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
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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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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.,
9
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.
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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).
77
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.
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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.