human multidrug and toxin extrusion protein 1 ...final approval and acceptance of this dissertation...
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Human Multidrug and Toxin ExtrusionProtein 1: Symmetry of substrate fluxes
Item Type Electronic Dissertation; text
Authors Dangprapai, Yodying
Publisher The University of Arizona.
Rights Copyright © is held by the author. Digital access to this materialis made possible by the University Libraries, University of Arizona.Further transmission, reproduction or presentation (such aspublic display or performance) of protected items is prohibitedexcept with permission of the author.
Download date 08/09/2021 18:45:18
Link to Item http://hdl.handle.net/10150/145435
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HUMAN MULTIDRUG AND TOXIN EXTRUSION PROTEIN 1:
SYMMETRY OF SUBSTRATE FLUXES
by
Yodying Dangprapai
Copyright © Yodying Dangprapai 2011
A Dissertation Submitted to the Faculty of the
PHYSIOLOGICAL SCIENCES
GRADUATE INTERDISCIPLINARY PROGRAM
In Partial Fulfilment of the Requirements for the Degree of
DOCTOR OF PHILOSOPHY
In the Graduate College
THE UNIVERSITY OF ARIZONA
2011
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THE UNIVERSITY OF ARIZONA GRADUATE COLLEGE
As members of the Dissertation Committee, we certify that we have read the dissertation prepared by Yodying Dangprapai entitled Human Multidrug and Toxin Extrusion protein 1: Symmetry of substrate fluxes and recommend that it be accepted as fulfilling the dissertation requirement for the Degree of Doctor of Philosophy
Date: 6/17/2011 Stephen H. Wright
Date: 6/17/2011 William H. Dantzler
Date: 6/17/2011 Nicholas A. Delamere
Date: 6/17/2011 Heddwen L. Brooks Final approval and acceptance of this dissertation is contingent upon the candidate’s submission of the final copies of the dissertation to the Graduate College. I hereby certify that I have read this dissertation prepared under my direction and recommend that it be accepted as fulfilling the dissertation requirement.
Date: 6/17/2011 Dissertation Director: Stephen H. Wright
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STATEMENT BY AUTHOR
This dissertation has been submitted in partial fulfillment of requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library. Brief quotations from this dissertation are allowable without special permission, provided that accurate acknowledgment of source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the copyright holder.
SIGNED: Yodying Dangprapai
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ACKNOWLEDGEMENTS
To Dr. Wattana Watanapa (Department of Physiology, Faculty of Medicine Siriraj hospital, Mahidol University, Bangkok, Thailand) who introduced me to the Physiological Sciences (PS) program at the University of Arizona. She also introduced me to Dr. William H. Dantzler who facilitated my admission to the PS program. To all faculty members who shared their knowledge and scientific skills during the years of coursework and research. Particularly, I would like to thank my committee members for the qualifying examination, including Dr. William H. Dantzler, Dr. Janis M. Burt, Dr. Heddwen L. Brooks, Dr. Alexander M. Simon, and Dr. Stephen H. Wright for your patience and kindness. To friends and colleagues in Tucson. Ryan; I would graduate earlier if I did not take your advice (I know that I asked for it), but working with you was so much fun. Taka; I wish you could have stayed here longer. Bethzaida; I had a good time sharing (political) opinions with you. Xiaohong; It was amazing how you cloned all those fifty-something mutants in no time (It would take me forever). I appreciate your help, including many trips to Costco. Theresa; thank you for keeping our lab organized. I know doing that can drive one crazy, so I wish you a peaceful retirement. Mark; although I prefer a nice and quiet lab, I have to (unwillingly) admit that our lab is too quiet when you are not around. Also, I would like to thank Kris for her kindness and Amrit for his assistance in Dr. Delamere’s lab. To friends back in Thailand for ongoing encouragement, especially to Dr. Suthammarak, Dr. Sewatanon, and Dr. Srisuma for all the phone conversations we shared. Additionally, I would like to thank Mr. Wiggins for being a generous landlord. You are very brave to let me take care of your house and your puppy. Susie is such wonderful company. To the Thai community in the University of Arizona for all the delicious lunches and dinners we shared. Importantly, I would like to express my gratitude to Dr. Supapan Seraphin (Faculty of Engineering, University of Arizona) for her kindness and her wisdom that kept me moving forward during a difficult time. “��������ก���” To the Department of Physiology, Faculty of Medicine Siriraj hospital and to the PS GIDP program, especially Holly A. Lopez, for this invaluable opportunity, for the financial support and for all administrative assistance. To my family for your trust in me, all along, and last but not least, ‘Thank you very much, Dr. Wright!’ Yodying Dangprapai June 2011
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TABLE OF CONTENTS
LIST OF FIGURES.................................................................................................8
LIST OF TABLES.................................................................................................10
LIST OF ACRONYMS.........................................................................................11
ABSTRACT...........................................................................................................12
CHAPTER 1: INTRODUCTION
1.1 Organic cations and the human body...........................................................13
1.2 Organic cations and the kidney....................................................................15
1.3 A cellular model of renal organic cation secretion......................................15
1.4 Study of organic cation transport in the kidney...........................................16
1.5 Cloning of the basolateral organic cation transporter..................................18
1.6 hOCT2 as a major contributor for renal basolateral OC uptake.................19
1.7 A molecular identity for the apical organic cation efflux transporter..........21
1.8 The Multidrug And Toxin Extrusion (MATE) transporters........................24
1.9 Cloning of MATE proteins in humans and other mammals........................25
1.10 The physiological fingerprint of MATE-mediated transport.....................26
1.11 Structure of MATE transporters................................................................27
1.12 Kinetics and Selectivity of MATEs...........................................................28
1.13 Kinetics and Selectivity of MATE-mediated transport:
uptake vs. efflux.........................................................................................28
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TABLE OF CONTENTS – Continued
CHAPTER 2: MATERIALS AND METHODS
2.1 Chemicals...................................................................................................31
2.2 Cloning of the hMATE1 gene....................................................................31
2.3 Expression of hMATE1 in CHO Flp-In cells.............................................31
2.4 Measurement of [3H]MPP uptake (for time course and kinetic studies)
into CHO hMATE1 cells............................................................................32
2.5 Measurement of MPP efflux from CHO hMATE1 cells............................33
2.6 Manipulation of intracellular pH................................................................35
2.7 Measurement of CHO cell volume.............................................................36
2.8 Statistics......................................................................................................37
CHAPTER 3: SYMMETRY OF INTERACTION FOR H+ AND hMATE1
3.1 Background.................................................................................................38
3.2 Cis-inhibition by extracellular H+ of hMATE1-mediated OC uptake........40
3.3 Characterization of hMATE1-mediated OC efflux....................................43
3.4 Interaction of intracellular H+ with the cytoplasmic face of hMATE1......48
3.5 Interaction between H+ and hMATE1: relevance to renal OC secretion....51
CHAPTER 4: AN APPLICATION OF THE EFFLUX PROTOCOL TO THE
STUDY OF hMATE1-MEDIATED OC INFLUX
4.1 Background................................................................................................56
4.2 Trans-stimulation of hMATE1-mediated OC efflux by MPP...................57
4.3 Trans-stimulation of hMATE1-mediated OC efflux by TEMA................59
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TABLE OF CONTENTS – Continued
CHAPTER 5: FUTURE DIRECTION................................................................61
REFERENCES....................................................................................................84
APPENDIX A: INTERACTION OF H+ WITH THE EXTRACELLULAR AND
INTRACELLULAR ASPECTS OF hMATE1....................................................91
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LIST OF FIGURES
Figure 1: Chemical structure of selected organic cations (OCs)..........................64
Figure 2: Molecular mechanism of renal organic cation secretion ......................65
Figure 3: Predicted membrane topology of hOCT2.............................................66
Figure 4: Predicted membrane topology of rbMATE1.........................................67
Figure 5: hMATE1-mediated MPP transport: uptake vs. efflux...........................68
Figure 6: The culture dish insert...........................................................................69
Figure 7: Time course and kinetics of [3H]MPP uptake in CHO hMATE1 cells..70
Figure 8: IC50 of extracellular H+ as an inhibitor of hMATE1-mediated MPP
uptake....................................................................................................71
Figure 9: Time course of MPP loss from CHO hMATE1 cells............................72
Figure 10: Time course of NBD-TMA efflux in CHO hMATE1 cells.................73
Figure 11: Effect of extracellular H+ on hMATE1-mediated [3H]MPP efflux......74
Figure 12: Effect of increasing extracellular [H+] on the kinetics of
hMATE1-mediated [3H]MPP efflux...................................................75
Figure 13: Intracellular pH (pHi) in CHO hMATE1 cells....................................76
Figure 14: Effect of NH4Cl and MPP preload on hMATE1-mediated [3H]MPP
uptake..................................................................................................77
Figure 15: Effect of NH4Cl on hMATE1-mediated [3H]MPP efflux...................78
Figure 16: Kinetics of interaction between inward-facing hMATE1 and
intracellular H+...................................................................................79
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LIST OF FIGURES – Continued
Figure 17: Effect of increasing extracellular [MPP] on the kinetics of
hMATE1-mediated [3H]MPP efflux..................................................80
Figure 18: Effect of extracellular TEMA on hMATE1-mediated [3H]MPP
efflux..................................................................................................81
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LIST OF TABLES
Table 1: Selected organic cations of clinical or environmental significance…..82
Table 2: Effect of extracellular pH on the kinetics of hMATE1-mediated MPP
uptake...................................................................................................83
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LIST OF ACRONYMS
BBMV brush border membrane vesicle
CHO cells Chinese hamster ovary cells
CHO hMATE1 cells Chinese hamster ovary cells expressing hMATE1
MATE multidrug and toxin extrusion protein
MDR multidrug resistance protein
MPP 1-methyl-4-phenylpyridinium, a prototypic OC
NBD-TMA [2-(4-nitro-2,1,3-benzoxadiazol-7-yl)
aminoethyl]trimethylammonium, a fluorescent OC
NHE sodium-hydrogen exchanger
NMN N-methylnicotinamide
OC organic cation
OCT organic cation transporter
OCTN organic cation transporter novel
RPT renal proximal tubule
RPTEC renal proximal tubule epithelial cell
T2O tritiated water
TBuA tetrabutylammonium
TEA tetraethylammonium, a prototypic OC
TEMA triethylmethylammonium
TPeA tetrapentylammonium
WB Waymouth’s buffer
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ABSTRACT
Human multidrug and toxin extrusion 1 (hMATE1) is a major candidate for being
the molecular identity of organic cation/proton (OC/H+) exchange activity in the luminal
membrane of renal proximal tubules (RPT). Although physiological function of
hMATE1 supports luminal OC efflux, the kinetics of hMATE1-mediated OC transport
have typically been characterized through measurement of uptake i.e., the interaction
between outward-facing hMATE1 and OCs. To examine kinetics of hMATE1-mediated
transport in a more physiologically relevant direction i.e., an interaction between inward-
facing hMATE1 and cytoplasmic substrates, I measured the time course of hMATE1-
mediated efflux of the prototypic MATE1-substrate, [3H]1-methyl-4-phenylpyridinium
([3H]MPP), under a variety of conditions, including different values for intra- and
extracellular pH, from CHO cells that stably expressed hMATE1. I showed that an
IC50/K i for interaction between extracellular H+ and outward-facing hMATE1 determined
from conventional uptake experiments [12.9 ± 1.23 nM (pH 7.89); n = 9] and from the
efflux protocol [14.7 ± 3.45 nM (pH 7.83); n = 3] were not significantly different (P =
0.6). To test a hypothesis that H+ interacts symmetrically with each face of hMATE1,
kinetics of interaction between intracellular H+ and inward-facing hMATE1 were
determined using the efflux protocol. The IC50 for interaction with H+ was 11.5 nM (pH
7.91), consistent with symmetrical interactions of H+ with the inward-facing and
outward-facing aspects of hMATE1. The efflux protocols demonstrated in this study are
a potential means to examine kinetics at cytoplasmic face of hMATE1 and also a
practical tool to screen uptake of substrates at extracellular face of hMATE1.
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CHAPTER 1
INTRODUCTION
The human body synthesizes and consumes organic electrolytes on a daily basis.
Once distributed in the body at physiological pH, each of these organic electrolytes
(OEs), as indicated by the name, will typically behave as organic cations (OCs) or
organic anions (OAs). Although various endogenous OEs have significant roles in
homeostasis of the body, my interest is on how the body handles various exogenous i.e.,
xenobiotic, and toxic OEs. Within the scope of this dissertation, the focus is on renal
handling of OCs and, in particular, on an OC efflux transporter at the apical membrane of
the renal proximal tubule (RPT), namely, the Multidrug And Toxin Extrusion protein,
MATE1. Following is an overview of topics related to this issue:
1.1 Organic cations and the human body
Organic cations (OCs) are typically nitrogenous compounds (Fig. 1) that carry net
positive charge(s) at physiological pH, i.e., pH 7.35 to pH 7.45. The human body
synthesizes a wide array of physiologically important OCs, including choline, dopamine,
norepinephrine, and epinephrine. Nevertheless, it is likely that most OCs to which the
body is exposed are exogenous i.e., xenobiotic, and many of these can be toxic.
Importantly, the human body consumes toxic OCs on a daily basis, mostly in the form of
various plant alkaloids (71). This can be viewed from an evolutionary perspective as an
example of “chemical warfare” between plants and animals. In order to survive and to
be able to pass their genes to the next generation, plants synthesize various chemicals that
are toxic to animals, especially to those not participating in pollination. As an adaptation
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to the efforts of plants to dissuade the animals that eat them, only animals that are
equipped with a powerful system to handle botanical toxins will survive and be able to
produce offspring (1).
Besides plant products, OCs increasingly appear in modern daily life in the form
of drugs. In fact, approximately 40% of current prescribed medications are OCs (23)
(Table 1). Furthermore, there are increasing numbers of unwanted drug-drug interactions
(DDIs) with mild to fatal results in patients under treatment with multiple OC drugs (40).
Thus, it is crucial to understand how the body is cleared of toxic OCs. In this regard, the
two major organs responsible for the challenging task of OC secretion are the liver and
the kidney. For this dissertation, the focus is on handling of OCs by the kidney. The
interested reader is directed to recent reviews that describe OC transport by hepatocytes
(e.g., (24).
Before further discussion on renal handling of OCs, it is important to address a
definition of ‘OCs’ used in this study. It is very useful to classify OCs into two subtypes
based on structure and physicochemical properties (71). First, ‘type I’ OCs refer to
comparatively small (MW< 400 Da), and generally monovalent compounds, such as
tetraethylammonium (TEA) and 1-methyl-4-phenylpyridinium (MPP). Second, ‘type II’
OCs refer to larger, and frequently polyvalent, compounds that are also typically
comparatively hydrophobic. In this study, unless otherwise indicated, the focus is on a
renal handling of ‘type I’ OCs.
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1.2 Organic cations and the kidney
The role of the kidney as a site for OC secretion was first demonstrated in 1947.
Sperber (45) showed evidence in birds for a renal secretion of N-methylnicotinamide
(NMN), an organic cation by-product from metabolism of nicotinamide (vitamin B3). In
the same year, Rennick et al. (47) demonstrated in dogs that renal secretion has a role in
excretion of tetraethylammonium (TEA), a prototypic OC, by the kidney. After these
early reports of a role for the kidney in OC secretion, subsequent studies on renal OC
transport were conducted using a variety of experimental systems, including whole
animals, kidney slices, isolated perfused renal tubules, and renal membrane vesicles.
Results from the studies between the 1950s and the 1970s (reviewed in (45) advanced the
knowledge of renal OC handling toward an understanding of the tubular mechanism for
renal OC transport. The major findings included: (i) renal OC transport is mediated by
unique transporter protein(s) distinct from a previously described transport system for
para-aminohippurate (PAH), a prototypic organic anion (OA); and (ii) a major site for
renal OC secretion is the early portion of the renal proximal tubule (RPT).
1.3 A cellular model of renal organic cation secretion
In 1980-81, based on their work with renal cortical membrane vesicles, Holohan
and Ross published two consecutive articles that described a mechanism for a basolateral
(peritubular) OC uptake that is distinct from a mechanism for an apical (luminal) OC
efflux (18, 19). Furthermore, they suggested that the transporter(s) responsible for apical
OC efflux is functionally coupled to activity of the apical Na+/H+ exchanger (NHE) to
facilitate OC secretion. Three years later, Holohan and Ross (48) proposed a cellular
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model for OC secretion in the RPT (Fig. 2). In summary, renal OC secretion is a two-
step process. First, at the basolateral membrane of renal proximal tubule epithelial cells
(RPTECs), OC uptake from the blood is mediated by a single process that operates as (i)
facilitated diffusion, driven by an inside negative membrane potential, or as (ii) OC
exchange with another cytoplasmic OC. Second, at the apical membrane, OC efflux into
the tubular filtrate is mediated by a transport process that exchanges an OC for H+ that is
supplied continuously from the filtrate (and thus influenced by activity of NHEs). It is
important to acknowledge that this cellular model of renal OC secretion proposed by
Holohan and Ross is still valid and widely referred to, despite the fact that the body of
knowledge on OC transport has grown significantly as a result of advanced molecular
biology techniques during the past 20 years.
1.4 Study of organic cation transport in the kidney
Between the mid-1980s and the early-1990s, after Holohan and Ross proposed the
model for renal OC secretion, most studies in this area focused mainly on testing
hypotheses concerning the mechanism of OC/H+ exchange because of its accepted role as
the rate-limiting step in renal OC secretion. Following is a summary (see Pritchard and
Miller, 1993) of observations from selected studies using renal BBMV that are
particularly pertinent to this dissertation and its focus on apical OC efflux in RPTEC:
● Apical OC/H+ exchange is an obligatory exchange process i.e., substrate (OC or
H+) is required for the conformational change(s) of the protein responsible for this
process in both directions across the cell membrane.
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● A gradient of H+ drives OC transport in the opposite direction and is capable of
net OC movement against an electrochemical gradient. For example, in renal BBMV, an
outwardly-directed H+ gradient can support concentrative vesicular OC uptake.
● Results from uptake studies of monovalent OCs in BBMV suggested that the
typical stoichiometry of this exchange process is one-to-one. In other words, OC/H+
exchange is electroneutral.
In addition to studies on the mechanism of OC secretion, some workers focused
on substrate selectivity of both basolateral OC uptake and apical OC efflux transport
processes. In summary, for the basolateral membrane, an OC with higher
hydrophobicity, stronger ionic strength, or with amino group(s) on the benzene ring will
penetrate into the cells more effectively than one without these characteristics (61). For
the apical membrane, higher hydrophobicity and basicity are correlated with better
transport (61).
Up until the early-1990s, models for studying renal OC transport, both basolateral
uptake and apical efflux, were limited to preparations from the kidney as mentioned
previously, e.g., renal membrane vesicles and isolated proximal tubules. Although data
from study of native kidney tissues are invaluable, conclusions regarding a role for a
specific transport protein in renal OC secretion could not be readily drawn using such
physiologically complex systems. Yet it was crucial to identify the specific proteins
responsible for OC transport in the renal proximal tubule to form the basis of
understanding the molecular mechanisms for renal handling of OCs. Below I provide a
brief overview of the molecular characteristics of the basolateral entry step in renal OC
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secretion, followed by a more extensive discussion of the molecular characteristics of the
apical transport process, MATE1, which is the focus of this dissertation.
1.5 Cloning of the basolateral organic cation transporter
In 1994, Gründemann et al. (12) successfully cloned, from rat kidney, a protein
capable of transporting structurally diverse OCs, and named the protein: organic cation
transporter 1 (rOct1; Slc22a1). Importantly, rOct1 demonstrated kinetic and selectivity
profiles similar to those of basolateral OC transport characterized in studies of tubules in
intact rat kidney, and basolateral membrane vesicles from rat kidney. For example,
affinity of NMN and tetramethylammonium (TMA) for rOct1 expressed in Xenopus
oocytes (12) was similar to affinity of each OC from kinetic studies in renal proximal
tubule (62). It is obvious that cloning of OCTs provided an opportunity for the direct
characterization of specific transport proteins both functionally (kinetically) and
structurally. The landmark study by Gründemann et al. led to the subsequent cloning of
several other organic cation transporter homologs and orthologs, including rOct2;
Slc22a2 (41) and the human (10) and rabbit orthologs (57) of these proteins.
Importantly, it is clear from studies of these different OCT orthologs that there are
marked species differences regarding substrate selectivity, kinetics, and sites of
expression (76). For example,the interaction between hOCT1 and tetraalkylammonium
compounds, such as TEA, is significantly distinct from the interaction between rodent
and rabbit Oct1 orthologs and these compounds (76). Also, whereas both Oct1 and Oct2
are expressed in rat and mouse kidney (71), expression in human kidney is effectively
restricted to OCT2 (39). Thus, to clearly understand handling of OCs in human, it is
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important to study hOCTs directly, or to (at least) be aware of inter-species differences
when interpreting kinetic information gained in studies with orthologs of hOCTs.
1.6 hOCT2 as a major contributor to renal basolateral OC uptake
Human OCTs e.g., hOCT1 (SLC22A1) and hOCT2 (SLC22A2) were successfully
cloned in 1997. First, Koehler et al. reported a location of genes coding for hOCT1 and
hOCT2 on chromosome 6 (25), and later, both transporters were successfully cloned (10)
and characterized in various heterologous expression systems, including Xenopus oocytes
(10), HeLa cells (76), HEK-293 cells (16) , CHO cells (3), MDCK cells (63), and
proteoliposomes (21). As a result of the cloning of the OCTs, our understanding of
basolateral OC uptake increased significantly during the late-1990s to the mid-2000s.
Following is a list of selected information regarding expression profile, structure and
kinetics of OCTs:
● OCTs display comparatively high levels of expression in many epithelial
tissues, including renal proximal tubule and liver. In the human, OCT2 is the major OCT
expressed at the basolateral membrane of RPTECs, based on expression of both mRNA
and protein (39). Expression of OCT1 in human kidney is less than 1% of the level for
hOCT2 (72). OCT1 is, however, the major OCT expressed at the sinusoidal membrane
of the human liver (32). In rat, mouse, and rabbit, the kidney expresses both Oct1 and
Oct2 (71). Importantly, from a comparison of renal TEA clearance in wild-type mice
and in mice lacking both homologs of Oct, it is evident that renal secretion of TEA is
accounted for quantitatively by the combined activity of mOct1 and mOct2 (20).
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● When expressed in various heterologous expression systems, OCTs
demonstrate an electrogenic OC transport across the plasma membrane (8). Additionally,
OC uptake mediated by OCTs is driven by an inside-negative membrane potential (7).
Furthermore, OCTs can operate as an OC/OC exchanger (8), but an interaction with a
substrate is not required for a conformational change of OCTs. In other words, OCTs do
not operate as an obligatory exchanger (8). Importantly, changes in membrane potential,
but not in the transmembrane gradient of Na+ or H+, affects kinetics of OCT-mediated
transport.
● OCTs interact with a structurally diverse array of substrates and inhibitors, in
agreement with the hallmark of renal OC secretion as a polyspecific transport system
(61). Computationally derived pharmacophores for OCT1 (5) and OCT2 (54) identified
several key molecular determinants of effective substrate/inhibitor interaction with OCTs,
including the placement of hydrophobic mass relative to a basic nitrogen, and the
influence of specific molecular shape and size of a substrate.
● OCTs are members of the Major Facilitator Superfamily of the transport
proteins. They contain 550 to 560 amino acid residues with a membrane topology of
twelve transmembrane spanning domains (TMDs), which includes intracellular N- and C-
termini, and a long cytoplasmic loop between TMDs 6 and 7, and a very long
extracellular loop between TMDs 1 and 2 (12). Figure 3 illustrates proposed secondary
and tertiary structures of hOCT2 (44).
In summary, based on its expression and selectivity profiles (71), OCT2 is widely
regarded as the major contributor to OC uptake at the basolateral membrane of RPTEC in
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human kidneys. hOCT2-mediated OC uptake is driven by an inwardly-directed
electrochemical gradient for these cationic substrates, reflecting the inside-negative
membrane potential of RPTECs. Although this basolateral OC uptake is a passive
process, at steady state, OC uptake into the RPTEC, mediated by hOCT2, is capable of
accumulating an OC to a level higher (some 10- to 15-fold) than its concentration in the
peritubular capillary. As discussed in an upcoming section, this fact will prove to play a
critical role in development of an integrated model of renal OC secretion that involves
basolateral OC uptake and MATE-mediated OC efflux in RPTECs.
1.7 A molecular identity for the apical organic cation efflux transporter
The molecular identity for a transporter protein(s) responsible for apical OC
efflux proved to be more elusive than that for basolateral OC entry (the OCTs, described
previously). The physiological characteristics of apical OC transport were known from
studies performed in renal brush border membrane vesicles (BBMV), or in isolated
perfused proximal tubules prepared from the kidney. This “physiological fingerprint”
can be summarized as follows:
● Apical OC efflux is the active and a rate-limiting step for the net OC secretion
across the RPT epithelium (72), based on observations from isolated rabbit RPT that
showed that (i) the rate of basolateral OC uptake is similar along the length of the RPT,
whereas the rate of apical OC efflux decreases (51), and that (ii) BBMV from the outer
renal cortex have a higher rate for OC uptake than BBMV from the inner medulla have
(34).
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● Apical OC/H+ exchange operates as an obligatory exchanger with 1:1
stoichiometry for its substrates (OCs or H+) in each translocation cycle resulting in a net
electroneutral substrate exchange across the cell membrane (68).
● Effects of H+ on OC transport i.e., trans-stimulation (via an increase in the
turnover rate) and cis-inhibition (via a competitive inhibition) were demonstrated in renal
brush border membrane vesicles and supported the model of OC efflux as an OC/H+
exchange process (69).
● The apical exit step in renal OC secretion displays a multispecificity
comparable to that of the basolateral entry step for this process (60).
There were a few early candidates for the molecular identity of the apical OC/H+
exchanger, particularly organic cation transporter novel 1 (OCTN1; SLC22A4), OCTN2
(SLC22A5), and multidrug resistance transporter 1 (MDR1; ABCB1) i.e., P-glycoprotein
(P-gp). However, when expressed in various heterologous expression systems and tested
with common OCs, neither of these transport proteins displayed a comparable
kinetic/selectivity profile to that of the OC/H+ exchanger characterized in renal brush
border membrane vesicles.
OCTN1 was an early favorite in large part because of a report showing that it
supported TEA/H+ exchange when expressed in oocytes (73). Nevertheless, an
expression study in the human kidney showed no significant mRNA expression for
hOCTN1 (39). This observation attenuated the potential of hOCTN1 for being the apical
OC/H+ exchanger. A subsequent study in HEK-293 cells expressing hOCTN1 showed
that transport efficiencies (‘TEs’; i.e., Jmax/Kt) ((13) for TEA and other‘type I’ OCs are so
23
small that it is unlikely that this process could play a significant role in the multispecific
OC secretion. Instead, it is more likely that OCTN1 is largely dedicated to the transport
of the antioxidant, ergothioneine (ET) (the TE for which is approximately 100 times
higher than that for TEA transport) (14). Interestingly, mutations in hOCTN1 gene are
found to associate with autoimmune diseases such as rheumatoid arthritis and Crohn’s
disease (26).
Another candidate for an apical OC efflux transporter was OCTN2, based on its
expression at the apical membrane of the RPTEC (39) and the observation that it can
mediate transport of prototypic OCs such as TEA. However, like its congener, hOCTN1,
hOCTN2-mediated OC transport has different kinetic/selectivity profiles from those
observed for OC/H+ exchange in renal BBMV. Interestingly, subsequent studies also
revealed that the transport activity of OCs by hOCTN2 probably has no physiological
significance (given the very low transport efficiencies for OC substrates). Instead,
hOCTN2 demonstrates high transport efficiency for carnitine that is required for normal
cellular lipid metabolism (55).
The final, ‘early’ candidate for apical OC export was MDR1. However, its
influence in renal OC secretion has long been problematic. Kinetic profiles for selected
prototypic OCs such as TEA for MDR1 are very different from those observed in renal
BBMV (72). Currently, there is a broad agreement that MDR1 is more likely to have a
role in an efflux of ‘type II’ OCs at the apical membrane of RPTEC (72). These
polyvalent and bulky OCs are proposed to passively diffuse across the basolateral
24
membrane into the RPTECs. Subsequently, an apical MDR1 helps to accelerate an ATP-
dependent secretion of the ‘type II’ OCs.
Because of these unsuccessful efforts to identify a transporter protein(s)
responsible for the OC/H+ exchange activity at the apical membrane of the RPTEC,
knowledge on the apical OC transport step lagged behind that for the basolateral element
in renal secretion of OCs.
1.8 The Multidrug And Toxin Extrusion (MATE) transporters
While students of renal OC transport were searching for the molecular identity of
the apical OC/H+ exchanger, a group of microbiologists also searched for an explanation
behind a resistance to cationic, quinolone-based antibiotics such as norfloxacin in Vibrio
parahaemolyticus. In 1998, Morita et al. demonstrated that a novel protein, NorM, is
responsible for norfloxacin resistance in V. parahaemolyticus, and suggested that NorM
represents a new group of energy-dependent, secondary active, multidrug efflux proteins
(36), that were later categorized as a new family of transporters named Multidrug And
Toxic compound Extruders, or MATEs (6). Mechanistically, NorM was proposed to
function as an Na+/OC exchanger (35). Since the description of NorM, more than 1,000
MATE family members in bacteria, plants, and fungi have been identified (28).
To introduce human MATE proteins (hMATEs) as the putative molecular identity
responsible for OC efflux at the apical membrane of RPTECs, I provide key information
in the following paragraphs on (i) discovery of hMATEs, (ii) functional characteristics of
hMATE1, (iii) structure of MATEs, and (iv) the kinetics and selectivity of hMATE1.
25
1.9 Cloning of MATE proteins in humans and other mammals
In 2005, Otsuka et al. successfully cloned transporters now considered as the
major molecular candidates for being the long sought apical OC/H+ exchange activity of
RPTECs (42). Briefly, they used the DNA sequence encoding NorM, a prokaryotic
Na+/OC exchanger from V. parahaemolyticus, as a template, and then searched the
human genome for similar DNA sequences. They found two similar genes located on the
short arm of chromosome 17 and later named the product of each gene as a human
multidrug and toxin extrusion protein, i.e., hMATE1 (SLC47A1) and hMATE2
(SLC47A2).
Orthologs of hMATE1 were also successfully cloned in rabbit, rat and mouse.
MATE2 was found, in the human, to have two additional splice variances, hMATE2-K
and hMATE2-B. hMATE2-K is a functionally active product of SLC47A2, expression
of which is restricted to the kidney (in humans). On the other hand, functional roles for
both hMATE2 and hMATE2-B (so-called because expression appears to be restricted to
the brain) are still to be elucidated. A functional ortholog of hMATE2-K is also
expressed in rabbit kidney as rbMate2-K (77). In mouse, there is an expression of a
mMate1 homolog that is currently named mMate2. However, mMate2 is not an ortholog
of hMATE2 or of rbMate2 (74), and phylogenetic evidence suggests mMate2 should be
called mMate3 (74). Interestingly, rodent ‘MATE2’ is not expressed in the kidney and is
largely restricted to the testes (29).
26
1.10 The physiological fingerprint of MATE-mediated transport
Several observations highlighted the potential role of hMATE proteins in renal
OC secretion. First, mRNA and protein for hMATE1 are highly expressed in the human
kidney (and human liver, the other major location for functional expression of OC/H+
exchange activity (38, 37). Next, from immunohistochemistry, hMATE1 is detected at
the apical region of the renal tubule including the proximal tubule (42). Importantly,
unlike other previous candidates for an apical OC/H+ exchanger, when expressed in a
heterologous expression system, hMATE1 revealed selectivity profiles (56) for various
OCs similar to the apical OC/H+ exchanger in renal BBMV and isolated renal proximal
tubules (71). Furthermore, significance of MATE1 on renal clearance of clinically
important substrates has been demonstrated in mice lacking mMate1, the only MATE
homolog expressed in mouse kidney (74). Renal clearance of the antidiabetic drug,
metformin (59) and the antibiotic, cephalexin (66), were significantly impaired in Mate1-
knockout mice.
As noted earlier, hMATE2-K is effectively restricted to the kidney (74) and has a
profile of substrate selectivity similar to that demonstrated for hMATE1 (74). However,
it is not known (i) if MATE1 and MATE2-K are coexpressed in the same RPTECs; or
(ii) if expression of either MATE homolog varies along the length of the RPT. It is
important to emphasize, therefore, that both hMATE1 and hMATE2-K are potential
contributors to the apical OC/H+ exchange activity, based on their expression profiles and
substrate interaction patterns, as discussed above. However, the aforementioned
evidence from studies of the importance of MATE1 in renal secretion of OCs in mouse
27
kidney clearly implicates this process as a likely contributor of OC secretion in human
kidney. Consequently, my study focused on characterization of hMATE1-mediated OC
transport.
1.11 Structure of MATE transporters
Since its discovery in 2005, primary attention has been directed toward
understanding the function of mammalian MATE transporters. Comparatively little
attention has been given to understanding the structure of these proteins. Hydropathy
analyses suggest that MATE family members typically have a secondary structure with
12 TMDs. However, all the mammalian MATEs appear to have 13 TMDs, the result of
an extracellular C-terminal hydrophobic domain (78). Studies using epitope-tagging and
impermeant thiol-reactive probes have confirmed the presence of this 13th TMD in
human, rabbit and mouse MATE1 (78); and unpublished observations). But in order to
appreciate the molecular mechanism of MATE transporters, a detailed molecular
structure at the atomic level is required.
In 2010, He et al. (17) reported an x-ray crystal structure of the prototypic
bacterial MATE family member, NorM, with a resolution of 3.6 Å. The structure
confirmed the presence of 12 TMDs and provided the template for development of a
homology model of the 3D structure of mammalian MATE1 (Fig. 4) that serves as the
basis of ongoing studies of structure/function relationships for the mammalian MATEs.
28
1.12 Kinetics and Selectivity of MATEs
Despite the acknowledged importance of MATE transporters in mediating renal
(and hepatic) OC secretion, comparatively little attention has been given to kinetics and
substrate selectivity of MATEs. Previous studies on MATEs have provided qualitative
information on their substrates; e.g., drug A interacts with hMATE1, whereas drug B
does not. Only a limited number of studies have been focused on the kinetic
characterization of MATE selectivity.
1.13 Kinetics and Selectivity of MATE-mediated transport: uptake vs. efflux
As noted above, there are comparatively few quantitative studies on the kinetics
of MATE-mediated transport. Importantly, what information is available is all directed
toward understanding the characteristics of MATE-mediated substrate ‘uptake.’ Under
physiological conditions at the apical membrane of the RPTEC, hMATE1 mediates OC
efflux i.e., an interaction between OCs and the cytoplasmic face of hMATE1, yet with
very few exceptions, what is known about hMATE1-mediated OC transport has been
obtained from uptake (influx) experiments, i.e., an interaction between the extracellular
face of hMATE1 and OC (Fig. 5). Results from hMATE1-mediated OC uptake have
been widely accepted to represent kinetics of hMATE1-mediated OC efflux based on an
assumption that interaction between each conformation of hMATE1 and substrates is
symmetrical (e.g., 31). Importantly, there is no experimental information on symmetry of
hMATE1-mediated transport.
There is, however, information on the symmetry of a poly-specific OC
transporter, namely, OCT2. Volk et al. (63) used the patch-clamp technique to examine
29
symmetry of rat organic cation transporter 2 (rOct2, slc22a2) and showed that binding
affinity of TBuA (tetrabutylammonium) and corticosterone to each face of rOct2 is
significantly different. More interestingly, while TBuA has 4-fold higher affinity to
external over cytoplasmic surface of rOct2, corticosterone has binding preference to
cytoplasmic (20-fold) over external surface of the protein (64). This study suggests not
only that an interaction between a substrate to each conformation of a poly-specific
transporter can be significantly different, but also that differences in affinity for a given
substrate at the two sides can vary widely among different substrates. Therefore, it is
critical to examine symmetry of substrate interaction in a poly-specific transporters,
including hMATE1.
To characterize hMATE1-mediated transport in a physiologically relevant
direction i.e., hMATE1 as an OC efflux transporter, I developed a method to
quantitatively examine the interaction between inward-facing hMATE1 and its substrates,
including H+. The major hypothesis of this study was that H+, as a substrate for hMATE1,
interacts symmetrically with each conformation of hMATE1. To test this hypothesis,
hMATE1 was stably expressed in Chinese Hamster Ovary (CHO) cells, and kinetics of
interaction between H+ and the inward and outward-facing faces of hMATE1 and H+
were examined.
30
The organization of upcoming Chapters
• Chapter 2 describes the general methods used throughout the studies that
comprise this dissertation.
• Chapter 3 presents the background, results and discussion of work described in
the research paper: Dangprapai Y, Wright SH. Interaction of H+ with the
extracellular and intracellular aspects of hMATE1. Am. J. Physiol. Renal Physiol,
2011 (in press).
• Chapter 4 describes the results of a set of currently unpublished experiments that
explore the utility of efflux measurements to assess the kinetics of hMATE1-
mediated substrate uptake.
• Chapter 5 provides an integrated view of this body of work and discusses the
future direction of MATE studies in the light of my research.
31
CHAPTER 2
MATERIALS AND METHODS
2.1 Chemicals
[3H]1-methyl-4-phenylpyridinium ([3H]MPP; 80 Ci/mmol) and [2-(4-nitro-2,1,3-
benzoxadiazol-7-yl)aminoethyl]trimethylammonium [NBD-TMA (4)] were synthesized
by the Department of Chemistry and Biochemistry, University of Arizona. [14C]Mannitol
(58.8 mCi/mmol) was purchased from PerkinElmer. 2’,7’-bis-(2-carboxyethyl)-5-(and-
6)-carboxyfluorescein acetoxymethyl (BCECF-AM) and nigericin were obtained from
Invitrogen. Tetrapentylammonium (TPeA) bromide, saponin, and other chemicals
(unless otherwise noted) were purchased from Sigma.
2.2 Cloning of the hMATE1 gene
pcDNA5/FRT vector (Invitrogen) containing hMATE1 cDNA was provided by
Dr. Giacomini (UCSF). Briefly, cDNA of hMATE1 (GenBank accession no.
NM_018242) was amplified from a human kidney RNA sample using the reverse
transcription polymerase chain reaction (RT-PCR). Subsequently, the cDNA product
was cloned into the pcDNA5/FRT vector, and the sequence later confirmed with an
Applied Biosystems 3730xl DNA analyzer at the University of Arizona sequencing
facility.
2.3 Expression of hMATE1 in CHO Flp-In cells
Chinese Hamster Ovary cells containing a single FRT site (CHO Flp-In cells)
were purchased from Invitrogen. The cells were cultured in nutrient mixture F-12 Ham
Kaighn's Modification (Ham's F-12K) medium (Sigma) supplemented with 10% fetal
32
bovine serum (HyClone) and 0.1 mg/ml Zeocin (Invitrogen). The cells were maintained
at 370C in an incubator (NuAire) supplied with 5% CO2. CHO Flp-In cells grown in T-
75 flasks (Cellstar) at ~ 80 – 90% confluency were used for electroporation.
pcDNA5/FRT/hMATE1 and pOG44 vectors containing Flp recombinase (Invitrogen)
were co-introduced by electroporation into a suspension of CHO Flp-In cells (~ 4 x 106
cells/ml in Ham’s F-12K medium). Subsequently, the electroporated cells were
maintained in serum-supplemented Ham’s F-12K medium containing 0.1 mg/ml of
Hygromycin B (Invitrogen).
2.4 Measurement of [3H]MPP uptake (for time course and kinetic studies) into CHO
hMATE1 cells
CHO hMATE1 cells were seeded in 24-well cell culture plate (Greiner Bio-one)
and allowed to reach ~100% confluency (typically 48 hours). The cells were rinsed twice
with WB pH 7.4 before being incubated in WB pH 8.5 containing [3H]MPP (~13 nM) at
room temperature (RT; 20 – 250C). For a time course study, [3H]MPP solution was
aspirated at various time points from each well and the cells were rinsed three times with
1 ml of ice-cold WB pH 8.5. For a kinetic study, the cells were incubated with WB pH
8.5 containing [3H]MPP with various concentrations of non-labeled MPP, and all
incubations were terminated at 5 minutes by removing the buffer and rinsing with ice-
cold WB as mentioned in the time course study. To release accumulated [3H]MPP, CHO
hMATE1 cells were lysed with 0.5 N NaOH containing 1% SDS and, subsequently,
neutralized with 1 N HCl. Cell lysate from each sample was aliquoted to a plastic vial
33
and mixed with scintillation cocktail (MP Biomedicals), and radioactivity in the lysate
was determined by liquid scintillation spectroscopy (Beckman).
Two approaches were used to study the kinetics of cis-inhibition by hydrogen ions
(H+) of hMATE1-mediated MPP uptake . For the first protocol, CHO hMATE1 cells
were incubated for 5 minutes with ~13 nM of [3H]MPP in WB of various H+
concentrations. The second approach was based on measuring the kinetics of hMATE1-
mediated MPP uptake at two different values of extracellular pH, as will be described
later in this dissertation.
2.5 Measurement of MPP efflux from CHO hMATE1 cells
Two similar approaches were developed to characterize hMATE1-mediated MPP
efflux. Both involved a two-step process that started with loading cells with [3H]MPP,
followed by assessment of [3H]MPP efflux from the preloaded cells. The first approach
was applied to CHO hMATE1 cells grown to complete confluence in 48-well cell culture
plates, whereas the second one was performed using a laminar flow perfusion chamber
created by assembly of a dish insert (Warner Instruments, Fig. 6) into a 35-mm cell
culture dish (Falcon) containing CHO hMATE1 cells.
For efflux studies using 48-well plates, cells were first incubated with WB pH 7.4
for 30 minutes. After aspirating this medium, the cells were loaded with labeled substrate
by adding WB pH 8.5 containing ~13 nM of [3H]MPP, followed by a 20 minute
incubation. After the buffer was aspirated, the cells were rinsed quickly (within 1 – 2
seconds) with WB pH 8.5 at room temperature. Efflux of the labeled substrate from
CHO hMATE1 cells was initiated by adding WB of various composition, according to
34
the purpose of each specific experiment. The time course of efflux under these test
conditions was determined by aspirating the efflux medium at desired time intervals and
then determining the amount of labeled substrate remaining in the cells (as described
earlier).
For experiments that used the perfusion chamber approach, CHO hMATE1 cells
were grown in a 35-mm cell culture dish. When reaching confluence, cell medium was
discarded and the cells were incubated with WB pH 7.4 for 30 minutes at room
temperature. After the WB was completely aspirated, a dish insert (Fig. 6) was assembled
on top of the layer of CHO hMATE1 cells in the cell culture dish. The flat surfaces of
the dish insert were covered with a thin layer of grease to create a sealed chamber
containing CHO hMATE1 cells (Fig. 6). Spring clips were used to firmly stabilize the
chamber. The exposed cells in the chamber were loaded with labeled substrate by
introducing WB pH 8.5 containing ~39 nM of [3H]MPP to the chamber via polyethylene
(PE) tubing. Additionally, this loading buffer also contained ~10 µM of [14C]Mannitol,
which was used to correct for extracellular volume during subsequent calculations. At
the end of the loading step (~20 minutes), the radioactive buffer was cleared from the
chamber by purging it with air (which cleared the label-containing medium via the
second PE tubing line; Fig. 6).
To initiate MPP efflux, WB was introduced into the chamber at a constant rate of
5 ml/minutes by a syringe pump (Harvard apparatus). The flow characteristics of the
chamber resulted in a laminar flow of medium through the chamber. The perfusate was
simultaneously collected into a sample vial from the distal end of the chamber at a rate of
35
6 seconds per sample. At the end of efflux duration, amount of radioactivity from each
radionuclide in each perfusate sample was determined, as was the amount of each label
remaining in the cells in the chamber, using a dual-label counting protocol.
2.6 Manipulation of intracellular pH
For measurement of intracellular pH (pHi), CHO hMATE1 cells were grown to
approximately 50% confluence in 35-mm cell culture dishes. The low level of cell
confluency facilitated monitoring pHi of a selected single cells during the study. To
measure pHi, the cells were first rinsed twice with WB pH 7.4 at room temperature, and
then incubated in the dark at room temperature in WB containing 5 µM of the pH-
sensitive fluorescent probe BCECF-AM (membrane permeable). Subsequently, the cells
were rinsed at least five times with WB pH 7.4 to remove excess probe. After the last
rinse, the cells were incubated in WB pH 7.4 for at least 10 minutes to allow the complete
enzymatic cleavage of intracellular BCECF-AM (less fluorescent) to membrane
impermeable BCECF (strongly fluorescent).
To visualize the cells, the 35-mm cell culture dish was securely placed on the
stage of a fluorescence microscope (Nikon TS100) equipped with PE tubing that was
connected to a peristaltic pump to provide a suction of test buffer during the
measurement. BCECF was excited alternately at 460 nm and 488 nm, with an emission
wavelength of 530 nm. Under the microscope, a visual field was selected that contained
cells with a homogeneous fluorescence signal. From this field, the pHi of at least 20 cells
was monitored. The fluorescence signal during each measurement was recorded using an
InCyt Im2 imaging system (Intracellular Imaging, Inc.). A value of pHi for each
36
fluorescence ratio (488/460 nm) was calculated from a calibration curve performed at the
end of each experiment. A calibration curve was determined by exposing cells to various
standard pH buffers mimicking fluid composition of the cytoplasm. Nigericin (10 µM,
freshly prepared) was added into each standard pH buffer to facilitate an equilibration of
pHi to pH of the buffer. Note that only cells that gave a stable signal throughout a course
of the measurement (10 – 15 minutes) were included in a final analysis for calculation of
the pHi.
2.7 Measurement of CHO cell volume
CHO hMATE1 cells were grown to complete confluence in a 48, 24, or 12-well
cell culture plate. For a time course study, the cells were incubated with WB pH 7.4
containing tritiated water (T2O) and [14C]Mannitol (approximately 60 mM and 4 µM,
respectively). The incubation was terminated after various durations by aspirating the
radioactive buffer (without any rinsing); steady state for T2O accumulation was reached
within 40 minutes. Amount of total T2O and [14C]Mannitol in the samples was
determined as described previously using a dual-labeled counting protocol. The
contribution of extracellular T2O to the total amount of T2O in each sample was
calculated from the amount of [14C]Mannitol, which represented extracellular volume
(separate experiments showed that mannitol was restricted to the extracellular space).
Note that due to the highly volatile nature of T2O, all incubations in this study were
performed with complete coverage of the cell culture wells using a plastic paraffin film
(Parafilm).
37
2.8 Statistics
Data were analyzed using Prism (GraphPad Software). Statistical tools such as
paired t-test or unpaired t-test were applied according to the nature of each data set.
38
CHAPTER 3
SYMMETRY OF H + INTERACTION WITH hMATE1 1
3.1 Background
The designation ‘organic cations’ (OCs) refers to a large group of organic
compounds that carry net positive charge(s) at physiological pH. The majority of OCs
are xenobiotic i.e., of exogenous origin, and many are toxic. On a regular basis,
individuals consume potentially toxic OCs, typically as plant products and/or prescribed
medicines. Currently, approximately 40 percent of commonly prescribed drugs can be
classified as OCs (23), including metformin (anti-diabetic), cimetidine (anti-peptic ulcer),
and clonidine (anti-hypertensive and analgesic). In the human body, the renal proximal
tubule (RPT) is the major site for OC secretion (71). In RPT epithelial cells (RPTECs),
OC secretion is a two-step process, with OC uptake across the peritubular (basolateral)
membrane by a process of facilitated diffusion, and OC efflux at the luminal (apical)
membrane, mainly by an OC/H+ exchange process (24).
There is broad agreement that the basolateral entry step in OC secretion by the
human RPT is dominated by human organic cation transporter 2 (hOCT2, SLC22A2),
which has been well characterized both structurally and functionally (71). There is also
growing consensus that the exit step, also the rate-limiting step (71), for OC secretion
involves the combined activity of two members of the Multidrug And Toxin Extrusion
(MATE) family, MATE1 (SLC47A1) and MATE2-K (SLC47A2) (24, 42). Although in
1 Content of this chapter was accepted for publication (9) as ‘Dangprapai Y, Wright SH. Interaction of H+ with the extracellular and intracellular aspects of hMATE1. Am. J. Physiol. Renal Physiol, 2011 May 25, in press.’ (Appendix A) during preparation of this dissertation.
39
their proposed physiological role MATEs mediate the efflux of OCs, the characterization
of MATE-mediated OC transport has, with few exceptions, relied on measurement of
uptake i.e., influx, with the tacit assumption that MATEs interact symmetrically with
their substrates, including H+ (31). Although this is a reasonable assumption with respect
to the catalytic and energetic mechanism of transport, i.e., mediated exchange of H+ for
OC (42, 58), it may well not be the case with respect to the kinetics and selectivity of
substrate interaction with these transporters. In that regard, it is significant that rat Oct2
(rOct2, slc22a2) has been shown to have markedly asymmetrical interactions with (at
least) selected OCs. For example, the affinity of tetrabutylammonium (TBuA) to
outward-facing rOct2 is four-fold higher than its affinity to inward-facing rOct2, whereas
the affinity of corticosterone to outward-facing rOct2 is twenty-fold lower than its
affinity to inward-facing rOct2 (64).
To study MATE-mediated OC transport in its physiologically relevant direction
i.e., hMATE1 as an OC efflux transporter, we quantified the MATE-mediated efflux of
the model OC, MPP. In this study, we show that kinetics of interaction between outward-
facing hMATE1 and H+ measured using an efflux protocol are not significantly different
from those obtained using conventional uptake experiments. Furthermore, for the first
time, we report kinetics of interaction between inward-facing hMATE1 and H+. These
results support the conclusion that, despite the likelihood of structurally distinct binding
regions exposed to the cytoplasmic vs. extracellular faces of hMATE1, the kinetics of H+
interaction with each face of the transporter are similar.
40
3.2 Cis-inhibition by extracellular H + of hMATE1-mediated OC uptake
We first characterized hMATE1-mediated transport of the test substrate, MPP, in
CHO cells that stably expressed the transporter. Uptake of [3H]MPP was linear for
approximately ten minutes (Fig. 7A) and five minute uptakes were selected for use in
subsequent kinetic studies as it covered the initial rate of MPP uptake in CHO hMATE1
cells. Figure 7B shows that unlabeled MPP blocked the uptake of labeled MPP with a
profile that was adequately described by the Michaelis-Menten equation for the
competitive interaction of labeled and unlabeled substrate (30):
max
t
J [MPP*]J* D[MPP*]
K [MPP*] [MPP]= +
+ +
where J* is the rate of [3H]MPP transport from a concentration equal to [MPP*]; Jmax is
the maximum rate of mediated MPP transport; Kt is the MPP concentration that resulted
in half-maximal transport (Michaelis constant); [MPP] is the concentration of unlabeled
MPP in the uptake reaction, and D is a constant that describes the non-saturable (first-
order) component of total substrate uptake (over the range of substrate concentrations
tested) that presumably reflected the combined influence of diffusive flux, nonspecific
binding, and/or incomplete rinsing of the cell layer. In four separate experiments, the
Jmax was 2.1 ± 0.18 pmol min-1 cm-2 and the apparent Kt (at pH 8.5) was 4.0 ± 0.29 µM
(Table 2).
The interaction between outward-facing hMATE1 and H+ was initially
characterized using two experimental approaches that each involved measurement of
uptake. First, the IC50 for H+ inhibition of hMATE1-mediated MPP uptake was
41
determined (Fig. 8). The decrease in MPP uptake resulting from an increase in
extracellular [H+] ([H+]o) was adequately described by the following relationship (11):
app
o
J [MPP*]J D[MPP*]
IC [H ]50
+= +
+
where J is the rate of [3H]MPP uptake; Japp is the product of the maximum rate of
[3H]MPP uptake (Jmax) and the ratio of the Ki of H+ and Kt for MPP transport; and IC50 is
the concentration of [H+]o that reduced mediated (i.e., blockable) [3H]MPP transport by
50%. The IC50 for H+-inhibited hMATE1-mediated MPP uptake was 12.4 ± 2.21 nM (pH
7.91, n = 5). Although this is the first kinetic assessment of the inhibitory effect of
extracellular H+ on hMATE1-mediated transport, several previous studies showed that
transport was reduced as extracellular pH was decreased below a value of 8.5 (42, 56),
(58). The most detailed of these (56) when replotted as the effect of [H+]o on MATE1-
mediated transport of [14C] tetraethylammonium (TEA, a prototypic OC) suggested an
IC50 of about 15 nM (pH ~7.8), similar to the value noted here.
It is also generally observed that hMATE1-mediated OC transport decreases as
pH values increase beyond 8.5 (42, 56) suggesting that H+ interaction with hMATE1 is
unlikely to be limited to a simple competitive interaction and probably includes indirect
effects of reduced [H+]o on the structure of hMATE1 and/or other proteins. Therefore,
we decided to use another approach to determine an ‘apparent Ki’ for hydrogen ion (KHi-
app), namely, the influence of extracellular [H+] on the kinetics of MPP transport.
Kinetics of hMATE1-mediated [3H]MPP uptake at pH 7.5 ([H+] = 31.6 nM) and pH 8.5
([H+] = 3.2 nM) are presented in Table 2. The increase in [H+]o resulted in a 3-fold
42
increase in the apparent Kt for MPP transport. This effect was anticipated given the
previous observation that extracellular H+ acts as a competitive inhibitor of OC/H+
exchange in isolated renal brush border membrane vesicles (BBMV) (69). However, in
our current experiments on [3H]MPP uptake in intact cells, it was also evident that the
increase in extracellular [H+] resulted in a modest (1.8-fold) but significant (P < 0.01)
increase in the Jmax for MPP transport, which is not consistent with a purely competitive
interaction of H+ with MPP at the external face of hMATE1. This result was, in fact,
seen in all four experiments that measured the kinetics of MPP transport at two different
pHs (Table 2). Nevertheless, although these data suggest that extracellular hydrogen ion
interacts with hMATE1 at several sites, resulting in a ‘mixed-type’ inhibitory profile, we
suggest that the predominant influence of H+ on apparent Kt reflected competition
between H+ and MPP for a common site or (more likely) a set of mutually exclusive sites
within a common binding surface.
The simplifying assumption that H+ acts predominantly as a competitive inhibitor
of hMATE1-mediated MPP transport permitted calculation of KHi-app using the
relationships (11):
max
t app
J [MPP]J
K [MPP]−
=+
, where ot app t H
i app
[H ]K K
K1 +
+
−−
= +
For these relationships, J and Jmax are as previously described; Kt is the Michaelis
constant for hMATE1-mediated MPP transport; and Kt-app is the apparent Kt for MPP
transport (reflecting the competitive influence of [H+]o). From the experiments shown in
Table 2, the KHi-app for the competitive interaction of H+ with extracellular aspect of
43
hMATE1 was 13.5 ± 0.84 nM (pH 7.87) which was not different from the apparent IC50
for H+ inhibition of hMATE1-mediated MPP transport, noted above (P = 0.73).
3.3 Characterization of hMATE1-mediated OC efflux
The intent of this study was development of a quantitative view of the kinetics of
hMATE1-mediated transport when operating in its ‘physiological’ direction, i.e.,
exporting OCs. To this end, we used two similar approaches to quantify hMATE1-
mediated efflux from cells stably expressing this protein. The first involved
measurement of the amount of substrate retained in cells over time after an initiation of
efflux. Figure 9A shows the 10-minute time course of [3H]MPP efflux from CHO
hMATE1 cells obtained using this method. As expected, there was a time-dependent
decrease in cell [3H]MPP and, as supported by observations described below, the loss of
[3H]MPP from the cells was effectively restricted to efflux via hMATE1. Because the
concentration of [3H]MPP in the cells at time zero was likely to be well below the Kt at
the cytoplasmic face of the transporter, the kinetics of efflux could have been expected to
be first-order2. This was not, however, the case, as shown by the non-linearity of the
semi-log presentation of these data (Fig. 9A). The curvilinearity of the plot suggested
that MPP efflux involved at least two compartments, arranged in series (e.g., an
2The volume of CHO cells, determined using tritiated water corrected for extracellular space with [14C]Mannitol, was approximately 0.3 µl/cm2 of CHO cell (confluent layer; data not shown), similar to previous reports on the volume of CHO cells (49). If accumulated [3H]MPP was distributed uniformly in this volume (which, as discussed in the text, is an oversimplification), the calculated ‘bulk’ concentration of [3H]MPP in CHO hMATE1 cells in this study was as high as ~0.4 µM at initiation of efflux. This level of accumulated MPP is about 35 times lower than the apparent Kt for interaction between MPP and external face of hMATE1 at pH 7.5 (12.3 ± 1.08 µM; n = 4). Although an apparent Kt for MPP interaction with cytoplasmic face of hMATE1 has not been characterized, we suggest that the [MPP]cell was sufficiently low to anticipate first-order behavior for hMATE1-mediated MPP efflux.
44
endosomal compartment and the cytoplasm) with the extracellular compartment.
Furthermore, the evident curvilinearity necessitates that passage from the first
(‘endosomal’) into the second (cytoplasm), is rate-limiting. This is not an unexpected
result in light of evidence that organic cations can, in fact, be sequestered within
intracellular compartments (33), including uptake of TEA into endosomal vesicles
isolated from rat renal cortex (46).
To assess, at least qualitatively, if such sequestration could account for the non-
first order efflux profile shown in Figure 9A, we examined the time course of efflux of a
fluorescent substrate for OC transporters, NBD-TMA (λex = 458 nm, λem = 530 nm; 4).
NBD-TMA blocked hMATE1-mediated MPP transport in CHO cells with an IC50 of 14.8
µM (data not shown), and though NBD-TMA may well display a quantitatively distinct
distribution profile in CHO cells than the structurally distinct MPP, it provided a useful
probe for visualizing the complexity of substrate distribution in these cells. CHO
hMATE1 cells were incubated in WB pH 8.5 containing 0.5 mM NBD-TMA for 15
minutes in an incubator with 5% CO2 at 37oC. The NBD-TMA loaded cells were then
transferred to the stage of an epifluorescence microscope (Olympus IX71). Intracellular
accumulation of fluorescent signal was only observed in cells that expressed hMATE1
(data not shown). After brief rinsing, NBD-TMA efflux was initiated by adding buffers
without substrate. Figure 10 provides a sequence of images showing the time course of
NBD-TMA efflux from CHO hMATE1 cells under several conditions.
Before the efflux was initiated, i.e., at t = 0, fluorescence within CHO hMATE1
cells could be categorized into two distinct patterns: a comparatively diffuse,
45
homogeneous signal and a punctate signal. The diffuse signal was observed throughout
the cells and probably represented free NBD-TMA in the cytoplasm. In contrast, the
punctate signal was scattered, more intense, and probably represented NBD-TMA
sequestered within cytoplasmic organelles. Our finding was similar to a result from rat
choroid plexus cells in primary culture showing that distribution of accumulated
quinacrine (another fluorescent OC) includes both a diffuse and a punctate fluorescent
signal (33). Furthermore, as noted earlier, endosomes isolated from the rat renal cortex
accumulate TEA via TEA/H+ exchange (68). Based on these results, we suggest that the
punctate distribution of NBD-TMA in CHO cells most likely represented a similar
sequestration in cytoplasmic endosomes.
But is efflux of sequestered OC from this intracellular compartment slow
compared to efflux from the cytoplasm across the plasma membrane? When efflux of
NBD-TMA was performed at extracellular pH 7.4, the diffuse signal rapidly diminished
and almost disappeared after 10 minutes, whereas the punctate signal was still largely
retained (Fig. 10, panel a). In contrast, when 0.1 mM TPeA, a non-transported inhibitor
of OC/H+ exchange in renal BBMV (67), was present in the extracellular medium during
the efflux period, both patterns of NBD-TMA signal were effectively retained for at least
10 minutes (Fig. 10, panel b). A brief exposure to 0.01% saponin can selectively
permeabilize cholesterol-rich membranes, such as the plasma membrane (52), and when
NBD-TMA loaded cells were exposed to saponin (co-exposed with 0.1 mM TPeA), the
diffuse signal cleared as early as 15 seconds while the punctate fluorescent signal still
remained after 10 minutes of efflux (Fig. 10, panel c).
46
These results support the contention that at the start of the efflux period, [3H]MPP
was distributed in at least two compartments: a ‘fast’ cytoplasmic compartment that
should readily access hMATE1 at the plasma membrane; and a ‘slow’ compartment
consisting of [3H]MPP sequestered in various cytoplasmic organelles. The solid line fit
to the data presented in Figure 3A assumed a two-phase model for the exponential loss of
[3H]MPP from CHO hMATE1 cells: a large compartment (77% of accumulated
[3H]MPP]) that turned over with a half-time of 1.6 minutes; and a smaller compartment
(23% of accumulated [3H]MPP) that turned over with a half-time of over ten minutes.
For this study, we focused on the early time course of [3H]MPP efflux that we suggest
was dominated by the fast, cytoplasmic compartment. We found that [3H]MPP in the
slow, sequestered compartment could be treated as a constant, permitting the data to be
analyzed using a one-compartment model that decayed to a plateau, using the
relationship: [3H]MPPt = ([3H]MPPt0 – C) e(-kt) + C, where [3H]MPPt is total labeled
MPP in the cells at any time t after initiation of efflux (t0); C is sequestered [3H]MPP; and
k is the first-order rate constant for hMATE1-mediated [3H]MPP efflux from the
cytoplasmic compartment. Correcting total [3H]MPP in the cells for the sequestered
compartment resulted in an efflux profile that was described by the relationship:
[3H]MPPt = ([3H]MPPt0) e(-kt)
The adequacy of this model to describe the efflux of [3H]MPP from CHO hMATE1 cells
is evident in the linearity of the semi log plot of these data presented in Figure 9B, and
the similarity in the rate constant of efflux (k) calculated after subtraction of sequestered
[3H]MPP (0.37 min-1) to the rate constant of efflux from the fast compartment of the two
47
compartment model (0.43 min-1; Fig. 9A). For the rest of this study, each efflux data set
was analyzed by a single-phase decay equation after subtraction of the estimated
sequestered pool of [3H]MPP, as shown above.
The analysis of [3H]MPP efflux from pre-loaded CHO hMATE1 cells also
permitted a direct test of the hypothesis that an inwardly-directed H+ gradient should
increase the rate of hMATE1-mediated MPP efflux. These experiments used the second
of the two efflux protocols described in the methods, the flow chamber that permitted
continuous monitoring of [3H]MPP that left the cells into the passing stream of buffer.
Within the limit of resolution of the method, the efflux of MPP from the cytoplasmic
compartment of CHO hMATE1 cells (i.e., total efflux corrected for the ‘slow’
compartment) was adequately described by first-order kinetics (Fig. 11). In these
experiments, the half-life (t0.5) of hMATE1-mediated MPP efflux in buffer of the
extracellular pH 8.5 was 9.8 ± 1.43 min (n = 4), and decreasing the external pH to 7.4
(i.e., increasing [H+]o from 3.2 to 39.8 nM) resulted in a 7-fold decrease (P = 0.0004) in
the t0.5, down to 1.4 ± 0.08 min (n = 5). These results confirmed previous observations
(57) that inwardly-directed H+ gradients trans-stimulate MATE1-mediated OC efflux
and, of particular importance to the present context, provided a validation for our
approach to measuring hMATE1-mediated OC efflux.
Quantification of MPP efflux also provided an alternative means for assessing the
kinetics of H+ interaction with the external face of hMATE1. Figure 12A shows the
effect of systematically increasing extracellular [H+] on the rate of MPP efflux. As
expected, hMATE1-mediated MPP efflux was stimulated by increasing the concentration
48
of extracellular H+, i.e., the exchangeable substrate. Furthermore, the rate constant for
MPP efflux supported by each extracellular [H+], when plotted as a function of [H+] (Fig.
12B), was adequately described by the Michaelis-Menten equation; the half-maximal
stimulation of MPP efflux occurred at an extracellular pH of 7.83 ([H+] = 14.7 ± 3.45 nM;
n = 3), and this result was not significantly different from either the IC50 (P = 0.79) or the
K i (P = 0.63) for extracellular H+ inhibition of hMATE1-mediated MPP uptake. Note
that hMATE1-mediated MPP efflux was also performed at pHo 6.5 and 6.0, but the rate
constants for efflux at these values of pHo (0.37 ± 0.013 and 0.36 ± 0.017 minute-1,
respectively) did not differ from that measured at pHo 7.0 (0.37 ± 0.02 minute-1) and, so
(for clarity) were not shown in Figure 12A.
3.4 Interaction of intracellular H + with the cytoplasmic face of hMATE1
To examine kinetics of interaction between H+ and the inward-facing aspect of
hMATE1, MPP efflux was determined as a function of intracellular H+ concentration.
From multiple measurements at room temperature in WB pH 7.4, CHO hMATE1 cells
demonstrated a baseline pHi of 7.57 ± 0.03 ([H+] i = 27.1 ± 1.79 nM; n = 21). This
baseline value varied by less than 0.2 pH units during a 15-minute exposure to WB of
different pH. Figure 13A shows a representative result of the change in pHi resulting
from an acute shift from an extracellular pH of 7.4 to one of 8.5; baseline pHi increased
by only 0.1 pH units (from pH 7.53 to pH 7.63). Manipulation of pHi was achieved
through acute exposure of cells to buffers containing increasing concentrations of NH4Cl.
Figure 13B shows the effect on the pHi of CHO hMATE1 cells of an acute exposure to
20 mM NH4Cl. Cytoplasmic pH shifted within 8 seconds from a steady-state value of 7.7
49
to 8.3, which was maintained up to 3 minutes. Acute exposure of these cells to 2.5, 5, 10,
or 20 mM NH4Cl effectively established cytoplasmic pH values of pH 7.8 ([H+] i = 15.8 ±
0.44 nM; n = 3), pH 7.9 (12.5 ± 1.42 nM; n = 6), pH 8.1 (8.7 ± 0.76 nM; n = 3), or pH 8.2
(5.8 ± 0.35 nM, n = 4), respectively.
We also considered it necessary to determine if extracellular NH4+ had a direct
effect on the transport process. The presence of 20 mM NH4Cl in an uptake buffer (pH
8.5) decreased the 5-minute uptake of [3H]MPP by about 70% (Fig. 14). An inhibition of
uptake was expected because of the associated decrease in exchangeable H+ in the
cytoplasm noted above. However, it could also have included a direct inhibitory effect of
NH4+ on the extracellular face of hMATE1. We reasoned that, if the observed decrease
in MPP uptake was due simply to a decrease in intracellular [H+], then providing an
alternative exchangeable substrate at the cytoplasmic face of hMATE1 should rescue the
inhibitory effect of extracellular NH4+ on MPP uptake. To test this hypothesis, CHO
hMATE1 cells were pre-loaded with a high concentration of non-labeled MPP (5 mM
MPP for 10 minutes) prior to measuring [3H]MPP uptake in WB containing 20 mM
NH4Cl. Results in Figure 8 demonstrated that [3H]MPP uptake inhibited by extracellular
NH4Cl was quantitatively rescued in the cells pre-loaded with non-labeled MPP,
suggesting that extracellular NH4+ did not directly inhibit hMATE1-mediated MPP
uptake and that observed effects of NH4Cl on transport reflected changes in the
cytoplasmic [H+].
The rate of MPP efflux should increase with decreasing intracellular H+
concentration because of the associated decrease in competition between the two
50
substrates at the cytoplasmic face of hMATE1. To test this, cells were pre-loaded with
[3H]MPP and efflux was initiated at extracellular pH 8.5 with no NH4Cl in the
extracellular medium. After one minute, buffer was switched to WB pH 7.4 (to stimulate
efflux; see Fig. 11) containing different concentrations of NH4Cl to manipulate
intracellular pH. Efflux was then continuously monitored for another 2 minutes. As
shown in Figure 15, decreasing the pH from 8.5 to 7.4 (i.e., increasing the concentration
of extracellular H+) produced the anticipated increase (approximately 7 fold) in rate of
[3H]MPP efflux. In this condition, with no extracellular NH4Cl, the pHi of CHO cells
was 7.57 ([H+] i = 27.1 nM). As expected, alkalinization of the cytoplasm by addition of
10 mM NH4Cl (pHi = 8.1, [H+] i = 8.7 nM) or 20 mM NH4Cl (pHi = 8.2, [H+] i = 5.8 nM)
did progressively increase the rate of [3H]MPP efflux.
To determine the kinetics of interaction between inward-facing hMATE1 and
intracellular H+, hMATE1-mediated MPP efflux was performed at an extracellular pH of
7.4 with no NH4Cl, and with 2.5, 5, 10, or 20 mM NH4Cl. As shown in Figure 16, the
efflux rate constant (k) increased in a concentration-dependent manner as the cells were
exposed to increasing concentrations of NH4Cl, i.e., as intracellular [H+] was
systematically decreased from 27.1 nM (pH 7.57) to 5.8 nM (pH 8.2), see inset to Fig. 16.
The IC50 for cytoplasmic H+ on hMATE1-mediated MPP efflux was at pH 7.94 ([H+] i =
11.5 nM). This value did not differ substantially from the range of values (12 nM to 15
nM) for the apparent Ki/IC50 of H+ interaction with the extracellular face of hMATE1
described earlier. Acknowledging that the pH range upon which this estimate of the
cytoplasmic IC50 was based was, of necessity, narrower than that used to study the
51
interaction of H+ with the extracellular aspect of hMATE1, we suggest that these data
support the conclusion that, within the limits of resolution of currently available methods,
the intra- and extracellular faces of hMATE1 display kinetically symmetrical interactions
with H+.
3.5 Interaction between H+ and hMATE1: relevance to renal OC secretion
It is relevant to place the kinetic profile of H+ interaction with hMATE1 into the
context of OC secretion in intact RPT. In the ‘early’ proximal tubule, pH of the
glomerular filtrate is moderately alkaline, ~7.1 (15), reflecting a plasma pH of ~7.4. The
current results suggest that, at this pH, the extracellular face of hMATE1 is ~ 85%
occupied by H+ in the filtrate. These protons not only facilitate an exchange for
cytoplasmic OCs, but would also limit interaction between luminal OCs and outward-
facing hMATE1, i.e., luminal H+ should reduce ‘re-uptake’ of secreted OCs via
hMATE1. Furthermore, as a result of H+ secretion, and bicarbonate and water
reabsorption, along the length of the proximal tubule, the concentration of luminal H+
increases. In the late proximal tubule, pH of the filtrate can be as acidic as pH 6.8 to pH
6.7 (2), representing a concentration of H+ that should occupy at least 92% of available
hMATE1 transporters. This increase in luminal H+ would continuously reduce a ‘re-
uptake’ of luminal OCs, the concentration of which can also be expected to increase
along the length of the RPT (due to the combined influence of MATE secretory activity
and water reabsorption).
The interaction of H+ with hMATE1 also underscores its role in providing the
‘driving force’ for OC secretion in RPT. As a secondary active H+ exchanger, MATE1
52
can mediate export of OCs against their electrochemical gradient driven by the coupled
influx of H+ moving down its electrochemical gradient. The active OC transport via the
OC/H+ exchanger is frequently described as being driven by an inward H+ gradient and,
indeed, is typically assessed in this way in studies using isolated renal BBMV – including
our own studies (68, 69). It is, however, evident from the previous discussion that this
can give a false impression of the energetic basis of OC export mediated by MATEs in
RPTEC. In the early RPT, which appears to be the primary site of OC secretion (51), the
inwardly-directed chemical gradient for H+ is very modest, about 1.25 fold, reflecting a
luminal pH of about 7.1 vs. a cytoplasmic pH of about 7.2 (75). Nevertheless, the tubule
is still poised to support a robust active transepithelial OC secretion because uptake of
OC from the blood into the RPTEC across the basolateral membrane is mediated by the
electrogenic activity of OCT2. Consequently, the inside negative membrane potential of
these cells, -60 to -70 mV, can be expected to support concentrative (yet passive)
accumulation of OCs to levels 10-15 times that in the blood. Importantly, the luminal
OC export step, mediated by MATE1 (and/or MATE2-K) involves obligatory,
electroneutral exchange (71). Thus, even in the absence of an inwardly-directed H+
chemical gradient, exchange of a proton in the filtrate for OC in the cytoplasm can permit
development of a luminal OC concentration that approaches the elevated level (compared
to the blood) of OC in the cytoplasm, with the result being active transepithelial
secretion. The MATE1-mediated step is still the ‘active’ element in this process because,
even in the absence of a chemical gradient for H+, protons are still far from
electrochemical equilibrium and kept that way through the activity of Na+/H+ exchange.
53
The increasing acidity of the tubular filtrate along the length of the RPT further enhances
the energetic effectiveness of MATE-mediated OC secretion into the tubular filtrate.
It is also relevant to discuss the potential impact on renal OC secretion of the
comparatively high affinity of the cytoplasmic face of hMATE1 for H+. With an
apparent Ki of 11.5 nM (pH 7.94), the cytoplasmic pH in RPTEC of pH 7.2 ([H+] = 63.1
nM) (75) suggests that the cytoplasmic face of MATE1 should be ~85% occupied by
hydrogen ion. Based on selectivity studies in our lab and others (42, 58), the affinity for
OCs of the outward-facing aspect of hMATE1 is typically 10 to 1000-times lower than
that for H+, and at least in general terms this is likely to be the case for the inwardly-
directed face of MATE1, as well. This would seem to ‘set the stage’ for rather brisk
competition between accumulated OCs and near-saturating levels of cytoplasmic H+. But
that scenario assumes that the ‘bulk’ pH of the cytoplasm (which is what the BCECF
measurements represent) is an accurate indicator of the hydrogen ion concentration
within the unstirred ‘microdomain’ at the cytoplasmic face of the cell membrane. While
this is arguably the case in the current experiments with CHO cells, in which cytoplasmic
pH values were 7.5 and higher [well below the ‘set-point’ for NHEs (43)], the same may
not be the case in salt transporting epithelia like the renal proximal tubule. Indeed, it is
worth noting that there is experimental evidence that the activity of Na+/H+ exchange
creates a markedly acidic microenvironment at the extracellular face of the luminal
membrane of enterocytes (2). We suggest, therefore, that the activity of the major
Na+/H+ exchangers (NHEs) in the luminal membrane of RPTEC i.e., NHE3 and NHE8,
could create local alkaline conditions near the cytoplasmic face of the membrane, thereby
54
facilitating the effective interaction of accumulated OCs with the inward face of MATE1.
Interestingly, results from a mathematical model of the renal proximal tubule brush
border membrane indicated that Na+/H+ exchange activity is (in principle) capable of
establishing a gradient of luminal H+ along the length of each microvillus, with the
cytoplasmic tip of the microvillus as alkaline as pH 8.0 (27). Another consequence of the
differential impact of NHE activity, i.e., local acidification of the extracellular face of the
membrane vs. local alkalinization of the cytoplasmic face of the membrane, would be an
effective increase in the inwardly-directed electrochemical gradient for H+ that would
enhance the secretory ‘power’ of MATE1-mediated OC export. Further study regarding
the expression profile of hMATE1 at the luminal membrane of the hRPTEC and the
influence of NHEs in hMATE1-mediated OC efflux are necessary for a better
understanding of this physiologically, pharmacologically, and toxicologically important
process.
Although our current results suggest that hMATE1 is symmetrical with respect to
its kinetic interactions with intra- and extracellular H+, it is premature to assume that
organic substrates will display a similar symmetry. Given the intrinsic structural
asymmetry of transport proteins, in conjunction with the ‘alternating access’ model of
transporter function, it is unlikely that binding surfaces of the ‘translocation pathway’
will be identical when accessed from the cytoplasmic vs. extracellular faces of the
membrane. Whereas protons titrate single carboxyl residues, the transient, weak
interactions of an organic substrate reflect interactions with multiple residues (65) and
even modest differences in the 3D structure of binding surfaces are likely to influence
55
binding kinetics. As noted previously, the affinities of the inner vs. outer faces of rOct2
for corticosterone and TBuA differ markedly. Importantly, the differences are in the
opposite direction; whereas the inward face has a higher (20x) affinity for corticosterone,
the outward face has a higher affinity (4x) for TBuA (64). We suggest that a similar
degree of kinetic asymmetry can be expected for other multispecific transporters,
including MATE1. In this regard, it is also appropriate to note that the kinetic profiles
reported here reflect the interaction of a single substrate with MATE1, i.e., MPP.
Although MPP is a widely used to characterize organic cation transporters, both OCTs
(10) and MATEs (42), structurally distinct substrates for the multiselective MATE
transporters could display a different kinetic interaction with H+.
In summary, the interaction between H+, both intracellular and extracellular, and
the MATE1-mediated transport of MPP was examined, and the results suggested that the
kinetics of interaction between inward-facing hMATE1 and intracellular H+ are not
significantly different from the kinetics of the interaction between outward-facing
hMATE1 and extracellular H+. Nevertheless, despite this apparent symmetry of H+
interaction with hMATE1, it can be anticipated that at least some OCs will display
asymmetrical interactions with intra- vs. extracellular faces of hMATE1. The approach
for assessing the kinetics of OC efflux from intact cultured cells introduced in the present
study offers a potential means to address this issue.
56
CHAPTER 4
AN APPLICATION OF THE EFFLUX PROTOCOL TO THE STUDY OF
hMATE1-MEDIATED OC INFLUX
4.1 Background
Since the successful cloning of mammalian MATEs in 2005, a number of OCs,
including prescribed drugs such as metformin (56), have been demonstrated to be
substrates for hMATE1. The typical approach to determine if a test compound is a
substrate for a defined transport protein, including MATEs, is direct measurement of
uptake of that compound into cells that heterologously express the protein, such as CHO
hMATE1 cells. An advantage of such uptake experiments is that they are comparatively
simple to do and straightforward to interpret. However, a major limitation of this
approach is the very limited access to radiolabeled compounds; few are commercially
available and they are very expensive to have synthesized. Measurement of unlabeled
compounds using mass spectrophotometry (MS) is an alternative approach, but also
expensive and time consuming. Consequently, the interaction of most compounds with
MATEs has been limited to measurement of their inhibitory effect on MATE-mediated
transport of a radiolabeled prototypic OC such as TEA, or MPP. For example, an
inhibition of [3H]MPP uptake into CHO hMATE1 cells by drug Y is generally viewed as
being sufficient to infer that drug Y is (probably) a substrate for hMATE1. A major
disadvantage of the inhibitory approach in uptake experiments is that both substrates and
non-transported inhibitors for hMATE1 can inhibit uptake of a radiolabeled OC into
CHO hMATE1 cells. An example of a non-transported inhibitor for apical OC/H+
57
exchange in renal BBMV is tetrapentylammonium (TPeA) (70), which was also shown in
my studies (Chapter 3) to block hMATE1 transport activity (Fig. 10).
hMATE1 is an obligatory exchanger, and the trans-stimulation effect of
hMATE1-mediated efflux was clearly demonstrated in Chapter 3. Thus, this efflux
protocol, when compared to using radiolabeled compounds or MS, has the potential to be
a more practical (simple, straightforward, and economical) approach to distinguish
substrates from non-transported inhibitors for MATEs-mediated transport. For example,
at low extracellular [H+], drug Y will not trans-stimulate [3H]MPP efflux from CHO
hMATE1 cells if drug Y is a non-transported inhibitor. The experiments in this chapter
provide a ‘proof-of-concept’ for the hypothesis that known substrates for hMATE1, MPP
and triethylmethylammonium (TEMA), will trans-stimulate hMATE1-mediated
[3H]MPP efflux in a manner consistent with their directly determined characteristics of
hMATE1-mediated uptake.
To test the hypothesis, [3H]MPP efflux from CHO hMATE1 cells was measured
using the flow chamber protocol, as described in Chapter 3. The main difference was
that, in this chapter, the focus was on the effect of extracellular MPP or TEMA, not H+,
on hMATE1-mediated [3H]MPP efflux. As shown below, both MPP and TEMA trans-
stimulated [3H]MPP efflux in a dose-dependent manner, and the trans-effect reached a
maximal limit at saturating concentrations of these substrates.
4.2 Trans-stimulation of hMATE1-mediated OC efflux by MPP
CHO hMATE1 cells grown to confluence in a 35-mm cell culture dish were pre-
loaded with [3H]MPP using the protocol described in Chapter 3. A dish insert was placed
58
inside the cell culture dish to create a closed chamber containing the cells. To examine
kinetics of interaction between outward-facing hMATE1 and extracellular MPP,
[3H]MPP efflux from CHO hMATE1 cells was initiated by introducing into the cell
chamber a continuous flow of efflux buffer (pH 8.5) containing various concentrations of
MPP.
Figure 17 shows that extracellular MPP trans-stimulated hMATE1-mediated
[3H]MPP efflux in a dose-dependent manner, and this stimulatory effect was not different
when extracellular [MPP] was increased from 100 µM to 200 µM. Based on the Kt value
of 5 µM for MPP with outward-facing hMATE1 (Fig. 7B), extracellular [MPP] of 100
µM should occupy ~ 95% of available, outwardly-directed hMATE1, and increasing
[MPP] to 200 µM would increase this to ~98%. In other words, MPP at 100 µM
concentration effectively saturated hMATE1 and increasing [MPP] to 200 µM would not
be expected to, and did not, provide a further stimulatory effect. Interestingly, the rate of
MPP efflux achieved by a saturating concentration of extracellular [MPP] was less than
that associated with a near saturating level of H+ (Fig. 17, pHo = 7.4). This implies that
the turnover number for hMATE1 occupied by H+ is greater than when occupied by
MPP.
To calculate the K50 for the stimulatory effect of MPP on hMATE1-mediated
[3H]MPP efflux, efflux rate constants (k) for [3H]MPP efflux from CHO hMATE1 cells
at each extracellular [MPP] were plotted as a function of extracellular [MPP] (Fig. 17,
inset). Analysis with the Michaelis-Menten equation revealed a K50 value for
extracellular MPP of 1.7 µM. This K50 value was similar to the Kt (4.00 ± 0.29 µM ) for
59
interaction between MPP and the external surface of hMATE1 determined from direct
measurement of uptake (Table 2). This observation emphasizes the potential of efflux as
a means for examining kinetics of OC interaction with the external surface of hMATE1.
4.3 Trans-stimulation of hMATE1-mediated OC efflux by TEMA
Triethylmethylammonium (TEMA) is a quaternary ammonium OC. The
molecular structure of TEMA is similar to that of TEA, the only difference being that an
ethyl group is replaced by a methyl group. Because it is comparatively easy to synthesize
3H-labeled TEMA, it has proven to be a useful test compound to characterize OC
transporters (Wright SH, unpublished observations). Furthermore, because the structure
of TEMA is very different from that of the planar, heterocyclic MPP, it provides a
structurally distinct test substrate for studying multispecific OC transporters. Not
surprisingly, the kinetics of TEMA (IC50 = 51.6 ± 4.57 µM ) are similar to those for TEA
(IC50 = 40.6 ± 4.57 µM) when presented as an inhibitor for hMATE1-mediated [3H]MPP
uptake (Astorga B., unpublished results).
The influence of extracellular TEMA on [3H]MPP efflux from CHO hMATE1
cells was studied using the chamber protocol as described earlier. Although I only
performed a single experiment, I consider the result worth reporting for two reasons.
First, as expected, TEMA trans-stimulated hMATE1-mediated [3H]MPP efflux in a
concentration-dependent manner, thereby extending the ‘proof-of-concept’ evident in the
more extensive studies with MPP reported above. Second, the maximum rate of TEMA-
stimulated MPP efflux was comparable to that supported by a saturating concentration of
MPP, suggesting that the turnover number of a TEMA-loaded transporters was (at least)
60
comparable to that of an MPP-loaded transporter. To provide kinetics of interaction
between TEMA and outward-facing hMATE1, more experiments are needed.
Nevertheless, I suggest that these limited observations support the view that measurement
of hMATE1-mediated efflux can extend information gained by determining the inhibitory
effect of a test compound on hMATE1 transport activity by assessing the ability of that
compound to stimulate hMATE1-mediated efflux.
61
CHAPTER 5
FUTURE DIRECTION
Apical OC/H+ exchange is the active and a rate-limiting step for renal secretion of
a structurally diverse array of xenobiotic and toxic OCs. Based on currently available
information, the major candidate for the molecular identity of apical OC/H+ exchange
activity in RPTECs is hMATE1 (along with its functionally similar congener, hMATE2-
K). Indeed, evidence in mice lacking Mate1 suggests that Mate1 is necessary for renal
secretion of the cationic drugs, metformin (anti-diabetic) and cephalexin (antibiotic).
Although study on MATEs has steadily increased since 2005 after the successful cloning
of hMATE1, data on kinetics of MATE-mediated transport are limited. Most reports
have focused on qualitative aspects of OC transport by MATEs, such as whether or not a
particular class of drug is a substrate for MATEs. To examine the molecular mechanism
of MATE-mediated transport, not only qualitative data on the process, but also
quantitative (kinetic) information is required.
In a native renal proximal tubule, hMATE1 functions as an OC efflux transporter
at the apical membrane. Significantly, even the limited information available for the
kinetic profiles of MATEs has been derived from uptake experiments, rather than from
more physiologically relevant efflux experiments. In a heterologous expression system,
data from an uptake study reflect an interaction between a substrate and the extracellular
surface of a transporter. On the other hand, data from an efflux experiment represent an
interaction between a substrate and cytoplasmic face of a transporter protein. The
emphasis on measurement of ‘uptake’ reflects the fact that such experiments are easier to
62
perform than are efflux studies. Although it is clear that characterization of hMATE1-
mediated transport in an efflux setting, compared to an uptake experiment, is a better
model to study hMATE1-mediated transport, prior to the current study, no kinetic profile
of hMATE1-mediated efflux has been reported.
In this study, for the first time, kinetics of interaction between H+ and the
cytoplasmic face of hMATE1 were examined and compared to H+ interaction at the
extracellular face. The results suggested a quantitatively symmetrical interaction of
hMATE1 with H+. However, it would be premature to assume that each face of
hMATE1 also interacts symmetrically with OCs, because of their structural complexity
compared to H+. Thus, I think it is critical that the kinetics of interaction between
cytoplasmic OC and hMATE1 be determined. Theoretically, the kinetics of OC efflux
(reflecting interaction with the intracellular face of hMATE1) can be calculated from a
time course of hMATE1-mediated OC efflux that begins with a near-saturating
intracellular OC concentration. The subsequent decline in the rate of efflux (as substrate
concentration falls) can be analyzed using the integrated Michaelis-Menten equation.
This approach was used successfully to determine the kinetics of fluorescein efflux from
pre-loaded renal proximal tubules (53). I have made preliminary attempts to conduct
such experiments with CHO hMATE1 cells. However, my attempt to saturate the
cytoplasmic face of hMATE1 with MPP has been challenging. Although these
observations could be interpreted as suggesting that the affinity of the cytoplasmic face of
hMATE1 for MPP is lower than that of the extracellular face, much more work needs to
be done to support such a conclusion. An optimized efflux protocol that assesses the first
63
seconds of efflux more efficiently, or a protocol that uses OCs other than MPP, have the
potential to more fully explore this topic. Alternatively, a study focusing on the
cytoplasmic face of a transporter protein, including MATEs, could perhaps be performed
using isolated secretory vesicles, which expose the intracellular face of a heterologously
expressed membrane transporter. Yeast have been used to generate such preparations
(50). Another approach could involve purified transport protein in reconstituted in
artificial proteoliposomes (22).
The current work also provides the way for addressing another critical issue,
namely, the extent to which inhibitors of MATE-mediated activity are also transported
substrates. Novel labeled, e.g., radioactive or fluorescent, substrates are expensive, and
the efflux method can be developed to provide high-throughput screening of the extent to
which unlabeled substrates stimulate efflux of inexpensive test probes such as [3H]MPP.
Finally, the need to understand the molecular basis of selectivity underscores the need to
address more completely the structure of MATE proteins. The results of empirical
measures of selectivity for test substrates should be compared to developing models of
the structure of MATE proteins to provide predictive models of substrate/inhibitor
interaction, with both faces, of these critically important elements of the renal pathway
for drug secretion.
64
Figure 1. Chemical structure of selected organic cations (OCs). This example
emphasizes that OCs are a group of structurally diverse chemicals. Thus, it is necessary
for organic cation transporters including MATEs to display a functional ‘multispecificity’
for OCs.
N+
CH3 CH3
CH3
CH3
N+ CH3
CH3
N
N
NHNH
N
CH3
S
NH
NH2
NH
NH
NH
NH2
Tetraethylammonium (TEA)
1-methyl-4-phenylpyridinium (MPP)
1-cyano-2-methyl-3-[2-[(5-methyl-1H-imidazol-4-yl)methylsulfanyl]ethyl]guanidine
(cimetidine)
1-(diaminomethylidene)guanidine (metformin)
65
Figure 2. Molecular mechanism of renal organic cation secretion. A model for
organic cation secretion at the renal proximal tubule epithelium, as discussed in the text,
was modified from the original model proposed by Holohan and Ross (48) with
additional detail regarding molecular identity of transporter proteins from currently
available information on renal OC secretion. (OC+ = organic cation, NKA = sodium-
potassium ATPase, NHE = sodium-hydrogen exchanger)
Peritubular
capillary
Renal
proximal tubule
66
Figure 3. Predicted membrane topology of hOCT2. Secondary (A) and tertiary (B)
structure of hOCT2 was proposed based on an atomic structure of a related bacterial
glycerol-3-phosphate transporter (GlpT). (This figure was modified from Figure 1 of
reference (44).
A B
67
Figure 4. Predicted membrane topology of rbMATE1. A homology model of
rbMATE1 (Chang and Wright, unpublished observation), without the 13th transmembrane
domain, was constructed from the atomic structure of bacterial NorM (17).
68
Figure 5. hMATE1-mediated MPP transport: uptake vs. efflux. A: hMATE1-
mediated MPP efflux from renal proximal tubular epithelial cells (RPTEC) is an
interaction between the cytoplasmic face of hMATE1 and MPP. B: hMATE1-mediated
MPP uptake in CHO hMATE1 cells is an interaction between extracellular face of
hMATE1 and MPP.
A OC efflux
RPTEC
B OC uptake
CHO cell
69
Figure 6. The culture dish insert. A top-view diagram of the closed bath chamber
insert (RC-37FC) with perfusion ports for 35-mm cell culture dish was modified from a
picture shown in a product brochure of Warner Instruments.
70
Figure 7. Time course and kinetics of [3H]MPP uptake in CHO hMATE1 cells. A:
[3H]MPP accumulated linearly in CHO hMATE1 cells up to approximately 10 minutes
(solid circles). [3H]MPP uptake was inhibited by co-incubating the cells with 1 mM
nonlabeled MPP (open circles). Buffer pH was 8.5. Each point is the mean (±SEM) of
uptake measured in two wells of a 24-well plate. B: Uptake (5 min; pH 8.5) of [3H]MPP
transport measured in the presence of increasing concentrations of unlabeled MPP (from
which the kinetics of MPP transport were determined). Each point is the mean (±SEM)
of uptakes measured in three wells of a 24-well plate.
0 5 10 15 20 25 300
30
60
90
1 µCi/ml [3H]MPP
1 µCi/ml [3H]MPPwith 1 mM MPP
CHO hMATE1 cells
A
Time (minutes)
[3H
]MP
P (
fmol
cm
-2)
0
5
10
15
20
1 10 100 1000
Jmax = 2.15 pmol min-1 cm-2
0
Kt = 4.89 µM
B
[MPP] (µM)
[3 H]M
PP
(fm
ol m
in-1
cm-2
)
71
Figure 8. IC50 of extracellular H+ as an inhibitor of hMATE1-mediated MPP
uptake. Five-minute uptakes of 27 nM [3H]MPP into CHO hMATE1 cells were plotted
as a function of extracellular [H+]. Each point is the mean (±SEM) of uptakes determined
in five separate experiments (n = 5).
10 - 210 - 10
100
200
300
400
100 101 102 1031
IC50H+ = 12.4 ± 2.21 nM(pH 7.91)
0
[H+] (nM)
[3 H]M
PP
(%
of
pH 7
.5)
72
Figure 9. Time course of MPP loss from CHO hMATE1 cells. A: Following a 20-
minute preload of 13.5 nM [3H]MPP, the amount of labeled MPP remaining in the cells
was measured as a function of time after efflux was initiated in an MPP-free buffer of pH
7.4 ([H+]o = 39.8 nM). Each point is the mean [3H]MPP content (±1/2 the range) of cells
in two wells of a 48-well plate in a single representative experiment. The semi-log plot
reveals that there were multiple compartments involved in [3H]MPP efflux from CHO
hMATE1 cells as discussed in the text. The solid line represents two-phase exponential
decline in cell [3H]MPP (kinetic parameters listed in the figure). B: Time course of the
first-order (single phase) decline in cell [3H]MPP, reflecting correction for sequestered
[3H]MPP (see text for discussion). The dotted lines in both graphs represent the 95%
confidence interval of the fitted values (solid line).
0 5 101
10
100
kfast = 0.37 ± 0.01 min-1
t0.5, fast = 1.88 min
B
Efflux duration (minutes)
Cel
l [3 H
]MP
P (
% o
f t
= 0
)
0 5 1010
100
kfast = 0.43 ± 0.12 min-1
t0.5, fast = 1.62 min
kslow = 0.06 ± 0.09 min-1
t0.5, slow = 10.7 min
A
Efflux duration (minutes)
Cel
l [3H
]MP
P (
% o
f t
= 0
)
73
Figure 10. Time course of NBD-TMA efflux in CHO hMATE1 cells. CHO hMATE1
cells were pre-loaded with 0.5 mM NBD-TMA at 370C for 15 minutes. Following
aspiration of the medium the cells were transferred to a fluorescent microscope, rinsed
with WB pH 8.5 at which point an image was collected representing ‘t = 0’. Subsequent
images were collected at the indicated time points following introduction of test media.
For each condition, the cells were illuminated only during collection of the image. The
10-minute images were from different fields of cells (to avoid the influence of
photobleaching).
a. pH 7.4
b. pH 7.4 + 0.1 mM TPeA
c. pH 8.5 + 0.1 mM TPeA + 0.01% saponin
t = 0 1 min. 10 min.
t = 0
t = 0 1 min.
1 min.
10 min.
10 min.
0.5 min.
0.5 min.
0.25 min.
74
Figure 11. Effect of extracellular H+ on hMATE1-mediated [3H]MPP efflux. Efflux
was performed at extracellular pH 8.5 ([H+]out = 3.2 nM) and at pH 7.4 ([H+]out = 39.8
nM) using the efflux chamber protocol (see text).
0.0 0.5 1.0 1.5 2.010
100
pH 8.5
pH 7.4
Efflux duration (minutes)
Cel
l [3H
]MP
P (
% o
f t =
0)
75
Figure 12. Effect of increasing extracellular [H+] on the kinetics of hMATE1-
mediated [3H]MPP efflux. A: Time course of [3H]MPP efflux from the ‘fast’
cytoplasmic compartment (as defined in the text) was determined in buffers of decreasing
external pH. The value of the slow compartment used for each condition was determined
from the efflux profile at pHo 7.0. Each point is the mean of results obtained in three
separate experiments. B: The rate constants for [3H]MPP efflux from the fast
compartment (the slopes of the lines fit to the data in Fig. 12A) plotted as a function of
extracellular [H+] (line fit using Michaelis-Menten equation).
0 25 50 75 1000.00
0.15
0.30
0.45
B
K50 = 14.67 ± 3.45 nM(pH 7.83)
[H+] (nM)
Effl
ux
rate
con
stan
t (k
; m
in-1
)
0 1 2 3 4 510
100
A
pH 8.0
pH 7.7
pH 8.2
pH 7.0pH 7.4
pH 8.5
Efflux duration (minutes)
Cel
l [3H
]MP
P (
% o
f t =
0)
76
Figure 13. Intracellular pH (pH i) in CHO hMATE1 cells. A: A representative
experiment showing the modest change in pHi associated with acute change of pHo; in
this case from pHo of 7.4 to pHo of 8.5 shifted pHi from 7.53 to 7.63. B: Upon acute
exposure to 20 mM NH4Cl the pHi of CHO hMATE1 cells increased rapidly (within 8
seconds) from its baseline pHi value (pHi of 7.57, i.e., [H+] i = 27.1 ± 1.79 nM; n = 21) to
a new value (pHi of 8.2, i.e., [H+] i = 5.8 ± 0.35 nM; n = 4) that was relatively stable for at
least three minutes. Data shown are from a single representative experiment. [Each point
of the fluorescence ratio in both A. and B. was an average value measured from twenty
eight CHO hMATE1 cells.]
0 3 6 9
1.1
1.2
1.3
pHo 7.4 pHo 8.5
pHi 7.53
pHi 7.63
A
Time (minutes)
Flu
ores
cenc
e ra
tio (
488
/460
nm
)
7.0
7.2
7.4
7.6
7.8
Intr
acel
lula
r p
H (
pH
i)
0 3 6 9 12
1.2
1.3
1.4
1.5
20 mMNH4Clcontrol control
pHi 7.7
pHi 8.3B
Time (minutes)
Flu
ores
cenc
e ra
tio (
488/
460
nm)
7.5
8.0
8.5
Intr
acel
lula
r p
H (
pHi)
77
Figure 14. Effect of NH4Cl and MPP preload on hMATE1-mediated [3H]MPP
uptake. The time course of hMATE1-mediated uptake of [3H]MPP (13.5 nM) was
measured under the conditions indicated in the figure. In every case, pHo was 8.5. Each
point is the mean (±SEM) of uptake values (measured in triplicate) from three separate
experiments (n = 3).
0 1 2 3 4 50
40
80
120
160 no preload, no NH4Cl (control)5 mM MPP preload, no NH4Clno preload, 20 mM NH4Cl during uptake5 mM MPP preload and20 mM NH4Cl during uptake
Duration (minutes)
[3 H]M
PP
(%
of
upta
ke a
t 5
min
.)
78
Figure 15. Effect of NH4Cl on hMATE1-mediated [3H]MPP efflux. Cells were
preloaded with [3H]MPP (36.8 nM) for 15 min. Efflux into a buffer of pH 8.5 was
measured for one minute, at which point the cells were exposed to a buffer of pH 7.4
containing the indicated concentrations of NH4Cl (to decrease [H+] i). Efflux into these
new solutions was then determined for two minutes. Single representative examples are
shown for each condition.
0 1 2 3
10
100
pH 8.5
[NH4Cl](mM)
20
pH 7.4
10
0
no NH4Cl
Efflux duration (minutes)
Cel
l [3 H
]MP
P (
% o
f am
ount
at
t =
0)
79
Figure 16. Kinetics of interaction between inward-facing hMATE1 and
intracellular H +. The time course of hMATE1-mediated [3H]MPP efflux was measured
for each of the indicated values of intracellular pH. Each point is the mean of values
determined in 3-6 separate experiments. The inset shows the relationship between
cytoplasmic [H+] and the calculated rate constants (mean ±SEM) for hMATE1-mediated
[3H]MPP efflux.
0.0 0.5 1.0 1.5 2.0
10
100
7.67.8
7.9
8.2
8.1
pHi
Duration (minutes)
Cel
l [3H
]MP
P (
% o
f t =
0)
0 10 20 300.0
0.5
1.0
1.5
2.0
2.5
IC50H+ = 11.47 nM (pH 7.94)
Intracellular [H+] (nM)
Eff
lux
rate
con
stan
t (m
in-1
)
80
Figure 17. Effect of increasing extracellular [MPP] on the kinetics of hMATE1-
mediated [3H]MPP efflux. Time course of [3H]MPP efflux from the ‘fast’ cytoplasmic
compartment (as defined in the text) was determined in buffers (pH 8.5) of increasing
extracellular concentration of unlabeled MPP. The value of the slow compartment used
for each condition was determined from the efflux profile at pHo 7.4. Each point is the
mean of results obtained in one or two separate experiments. The rate constants for
[3H]MPP efflux from the fast compartment plotted as a function of extracellular [MPP]
is shown in the inset. The line was created from the Michaelis-Menten equation.
0.0 0.5 1.0 1.5 2.0
32
40
50
63
79
100
00.1110
100200
0(pH 7.4)
[MPP](mM)
Efflux duration (minutes)
Cel
l [3 H
]MP
P (
% o
f t =
0)
0 50 100 150 2000.0
0.1
0.2
0.3
[MPP] (µµµµM)
K (
min
-1)
K50 = 1.7 µM
[MPP] (µM)
81
Figure 18. Effect of extracellular TEMA on hMATE1-mediated [3H]MPP efflux.
Efflux was performed using chamber protocol at extracellular pH 8.5 ([H+]out = 3.2 nM)
with various concentration of TEMA, and at extracellular pH 7.4 ([H+]out = 39.8 nM) as a
positive control.
0.0 0.5 1.0 1.5 2.0
32
40
50
63
79
100
0
0(pH 7.4)
0.05
0.11
100
[TEMA](mM)
Efflux duration (minutes)
Cel
l [3 H
]MP
P (
% o
f t =
0)
10
82
Antidiabetic – metformin
Antiarrhythmic – procainamide, quinidine
Antibiotic – levofloxacin
Antiviral – acyclovir, ganciclovir, tenofovir
Antiparasitic – quinine, pyrimethamine
Chemotherapeutic – platinum-containing agents such as cisplatin, oxaliplatin
Recreational – nicotine
Misc. – cimetidine (anti-peptic ulcer), clonidine (anti-hypertensive and analgesic)
(Endogenous – choline, dopamine)
Table 1. Selected organic cations of clinical or environmental significance.
83
Table 2. Effect of extracellular pH on the kinetics of hMATE1-mediated MPP
uptake. Five-minute uptakes of [3H]MPP (~13 nM) were determined as a function of
increasing concentration of unlabeled MPP at two values for external pH (pHo), pH 7.5 ±
0.1 (n = 4) and pH 8.5 ± 0.02 (n = 4). Both Jmax and Kt of MPP uptake in CHO hMATE1
cells from these two pH values were significantly different (* p = 0.01).
pHo
7.5 ± 0.1
8.5 ± 0.02 4.0 ± 0.29
12.3 ± 1.08* 3.7 ± 0.36*
2.1 ± 0.18
Kt
(µM)
Jmax
(pmol min-1 cm-2)
84
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APPENDIX A
INTERACTION OF H+ WITH THE EXTRACELLULAR AND
INTRACELLULAR ASPECTS OF hMATE1
Dangprapai Y, Wright SH. Interaction of H+ with the extracellular and intracellular
aspects of hMATE1. Am J Physiol Renal Physiol (May 25, 2011). doi:
10.1152/ajprenal.00075.2011
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93
Interaction of H+ with the extracellular and intracellular aspects of hMATE1
Authors: Yodying Dangprapai and Stephen H. Wright Affiliation: Department of Physiology, University of Arizona, Tucson, AZ 85724 Running head: Symmetry of H+ interaction with hMATE1 Corresponding author: Stephen H. Wright University of Arizona College of Medicine Department of Physiology Address: P.O. Box 245051 Tucson, AZ 85724 Phone: (520) 626-4253 Fax: (520) 626-2382 E-Mail: [email protected]
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Abstract
Interaction of H+ with the extracellular and intracellular aspects of hMATE1– Human
multidrug and toxin extrusion 1 (hMATE1, SLC47A1) is a major candidate for being the
molecular identity of organic cation/proton (OC/H+) exchange activity in the luminal
membrane of renal proximal tubules (RPT). Although physiological function of
hMATE1 supports luminal OC efflux, the kinetics of hMATE1-mediated OC transport
have typically been characterized through measurement of uptake i.e., the interaction
between outward-facing hMATE1 and OCs. To examine kinetics of hMATE1-mediated
transport in a more physiologically relevant direction i.e., an interaction between inward-
facing hMATE1 and cytoplasmic substrates, we measured the time course of hMATE1-
mediated efflux of the prototypic MATE1-substrate, [3H]1-methyl-4-phenylpyridinium
([3H]MPP), under a variety of intra- and extracellular pH conditions, from CHO cells that
stably expressed the transporter. In this study, we showed that an IC50/K i for interaction
between extracellular H+ and outward-facing hMATE1 determined from conventional
uptake experiments [12.9 ± 1.23 nM (pH 7.89); n = 9] and from the efflux protocol [14.7
± 3.45 nM (pH 7.83); n = 3] were not significantly different (P = 0.6). Furthermore,
kinetics of interaction between intracellular H+ and inward-facing hMATE1 determined
using the efflux protocol revealed an IC50 for H+ of 11.5 nM (pH 7.91), consistent with
symmetrical interactions of H+ with the inward-facing and outward-facing aspects of
hMATE1.
Key Words: organic cation, transport, secretion, renal proximal tubule, pH
95
The designation ‘organic cations’ (OCs) refers to a large group of organic
compounds that carry net positive charge(s) at physiological pH. The majority of OCs
are xenobiotic i.e., of exogenous origin, and many are toxic. On a regular basis,
individuals consume potentially toxic OCs, typically as plant products and/or prescribed
medicines. Currently, approximately 40 percent of commonly prescribed drugs can be
classified as OCs (7), including metformin (anti-diabetic), cimetidine (anti-peptic ulcer),
and clonidine (anti-hypertensive and analgesic). In the human body, the renal proximal
tubule (RPT) is the major site for OC secretion (24). In RPT epithelial cells (RPTECs),
OC secretion is a two-step process, with OC uptake across the peritubular (basolateral)
membrane by a process of facilitated diffusion, and OC efflux at the luminal (apical)
membrane, mainly by an OC/H+ exchange process (8).
There is broad agreement that the basolateral entry step in OC secretion by the
human RPT is dominated by human organic cation transporter 2 (hOCT2, SLC22A2),
which has been well characterized both structurally and functionally (24). There is also
growing consensus that the exit step, also the rate-limiting step (24), for OC secretion
involves the combined activity of two members of the Multidrug And Toxin Extrusion
(MATE) family, MATE1 (SLC47A1) and MATE2-K (SLC47A2) (8, 12). Although in
their proposed physiological role MATEs mediate the efflux of OCs, the characterization
of MATE-mediated OC transport has, with few exceptions, relied on measurement of
uptake i.e., influx, with the tacit assumption that MATEs interact symmetrically with
their substrates, including H+ (10). While this is a reasonable assumption with respect to
the catalytic and energetic mechanism of transport, i.e., mediated exchange of H+ for OC
96
(12, 19), it may well not be the case with respect to the kinetics and selectivity of
substrate interaction with these transporters. In that regard, it is significant that rat Oct2
(rOct2, slc22a2) has been shown to have markedly asymmetrical interactions with (at
least) selected OCs. For example, the affinity of tetrabutylammonium (TBuA) to
outward-facing rOct2 is four-fold higher than its affinity to inward-facing rOct2, whereas
the affinity of corticosterone to outward-facing rOct2 is twenty-fold lower than its
affinity to inward-facing rOct2 (20).
To study MATE-mediated OC transport in its physiologically relevant direction
i.e., hMATE1 as an OC efflux transporter, we quantified the MATE-mediated efflux of
the model OC, MPP. In this study, we show that kinetics of interaction between outward-
facing hMATE1 and H+ measured using an efflux protocol are not significantly different
from those obtained using conventional uptake experiments. Furthermore, for the first
time, we report kinetics of interaction between inward-facing hMATE1 and H+. These
results support the conclusion that, despite the likelihood of structurally distinct binding
regions exposed to the cytoplasmic vs. extracellular faces of hMATE1, the kinetics of H+
interaction with each face of the transporter are similar.
Materials and Methods
Chemicals. [3H]1-methyl-4-phenylpyridinium ([3H]MPP; 80 Ci/mmol) and [2-(4-nitro-
2,1,3-benzoxadiazol-7-yl)aminoethyl]trimethylammonium [NBD-TMA (2)] were
synthesized by the Department of Chemistry and Biochemistry, University of Arizona.
[14C]Mannitol (58.8 mCi/mmol) was purchased from PerkinElmer. 2’,7’-bis-(2-
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carboxyethyl)-5-(and-6)-carboxyfluorescein acetoxymethyl (BCECF-AM) and nigericin
were obtained from Invitrogen. Tetrapentylammonium (TPeA) bromide, saponin, and
other chemicals (unless otherwise noted) were purchased from Sigma.
Cell culture. CHO Flp-In cells (Invitrogen) that stably expressed hMATE1 (CHO
hMATE1 cells), created in our lab as previously described (26), were grown in Ham’s
F12 Kaighn’s modification medium (Sigma) supplemented with 10% fetal bovine serum
(HyClone) and 0.1 mg/ml of Hygromycin B (Invitrogen). These cells were maintained in
an incubator (NuAire) at 37oC supplied with 5% CO2. Subculture of the cells was
performed every 3 to 4 days. For transport studies, the CHO hMATE1 cells were seeded
in multi-well cell culture plates (Cellstar) or in 35-mm cell culture dishes (Falcon).
hMATE1-mediated MPP uptake. CHO hMATE1 cells were grown to confluence in 12-
well or 24-well cell culture plates. Before each uptake, cells were incubated in
Waymouth buffer (WB; in mM, NaCl 135, Hepes 13, CaCl2-2H2O 2.5, MgCl2-6H2O 1.2,
MgSO4-7H2O 0.8, and D-glucose 28) pH 7.4 at room temperature (between 20oC to
25oC) for 30 minutes. After removal of WB pH 7.4, MPP uptake was initiated by adding
WB pH 8.5 containing [3H]MPP (approximately 13 nM). For a time course study,
[3H]MPP uptake was terminated at specific times by aspirating and rinsing each well with
ice-cold (4oC) WB pH 8.5. Cells were lysed by adding 0.5 N NaOH/1% SDS, and
neutralized with 1 N HCl. After being mixed with scintillation cocktail (MP
Biomedicals), lysate from each sample was transferred to a liquid scintillation counter
(Beckman LS6000IC) to determine the amount of [3H]MPP.
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hMATE1-mediated MPP efflux. Two (similar) protocols were used. The first used
CHO hMATE1 cells grown in 48-well cell culture plates. Cells grown to confluence
were incubated in WB pH 7.4 for 30 minutes at room temperature. Then, WB pH 7.4
was replaced by WB pH 8.5 containing [3H]MPP (~13 nM) for 20 minutes to allow
accumulation of labeled MPP into the cells. After the solution containing [3H]MPP was
removed, the cells were briefly rinsed with WB pH 8.5 at room temperature. Efflux of
the labeled MPP was initiated by adding various buffers, according to the purpose of each
experiment. The efflux was terminated by removing the buffer at a specific time, and the
remaining intracellular [3H]MPP was determined by lysing the cells with 0.5 N NaOH
containing 1% SDS. Radioactivity in each cell lysate was determined as described
above. The amount of [3H]MPP remaining in the cells at each time point was expressed
as a percentage of the amount of accumulated [3H]MPP before efflux was initiated.
The second efflux protocol measured continuous efflux from CHO hMATE1 cells
grown in a 35-mm cell culture dish. After the cells were incubated in WB pH 7.4 at room
temperature for 30 minutes, a dish insert (Warner Instruments, RC-37FC; Supplemental
Fig. S1) was assembled within the 35-mm dish, creating a closed chamber with a growth
surface area of 1.85 cm2 and a volume of 185 µl. WB pH 8.5 containing [3H]MPP and
[14C]mannitol (approximately 39 nM and 10 µM, respectively) was introduced into the
chamber via polyethylene (PE) tubing and [3H]MPP was allowed to accumulate for 20
minutes within the CHO hMATE1 cells (the mannitol served as a marker for extracellular
volume). Subsequently, the buffer containing labeled solutes was removed from the
chamber through PE tubing at the distal end of the chamber and hMATE1-mediated
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[3H]MPP efflux was initiated by introducing laminar flow of various buffers through the
chamber using a syringe pump (Harvard Apparatus) at a constant rate (5 ml/minute). A
fraction collector was used to continuously collect buffer at the distal end of the chamber;
fractions were typically collected every six seconds (usually for 2 to 2.5 minutes). The
amounts of [3H]MPP and [14C]mannitol were measured in each sample as described
earlier using a double-label counting protocol. A representative result of [3H]MPP efflux
measured using the chamber protocol is shown in Supplemental Figure S2.
Intracellular pH measurement. CHO hMATE1 cells were incubated in WB containing
5 µM BCECF-AM at room temperature for 15 minutes. Using excitation wavelengths of
488 nm and 460 nm, the fluorescence signal from intracellular BCECF was measured at
an emission wavelength of 535 nm. The ratio of fluorescence from the two excitation
wavelengths was recorded over time using an InCyt Im2 imaging system (Intracellular
Imaging, Inc.). Intracellular pH (pHi) of CHO hMATE1 cells was calculated from a
calibration curve performed after each measurement in standard pH buffers containing 10
µM nigericin (see Supplemental Fig. S3 for a representative example).
Statistics. Data were analyzed using Prism (GraphPad Software). Statistical tools such
as paired t-test or unpaired t-test were applied according to the nature of each data set.
Results and Discussion
Cis-inhibition by extracellular H+ of hMATE1-mediated OC uptake. We first
characterized hMATE1-mediated transport of the test substrate, MPP, in CHO cells that
stably expressed the transporter. Uptake of [3H]MPP was linear for approximately ten
100
minutes (Fig. 1A) and five minute uptakes were selected for use in subsequent kinetic
studies as it covered the initial rate of MPP uptake in CHO hMATE1 cells. Figure 1B
shows that unlabeled MPP blocked the uptake of labeled MPP with a profile that was
adequately described by the Michaelis-Menten equation for the competitive interaction of
labeled and unlabeled substrate (Malo and Berteloot, 1991):
max
t
J [MPP*]J* D[MPP*]
K [MPP*] [MPP]= +
+ +
where J* is the rate of [3H]MPP transport from a concentration equal to [MPP*]; Jmax is
the maximum rate of mediated MPP transport; Kt is the MPP concentration that resulted
in half-maximal transport (Michaelis constant); [MPP] is the concentration of unlabeled
MPP in the uptake reaction, and D is a constant that describes the non-saturable (first-
order) component of total substrate uptake (over the range of substrate concentrations
tested) that presumably reflected the combined influence of diffusive flux, nonspecific
binding, and/or incomplete rinsing of the cell layer. In four separate experiments, the
Jmax was 2.1 ± 0.18 pmol min-1 cm-2 and the apparent Kt (at pH 8.5) was 4.0 ± 0.29 µM
(Table 1).
The interaction between outward-facing hMATE1 and H+ was initially
characterized using two experimental approaches that each involved measurement of
uptake. First, the IC50 for H+ inhibition of hMATE1-mediated MPP uptake was
determined (Fig. 2). The decrease in MPP uptake resulting from an increase in
extracellular [H+] ([H+]o) was adequately described by the following relationship (4):
app
o
J [MPP*]J D[MPP*]
IC [H ]50
+= ++
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where J is the rate of [3H]MPP uptake; Japp is the product of the maximum rate of
[3H]MPP uptake (Jmax) and the ratio of the Ki of H+ and Kt for MPP transport; and IC50 is
the concentration of [H+]o that reduced mediated (i.e., blockable) [3H]MPP transport by
50%. The IC50 for H+-inhibited hMATE1-mediated MPP uptake was 12.4 ± 2.21 nM (pH
7.91, n = 5). Although this is the first kinetic assessment of the inhibitory effect of
extracellular H+ on hMATE1-mediated transport, several previous studies showed that
transport was reduced as extracellular pH was decreased below a value of 8.5 (12, 18,
19). The most detailed of these (18), when replotted as the effect of [H+]o on MATE1-
mediated transport of [14C] tetraethylammonium (TEA, a prototypic OC) suggested an
IC50 of about 15 nM (pH ~7.8), similar to the value noted here.
It is also generally observed that hMATE1-mediated OC transport decreases as
pH values increase beyond 8.5 (12, 18), suggesting that H+ interaction with hMATE1 is
unlikely to be limited to a simple competitive interaction and probably includes indirect
effects of reduced [H+]o on the structure of hMATE1 and/or other proteins. Therefore,
we decided to use another approach to determine an ‘apparent Ki’ for hydrogen ion (KHi-
app), namely, the influence of extracellular [H+] on the kinetics of MPP transport.
Kinetics of hMATE1-mediated [3H]MPP uptake at pH 7.5 ([H+] = 31.6 nM) and pH 8.5
([H+] = 3.2 nM) are presented in Table 1. The increase in [H+]o resulted in a 3-fold
increase in the apparent Kt for MPP transport. This effect was anticipated given the
previous observation that extracellular H+ acts as a competitive inhibitor of OC/H+
exchange in isolated renal brush border membrane vesicles (BBMV) (23). However, in
our current experiments on [3H]MPP uptake in intact cells, it was also evident that the
102
increase in extracellular [H+] resulted in a modest (1.8-fold) but significant (P < 0.01)
increase in the Jmax for MPP transport, which is not consistent with a purely competitive
interaction of H+ with MPP at the external face of hMATE1. This result was, in fact,
seen in all four experiments that measured the kinetics of MPP transport at two different
pHs (Table 1). Nevertheless, although these data suggest that extracellular hydrogen ion
interacts with hMATE1 at several sites, resulting in a ‘mixed-type’ inhibitory profile, we
suggest that the predominant influence of H+ on apparent Kt reflected competition
between H+ and MPP for a common site or (more likely) a set of mutually exclusive sites
within a common binding surface.
The simplifying assumption that H+ acts predominantly as a competitive inhibitor
of hMATE1-mediated MPP transport permitted calculation of KHi-app using the
relationships (4):
max
t app
J [MPP]J
K [MPP]−
=+
, where ot app t H
i app
[H ]K K
K1 +
+
−−
= +
For these relationships, J and Jmax are as previously described; Kt is the Michaelis
constant for hMATE1-mediated MPP transport; and Kt-app is the apparent Kt for MPP
transport (reflecting the competitive influence of [H+]o). From the experiments shown in
Table 1, the KHi-app for the competitive interaction of H+ with extracellular aspect of
hMATE1 was 13.5 ± 0.84 nM (pH 7.87) which was not different from the apparent IC50
for H+ inhibition of hMATE1-mediated MPP transport, noted above (P = 0.73).
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Characterization of hMATE1-mediated OC efflux. The intent of this study was
development of a quantitative view of the kinetics of hMATE1-mediated transport when
operating in its ‘physiological’ direction, i.e., exporting OCs. To this end, we used two
similar approaches to quantify hMATE1-mediated efflux from cells stably expressing this
protein. The first involved measurement of the amount of substrate retained in cells over
time after an initiation of efflux. Figure 3A shows the 10-minute time course of [3H]MPP
efflux from CHO hMATE1 cells obtained using this method. As expected, there was a
time-dependent decrease in cell [3H]MPP and, as supported by observations described
below, the loss of [3H]MPP from the cells was effectively restricted to efflux via
hMATE1. Because the concentration of [3H]MPP in the cells at time zero was likely to
be well below the Kt at the cytoplasmic face of the transporter, the kinetics of efflux
could have been expected to be first-order1. This was not, however, the case, as shown
by the non-linearity of the semi-log presentation of these data (Fig. 3A). The
curvilinearity of the plot suggested that MPP efflux involved at least two compartments,
arranged in series (e.g., an endosomal compartment and the cytoplasm) with the
extracellular compartment. Furthermore, the evident curvilinearity necessitates that
passage from the first (‘endosomal’) into the second (cytoplasm), is rate-limiting. This is
not an unexpected result in light of evidence that organic cations can, in fact, be
sequestered within intracellular compartments (11), including uptake of TEA into
endosomal vesicles isolated from rat renal cortex (14).
To assess, at least qualitatively, if such sequestration could account for the non-
first order efflux profile shown in Figure 3A, we examined the time course of efflux of a
104
fluorescent substrate for OC transporters, NBD-TMA (λex = 458 nm, λem = 530 nm) (2).
NBD-TMA blocked hMATE1-mediated MPP transport in CHO cells with an IC50 of 14.8
µM (data not shown), and though NBD-TMA may well display a quantitatively distinct
distribution profile in CHO cells than the structurally distinct MPP, it provided a useful
probe for visualizing the complexity of substrate distribution in these cells. CHO
hMATE1 cells were incubated in WB pH 8.5 containing 0.5 mM NBD-TMA for 15
minutes in an incubator with 5% CO2 at 37oC. The NBD-TMA loaded cells were then
transferred to the stage of an epifluorescence microscope (Olympus IX71). Intracellular
accumulation of fluorescent signal was only observed in cells that expressed hMATE1
(data not shown). After brief rinsing, NBD-TMA efflux was initiated by adding buffers
without substrate. Figure 4 provides a sequence of images showing the time course of
NBD-TMA efflux from CHO hMATE1 cells under several conditions.
Before the efflux was initiated, i.e., at t = 0, fluorescence within CHO hMATE1
cells could be categorized into two distinct patterns: a comparatively diffuse,
homogenous signal and a punctate signal. The diffuse signal was observed throughout
the cells and probably represented free NBD-TMA in the cytoplasm. In contrast, the
punctate signal was scattered, more intense, and probably represented NBD-TMA
sequestered within cytoplasmic organelles. Our finding was similar to a result from
primary rat choroid plexus cells showing that distribution of accumulated quinacrine
(another fluorescent OC) includes both a diffuse and a punctate fluorescent signal (11).
Furthermore, as noted earlier, endosomes isolated from the rat renal cortex accumulate
TEA via TEA/H+ exchange (14). Based on these results, we suggest that the punctate
105
distribution of NBD-TMA in CHO cells most likely represented a similar sequestration in
cytoplasmic endosomes.
But is efflux of sequestered OC from this intracellular compartment slow
compared to efflux from the cytoplasm across the plasma membrane? When efflux of
NBD-TMA was performed at extracellular pH 7.4, the diffuse signal rapidly diminished
and almost disappeared after 10 minutes, whereas the punctate signal was still largely
retained (Figure 4, panel a). In contrast, when 0.1 mM TPeA, a non-transported inhibitor
of OC/H+ exchange in renal BBMV (22), was present in the extracellular medium during
the efflux period, both patterns of NBD-TMA signal were effectively retained for at least
10 minutes (Figure 4, panel b). A brief exposure to 0.01% saponin can selectively
permeabilize cholesterol-rich membranes, such as the plasma membrane (17), and when
NBD-TMA loaded cells were exposed to saponin (co-exposed with 0.1 mM TPeA), the
diffuse signal cleared as early as 15 seconds while the punctate fluorescent signal still
remained after 10 minutes of efflux (Fig. 4, panel c).
These results support the contention that at the start of the efflux period, [3H]MPP
was distributed in at least two compartments: a ‘fast’ cytoplasmic compartment that
should readily access hMATE1 at the plasma membrane; and a ‘slow’ compartment
consisting of [3H]MPP sequestered in various cytoplasmic organelles. The solid line fit
to the data presented in Figure 3A assumed a two-phase model for the exponential loss of
[3H]MPP from CHO hMATE1 cells: a large compartment (77% of accumulated [3H]
MPP]) that turned over with a half-time of 1.6 minutes; and a smaller compartment (23%
of accumulated [3H]MPP) that turned over with a half-time of over ten minutes. For this
106
study, we focused on the early time course of [3H]MPP efflux that we suggest was
dominated by the fast, cytoplasmic compartment. We found that [3H]MPP in the slow,
sequestered compartment could be treated as a constant, permitting the data to be
analyzed using a one-compartment model that decayed to a plateau, using the
relationship: [3H]MPPt = ([3H]MPPt0 – C) e(-kt) + C, where [3H]MPPt is total labeled
MPP in the cells at any time t after initiation of efflux (t0); C is sequestered [3H]MPP; and
k is the first-order rate constant for hMATE1-mediated [3H]MPP efflux from the
cytoplasmic compartment. Correcting total [3H]MPP in the cells for the sequestered
compartment resulted in an efflux profile that was described by the relationship:
[3H]MPPt = ([3H]MPPt0) e(-kt)
The adequacy of this model to describe the efflux of [3H]MPP from CHO hMATE1 cells
is evident in the linearity of the semi log plot of these data presented in Figure 3B, and
the similarity in the rate constant of efflux (k) calculated after subtraction of sequestered
[3H]MPP (0.37 min-1) to the rate constant of efflux from the fast compartment of the two
compartment model (0.43 min-1; Fig. 3A). For the rest of this study, each efflux data set
was analyzed by a single-phase decay equation after subtraction of the estimated
sequestered pool of [3H]MPP, as shown above.
The analysis of [3H]MPP efflux from pre-loaded CHO hMATE1 cells also
permitted a direct test of the hypothesis that an inwardly-directed H+ gradient should
increase the rate of hMATE1-mediated MPP efflux. These experiments used the second
of the two efflux protocols described in the methods, the flow chamber that permitted
continuous monitoring of [3H]MPP that left the cells into the passing stream of buffer.
107
Within the limit of resolution of the method, the efflux of MPP from the cytoplasmic
compartment of CHO hMATE1 cells (i.e., total efflux corrected for the ‘slow’
compartment) was adequately described by first-order kinetics (Fig. 5). In these
experiments, the half-life (t0.5) of hMATE1-mediated MPP efflux in buffer of the
extracellular pH 8.5 was 9.8 ± 1.43 min (n = 4), and decreasing the external pH to 7.4
(i.e., increasing [H+]o from 3.2 to 39.8 nM) resulted in a 7-fold decrease (P = 0.0004) in
the t0.5, down to 1.4 ± 0.08 min (n = 5). These results confirmed previous observations
(12, 19) that inwardly-directed H+ gradients trans-stimulate MATE1-mediated OC efflux
and, of particular importance to the present context, provided a validation for our
approach to measuring hMATE1-mediated OC efflux.
Quantification of MPP efflux also provided an alternative means for assessing the
kinetics of H+ interaction with the external face of hMATE1. Figure 6A shows the effect
of systematically increasing extracellular [H+] on the rate of MPP efflux. As expected,
hMATE1-mediated MPP efflux was stimulated by increasing the concentration of
extracellular H+, i.e., the exchangeable substrate. Furthermore, the rate constant for MPP
efflux supported by each extracellular [H+], when plotted as a function of [H+] (Fig. 6B),
was adequately described by the Michaelis-Menten equation; the half-maximal
stimulation of MPP efflux occurred at an extracellular pH of 7.83 ([H+] = 14.7 ± 3.45 nM;
n = 3), and this result was not significantly different from either the IC50 (P = 0.79) or the
K i (P = 0.63) for extracellular H+ inhibition of hMATE1-mediated MPP uptake. Note
that hMATE1-mediated MPP efflux was also performed at pHo 6.5 and 6.0, but the rate
constants for efflux at these values of pHo (0.37 ± 0.013 and 0.36 ± 0.017 minute-1,
108
respectively) did not differ from that measured at pHo 7.0 (0.37 ± 0.02 minute-1) and, so
(for clarity) were not shown in Figure 6A.
Interaction of intracellular H+ with the cytoplasmic face of hMATE1. To examine
kinetics of interaction between H+ and the inward-facing aspect of hMATE1, MPP efflux
was determined as a function of intracellular H+ concentration. From multiple
measurements at room temperature in WB pH 7.4, CHO hMATE1 cells demonstrated a
baseline pHi of 7.57 ± 0.03 ([H+] i = 27.1 ± 1.79 nM; n = 21). This baseline value varied
by less than 0.2 pH units during a 15-minute exposure to WB of different pH. Figure 7A
shows a representative result of the change in pHi resulting from an acute shift from an
extracellular pH of 7.4 to one of 8.5; baseline pHi increased by only 0.1 pH units (from
pH 7.53 to pH 7.63). Manipulation of pHi was achieved through acute exposure of cells
to buffers containing increasing concentrations of NH4Cl. Figure 7B shows the effect on
the pHi of CHO hMATE1 cells of an acute exposure to 20 mM NH4Cl. Cytoplasmic pH
shifted within 8 seconds from a steady-state value of 7.7 to 8.3, which was maintained up
to 3 minutes. Acute exposure of these cells to 2.5, 5, 10, or 20 mM NH4Cl effectively
established cytoplasmic pH values of pH 7.8 ([H+] i = 15.8 ± 0.44 nM; n = 3), pH 7.9
(12.5 ± 1.42 nM; n = 6), pH 8.1 (8.7 ± 0.76 nM; n = 3), or pH 8.2 (5.8 ± 0.35 nM, n = 4),
respectively.
We also considered it necessary to determine if extracellular NH4+ had a direct
effect on the transport process. The presence of 20 mM NH4Cl in an uptake buffer (pH
8.5) decreased the 5-minute uptake of [3H]MPP by about 70% (Fig. 8). An inhibition of
109
uptake was expected because of the associated decrease in exchangeable H+ in the
cytoplasm noted above. However, it could also have included a direct inhibitory effect of
NH4+ on the extracellular face of hMATE1. We reasoned that, if the observed decrease
in MPP uptake was due simply to a decrease in intracellular [H+], then providing an
alternative exchangeable substrate at the cytoplasmic face of hMATE1 should rescue the
inhibitory effect of extracellular NH4+ on MPP uptake. To test this hypothesis, CHO
hMATE1 cells were pre-loaded with a high concentration of non-labeled MPP (5 mM
MPP for 10 minutes) prior to measuring [3H]MPP uptake in WB containing 20 mM
NH4Cl. Results in Figure 8 demonstrated that [3H]MPP uptake inhibited by extracellular
NH4Cl was quantitatively rescued in the cells pre-loaded with non-labeled MPP,
suggesting that extracellular NH4+ did not directly inhibit hMATE1-mediated MPP
uptake and that observed effects of NH4Cl on transport reflected changes in the
cytoplasmic [H+].
The rate of MPP efflux should increase with decreasing intracellular H+
concentration because of the associated decrease in competition between the two
substrates at the cytoplasmic face of hMATE1. To test this, cells were pre-loaded with
[3H]MPP and efflux was initiated at extracellular pH 8.5 with no NH4Cl in the
extracellular medium. After one minute, buffer was switched to WB pH 7.4 (to stimulate
efflux; see Fig. 5) containing different concentrations of NH4Cl to manipulate
intracellular pH. Efflux was then continuously monitored for another 2 minutes. As
shown in Figure 9, decreasing the pH from 8.5 to 7.4 (i.e., increasing the concentration of
extracellular H+) produced the anticipated increase (approximately 7 fold) in rate of
110
[3H]MPP efflux. In this condition, with no extracellular NH4Cl, the pHi of CHO cells
was 7.57 ([H+] i = 27.1 nM). As expected, alkalinization of the cytoplasm by addition of
10 mM NH4Cl (pHi = 8.1, [H+] i = 8.7 nM) or 20 mM NH4Cl (pHi = 8.2, [H+] i = 5.8 nM)
did progressively increase the rate of [3H]MPP efflux.
To determine the kinetics of interaction between inward-facing hMATE1 and
intracellular H+, hMATE1-mediated MPP efflux was performed at an extracellular pH of
7.4 with no NH4Cl, and with 2.5, 5, 10, or 20 mM NH4Cl. As shown in Figure 10, the
efflux rate constant (k) increased in a concentration-dependent manner as the cells were
exposed to increasing concentrations of NH4Cl, i.e., as intracellular [H+] was
systematically decreased from 27.1 nM (pH 7.57) to 5.8 nM (pH 8.2), see inset to Fig. 10.
The IC50 for cytoplasmic H+ on hMATE1-mediated MPP efflux was at pH 7.94 ([H+] i =
11.5 nM). This value did not differ substantially from the range of values (12 nM to 15
nM) for the apparent Ki/IC50 of H+ interaction with the extracellular face of hMATE1
described earlier. Acknowledging that the pH range upon which this estimate of the
cytoplasmic IC50 was based was, of necessity, narrower than that used to study the
interaction of H+ with the extracellular aspect of hMATE1, we suggest that these data
support the conclusion that, within the limits of resolution of currently available methods,
the intra- and extracellular faces of hMATE1 display kinetically symmetrical interactions
with H+.
It is relevant to place the kinetic profile of H+ interaction with hMATE1 into the
context of OC secretion in intact RPT. In the ‘early’ proximal tubule, pH of the
glomerular filtrate is moderately alkaline, ~7.1 (6) reflecting a plasma pH of ~7.4. The
111
current results suggest that, at this pH, the extracellular face of hMATE1 is ~ 85%
occupied by H+ in the filtrate. These protons not only facilitate an exchange for
cytoplasmic OCs, but would also limit interaction between luminal OCs and outward-
facing hMATE1, i.e., luminal H+ should reduce ‘re-uptake’ of secreted OCs via
hMATE1. Furthermore, as a result of H+ secretion, and bicarbonate and water
reabsorption, along the length of the proximal tubule, the concentration of luminal H+
increases. In the late proximal tubule, pH of the filtrate can be as acidic as pH 6.8 to pH
6.7 (1), representing a concentration of H+ that should occupy at least 92% of available
hMATE1 transporters. This increase in luminal H+ would continuously reduce a ‘re-
uptake’ of luminal OCs, the concentration of which can also be expected to increase
along the length of the RPT (due to the combined influence of MATE secretory activity
and water reabsorption).
The interaction of H+ with hMATE1 also underscores its role in providing the
‘driving force’ for OC secretion in RPT. As a secondary active H+ exchanger, MATE1
can mediate export of OCs against their electrochemical gradient driven by the coupled
influx of H+ moving down its electrochemical gradient. The active OC transport via the
OC/H+ exchanger is frequently described as being driven by an inward H+ gradient and,
indeed, is typically assessed in this way in studies using isolated renal BBMV – including
our own studies (23). It is, however, evident from the previous discussion that this can
give a false impression of the energetic basis of OC export mediated by MATEs in
RPTEC. In the early RPT, which appears to be the primary site of OC secretion (16), the
inwardly-directed chemical gradient for H+ is very modest, about 1.25 fold, reflecting a
112
luminal pH of about 7.1 vs. a cytoplasmic pH of about 7.2 (25). Nevertheless, the tubule
is still poised to support a robust active transepithelial OC secretion because uptake of
OC from the blood into the RPTEC across the basolateral membrane is mediated by the
electrogenic activity of OCT2. Consequently, the inside negative membrane potential of
these cells, -60 to -70 mV (25), can be expected to support concentrative (yet passive)
accumulation of OCs to levels 10-15 times that in the blood. Importantly, the luminal
OC export step, mediated by MATE1 (and/or MATE2-K) involves obligatory,
electroneutral exchange (24). Thus, even in the absence of an inwardly-directed H+
chemical gradient, exchange of a proton in the filtrate for OC in the cytoplasm can permit
development of a luminal OC concentration that approaches the elevated level (compared
to the blood) of OC in the cytoplasm, with the result being active transepithelial
secretion. The MATE1-mediated step is still the ‘active’ element in this process because,
even in the absence of a chemical gradient for H+, protons are still far from
electrochemical equilibrium and kept that way through the activity of Na+/H+ exchange.
The increasing acidity of the tubular filtrate along the length of the RPT further enhances
the energetic effectiveness of MATE-mediated OC secretion into the tubular filtrate.
It is also relevant to discuss the potential impact on renal OC secretion of the
comparatively high affinity of the cytoplasmic face of hMATE1 for H+. With an
apparent Ki of 11.5 nM (pH 7.94), the cytoplasmic pH in RPTEC of pH 7.2 ([H+] = 63.1
nM) (25) suggests that the cytoplasmic face of MATE1 should be ~85% occupied by
hydrogen ion. Based on selectivity studies in our lab and others (12, 19), the affinity for
OCs of the outward-facing aspect of hMATE1 is typically 10 to 1000-times lower than
113
that for H+, and at least in general terms this is likely to be the case for the inwardly-
directed face of MATE1, as well. This would seem to ‘set the stage’ for rather brisk
competition between accumulated OCs and near-saturating levels of cytoplasmic H+. But
that scenario assumes that the ‘bulk’ pH of the cytoplasm (which is what the BCECF
measurements represent) is an accurate indicator of the hydrogen ion concentration
within the unstirred ‘microdomain’ at the cytoplasmic face of the cell membrane. While
this is arguably the case in the current experiments with CHO cells, in which cytoplasmic
pH values were 7.5 and higher [well below the ‘set-point’ for NHEs (13)], the same may
not be the case in salt transporting epithelia like the renal proximal tubule. Indeed, it is
worth noting that there is experimental evidence that the activity of Na+/H+ exchange
creates a markedly acidic microenvironment at the extracellular face of the luminal
membrane of enterocytes (1). We suggest, therefore, that the activity of the major
Na+/H+ exchangers (NHEs) in the luminal membrane of RPTEC i.e., NHE3 and NHE8,
could create local alkaline conditions near the cytoplasmic face of the membrane, thereby
facilitating the effective interaction of accumulated OCs with the inward face of MATE1.
Interestingly, results from a mathematical model of the renal proximal tubule brush
border membrane indicated that Na+/H+ exchange activity is (in principle) capable of
establishing a gradient of luminal H+ along the length of each microvillus, with the
cytoplasmic tip of the microvillus as alkaline as pH 8.0 (9). Another consequence of the
differential impact of NHE activity, i.e., local acidification of the extracellular face of the
membrane vs. local alkalinization of the cytoplasmic face of the membrane, would be an
effective increase in the inwardly-directed electrochemical gradient for H+ that would
114
enhance the secretory ‘power’ of MATE1-mediated OC export. Further study regarding
the expression profile of hMATE1 at the luminal membrane of the hRPTEC and the
influence of NHEs in hMATE1-mediated OC efflux are necessary for a better
understanding of this physiologically, pharmacologically, and toxicologically important
process.
Although our current results suggest that hMATE1 is symmetrical with respect to
its kinetic interactions with intra- and extracellular H+, it is premature to assume that
organic substrates will display a similar symmetry. Given the intrinsic structural
asymmetry of transport proteins, in conjunction with the ‘alternating access’ model of
transporter function, it is unlikely that binding surfaces of the ‘translocation pathway’
will be identical when accessed from the cytoplasmic vs. extracellular faces of the
membrane. Whereas protons titrate single carboxyl residues, the transient, weak
interactions of an organic substrate reflect interactions with multiple residues (5, 21) and
even modest differences in the 3D structure of binding surfaces are likely to influence
binding kinetics. As noted previously, the affinities of the inner vs. outer faces of rOct2
for corticosterone and TBuA differ markedly. Importantly, the differences are in the
opposite direction; whereas the inward face has a higher (20x) affinity for corticosterone,
the outward face has a higher affinity (4x) for TBuA (20). We suggest that a similar
degree of kinetic asymmetry can be expected for other multispecific transporters,
including MATE1. In this regard, it is also appropriate to note that the kinetic profiles
reported here reflect the interaction of a single substrate with MATE1, i.e., MPP.
Although MPP is a widely used to characterize organic cation transporters, both OCTs
115
(3) and MATEs (12), structurally distinct substrates for the multiselective MATE
transporters could display a different kinetic interaction with H+.
In summary, the interaction between H+, both intracellular and extracellular, and
the MATE1-mediated transport of MPP was examined, and the results suggested that the
kinetics of interaction between inward-facing hMATE1 and intracellular H+ are not
significantly different from the kinetics of the interaction between outward-facing
hMATE1 and extracellular H+. Nevertheless, despite this apparent symmetry of H+
interaction with hMATE1, it can be anticipated that at least some OCs will display
asymmetrical interactions with intra- vs. extracellular faces of hMATE1. The approach
for assessing the kinetics of OC efflux from intact cultured cells introduced in the present
study offers a potential means to address this issue.
116
Footnote
1The volume of CHO cells, determined using tritiated water corrected for extracellular
space with [14C]mannitol, was approximately 0.3 µl/cm2 of CHO cell (confluent layer;
data not shown), similar to previous reports on the volume of CHO cells (15). If
accumulated [3H]MPP was distributed uniformly in this volume (which, as discussed in
the text, is an oversimplification), the calculated ‘bulk’ concentration of [3H]MPP in
CHO hMATE1 cells in this study was as high as ~0.4 µM at initiation of efflux. This
level of accumulated MPP is about 35 times lower than the apparent Kt for interaction
between MPP and external face of hMATE1 at pH 7.5 (12.3 ± 1.08 µM; n = 4). Although
an apparent Kt for MPP interaction with cytoplasmic face of hMATE1 has not been
characterized, we suggest that the [MPP]cell was sufficiently low to anticipate first-order
behavior for hMATE1-mediated MPP efflux.
117
Acknowledgments
The authors would like to thank Dr. Amritlal Mandal for his assistance on intracellular
pH measurement. The work presented here was supported, in part, by NIH grants
DK58251, DK080801, and ES06694. During the course of this study, funding for Y.
Dangprapai was partially supported by the Department of Physiology, Faculty of
Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand.
118
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121
Figure legends
Figure 1. Time course and kinetics of [3H]MPP uptake in CHO hMATE1 cells. A:
[3H]MPP accumulated linearly in CHO hMATE1 cells up to approximately 10 minutes
(solid circles). [3H]MPP uptake was inhibited by co-incubating the cells with 1 mM non-
labeled MPP (open circles). Buffer pH was 8.5. Each point is the mean (±SEM) of
uptake measured in two wells of a 24-well plate. B: Uptake (5 min; pH 8.5) of [3H]MPP
transport measured in the presence of increasing concentrations of unlabeled MPP (from
which the kinetics of MPP transport were determined). Data shown are from a single
representative experiment. Each point is the mean (±SEM) of uptakes measured in three
wells of a 24-well plate.
Figure 2. IC50 of extracellular H+ as an inhibitor of hMATE1-mediated MPP
uptake. Five-minute uptakes of 13 nM [3H]MPP into CHO hMATE1 cells were plotted
as a function of extracellular [H+]. Each point is the mean (±SEM) of uptakes determined
in five separate experiments (n = 5).
Figure 3. Time course of MPP loss from CHO hMATE1 cells. A: Following a 20-
minute preload of 13.5 nM [3H]MPP, the amount of labeled MPP remaining in the cells
was measured as a function of time after efflux was initiated in an MPP-free buffer of pH
7.4 ([H+]o = 39.8 nM). Each point is the mean [3H]MPP content (±SEM) of cells in two
wells of a 48-well plate in a single representative experiment. The semi-log plot reveals
that there were multiple compartments involved in [3H]MPP efflux from CHO hMATE1
122
cells as discussed in the text. The solid line represents two-phase exponential decline in
cell [3H]MPP (kinetic parameters listed in the figure). B: Time course of the first-order
(single phase) decline in cell [3H]MPP, reflecting correction for sequestered [3H]MPP
(see text for discussion). The dotted lines in both graphs represent the 95% confidence
interval of the fitted values (solid line).
Figure 4. Time course of NBD-TMA efflux in CHO hMATE1 cells. CHO hMATE1
cells were pre-loaded with 0.5 mM NBD-TMA at 370C for 15 minutes. Following
aspiration of the medium the cells were transferred to a fluorescent microscope, rinsed
with WB pH 8.5 at which point an image was collected representing ‘t = 0.’ Subsequent
images were collected at the indicated time points following introduction of test media.
For each condition, the cells were illuminated only during collection of the image. The
10-minute images were from different fields of cells (to avoid the influence of
photobleaching).
Figure 5. Effect of extracellular H+ on hMATE1-mediated [3H]MPP efflux. Efflux
was performed at extracellular pH 8.5 ([H+]out = 3.2 nM) and at pH 7.4 ([H+]out = 39.8
nM) using the efflux chamber protocol (see text).
Figure 6. Effect of increasing extracellular [H+] on the kinetics of hMATE1-
mediated [3H]MPP efflux. A: Time course of [3H]MPP efflux from the ‘fast’
cytoplasmic compartment (as defined in the text) was determined in buffers of decreasing
123
external pH. The value of the slow compartment used for each condition was determined
from the efflux profile at pHo 7.0. Each point is the mean of results obtained in three
separate experiments. B: The rate constants for [3H]MPP efflux from the fast
compartment (the slopes of the lines fit to the data in Fig. 6A) plotted as a function of
extracellular [H+] (line fit using Michaelis-Menten equation).
Figure 7. Intracellular pH (pH i) in CHO hMATE1 cells. A: A representative
experiment showing the modest change in pHi associated with acute change of pHo; in
this case a shift from pHo of 7.4 to pHo of 8.5 increased pHi from 7.53 to 7.63. B: Upon
acute exposure to 20 mM NH4Cl the pHi of CHO hMATE1 cells increased rapidly
(within 8 seconds) from a baseline pHi value (pHi of 7.57, i.e., [H+] i = 27.1 ± 1.79 nM; n
= 21) to a new value (pHi of 8.2, i.e., [H+] i = 5.8 ± 0.35 nM; n = 4) that was relatively
stable for at least three minutes. Data shown are from a single representative experiment.
(Each point of the fluorescence ratio in both A. and B. was an average value measured
from twenty eight CHO hMATE1 cells.)
Figure 8. Effect of NH4Cl and MPP preload on hMATE1-mediated [3H]MPP
uptake. The time course of hMATE1-mediated uptake of [3H]MPP (13.5 nM) was
measured under the conditions indicated in the figure. In every case, pHo was 8.5. Each
point is the mean (±SEM) of uptake values (measured in triplicate) from three separate
experiments (n = 3).
124
Figure 9. Effect of NH4Cl on hMATE1-mediated [3H]MPP efflux. Cells were
preloaded with [3H]MPP (36.8 nM) for 15 min. Efflux into a buffer of pH 8.5 was
measured for one minute, at which point the cells were exposed to a buffer of pH 7.4
containing the indicated concentrations of NH4Cl (to decrease [H+] i). Efflux into these
new solutions was then determined for two minutes. Single representative examples are
shown for each condition.
Figure 10. Kinetics of interaction between inward-facing hMATE1 and
intracellular H +. The time course of hMATE1-mediated [3H]MPP efflux was measured
for each of the indicated values of intracellular pH. Each point is the mean of values
determined in 3-6 separate experiments. The inset shows the relationship between
cytoplasmic [H+] and the calculated rate constants (mean ±SEM) for hMATE1-mediated
[3H]MPP efflux.
125
Table
Table 1. Effect of extracellular pH on the kinetics of
hMATE1-mediated MPP uptake Five-minute uptakes of [3H]MPP (~13 nM) were determined
as a function of increasing concentration of unlabeled MPP at two values for external pH (pHo), pH 7.5 ± 0.1 (n = 4) and pH 8.5 ± 0.02 (n = 4). Both Jmax and Kt of MPP uptake in CHO hMATE1 cells from these two pH values were significantly different (* p = 0.01).
pHo
K t
(µM)
Jmax
(pmol min-1 cm-2)
7.5 ± 0.1
8.5 ± 0.02 4.0 ± 0.29
12.3 ± 1.08* 3.7 ± 0.36*
2.1 ± 0.18
126
Figure 1
0 5 10 15 20 25 300
30
60
90
1 µCi/ml [3H]MPP
1 µCi/ml [3H]MPPwith 1 mM MPP
CHO hMATE1 cells
A
Time (minutes)
[3 H]M
PP
(fm
ol c
m-2
)
0
5
10
15
20
1 10 100 1000
Jmax = 2.1 ± 0.18 pmol min-1 cm-2
0
Kt = 4.0 ± 0.29 µM
B
[MPP] (µM)[3 H
]MP
P (
fmol
min
-1cm
-2)
127
Figure 2
10 - 210 - 10
100
200
300
400
100 101 102 1031
IC50H+ = 12.4 ± 2.21 nM(pH 7.91)
0
[H+] (nM)
[3H
]MP
P (
% o
f pH
7.5
)
128
Figure 3
0 5 101
10
100
kfast = 0.37 ± 0.01 min-1
t0.5, fast = 1.88 min
B
Efflux duration (minutes)C
ell [
3 H]M
PP
(%
of
t =
0)
0 5 1010
100
kfast = 0.43 ± 0.12 min-1
t0.5, fast = 1.62 min
kslow = 0.06 ± 0.09 min-1
t0.5, slow = 10.7 min
A
Efflux duration (minutes)
Cel
l [3H
]MP
P (
% o
f t
= 0
)
129
Figure 4
a. pH 7.4
b. pH 7.4 + 0.1 mM TPeA
c. pH 8.5 + 0.1 mM TPeA + 0.01% saponin
t = 0 1 min. 10 min.
t = 0
t = 0 1 min.
1 min.
10 min.
10 min.
0.5 min.
0.5 min.
0.25 min.
130
Figure 5
0.0 0.5 1.0 1.5 2.010
100
pH 8.5
pH 7.4
Efflux duration (minutes)
Cel
l [3H
]MP
P (
% o
f t
= 0
)
131
Figure 6
0 25 50 75 1000.00
0.15
0.30
0.45
B
K50 = 14.67 ± 3.45 nM(pH 7.83)
[H+] (nM)E
fflux
rat
e co
nst
ant (
k; m
in-1
)
0 1 2 3 4 510
100
A
pH 8.0
pH 7.7
pH 8.2
pH 7.0pH 7.4
pH 8.5
Efflux duration (minutes)
Cel
l [3H
]MP
P (
% o
f t =
0)
132
Figure 7
0 3 6 9
1.1
1.2
1.3
pHo 7.4 pHo 8.5
pHi 7.53
pHi 7.63A
Time (minutes)
Flu
ores
cen
ce r
atio
(4
88/4
60
nm)
7.0
7.2
7.4
7.6
7.8
Intr
acel
lula
r p
H (
pHi)
0 3 6 9 12
1.2
1.3
1.4
1.5
20 mMNH4Clcontrol control
pHi 7.7
pHi 8.3B
Time (minutes)
Flu
ores
cenc
e ra
tio (
488/
460
nm)
7.5
8.0
8.5
Intr
acel
lula
r p
H (
pH
i)
133
Figure 8
0 1 2 3 4 50
40
80
120
160 no preload, no NH4Cl (control)5 mM MPP preload, no NH4Clno preload, 20 mM NH4Cl during uptake5 mM MPP preload and20 mM NH4Cl during uptake
Duration (minutes)
[3 H]M
PP
(%
of
upta
ke a
t 5
min
.)
134
Figure 9
0 1 2 3
10
100
pH 8.5
[NH4Cl](mM)
20
pH 7.4
10
0
no NH4Cl
Efflux duration (minutes)
Cel
l [3 H
]MP
P (
% o
f am
ount
at
t =
0)
135
Figure 10
0.0 0.5 1.0 1.5 2.0
10
100
7.67.8
7.9
8.2
8.1
pHi
Duration (minutes)
Cel
l [3H
]MP
P (
% o
f t =
0)
0 10 20 300.0
0.5
1.0
1.5
2.0
2.5
IC50H+ = 11.47 nM (pH 7.94)
Intracellular [H+] (nM)
Eff
lux
rate
con
stan
t (m
in-1
)
136
Supplemental Figure Legends
Figure S1. The culture dish insert. A top-view diagram of the closed bath chamber
insert (RC-37FC) with perfusion ports for 35-mm cell culture dish was modified from a
picture shown in a product brochure of Warner Instruments.
Figure S2. Efflux of [ 3H]MPP from CHO hMATE1 cells (determined in three separate
experiments using the chamber protocol described in the text). After preloading the cells
with [3H]MPP (and [14C]mannitol) at pH 8.5, the chamber was perfused with WB pH 7.4.
Total [3H]MPP and [14C]mannitol was determined in perfusate samples collected every 6
seconds. A: The amount of [3H]MPP in each perfusate sample that effluxed from the
cells (solid circles) was calculated by subtracting the amount of residual [3H]MPP from
the pre-loading solution that was in the extracellular volume (calculated from the level of
[14C]mannitol in each sample) from total [3H]MPP in each sample (open circles). B:
Amount of [3H]MPP remaining in the cells at each time point during the efflux period
(solid circles) was calculated from the total amount of [3H]MPP lost from the cells, plus
the amount of [3H]MPP left in the cells at the end of the efflux period. Here, the cellular
MPP is expressed as a percentage of total [3H]MPP (i.e., cellular plus extracellular)
present at the start of the efflux period, to provide insight into (i). the relative amounts of
the compound in each compartment; and (ii). the precision of the measurement. Only the
corrected amount of cellular [3H]MPP was used for further analysis. [Dashed lines
represented a 95% confidence interval resulted from analyzing the data with a single-
phase decay (Prism).]
137
Figure S3. Intracellular pH (pH i) in CHO hMATE1 cells. CHO hMATE1 cells were
pre-loaded with BCECF-AM, and fluorescence of intracellular BCECF was measured at
room temperature as described in the Methods section. A representative result shown in
this figure demonstrated the effect of extracellular NH4Cl on pHi. The inset shows the
calibration of intracellular pH using 10 µM nigericin and standardized pH buffers
(performed after each experiment).
138
Figure S1
139
Figure S2
0.0 0.5 1.0 1.5 2.0 2.50
20
40
60
100
total [3H]MPPcorrected [3H]MPP
A
Efflux duration (minutes)
[3 H]M
PP
(%
eff
lux
at t
= 0
.1 m
in)
0.0 0.5 1.0 1.5 2.0 2.510
100
total
corrected
B
Efflux duration (minutes)
[3 H]M
PP
(%
am
ount
at
t =
0)
140
Figure S3
0 10 20 30 40 50 60 700.0
0.5
0.8
1.0
1.2
1.4
1.65 mMNH4Cl
10 mMNH4Cl
WBpHo7.4
WBpHo7.4
WBpHo7.4
10µµµµM NigericinpH 4.3 5.0 6.0 7.47.0 8.2 9.0
Time (minutes)
Ave
rage
rat
io (
488
nm/4
60 n
m)
00
0.75 1.00 1.25 1.50
4.0
6.0
8.0
10.0
R2 = 0.98
Ratio 488 nm/460 nm
Intr
acel
lula
r pH