGCCAGATGCTGAGCTGAGTATCAGGAGGAAAGTAGAGGAACCAGGAGATGGTTACCACGT 3710

KO CCTTCGATGAGCCAGATGCTGAGCTGAGTATCAGGAGGAAAGTAGAGGAACCAGGAGATGGTTACCACGT 3286

WT GAGCGCCCGGCATCTCCTCTATCCCAATGCCCGCATCATGCGCTTCCCCGTGCCTAACGAGAAGGTGCCT 3780

KO GAGCGCCCGGCATCTCCTCTATCCCAATGCCCGCATCATGCGCTTCCCCGTGCCTAACGAGAAGGTGCCT 3356

WT TGGGCGGCAGAGTTTCTGATCTACGATCCTCCCTTTTACACCGCTGAGAAGGATGTGGCTCTCACAGACC 3850

KO TGGGCGGCAGAGTTTCTGATCTACGATCCTCCCTTTTACACCGCTGAGAAGGATGTGGCTCTCACAGACC 3426

WT CCGTGGGAGACACTGCAGAACCTCTGTCTAAGATCAGTTACAACGTCGTGGATGGACCGACGGACCGTCG 3920

74

KO CCGTGGGAGACACTGCAGAACCTCTGTCTAAGATCAGTTACAACGTCGTGGATGGACCGACGGACCGTCG 3496

WT CAGCTTCCATGGAGTCTACGTGGTCGAGTATGGGTTCCCGTTGAACCCCATGGGCCGCACAGGGTTGCGT 3990

KO CAGCTTCCATGGAGTCTACGTGGTCGAGTATGGGTTCCCGTTGAACCCCATGGGCCGCACAGGGTTGCGT 3566

WT GGTCGTGGGAGCCTCAGCTGGTTTGGTCCCAACCACACTCTGCAGCCAGTTGTCACCCGGTGGAAGAGGA 4060

KO GGTCGTGGGAGCCTCAGCTGGTTTGGTCCCAACCACACTCTGCAGCCAGTTGTCACCCGGTGGAAGAGGA 3636

WT ACCAGGGTGGAGCCATCTGCCGGAAGAGTGTCAGGAAGATGCTGGAGGTGCTAGTGATGAAGCTGCCTCG 4130

KO ACCAGGGTGGAGCCATCTGCCGGAAGAGTGTCAGGAAGATGCTGGAGGTGCTAGTGATGAAGCTGCCTCG 3706

WT CTCTGAGCACTGGGCCTTGCCTGGGGGCTCTAGGGAGCCAGGGGAGATGCTACCACGGAAGCTGAAACGG 4200

KO CTCTGAGCACTGGGCCTTGCCTGGGGGCTCTAGGGAGCCAGGGGAGATGCTACCACGGAAGCTGAAACGG 3776

WT GTCCTCCGGCAGGAGTTCTGGGTGGCCTTTGAGACCTTGCTGATGCAAGGTACAGAGGTATACAAAGGGT 4270

KO GTCCTCCGGCAGGAGTTCTGGGTGGCCTTTGAGACCTTGCTGATGCAAGGTACAGAGGTATACAAAGGGT 3846

WT ACGTGGATGACCCAAGGAACACAGACAATGCCTGGATCGAGACAGTGGCTGTCAGCATCCATTTTCAGGA 4340

KO ACGTGGATGACCCAAGGAACACAGACAATGCCTGGATCGAGACAGTGGCTGTCAGCATCCATTTTCAGGA 3916

WT CCAGAATGATATGGAGCTGAAGAGGCTGGAAGAGAACCTGCACACTCATGATCCAAAGGAGTTGACCCGT 4410

KO CCAGAATGATATGGAGCTGAAGAGGCTGGAAGAGAACCTGCACACTCATGATCCAAAGGAGTTGACCCGT 3986

WT GACCTGAAGCTGTCTACTGAATGGCAGGTGGTAGACCGGCGGATCCCTCTGTATGCGAACCACAAGACCA 4480

KO GACCTGAAGCTGTCTACTGAATGGCAGGTGGTAGACCGGCGGATCCCTCTGTATGCGAACCACAAGACCA 4056

WT TCCTCCAGAAGGTGGCCTCACTGTTTGGAGCTCACTTCTGA 4521

KO TCCTCCAGAAGGTGGCCTCACTGTTTGGAGCTCACTTCTGA 4097

Figure 2. Sequence similarity between wild and Trpm2 knockout mouse as synthesized by commercial gene

synthesis. The synthesized DNA sequence includes the open reading frame of WT mouse which consisted of

4521bp, and Trpm2 KO mouse which consisted of 4097 bp. Start and stop codon are highlighted in red and

deleted region of knockout are highlighted in blue.

Trpm2 knockout mice were protected against CAIA

To establish the pathological significance in vivo of TRPM2 in RA, we employed the CAIA

model. Anti-CII Ab was injected into both wild-type and Trpm2 knockout mice on day 0, and LPS

was subsequently injected on day 3. In wild-type mice, swelling of right front paws started after day

4, reached a maximum on day 7–10, and decreased gradually after that until day 21 (Figure 3A,B).

The swelling was dramatically suppressed in Trpm2 knockout mice (Figure 3A,B). In agreement

with the results from the right front paws, swelling of left front paws, right hind paws, and left hind

paws was observed in wild-type mice, whereas this was extensively suppressed in Trpm2 knockout

mice (Figure 3C). In Trpm2 knockout mice, paw swelling was prominently suppressed, but it was

unclear whether anti-CII Ab reached the cartilage in the joint. To evaluate localization of anti-CII Ab

in the joint, IgG of anti-CII Ab was assessed by anti-mouse IgG antibody. Like wild-type, anti-CII

Ab was bound to cartilage in Trpm2 knockout mice (Figure 3D). These results suggest that TRPM2

has major roles in the progressive severity of CAIA after localization of the anti-CII Ab in the joint.

75

Figure 3. Trpm2 knockout mice are protected against CAIA. (A–C) Changes induced by CAIA in paw

thickness in wild-type and Trpm2 knockout (KO) mice with CAIA. Wild-type and Trpm2 KO mice were

injected with anti-CII Ab (day 0) and LPS (day 3), and changes in right front paw thickness were recorded over

3 weeks (each n = 6) (A). Representative photos of right front paws on day 0 (before the injection of anti-CII

Ab) and 7 are shown (B). Scale bar, 5 mm. Changes in left front, right hind, and left hind paw thickness were

also recorded over 3 weeks (each n = 6) (C). **P < 0.01 and ***P < 0.001. Data points are mean ± S.E.M. (D)

DAB staining of joint section of right front paw is performed with HRP-conjugated mouse IgG-specific

antibody (anti-IgG, brown). Anti-CII Ab and LPS-treated and only LPS-treated wild-type or Trpm2 KO joints

on day 4 were used. Scale bar, 20 μm.

76

For histological assessment, the paraffin-embedded sections of the metacarpophalangeal joints

of right front paw fingers were stained with H&E. On day 7, massive infiltration of immune cells

into the joint cavity, synovitis, swelling of joint cavity, and cellular debris were observed in wild-

type mice with CAIA (Figure 4). On day 14 and 21, bone erosion and remodeling also emerged in

wild-type mice. In contrast, these features were not observed in Trpm2 knockout mice on day 7, 14,

and 21.

Left front paws were scanned using a CT scanner. In wild-type mice with CAIA, sever bone

erosion and remodeling were induced around metacarpophalangeal joints on day 14 and 21 as shown

in the above histological assessment, whereas Trpm2 KO mice showed no significance signs of bone

damage (Figure 5).

Figure 4. Histological assessment of joints. H&E staining of metacarpophalangeal joint sections of right front

paw fingers isolated from wild-type and Trpm2 KO mice with CAIA on day 0 (before the injection of anti-CII

Ab), 7, 14, and 21. Scale bar, 300 μm.

Trpm2 knockout disrupts inflammatory cytokine production in joints

To understand cellular mechanisms underlying symptoms observed by H&E staining and CT, we

measured the levels of inflammatory cytokines in the hind paws of mice. Inflammatory cytokines

(IL-1β, IL-6, and TNF-α) and chemotactic cytokines known as the chemokines (CCL2, CXCL1, and

CXCL2) were robustly produced on day 7 in wild-type mice by CAIA, whereas their production was

prominently reduced in Trpm2 KO mice (Figure 6A,B). The Levels of other cytokines such as IL-

17, IL-10, IL-12p70, and interferon (IFN)-γ were not critical for CAIA (data not shown). It has been

reported that receptor activator of nuclear factor-κB ligand (RANKL) is necessary for the formation

of osteoclast, which is a specialized type of cell in its capacity to resorb bone (Boyle et al., 2003).

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RANKL production was robustly induced in wild-type mice with CAIA on day 7, whereas it was

prominently reduced in Trpm2 KO mice (Figure. 6C), suggesting that osteoclast formation started

after day 7, and subsequent bone absorption was shown on day 14 indicated in Fig. 2 and 3. These

results suggest that inflammatory cytokines and chemokines are important in the development of

CAIA, and are attenuated by Trpm2 disruption.

Figure 5. Suppression of bone damage in Trpm2 KO mice. Representative μCT photographs of left front paws

isolated from wild-type and Trpm2 KO mice with CAIA on day 0 (before the injection of anti-CII Ab), 7, 14,

and 21. Scale bar, 500 μm.

78

Figure 6. Production of inflammatory mediators is suppressed in Trpm2 KO mice. (A–C) Expression levels

of IL-1β, IL-6, and TNF-α (A), CCL2, CXCL1, and CXCL2 (B), and RANKL (C) protein in hind paws isolated

from either wild-type or Trpm2 KO mice with CAIA on day 0 (before the injection of anti-CII Ab), 7, 14, and

21 (n = 6). *P < 0.05, **P < 0.01, and ***P < 0.001. Data points are mean ± S.E.M

.

79

DISCUSSION

In the present study, we found that the disruption of ROS-sensitive TRPM2 channels dramatically

reduces the severity of CAIA, which has been associated with the augmentation of ROS. Thus, our

data suggest that the activation of TRPM2 by ROS is important in CAIA development.

In Trpm2 KO mice, the inflammation in joints was almost completely inhibited on day 7 and the

swelling of paws was suppressed before day 7, despite the localization of collagen antibody on

cartilage surface on day 4. These results indicate that TRPM2 is essential for innate immune

processes before day 7. It is possible that TRPM2 is important for the infiltration of neutrophils in

the joint, because the infiltration of immune cells such as neutrophils was inhibited in Trpm2

knockout mice (Hara et al., 2002; Yamamoto et al., 2008). Furthermore, it is also possible that

TRPM2 mediates cell death in the joint, because the accumulation of cellular debris, which may be

resulted from cell death and calcification of synovial cells and neutrophils, was present in the joint

cavity only in wild-type mice treated with CAIA.

Biological roles induced by ROS-activated TRPM2 have been studied. Our group have identified

that treatment of monocytes with a sub lethal concentration of H2O2 triggers the production of

neutrophil chemokines through TRPM2-mediated Ca2+ entry (Yamamoto et al., 2008). Our group

have also shown that neutrophil TRPM2 plays an important role in neutrophil adhesion to endothelial

cells by the combination of H2O2 and leukotriene B4 (Hiroi et al., 2013). It is possible that roles of

TRPM2 for the chemotaxis of neutrophils described above are also critical in the development of

CAIA. Furthermore, it has been reported that ROS-activated TRPM2 can mediate cell death (Hara

et al., 2002; McNulty and Fonfria, 2005). Thus, TRPM2-mediated cell death of synovial cells or

infiltrated neutrophils may be critical in the development of CAIA.

It has been reported that TRPM2 is gated by adenosine diphosphoribose (ADPR) through the

binding of ADPR to the C-terminal domain, and the ROS sensitivity of TRPM2 can be mediated by

ADPR released from mitochondria and nucleus (Perraud et al., 2001; Kühn et al., 2005; Perraud et

al., 2005; Knowles et al., 2013). ADPR can be generated in the nucleus by a pathway involving

poly(ADPR) polymerase-1 (PARP-1), with a similar pathway putatively functional in mitochondria

(Du et al., 2003; Schreiber et al., 2006). The report that PARP-1-deficient mice are partially protected

against CAIA (Garcia et al., 2006) supports our idea that TRPM2 activation by ROS via PARP-1

pathway is involved in CAIA development.

It can be speculated that the activation of TRPM2 is involved in by at least two other pathways

in addition to the ROS pathway. The first pathway is immune complex (IC)-mediated signaling. In

80

CAIA, IC can be formed by the binding of anti-collagen antibody to type II collagen. IC can induce

neutrophil migration through FcγRs (Mayadas et al., 2009). TRPM2 may play a role in altering the

IC and FcγRs-mediated signaling processes. The second pathway is LPS signaling. It has been shown

that TRPM2 is involved in LPS-induced cytokine production in monocytes (Wehrhahn et al., 2010).

Our data, which demonstrate that LPS-evoked gaining access of the collagen antibody to the joint is

not affected by TRPM2, suggest that TRPM2 is not important in LPS-dependent processes, although

we cannot completely exclude the possibility that TRPM2 is involved in LPS signaling.

In conclusion, our findings that loss of TRPM2 function protects against mouse CAIA

development present the opportunity for developing a new therapeutic strategy for treating RA by

the inhibition of TRPM2. Our results also suggest that the therapeutic strategy to inhibit TRPM2

should be applicable to other autoimmune inflammatory diseases.

81

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Chapter 3

Functional and Structural Divergence in Human TRPV1 Channel

Subunits by Oxidative Cysteine Modification

ABSTRACT

Transient receptor potential vanilloid 1 (TRPV1) channel is a tetrameric protein that acts as a

sensor for noxious stimuli such as heat, and for diverse inflammatory mediators such as

oxidative stress to mediate nociception in a subset of sensory neurons. In TRPV1 oxidation

sensing, cysteine (Cys) oxidation has been considered as the principle mechanism; however, its

biochemical basis remains elusive. Here, we characterize the oxidative status of Cys residues in

differential redox environments and propose a model of TRPV1 activation by oxidation.

Through employing a combination of non-reducing SDS-PAGE, electrophysiology and mass

spectrometry, we have identified the formation of subunit dimers carrying a stable inter-

subunit disulfide bond between Cys258 and Cys742 of human TRPV1 (hTRPV1). C258S and

C742S hTRPV1 mutants have a decreased protein half-life, reflecting the role of the inter-

subunit disulfide bond in supporting channel stability. Interestingly, the C258S hTRPV1

mutant shows an abolished response to oxidants. Mass spectrometric analysis of Cys residues

of hTRPV1 treated with hydrogen peroxide shows that Cys258 is highly sensitive to oxidation.

Our results suggest that Cys258 residues are heterogeneously modified in the hTRPV1

tetrameric complex and comprise Cys258 with free thiol for oxidation sensing and Cys258,

which is involved in the disulfide bond for assisting subunit dimerization. Thus, the hTRPV1

channel has a heterogeneous subunit composition in terms of both redox status and function.

Reprinted with permission of the American Society for Biochemistry and Molecular Biology. All

rights reserved.

Copyright 2015 by the American Society for Biochemistry and Molecular Biology.

87

INTRODUCTION

Ion channels play important roles in sensory systems by detecting diverse changes in the

environment and inside the body. Drosophila melanogaster transient receptor potential (TRP)

channel and its homologs are putative six-transmembrane proteins that assemble into tetramers to

form non-selective cation channel biosensors (Clapham, 2003; Clapham et al., 2005; Liao et al.,

2013). One of the intriguing features of TRP channels is their ability to detect changes in the redox

environment (Kozai et al., 2014). Indeed, numerous TRP channels including TRPV1, TRPV3,

TRPV4, TRPC1, TRPC4, TRPC5, and TRPA1, sense alterations in the redox environment (Yoshida

etal., 2006; Susankova et al., 2006; Takahashi et al., 2008). The oxidative modification of the cysteine

(Cys) residues in these channels mediates their activation and plays an essential role in various

physiological and pathological conditions (Vyklicky et al., 2002; Yoshida et al., 2006; Susankova et

al., 2006; Andersson et al., 2008; Bessac et al., 2008; Xu et al., 2008; Liu et al.,2013).

TRPV1 is localized in a subset of sensory neurons such as C- and Aδ- fibers, which are associated

with acute and chronic inflammatory pain. TRPV1 is activated by diverse inflammatory mediators,

such as heat, acidity, pungent compounds, endogenous lipid signaling molecules, and oxidative stress,

to mediate nociception (Caterina et al., 1997; Tominage et al., 1998; Zygmunt et al., 1999; Caterina

et al., 2000; Davis et al., 2000; Hwang et al., 2000; Premkumar and Ahern et al., 2000; Xu et al.,

2005;Yoshida et al., 2006; Susankova et al., 2006). During inflammation, the redox environment

surrounding the cells changes due to rapid production of reactive oxygen species (ROS). Increased

levels of ROS are implicated in the etiology of inflammatory hyperalgesia (Tominage et al., 1998;

Wang et al., 2004; Ndengele et al., 2008; Khattab, 2006; Bezerra et al., 2007). Importantly, TRPV1

was found to be responsible for long sustained thermal hyper-sensitivity in inflammatory

hyperalgesia induced by increased levels of hydrogen peroxide (H2O2) (Keeble et al., 2009). It is,

therefore, possible that oxidative stress triggers hyperalgesia, at least in part, through activation of

TRPV1.

There have been multiple hypotheses proposed for the regulation of TRPV1 by oxidation. We

have previously reported that oxidizing agents activate rat TRPV1 (rTRPV1) by modifying the

extracellular Cys residues, where Cys modification acts as a “switch” for channel activation (Yoshida

et al., 2006). Another group alternatively hypothesized that H2O2 activates chicken TRPV1

(cTRPV1) via oxidizing multiple Cys residues located on the cytoplasmic side; each Cys residue

contributing to the activation in a “graded” fashion (Chuang and Lin, 2009). The same group also

claimed that C-terminal dimerization through disulfide bond formation is essential for oxidation-

induced cTRPV1 activation (Wang and Chuang, 2011). However, a unified view of how oxidizing

agents activate the channel has not yet been achieved, although there is a consensus that oxidative

88

modifications of multiple Cys residues are involved in this mode of TRPV1 activation.

There are two crucial caveats in the above hypotheses that limit the elucidation of the mechanism

underlying TRPV1 activation by oxidation. First, all of the studies concluding the involvement of

oxidation of Cys residues in the activation mechanism are heavily based on site-directed Cys

mutagenesis experiments, which are often regarded as an indirect approach to assess functional

importance of specific residues. Secondly, most of these mechanistic models are built based on the

simple assumption that Cys residues in TRPV1 have predominantly free thiols, both readily

accessible and modifiable, at cellular conditions. Although this is a fair assumption based on studies

implicating that the intracellular milieu is primarily reduced (Hansen et al., 2009), previous studies

have implicated otherwise. For instance, TRPA1 was demonstrated to possess a disulfide bond

between Cys residues facing the intracellular side (Wang et al., 2012). Moreover, TRPV1 has been

hypothesized to possess stable structural disulfide bonds because a reducing agent dithiothreitol

(DTT) alters the cooperative binding of TRPV1-specific agonist resiniferatoxin to rat dorsal root

ganglion (DRG) and regulates rTRPV1 activity when expressed in HEK293 cells (Szallasi and

Blumberg, 1993; Szallasi et al., 1993; Vyklicky et al., 2002; Susankova et al., 2006).

Here, we address these two caveats for the previous approaches to understand the role of Cys

residues, by characterizing the oxidative status of Cys residues in differential redox environments

and propose a refined mechanism of TRPV1 activation by oxidation. In particular, we focus on the

activation mechanism of human TRPV1 (hTRPV1), which has not been characterized previously.

89

EXPERIMENTAL PROCEDURES

Reagents. DTT was purchased from Nacalai Tesque (Kyoto, Japan). Capsaicin, cycloheximide, N-

ethylmaleimide (NEM), and iodoacetamide (IAA) were purchased from Sigma-Aldrich (St. Louis,

MO). Fura-2-AM ester, 5-nitro-2-pyridyl disulfide (5-nitro-2-PDS), and 2-pyridyl disulfide were

purchased from Dojindo (Kumamoto, Japan). 3-nitrophenyl disulfide, dipropyl disulfide, dimethyl

sulfoxide (DMSO), H2O2, and reduced glutathione were purchased from Wako (Osaka, Japan). 4-

chlorophenyl disulfide and 4-aminophenyl disulfide were purchased from Tokyo Chemical (Tokyo,

Japan). NEM, IAA, cycloheximide, and reactive disulfides were prepared as stock solutions in

DMSO and were diluted at working concentrations in aqueous solutions containing 0.01% or 0.1%

DMSO. DTT and H2O2 were directly dissolved in aqueous solutions at working concentrations.

Cell Culture. Human embryonic kidney (HEK) 293 and HEK293T cells were cultured in

Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal bovine serum, 30 units/ml

penicillin, and 30 μg/ml streptomycin at 37°C under 5% CO2. HaCaT cells were culture in DMEM

containing 10% fetal bovine serum, 30 units/ml penicillin, 30 μg/ml streptomycin and 2 mM L-

glutamate at 37°C under 5% CO2.

cDNA Cloning and Recombinant Plasmid Construction. Plasmids carrying hTRPV1 and

rTRPV1 cDNA were prepared as previously described (Yoshida et al., 2006). TRPV1 and hTRPV1

Cys mutants were subcloned into pCI-neo vector (Promega Corporation, Madison, WI), pEGFP-N

(Clontech Laboratories, Mountain View, CA) or pCMV-Tag2 (Stratagene, Palo Alto, CA).

cDNA Expression in Cells. HEK293 cells and HEK293T cells were transfected with recombinant

plasmids using SuperFect Transfection Reagent (Qiagen, Valencia, CA) and Lipofectamine 2000

transfection reagent (Invitrogen, Life Technologies Corporation, Grand Island, NY), respectively,

according to the manufacturer’s instructions. For intracellular Ca2+ concentration ([Ca2+]i)

measurement and electrophysiological measurements, recombinant plasmids were co-transfected

with pEGFP-F and pEGFP-N1, respectively, and HEK293 cells with green fluorescence were

analyzed. Transfected cells were grown for 30-40 h prior to [Ca2+]i measurement and

electrophysiological measurements, and were grown for 48 h prior to immunoprecipitation and

western blotting (WB).

Western Blots. Cells were lysed in RIPA buffer containing the followings 50 mM Tris (pH 7.4), 150

mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS, and 20 mM NEM. Protein

samples were fractioned by electrophoresis through 7.5% SDS-polyacrylamide gels (SDS-PAGE)

and analyzed by WB with anti-TRPV1 (Santa Cruz, CA, USA, SC-1248), anti-β-actin (Sigma,

90

A2228), and anti-α-tubulin (Sigma, T6074) antibodies. The chemiluminescence intensities of the

bands were measured by Multigauge version 3.0 (Fuji film, Tokyo, Japan).

Blue Native Polyacrylamide Gel Electrophoresis. Blue native polyacrylamide gel electrophoresis

was performed according to the method of Schägger and co-workers with some modification (Wittig

et al., 2006). HEK293 cells transfected with rTRPV1 or hTRPV1 were suspended in sucrose buffer

(83 mM sucrose, 6.6 mM imidazole/HCl, 20 mM NEM, pH 7.0) and homogenized with 20-40 strokes

using dounce homogenizer. The lysates were centrifuge at 15000 g and the pellet was solubilized in

35 µl of solubilization buffer (50 mM NaCl, 50 mM imidazole/HCL, 6-aminohexanoic acid, 1 mM

EDTA, 20 mM NEM, pH 7.0). Subsequently, 5 µl of 20% n-Dodecyl β-D-maltoside was added and

incubated for 10 min at 4°C. The sample was then centrifuged at 15000 g and supernatant was

collected, mixed with 5 µl of 50% glycerol, and 2.5 µl of 5% commassie brilliant blue G-250. The

supernatant was aliquoted into four samples with increasing concentration of SDS (0, 0.5, 1.0, and

1.5%) and incubated for 20 min at room temperature (RT). The samples were then subjected to non-

denaturing electrophoresis initially at 100 V until the samples were within the stacking gel and were

continued with a voltage and current limited to 500 V and 15 mA at 4°C. After separation, the gel

was de-stained for 3 min with 100% methanol and electro-transferred to PVDF membrane (Millipore,

Bedford, MA, USA). After the transfer, the PVDF membrane was incubated in denaturing solution

containing 50 mM Tris-HCl (pH 7.4) and 2% SDS at 50°C for 30 min. The blots were then analyzed

by anti-TRPV1.

Chemical Cross-Linking. HEK293 cells transfected with hTRPV1 were detached from the 6 mm

dish using trypsin and washed three times with phosphate buffer saline (PBS). The sample was

aliquoted into three samples with the concentrations of disuccinimidyl suberate (DSS) (Sigma) at

final concentration of 0, 25, and 75 µM for 20 min at RT. The cross linking reaction was quenched

with 1 M Tris-HCl pH 8.0 and samples were mix with SDS sample buffer and then analyzed by 3-

18% polyacrylamide SDS-PAGE and WB with an antibody to TRPV1.

Cell Surface Labeling Experiments. The cell surface hTRPV1 was measured by biotinylation as

previously described with some modifications (Yoshida et al., 2006). Cells grown in 60 mm dishes

were washed with HEPES-buffered saline (HBS) containing the following: 107 mM NaCl, 6 mM

KCl, 1.2 mM MgSO4, 2 mM CaCl2, 11.5 mM glucose, and 20 mM HEPES (pH adjusted to 7.4 with

NaOH). After incubation of the cells with 10 mM DTT, dishes were washed with HBS twice and

then incubated with sulfo-NHS-SS-biotin (Thermo Fisher Scientific, San Jose, CA, USA) for 30 min

at 4°C. The cells were washed with PBS containing 10 mM glycine three times to stop the

biotinylation reaction. The cells were then lysed with 300 µl of RIPA buffer with 20 mM NEM, and

rotated for 2 h at 4°C. After centrifugation, the supernatant was collected and incubated with

91

NeutrAvidin-Plus beads (Thermo Fisher Scientific) for 2 h at 4°C. The proteins were washed with

RIPA buffer containing 50 mM NEM 3 times and resuspended in SDS sample buffer and subjected

to SDS-PAGE and WB.

Protein Stability Assay. HEK293 cells were transfected with hTRPV1 or hTRPV1 Cys mutants and

cultured for 24 h. The cycloheximide was added to a final concentration of 50 µg/ml to culture media

and cells were washed twice with PBS and recovered with RIPA buffer after the indicated times.

The samples were mixed with reducing sample buffer and analyzed by SDS-PAGE and WB using

antibody to TRPV1 and β-actin. Quantification of hTRPV1 expression was achieved by

normalization with β-actin.

Isolation of Rat DRG Neurons. Dorsal Root Ganglion (DRG) neurons were prepared from 4 weeks

old male Sprague Dawley rats as described previously (Takahashi et al., 2008). The DRG neurons

were subjected to WB. All animal experiments were performed in accordance with protocols

approved by the Institutional Animal Care and Use Committees of the Graduate School of

Engineering, Kyoto University.

[Ca2+]i Measurement. Transfected HEK293 cells were subjected to [Ca2+]i measurement 3-16 h

after plating onto poly-L-lysine-coated glass coverslips. The fura-2 fluorescence was measured in

HBS. Fluorescence images of the cells were recorded and analyzed with the video image analysis

system AQUACOSMOS (Hamamatsu Photonics, Shizuoka, Japan) according to the manufacturer’s

instructions. Fura-2 measurements were carried out at RT in HBS, otherwise indicated. The

340:380-nm ratio images were obtained on a pixel-by-pixel basis and converted to Ca2+

concentrations by in vivo calibrations using 40 µM ionomycin (Nishida et al., 1999).

Electrophysiology. For electrophysiological measurements, coverslips with cells were placed in

dishes containing bath solutions. Currents from cells were recorded at RT using path-clamp

techniques of whole-cell mode with EPC-10 (Heka Elektronik, Lambrecht/Pfalz, Germany) patch-

clamp amplifier as previously described (Okada et al., 1999). The patch electrode prepared from

borosilicate glass capillaries had a resistance of 2-4 MΩ. Current signals were filtered at 2.9 kHz

with a four-pole Bessel filter and digitized at 10 kHz. Patchmaster (Heka Elektronik) software was

used for command pulse control, data acquisition, and data analysis. The series resistance was

compensated (50-70%) to minimize voltage errors. A holding potential of –80 mV was used for all

the experiment. The external solution contained the following: 100 mM NaCl, 5 mM KCl, 2 mM

BaCl2, 5 mM MgCl, 25 mM HEPES, 30 mM Glucose (pH 7.3 adjusted with NaOH, and osmolarity

adjusted to 320 mOsm with D-mannitol). The pipette solution contained the following: 140 mM CsCl,

4 mM MgCl2, 10 mM EGTA, 10 mM HEPES (pH 7.3 adjusted with CsOH and osmolarity adjusted

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to 300 mOsm with D-mannitol). For the DTT treatment, liquid junction potential was adjusted before

each experiment.

Liquid Chromatography-Mass Spectrometry. Flag-hTRPV1 or Flag-hTRPV1 Cys mutants were

transfected into HEK293T cells. After 48 h, the cells were lysed with buffer containing 50 mM Tris

(pH 7.4), 2% SDS, 50 mM NEM, 150 mM NaCl, 1% Nonidet P-40, and 0.5% sodium deoxycholate.

The cell lysates were sonicated, centrifuge at 15000 g and 10x diluted in RIPA buffer containing 50

mM NEM. Flag-hTRPV1 was immunoprecipitated with M2 monoclonal antibody conjugated to

protein A-agarose breads (Sigma) and was rocked overnight at 4°C. The complex was washed eight

times with RIPA buffer containing 50 mM NEM for 5 min at RT and re-suspended into SDS sample

buffer and rocked for 30 min. The protein samples were fractionated by 7.5% SDS-PAGE. The bands

corresponding to the molecular weight of dimeric or monomeric Flag-hTRPV1 were excised and

reduced with 10 mM DTT, alkylated with 55 mM iodoacetamide (IAA), and digested in-gel by

treatment with trypsin (12.5 ng/µl) or chymotrypsin (10 ng/µl) or glu-C (50 ng/µl) in 50 mM

ammonium bicarbonate for overnight at 37°C as previously described (Grønborg et al., 2004). The

samples were analyzed by reverse phase liquid chromatography tandem mass spectrometry using a

LTQ-orbitrap-XL hybrid mass spectrometer (Thermo Fisher Scientific) as follows.

A capillary reverse phase HPLC-MS/MS system composed of an Agilent 1100 series gradient

pump equipped with Valco C2 valves with 150 µm-pots, and LTQ-orbitrap XL hybrid mass

spectrometer equipped with an XYZ nano-electrospray ionization (NSI) source (AMR, Tokyo,

Japan) were employed. Samples were automatically injected using PAL system (CTC analytics,

Zwingen, Switzerland) into a peptide L-trap column (Chemical Evaluation Research Institute, Tokyo,

Japan) attached to an injector valve for desalinating and concentrating peptides. After washing the

trap with water containing 0.1% trifluoroacetic acid and 2% acetonitrile, the peptides were separated

on a NTCC—360/100-3-123 column (100 µM diameter, 3 µM C18 particles, Nikkyo Technos Co.

Ltd., Tokyo, Japan) using a gradient of A (0.1% formic acid) and B (80% acetonitrile, 0.1% formic

acid). The elution was carried out at 500 nl/min by running from 5% to 45% over 45 min (as shown

in Fig. 5 and 6) or 25 minutes (as shown in Fig. 8 and 9), 95% for 6 min, 5% for 9 min for selected

reaction monitoring (SRM) transitions for the peptide containing Cys158, 258, 363 and 742. The

elution was carried out from 25% to 80% over 20 min, 95% for 6 min, 5% for 9 min for SRM

transition for the peptides containing Cys127 (as shown in Fig. 8). SRM was carried out with

collision induced dissociation (CID) (35% normal collision energy), 1 microscan, 100 ms ion time

on LTQ linear ion trap. Peak area was calculated using Xcaliber 3.0 software version 2.2.0.48

(Thermo Fisher Scientific). For the shotgun proteomics experiment, the LTQ orbitrap XL was

operated in a data dependent mode, selecting 3 precursors for fragmentation by CID. The survey scan

93

was performed in the orbitrap at 30000 resolutions from 300-2000 m/z. Precursors were isolated with

a width of 2 m/z. Single and assigned charge states were ignored for precursor selection. Proteome

discoverer 1.3 with Sequest (Eng et al., 1994) was used for protein identification against protein

database containing hTRPV1 sequence (accession number Q8NER1). Precursor mass tolerance was

set to 10 ppm and 0.8 Da for orbitrap and linear ion-trap, respectively. Oxidized methionine (M),

carbamidomethylation (C) and NEM (C) were set as dynamic modification. Trypsin, chymotrypsin,

or trypsin/glu-C were chosen as the enzymes for digestion.

Structural Modeling. Three dimensional structure of rTRPV1 (PDB # 3J5P) was used to estimate

the distance between the Cys residues. Structure was visualized using CCP4mg software version

2.9.0 (McNicholas et al., 2011).

Statistical Analysis. All data are expressed as means ± standard error of the mean (SEM) or ±

standard deviation (SD). We accumulated the data for each condition from at least three independent

experiments. The statistical analyses were performed using Student’s t test. A value of P < 0.05 was

considered significant.

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RESULTS

Characterization of redox modification in TRPV1

Activation of cTRPV1 under oxidative stress via dimerization has been described previously

(Wang et al., 2011). To test whether oxidizing agents activate mammalian TRPV1 via dimerization

similar to cTRPV1, the protein multimerization of hTRPV1 and rTRPV1 expressed in HEK293 cells

treated with a congeneric series of reactive disulfides with differential redox potentials, was assessed

by non-reducing SDS-PAGE (Figure 1A) (Takahashi et al., 2011). Once the cells were lysed, sample

preparation was carried out in the presence of NEM, an irreversible Cys alkylating agent, which

prevents post-lysis oxidation and thiol-disulfide shuffling (Ryle and Sanger, 1954). hTRPV1 and

rTRPV1 showed significant differences in their mobility, even in the absence of exogenous oxidizing

agents. While the rTRPV1 band was observed at ~100 kDa as predicted from its molecular weight

(95 kDa), hTRPV1 was predominantly observed at ~245 kDa (Figure 1B). Recombinant expression

of hTRPV1 and rTRPV1 elicited robust [Ca2+]i increases in response to the congeneric series of

reactive disulfides in HEK293 cells, which were not observed in the vector-transected cells (Figure

1C). Notably, the reactive disulfides that activated the channels did not alter the band patterns of

human or rat TRPV1 on the non-reducing SDS-PAGE (Figure 1B). These data suggests that

oxidative modification elicited by reactive disulfides do not affect the multimeric patterns of

mammalian TRPV1 channels.

Incubation of hTRPV1-expressing HEK293 cells with DTT shifted the 245 kDa band to a

monomeric position in a dose-dependent manner (Figure 1D), suggesting that the formation of

disulfide bonds may be responsible for the band shift observed under the non-reducing conditions.

To examine the reversibility of the 245 kDa band in a cellular environment, we incubated the

hTRPV1-expressing HEK293 cells with DTT for 10 min followed by washes and re-formation of

the band was monitored. The 245 kDa band reappeared after 5 min, in a time dependent manner,

demonstrating that the monomeric hTRPV1 has high propensity to form a 245 kDa complex when

expressed in cells (Figure 1E). To confirm the identity of the 245 kDa band, a purified Flag-hTRPV1

245 kDa band was analyzed by mass spectrometry following SDS-PAGE and enzymatic digestions.

We obtained an 81% amino acid sequence coverage of hTRPV1, including the C- and N-terminus,

revealing that the dimeric band indeed consists of full length hTRPV1 (Figure 1F). Thus, hTRPV1

appears to possess a stable inter-subunit disulfide bond between two subunit proteins which is

unaffected by oxidative conditions.

95

Figure 1. Oxidative modification and its impacts on human and rat TRPV1. (A) Congeneric series of

reactive disulfides with their redox potentials (mV). Compounds with less negative redox potentials have

stronger electrophilicity. (B) Non-reducing SDS-PAGE and WB analysis of hTRPV1- or rTRPV1-transfected

HEK293 cells treated with congeneric series of reactive disulfides (10 μM) for 10 min at room temperature.

WB for β-actin are also shown (bottom). (C) Averaged time courses of [Ca2+]i rises evoked by reactive

disulfides with indicated redox potentials for 8 min in HEK293 cells expressing hTRPV1, rTRPV1, or vector

(left). Plots of maximal rise of Ca2+ (Δ[Ca2+]i) induced by 10 μM concentrations of reactive disulfides for 8

min in HEK293 cells expressing hTRPV1 or rTRPV1 (n = 10–52) (right). Data points are the mean ± S.EM.

*p < 0.01 compared with rTRPV1. (D) Non-reducing SDS-PAGE and WB analysis of hTRPV1-transfected

HEK293 cells treated with various concentrations of DTT for 10 min at 37°C. (E) hTRPV1-transfected

HEK293 cells were treated with 10 mM DTT for 10 min, washed with HBS twice, and recovered as lysates

after the indicated time points to be analyzed with non-reducing SDS-PAGE and WB. (F) Identification of the

FLAG-hTRPV1 245-kDa band isolated from HEK293T cells by mass spectrometric analysis. The excised 245-

kDa band was subjected to enzymatic digestion (combination of trypsin or chymotrypsin or trypsin/glu-C) and

analyzed with liquid chromatography coupled with tandem mass spectrometry. The amino acids in black were

identified.

To further test whether hTRPV1 forms a complex with other proteins or hTRPV1 multimer, we

co-expressed hTRPV1-GFP and hTRPV1 and assessed whether an intermediate band that consists

of hTRPV1-GFP+hTRPV1 could be observed under non-reducing conditions. We observed three

distinct bands that corresponded to the molecular weights of hTRPV1+hTRPV1, hTRPV1-

GFP+hTRPV1-GFP, and hTRPV1+hTRPV1-GFP complexes, suggesting that hTRPV1 forms an

96

inter-subunit disulfide bond (Figure 2A).To confirm the stoichiometry of disulfide-linked hTRPV1,

the band position of the disulfide-linked hTRPV1 complex was compared with DSS cross-linked

hTRPV1 bands. The position of the disulfide-linked hTRPV1 corresponded to two DSS cross-linked

hTRPV1, revealing the disulfide linkage between two hTRPV1 subunits (Figure 2B). To test whether

disulfide linked hTRPV1 subunits are incorporated into the tetramer, hTRPV1 was subjected to non-

reducing blue native PAGE. rTRPV1 was also analyzed in parallel as a control for disulfide-less

TRPV1. hTRPV1 and rTRPV1 showed a marked difference in the mobility; rTRPV1 showed a

tetramer to monomer transition as a function of increasing concentrations of SDS, while hTRPV1

showed an absence of monomeric band but increased intensity in the dimeric band (Figure 2C).

These results suggest that hTRPV1 has a unique inter-subunit disulfide bond.

Figure 2. The hTRPV1 channel has inter-subunit disulfide bond between two subunits. (A) hTRPV1 and

hTRPV1-GFP were transfected individually and co-transfected to HEK293 cells. The lysates were analyzed

with reducing and non-reducing SDS-PAGE and WB. (B) Cell lysates from hTRPV1-transfected HEK293 cells

with and without treatment of indicated concentrations of DSS were subjected to reducing SDS-PAGE and

WB. (C) Cell lysates of hTRPV1- or rTRPV1-expressing HEK293 cells were treated with or without various

concentration of SDS and subjected to non-reducing blue-native PAGE. (D-E) Reducing and non-reducing

SDS-PAGE and WB analysis of native hTRPV1 and rTRPV1 in HaCaT cells (D) and rat DRG neurons (E),

respectively.

To examine whether the inter-subunit disulfide bond of hTRPV1 could be observed in an

endogenous system, HaCaT cells, human keratinocytes known to express TRPV1 (Lee et al., 2008;

Boda et al., 2005; Toth et al., 2011), were subjected to non-reducing SDS-PAGE. TRPV1 expressed

in rat DRG was also examined as a negative control. Although hTRPV1 inter-subunit formation was

less pronounced than in the recombinant system, a DTT-sensitive hTRPV1 dimeric band was clearly

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observed in HaCaT cells (Figure 2D), whereas DTT-sensitive bands were not observed with rTRPV1

expressed in rat DRG (Figure 2E).

hTRPV1 has an inter-subunit disulfide bond between N- and C-terminal Cys residues at the

subunit interface.

An alignment of rat and human TRPV1 sequences revealed two differences in Cys residues

(Figure 3A). rTRPV1 has two extra Cys residues compared with hTRPV1, while the remaining Cys

residues are conserved among the two groups. To test whether substitution of these rTRPV1 Cys

residues into hTRPV1 alter the disulfide bond status of the hTRPV1, S31C and W616C hTRPV1

mutants were subjected to non-reducing SDS-PAGE. Neither S31C nor W616C substitution

disrupted the inter-subunit disulfide bond formation of hTRPV1 (Figure 3A). However, to our

surprise, we found that rTRPV1-mimicking double amino acid substitutions, S31C and W616C,

disrupted the inter-subunit disulfide bond of hTRPV1 (Figure 3B), suggesting that the difference in

the disulfide bond pattern between rat and human species is affected by the difference in Cys

homology. This is an intriguing case where presence, not absence, of additional Cys residues disrupts

the formation of disulfide bond.

Figure 3. Formation of inter-subunit disulfide bond between C- and N- termini in the hTRPV1 channel.

(A) Alignments of partial amino acid sequences of rat with human TRPV1 where Cys residues are not

conserved (GenBank accession number NM_018727 and NM_031982, respectively). Non-reducing SDS-

PAGE and WB analysis of HEK293 cells expressing WT, S31C, W616C, or S31C + W616C hTRPV1. (B)

Non-reducing SDS-PAGE and WB analysis of HEK293 cells expressing WT or double S31C + W616C

hTRPV1 mutant. (C) Non-reducing SDS-PAGE and WB analysis of hTRPV1 mutants with single Cys

substitution transiently expressed in HEK293T cells. The amounts of transfected vectors were adjusted to elicit

comparative expression level. The extent of inter-subunit disulfide bond formation was quantified by the ratio

between the dimer and monomer band (top) (n = 5 independent experiments). Quantitative densitometric

analysis of bands shown in the top panel (bottom). *P < 0.05 and **P < 0.01 compared to WT. (D) hTRPV1

Cys mutants with significant reductions in dimer/monomer ratio are indicated as black circles on a schematic

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topology of hTRPV1 (left). Shown is a ribbon structure model of rTRPV1 adapted from Liao et al. (3) showing

views from side (right). Cys residues are marked with green, and the residues implicated in inter-subunit

disulfide bond are marked black and indicated with arrows. Unresolved C-terminal electron density that is

predicted to contain Cys-742 is also highlighted in black.

To identify Cys residues that form inter-subunit disulfide bonds, 16 individual Cys residues in

hTRPV1 were mutated to serines and the dimer/monomer ratios were tested using non-reducing

SDS-PAGE. Mutation of Cys158, 258, 363, and 742 significantly reduced the dimer/monomer ratios

(Figure 3C). The former three Cys residues are within or vicinal to an ankyrin repeat domain (ARD)

and Cys742 was previously predicted to interface the neighboring subunit (Liao et al., 2013; Flynn et

al., 2014). ARD and the C-terminus were hypothesized to mediate subunit interactions based on the

high resolution structure (Liao et al., 2013), where ARD interfaces the ARD-S1(S1:the first

transmembrane region) linker region and the C-terminal β-strands of the neighboring subunit (Figure

3D).

To delineate the disulfide pairing of the hTRPV1 inter-subunit disulfide bond,

electrophysiological analysis was conducted to categorize the functional phenotypes of hTRPV1

mutants with disrupted inter-subunit disulfide bonds. First, a surface labeling assay confirmed the

inter-subunit disulfide bond formation of channel protein complexes at the plasma membrane

(Figure 4A). The presence of DTT, which efficiently disrupts dimerization of hTRPV1 (Figure 1D),

in the external solution caused an increase in capsaicin’s EC50 of hTRPV1, suggesting that the

disruption of disulfide bond formation reduced the sensitivity to capsaicin (Figure 4B,C). The C258S

and C742S mutations of hTRPV1 caused an increase in the EC50 to capsaicin, while C158S and

C363S mutations caused no such change (Figure 4D). Importantly, the C258S and C742S mutations

caused a similar change in the EC50 as the DTT treatment, suggesting a Cys258-Cys742 disulfide

bond pairing between hTRPV1 proteins.

Mass Spectrometric analysis of the hTRPV1 inter-subunit disulfide bond between Cys258 and

Cys742

To further delineate the disulfide bond pairing at the N- and C-termini interface, quantitative

mass spectrometric analysis was carried out. We reasoned that if a disulfide bond is formed between

Cys258 and Cys742, mutation of C742S should reduce the oxidative status of Cys258 because the

oxidizing partner Cys742 is not present in proximity. In contrast, we supposed that the oxidative

status of the other Cys residues that do not form a disulfide pair should remain relatively unaffected

(Figure 5A). The quantitation of Cys oxidation for Cys158, 258, and 363 were carried out by

masking the free (reduced) thiol with NEM and oxidized thiol with carbamidomethyl group (CAM)

and determining the peak area for the peptides with the respective chemical modifications using

selected reaction monitoring (SRM) (Figure 5B). Cys158 is spanned by proximal lysine (Lys)

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residues, which results in short peptides upon tryptic cleavage hindering identification and

quantitation by SRM. The Lys residue at position 156 was therefore mutated to glutamine (K156Q)

to optimize the protein sequence for quantitation by SRM. The inter-subunit disulfide bond of

hTRPV1 and the response against reactive disulfide were largely unchanged by the K156Q mutation

(Figure 5C,D).

Figure 4. Electrophysiology analysis of hTRPV1 with disrupted inter-subunit disulfide bond. (A) Cell

surface expression of hTRPV1-expressing HEK293 cells treated with or without 10 mM DTT for 10 min. Cell

lysates prepared after exposure to sulfo-NHS-SS-biotin were incubated with NeutrAvidin beads and subjected

to non-reducing SDS-PAGE and WB. (B) Whole-cell currents from hTRPV1-expressing HEK293 cells in

response to various concentration of capsaicin in the presence or absence of 10 mM DTT. (C) Relative

amplitudes of whole-cell currents of experiments shown in (B) (n = 3-4). EC50 are summarized on the right.

*P < 0.05 compared to DTT (–). (D) Relative amplitudes of whole-cell currents in hTRPV1 mutants-transfected

HEK293 cells in response to various doses of capsaicin (n = 3-7). EC50 are summarized on the right. *P < 0.05

and ***P < 0.001 compared to WT. The plots were fitted to the Hill equation ƒ(x) = A0 + (Amax –

A0)/(1+(EC50/x)n, where A0 is the basal response, Amax is the maximum response, x is the capsaicin

concentration, and n is the Hill coefficient. A holding potential of –80 mV was used and the current amplitudes

were normalized with the maximum current amplitudes obtained by capsaicin concentration that elicited the

largest response (I/I max). Data points are mean ± SEM.

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Following non-reducing SDS-PAGE, the dimeric hTRPV1 band and monomeric hTRPV1 C742S

band (or the respective K156Q mutants) were excised and subjected to trypsin digestion and

quantification of the CAM and NEM labeled peptides with SRM. The transitions for SRM were

selected based on MS/MS spectra for both CAM and NEM labeled peptides (Figure 5E,F) and the

largest peak area was quantified by multiple reaction monitoring (MRM) (Figure 5G,H). The

mutation of C742S substantially reduced the oxidation status of Cys258, as demonstrated by

decreased percentage of CAM labeled Cys258 containing peptides (Figure. 6A) whereas %CAM

labeled peptide for the Cys residues Cys363 and Cys158 (Figure 6B,C) were relatively unaffected

(Figure 6D). We selected to test the effect of C742S mutation on the oxidative status of Cys258

instead of the effect of C258S mutation on the oxidative status of Cys742 because the short tryptic

peptide containing Cys742 was not competent for optimization for detection by mass spectrometry

due to the disruption of hTRPV1 dimer following mutation of proximal arginine residue. These

results strongly suggest the inter-subunit disulfide bond formation occurs between Cys258 and

Cys742 of hTRPV1.

Figure 5. Quantitative mass spectrometric analyses of disulfide bond pairing. (A) Disulfide bond pairings

that are tested. (B) Schematic description of the protocol employed for elucidation of disulfide pairing. (C)

Non-reducing SDS-PAGE and WB analysis of WT and K156Q hTRPV1-expressing HEK293 cells. (D)

Maximal Ca2+ responses (∆[Ca2+]i) to 10 µM 3-nitrophenyl disulfide stimulation ( 19-33) were normalized to

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responses to 10 µM capsaicin (n = 28-42) for K156Q hTRPV1 mutant expressed in HEK293 cells. Differences

not statistically significant are labeled as (ns). (E,F) MS/MS spectra of CAM (E) and NEM (F) labeled

GRPGFYFGELPLSLAA258CTNQLGIVK peptide, which were generated from tryptic digestion of Flag-

hTRPV1 purified from HEK293T cells. The asterisks (*) denote peaks used in MRM transitions. G and H,

Representative MRM analysis of CAM (black) and NEM (grey) labeled Cys258 containing peptides.

Figure 6. An inter-subunit disulfide bond between Cys258 and Cys742 in hTRPV1 channel. (A,B) Representative SRM experiments of peptides containing Cys258 and Cys363, which were generated from

tryptic digestion of purified WT and C742S Flag-hTRPV1. (C) SRM experiments for peptides containing

Cys158, which were generated from tryptic digestion of purified Flag-hTRPV1 mutants, K156Q and

K156Q·C742S Flag-hTRPV1. The transitions used to quantify CAM (black) and NEM (grey) labeled peptides

containing Cys258, Cys363, and Cys158 are shown below. Mode of modification is followed by transitions for

the peptides containing the respective Cys residues: CAM (1354.72(z = +2) > 1124.55(b10+)) and NEM

(1388.73(z = +2) > 1124.55(b10+)) for Cys258, CAM (530.74(z = +2) > 561.25(y4+)) and NEM (564.75(z =

+2) > 629.27(y4+)) for Cys363, and CAM (749.36(z = +3) > 1146.58(y10+)) and NEM (772.04(z = +3) >

1214.61(y10+)) for Cys158. The K156Q mutation is labeled as lowercase q. The extent of Cys oxidation is

represented by %CAM labeled peptide, which is the peak area of CAM labeled peptide/(peak area of CAM

labeled peptide + peak area of NEM labeled peptide) (left). (D) SRM experiments shown in A-C are

summarized as % change in %CAM labeled peptide between WT and C742S for Cys258 and Cys363 (between

K156Q and K156Q·C742S for Cys158). Data points are mean ± SD of three analytical replicates.

Disruption of the inter-subunit disulfide bond reduces channel stability

The primary function of a disulfide bond is to provide structural support, increasing the stability

of a protein. The stability of the disulfide-less C258S and C742S hTRPV1 mutants were examined

by culturing the transfected HEK293T cells for various times in the presence of cycloheximide, an

inhibitor of protein synthesis (Whiffen, 1948; Kerridge, 1958), and monitoring the expression level of

the proteins. We found that the disulfide-less C258S and C742S hTRPV1 had shorter protein life-

spans than WT hTRPV1: protein half-life (t½ ) values for C258S, C742S, and WT were 67, 19, and

122 min, respectively (Figure 7A,B). This result suggests that the inter-subunit disulfide bond of

hTRPV1 enhance protein stability.

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Figure 7. Decreased protein stability of C258S and C742S hTRPV1 mutants. (A) HEK293T cells

expressing WT, C258S, and C742S hTRPV1 were cultured in the presence of cycloheximide for the indicated

time periods. The cell lysates were subjected to reducing SDS-PAGE and WB. The levels of hTRPV1 were

quantified through normalization by those of α-tubulin. (B) Quantitative densitometric analysis of the protein

levels of WT, C258S, and C742S normalized to those of α-tubulin (n = 3). Data points are mean ± SEM. The

plots were fitted to the exponential decay equation ƒ(t) = C + A0e-λt, where C is the constant, A0 is the initial

amount, x is the time after cycloheximide administration, and λ is the decay constant. The half-life (t1/2) was

calculated by ln(2)/ λ.

Cys residues involved in inter-subunit disulfide bond formation are also reactive to oxidation

Having elucidated the presence of the structural disulfide bond at the subunit interface, we next

asked whether this same region plays a role in sensing oxidation since the ARD and C-terminus of

TRPV1 is known to be a target of diverse modulators such as ATP and calmodulin (Lishko et al.,

2007). First, the accessibility of Cys residues localized at the subunit-interface was assessed. Flag-

hTRPV1-expressing HEK293 cells were treated with 500 µM 5-nitro-2-PDS, reduced, and modified

Cys residues were masked with NEM and CAM, respectively, followed by quantification by SRM.

We found that Cys127, 158, 258, 363, and 742 are oxidized upon treatment with 5-nitro-PDS,

suggesting that Cys residues localized at the cytoplasmic N- and C-terminal interface are at least in

part accessible by reactive disulfide oxidizing agents (Figure 8A-F). Considering that oxidizing

agents do not affect the inter-subunit disulfide bond formation of hTRPV1 (Figure 1B), these data

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also suggest that Cys258 and Cys742 of the hTRPV1 dimer possess free thiols on one subunit, and

inter-subunit disulfide bond on the adjacent subunit.

Figure 8. Cys residues at the interface are accessible by oxidants. (A-E) Cys oxidation of WT or K156Q

Flag-hTRPV1 induced by 500 µM 5-nitro-2-PDS treatment for 30 min at 37°C. Oxidized and reduced Cys of

WT or K156Q Flag-hTRPV1 (for Cys158 only) treated with or without 500 µM 5-nitro-2-PDS were masked

with CAM (black) and NEM (gray), respectively. The samples were subjected to SRM analysis following

purification from HEK293T cells and trypsinization. %CAM labeled peptides were determined for Cys127,

Cys158, Cys258, Cys363, and Cys742 using the following SRM transitions: Cys127 CAM (894.13(z = +3) >

1017.52(y172+)) and NEM (916.80(z = +3) > 1051.53(y172+)) for (A). Cys158 CAM (749.36(z = +3) >

1146.58(y10+)) and NEM (772.04(z = +3) > 1214.61(y10+)) for (B). Cys258 CAM (1354.72(z = +2) >

1124.55(b10+)) and NEM (1388.73(z = +2) > 1124.55(b10+)) for (C). Cys363 CAM (530.74(z = +2) >

561.25(y4+)) and NEM (564.75(z = +2) > 629.27(y4+)) for (D). Cys742 CAM (797.05(z = +3) > 968.43(y152+))

and NEM (819.73(z = +3) > 1002.44(y152+)) for (E). (F) Summary of %CAM labeled peptides for the

respective Cys residues. Data points are mean ± SD of three analytical replicates.

To examine whether Cys residues involved in inter-subunit disulfide bond formation are

implicated in sensing oxidation, the activity of C158S, C258S, C363S, and C742S hTRPV1 against

H2O2 was tested. C258S and C363S hTRPV1 mutants showed a significant reduction in Ca2+ rises

(∆[Ca2+]i) evoked by H2O2 when normalized to capsaicin responses, suggesting that Cys258 and

Cys363 play a role in sensing oxidation (Figure 9A). We next quantified the oxidation sensitivity of

these mutants by systematically comparing the responses of WT, C258S, and C363S hTRPV1 to a

congeneric series of reactive disulfides with differential redox potentials (Figure 1A). Consistent

with the response to H2O2, C258S and C363S mutants showed significant impairment in oxidation

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sensitivity, reflected by their activating redox potentials (redox potentials where the channel begins

to respond); ~ –2000 mV for WT, ~ –1300 mV for C363S and no response for C258S (Figure 9B).

To explore the hTRPV1 structure-function relationship, the oxidation sensitivity of Cys258 and

its neighboring Cys residues (Cys158 and Cys363) were assessed by determining the half-maximum

saturation (half-max) of H2O2 for each Cys residues by SRM analysis. The Cys258 and Cys363

showed dose-dependent relationship between H2O2 concentrations and %CAM labeled peptide,

revealing they are indeed oxidized upon treatment with H2O2 (Figure 9C,D). In contrast, Cys158

was not susceptible to oxidation at the same dosage of H2O2 as Cys258 or Cys363 (Figure 9E).

Notably, Cys258 showed the highest sensitivity to oxidation among the neighboring Cys residues,

suggesting Cys258 is the prime target of oxidation. These data are consistent with functional analysis

where C258S and C363S hTRPV1 showed impaired responses against oxidation while C158S

showed intact responses (Figure 9A,B). Moreover, these data may implicate that one of the Cys258

in the hTRPV1 disulfide-linked dimer is involved in the disulfide bond formation, while the other

Cys258 retains a free thiol that is modified by oxidizing agents. Furthermore, Cys258half-max and

Cys363half-max lie in the range of hTRPV1 activation by H2O2 and are oxidized differentially at varying

H2O2 concentration, confirming the hypothesis that the activation of hTRPV1 by oxidants is ‘graded’,

as previously proposed (Figure. 9F) (Chuang and Lin, 2009). These results suggest that Cys258 may

not only contribute to the stability of the ion channel by forming a structural disulfide bond but also

acts as an allosteric switch for channel activation (Figure 10).

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Figure 9. Quantification of hTRPV1 Cys oxidation induced by H2O2. (A) Maximal Ca2+ responses

(∆[Ca2+]i) to 1 mM H2O2 stimulation at 37°C (n = 22-100) were normalized to responses to 100 nM capsaicin

(n =12-46) for disulfide-less hTRPV1 mutants expressed in HEK293 cells. *P< 0.01 compared to WT. (B)

Oxidation sensitivity of disulfide-less hTRPV1 mutants. Plots of ∆[Ca2+]i evoked by 10 µM of the congeneric

series of reactive disulfides (See Fig. 1A) in HEK293 cells expressing hTRPV1 or hTRPV1 Cys mutants (n =

5-37). P< 0.05 compared to vector for hTRPV1 WT (*) and C363S mutant (#). Data points are mean ± SEM.

(C-E) Oxidized and reduced Cys of Flag-hTRPV1 (for Cys258 and Cys363) or Flag-hTRPV1 K156Q (for

Cys158) treated with various doses of H2O2 for 30 min at 37°C were masked with CAM and NEM, respectively.

The samples were subjected to SRM analysis following purification from HEK293T cells and trypsinization.

The half-maximal saturation (half-max) was derived by fitting the plots to the Hill equation (Cys258half-max =

0.7 mM, Cys363half-max = 2.2 mM). Hill equation ƒ(x) = A0 + (Amax – A0)/(1+(EC50/x)n, where A0 is then

basal %CAM labeled peptide, Amax is maximum %CAM labeled peptide, x is the H2O2 concentration, and n is

the Cyshalf-max. Data points are mean ± SD of three analytical replicates. F, The half-maximal saturation of H2O2

for Cys258 and Cys363 were indicated as dotted lines on a dose-response curve of H2O2 analyzed at 37°C

stimulated for 8 min (n = 41-115). Data points are mean ± SEM. *P< 0.01 compared to vector.

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Figure 10. Schematic representation of hTRPV1 activation by oxidants. Schematic of hTRPV1 tetrameric

complex viewed from the cytoplasmic side (assuming identical structure with the rTRPV1 structure reported

by Liao et al. (3)). Four subunits of hTRPV1 are shown in different colors (green, red, purple and blue) and the

respective Cys258 (black circle) and the unresolved C-terminal electron density (brown) predicted to contain

Cys742 (white circle) are indicated.

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DISCUSSION

Our combined approach of using non-reducing SDS-PAGE, electrophysiology, and mass

spectrometry consistently suggest disulfide bond formation between Cys258 and Cys742 from two

adjacent subunits. Non-reducing SDS-PAGE demonstrated that C158S, C258S, C363S, and C742S

hTRPV1 mutants have significantly impaired inter-subunit disulfide bond formation (Figure 3C).

We aimed to determine the details of Cys-based mechanisms of hTRPV1 subunits dimerization and

oxidation sensing through Cys258 and Cys742. C258S and C742S mutations in hTRPV1 caused a

similar increase in the EC50 of capsaicin as DTT treatment, which efficiently disrupts disulfide bonds

(Figure 4C,D). Quantitative mass spectrometric analysis revealed that insertion of the C742S

mutation into hTRPV1 substantially reduced the oxidative status of Cys258, corroborating that idea

that Cys258-Cys742 are the disulfide bond forming pair (Figure 6D). In addition, a high resolution

EM structure of rTRPV1 also supports disulfide bond formation between Cys258 and Cys742.

Cys258 and Cys742 in hTRPV1 subunits are likely to be in proximal to each other, considering that

C-terminal β-strands (predicted to be residues within 719-764) are interfacing the finger three and

four of the ARD where Cys257 (rTRPV1 counterpart of hTRPV1 Cys258) is positioned in rTRPV1

(Liao et al., 2013). It is highly unlikely that a disulfide bond could form between any combinations

of Cys157 (rTRPV1 counterpart of hTRPV1 Cys158), Cys257 and Cys362 (rTRPV1 counterpart of

hTRPV1 Cys363) from two adjacent subunits, where the proximity of 2.05 Å is required (Figure

3D) and the shortest distance, estimated from the near atomic structure of rTRPV1 among them is

~31.3 Å (Cys157-Cys362). However, we do not rule out the possibility of other inter-subunit

disulfide bonds formed by either Cys158 or Cys363. In fact, the mutation of any one of Cys158, 258,

363, and 742 do not eliminate the inter-subunit disulfide bond of hTRPV1, although a significant

reduction in the dimer/monomer ratios were observed (Figure 3C). Moreover, our stability data

(Figure 7B) indicates that C258S hTRPV1 mutant display reduced stability than WT hTRPV1 but

considerably higher stability than C742S hTRPV1 mutant, suggesting that an absence of Cys258

could be compensated through Cys742 interacting with other Cys residues, whereas Cys742 is

necessary for stable channel expression. Further work is necessary to fully appreciate the complexity

of oxidative modification in hTRPV1 at the subunit interface.

Interestingly, the C258S hTRPV1 mutant showed no response to oxidants (Figure 9A,B).

Moreover, mass spectrometric analysis of Cys residues of hTRPV1 treated with H2O2 showed that

Cys258 is highly sensitive to oxidation, demonstrating that Cys258 confers oxidation sensitivity to

hTRPV1 (Figure 9C). These results showed that Cys258 residues are heterogeneously modified in

the hTRPV1 tetrameric complex and comprise Cys258 with a free thiol for oxidation sensing and

Cys258 involved in the disulfide bond formation for assisting in subunit dimerization. Thus, the

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homotetrameric hTRPV1 channel is essentially a heterotetrameric channels in terms of both redox

status and function (Figure 10).

The TRPV1 tetrameric complex is thought to have a four-fold symmetry centering the channel

pore under the reducing conditions (Liao et al., 2013). If this is true, all four Cys258 on each TRPV1

subunits should have equal chance of being oxidized and thus should result in disulfide-linked

tetramer. However, our data show that TRPV1 forms disulfide-linked dimer. In order to obtain

tetrameric complex consisting of two disulfide-linked TRPV1 dimers, the inter-subunit disulfide

bonds must be arranged in the opposing position (Figure 10). Considering that TRPV1 do not form

disulfide-linked tetramer even under the extreme oxidizing environment (such as treatment with

oxidizing agent (Figure 1B), it is possible that four-fold symmetry of TRPV1 is not present in human

species and that TRPV1 possesses heterogeneity in subunit orientation. Although our current data is

limited in drawing definitive conclusion concerning the three dimensional structural orientation

underlying heterogeneous oxidation, we speculate that hTRPV1 is susceptible to heterogeneous

oxidation due to heterogeneity in orientation of Cys258 on each subunit. Ion channels are known to

possess disulfide bonds to provide a structural scaffold or redox sensitivity. These include ion

channels such as NMDA receptor, CLIC1, Kir2.2, cardiac ryanodine receptor, TWIK-1 channel, and

TRPC5 ( Sullivan et al., 1994; Lesage et al., 1996; Wo and Oswald, 1996; Cho et al., 2000; Littler et

al., 2004; Zissimopoulos et al., 2013; Hong et al., 2014). However, to our knowledge, this is the first

observation in which an ion channel uses the redox heterogeneity of a single specific Cys residue

(Cys258) to provide both oxidation sensitivity and protein stability. Because of the unexplored field

of Cys biochemistry in membrane proteins, we believe that this type of functional bifurcation is likely

to be found in other channels in the near future.

The inter-subunit disulfide bond of hTRPV1 was highly stable, demonstrated by rapid

reformation following reduction by DTT (Figure 1E). This may explain how cytoplasmic inter-

subunit disulfide bond may have survived the reductive intracellular milieu. Consistent with our

observation, Tao et al. reported that the preparation of Kir2.2 crystal structure in the presence of 20

mM DTT and 3 mM tris-(2-carboxyethyl) phosphine (TCEP) still yielded Kir2.2 with a disulfide

bond, suggesting that disulfide bonds can be a stable structural feature of ion channels (Tao et al.,

2009).

In our study, it is also important to note that the oxidative status of Cys residues was characterized

based on the reactivity of free Cys residues with NEM and the reversibility of oxidized Cys residues

with DTT. As a consequence, oxidative Cys modifications including sulfenic acid, disulfides,

glutathionylation and nitrosylation were not distinguished (Reisz et al., 2013). In addition, because

the irreversible oxidative modification such as sulfonic or sulfinic acid cannot be detected with the

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methodology employed, these modifications were also not accounted for in our analysis.

In the current study, we focused primarily on the elucidation of an oxidant induced activation

mechanism at the N- and C-terminal interface of hTRPV1. However, as described previously by

others for cTRPV1, other domains in the TPRV1 channels are likely to be involved in sensing

oxidation and the complete picture of oxidation sensing requires characterization of Cys residues in

these domains (Chuang and Lin, 2009). Although this study describes Cys modification as the

primary mechanism of activation by oxidants, it is highly likely that other residues, such as Lys

residue and methionine residue are also targeted by oxidants. In fact, a Lys residues in TRPA1 is a

known target of electrophiles and methionine residues in TRPM2 is implicated in sensing H2O2

(Hinman et al., 2006; Kashio et al., 2012).

We found that oxidative post-translational modifications between hTRPV1 and rTRPV1 are

significantly different, as rTRPV1 lacks the propensity to form an inter-subunit disulfide bond

(Figure 1B). This suggests that the structure and function between hTRPV1 and rTRPV1 is not

completely conserved for the oxidizing stimuli. Thus, one may need to be cautious when applying a

conclusion to both rTRPV1 and hTRPV1. This is also true for cTRPV1, where oxidation-induced

activation was correlated with dimerization (Wang and Chuang, 2011). Interestingly, hTRPV1

endogenously expressed in HaCaT cells is subjected to multiple redox modifications (as implicated

by multiple DTT-sensitive bands), in stark contrast with rTRPV1 expressed in DRG neurons where

only the monomeric protein was observed under the non-reducing SDS-PAGE (Figure 2D,E).

Functional characterization and clarification of the significance of unique redox modifications in

hTRPV1 are largely unexplored and appear to be a fertile field for further investigation.

To our surprise, we found that rTRPV1-mimicking double amino acid substitutions, S31C and

W616C, disrupted the inter-subunit disulfide bond of hTRPV1 (Figure 3B), suggesting that the

difference in the disulfide bond pattern between rat and human species is affected by the difference

in Cys homology. This is an intriguing case where presence, not absence, of additional Cys residues

disrupts the formation of disulfide bond. There could be two possibilities underlying this

phenomenon: 1) the presence of Cys31 and Cys616 alters the oxidative folding at the ER and

consequent disulfide bond patterns or 2) the Cys31 and Cys616 react with Cys residues that otherwise

form disulfide bond and prevents the formation of inter-subunit disulfide bond. In the near-atomic

EM structure of TRPV1, both Cys31 and Cys616 are not resolved, preventing us from drawing

inferences from the current available structural information (Liao et al., 2013). Further investigation

of the mechanism underlying the disruption of disulfide bond by insertion of Cys31 and Cys616 will

enhance our understanding of biochemistry of TRPV1. TRPV1 is a polymodal sensor that has diverse

physiological functions in addition to nociception, and general blockage of TRPV1 function for

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treatment of pain has caused serious side effects in vivo (Patapoutian et al., 2009). Detailed

understanding of the activation mechanisms is necessary to achieve selective regulation of TRPV1

in a modality specific manner. Our findings improve the understanding of activation of hTRPV1 by

oxidation and provide new insight to enable the design of modality-specific drugs for treating pain.

111

Acknowledgments: We thank H. Atomi, M. Hirano, M.X. Mori, R. Sakaguchi for experimental

advice and helpful discussions. This work was supported by a Grant-in-Aid for Scientific Research

on Innovative Areas “Oxygen biology: a new criterion for integrated understanding of life”

[26111004] of The Ministry of Education, Culture, Sports, Science and Technology, Japan; and a

Grant-in-Aid for Scientific Research (A) [24249017] of Japan Society for the Promotion of Science.

112

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118

General Conclusion

The findings in the first three chapters identified and add substantially to our knowledge about

the physiological, molecular, and pathological role of TRPC1, TRPV1, TRPM2, and TRPM7 among

oxidation status-sensitive TRP channels.

Different contribution of redox-sensitive transient receptor potential channels to

acetaminophen-induced death of human hepatoma cell line

The current work for the first time systematically and comprehensively compares the significance

of redox-sensitive TRP channels in APAP overdose-induced hepatocyte response, representing an

outstanding advance in our current understanding of the mechanism of APAP-induced liver toxicity.

We investigated the mechanism of acetaminophen (APAP)-induced liver damage. We investigated

also the contribution of reactive oxygen species, and the involvement of calcium entry via TRP

channels, to apoptotic cell death induced by relatively high concentrations of APAP. It was

demonstrated that the addition of ROS scavengers and pharmacological blockade or siRNA-

mediated reduction of TRP channel activity reduced cell death. Specifically, reduction of TRPV1

and TRPC1 activity was the most effective at reducing cell death. Based on biotinylation of these

channels by sulfhydryl-selective reagents, and reduction of biotinylation by APAP, it is suggested

that redox activation of these channels causes toxicity by influx of calcium.

TRPM2 regulates the development of rheumatoid arthritis in the anti-collagen-induced mouse

model

Rheumatoid arthritis (RA) is a good example of a ROS-related chronic inflammatory disease that

is difficult to treat. However, the molecular mechanisms that connect ROS and RA are unclear. This

study reveals that ROS-sensitive TRPM2 plays an essential role in the development of anti-type II

collagen antibody-induced arthritis (CAIA). TRPM2 is a member of the transient receptor potential

protein (TRP) family and its gene encodes a plasma membrane calcium ion channel. First we

identified the sequence of TRPM2 knockout mice and showing the deletion exons 17, 18 & 19 leads

to formation of stop codon. TRPM2 knockout mice is composed of 834 amino acid residues.

Compared with the wild-type mice with CAIA, paw swelling, inflammation in joints, bone erosions,

and inflammatory cytokine production were prominently suppressed in Trpm2-deficient mice,

despite the localization of collagen antibody on cartilage surface. These results implicate that

functional inhibition of TRPM2 is a novel therapeutic strategy for treating RA. We suggest TRPM2

knockout mice serve as a valuable animal model for drug discovery, in screening, developing and

clarifying mechanisms of various inflammatory diseases and possible roles of oxidative stress-

activated TRPM2 channels in the development of rheumatoid arthritis.

119

Functional and structural divergence in human TRPV1 channel subunits by oxidative cysteine

modification

TRPV1 channel is a tetrameric protein which acts as a sensor for noxious stimuli such as heat,

and for different inflammatory mediators such as oxidative stress to mediate nociception in a subset

of sensory neurons. In TRPV1 oxidation sensing, cysteine (Cys) oxidation has been considered as

the principle mechanism; however, its biochemical basis was previously known. We characterized

the oxidative status of Cys residues in differential redox environments and identified a formation of

subunit dimers carrying a stable inter-subunit disulfide bond between Cys258 and Cys742 of

hTRPV1. Our results suggest that Cys258 residues are heterogeneously altered in the hTRPV1

tetrameric complex and constitute Cys258 with free thiol for oxidation sensing and Cys258, which

is involved in the disulfide bond for assisting subunit dimerization. Thus, the hTRPV1 channel has

a heterogeneous subunit composition in terms of both redox status and function.

120

List of Publications

a) Articles

1. Different contribution of redox-sensitive transient receptor potential channels to

acetaminophen-induced death of human hepatoma cell line. (Chapter 1)

Heba Badr, Daisuke Kozai, Reiko Sakaguchi, Tomohiro Numata, and Yasuo Mori

Frontiers in Pharmacology Feb 2016. (Chapter 1)

2. TRPM2 regulates the development of rheumatoid arthritis in the anti-collagen-induced

mouse model.

Manuscript in Preparation. (Chapter 2).

3. Functional and structural divergence in human TRPV1 channel subunits by oxidative

cysteine modification.

Nozomi Ogawa, Tatsuki Kurokawa, Kenji Fujiwara, Onur Kerem Polat, Heba Badr, Nobuaki

Takahashi, and Yasuo Mori

Journal of Biological Chemistry Feb 2016. (Chapter 3)

b) International Conferences

1. Identification of Redox–Sensitive TRP Channels as a key Pathway in Paracetamol induced

Hepatocellular Damage.

Heba Badr, Tomohiro Numata, and Yasuo Mori

NIPS International Workshop, TRPs and SOCs ~ Unconventional Ca2+ Physiology. Nagoya,June

2015, oral presentation.

c) Japanese Conferences

1. Characterization of redox-sensitive TRP Channels in acetaminophen-induced death of

human hepatoma cell line.

Heba Badr, Tomohiro Numata, and Yasuo Mori

88th Annual Meeting of Japanese Pharmacological Society, Nagoya, March, 2015, poster

presentation.