Original Paper |
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Acta Biochim Biophys |
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doi:10.1111/j.1745-7270.2008.00400.x |
Activity assay of membrane
transport proteins
Hao Xie*
Department of Biological Science and
Biotechnology, Institute of Science, Wuhan University of Technology, Wuhan
430070, China
Received: October
8, 2007
Accepted: December
9, 2007
This work was
supported by a grant from the National Natural Science Foundation of China (No.
30600004)
*Corresponding
author: Tel, 86-27-63391990; Fax, 86-27-87875245; E-mail, [email protected]
Membrane transport proteins are integral
membrane proteins and considered as potential drug targets. Activity assay of
transport proteins is essential for developing drugs to target these proteins.
Major issues related to activity assessment of transport proteins include
availability of transporters, transport activity of transporters, and
interactions between ligands and transporters. Researchers need to consider the
physiological status of proteins (bound in lipid membranes or purified),
availability and specificity of substrates, and the purpose of the activity
assay (screening, identifying, or comparing substrates and inhibitors) before
choosing appropriate assay strategies and techniques. Transport proteins bound
in vesicular membranes can be assayed for transporting substrate across
membranes by means of uptake assay or entrance counterflow assay.
Alternatively, transport proteins can be assayed for interactions with ligands
by using techniques such as isothermal titration calorimetry, nuclear magnetic
resonance spectroscopy, or surface plasmon resonance. Other methods and
techniques such as fluorometry, scintillation proximity assay,
electrophysiological assay, or stopped-flow assay could also be used for
activity assay of transport proteins. In this paper the major strategies and
techniques for activity assessment of membrane transport proteins are reviewed.
Keywords membrane
transport protein; activity assay; protein-ligand interaction
Statistical analysis of available genome-wide sequences from
eubacterial, archaean, and eukaryotic organisms revealed that 20%–30% of all open
reading frames are predicted to encode integral membrane proteins (IMPs) [1]. A
recent report using different computational methods predicted that
approximately 15%–39% of human genes encoded IMPs [2]. These IMPs locate in cell
membranes and are involved in many biological processes, such as mediating the
flow of materials and information between the cytosol and the extracellular
environment, and energy generation and transformation. Receptors and
transporters play key roles in regulating the physiological state of cells and
are often the primary targets for pharmaceutical drugs. Actually more than half
of today’s drugs are targeted to these two IMP classes [3,4]. Research on
receptors and transport proteins is one of the hottest biological areas. To
date, many techniques and theories have been developed to facilitate structural
and functional investigations of the two IMP classes. For screening potential
drugs, techniques for assaying interaction between ligands and target proteins
are very important. For transport proteins, the main focus is on transport
assays of substrates and inhibitors.
All living cells are able to obtain nutrients and other essential
substrates for metabolism and excrete metabolic waste and other products. This
process is either energy-dependent or energy-independent and carried out by
different kinds of transport proteins. As recommended by the Nomenclature
Committee of the International Union of Biochemistry and Molecular Biology
(NC-IUBMB), the term “transport protein” refers to all membrane
proteins directly involved in different transport mechanisms (http://www.chem.qmul.ac.uk/iubmb/mtp/).
These proteins are responsible for non-specific permeation or specific
transport of materials [5] (Table 1). In bacteria, 3%–15% of genes are
predicted to encode membrane transport proteins [6–9].
Lipid membranes are essential for maintaining the structure and
functions of transport proteins and always among the important factors to be
considered in activity assay experiments. Other factors include energy supply,
protein expression, physiological form and state of proteins, and specificity
of substrates and inhibitors. There have been many publications about membrane
transport proteins and their structure, functions, and applications in medicine
[10–12].
The emphasis of this paper is the major strategies and techniques in
assaying the activity of transport proteins including pores, channels,
transporters, carriers, and translocators. Transport involving endocytosis and
exocytosis will not be discussed.
Availability of Transport
Proteins for Activity Assay
In nature, many membrane proteins are expressed at very low levels.
Expression and isolation is always the bottleneck for research in membrane
transport proteins, especially for structural investigation [13]. Some common
problems for the recombinant production of membrane proteins are: low overall
yields due to a lack of intrinsic stability or folding efficiency;
susceptibility to proteolytic degradation; toxicity of the recombinant product;
low translational efficiency caused by different codon usage; wrong or missing
post-translational modifications; and mistargeting, such as accumulation of
plasma membrane proteins in internal membrane.
In transport assays, only a small amount of proteins show detectable
activity. Thus, the challenge is to avoid interference from irrelevant
proteins. Homogeneity and quality are more important than the quantity of
proteins. For example, if one needs to examine the nucleoside transport in Escherichia
coli, researchers have to avoid interference from nucleoside transporters
such as NupC, NupG, and XapB. This can be achieved by using a
transporter-deficient strain [14] or purifying and assaying target transport
proteins [15].
Expression of transport proteins can be naturally occurring or
artificially controlled. Development of modern molecular biology has made
possible deliberate gene expression. Nowadays, the following strategy is most
effective for recombinant production of membrane transport proteins [13].
(1) Select the gene of interest, transfer it into an expression
vector, and construct a recombinant form of target protein containing an
affinity tag. The affinity tag can be used for monitoring the expression and
purification of the target protein. However, researchers need to test whether
the affinity tag affects the expression, structure, and activity of
transporters by changing and comparing the types and position of the affinity
tag. For example, Mohanty and Wiener analyzed the effects of polyhistidine tag
length and position on the expression of E. coli water channel AqpZ
[16]. They placed 6- or 10-histidine tags at the N- or C-termini of AqpZ and
found that expression levels were similar and tag length had a greater effect
than tag position upon yield.
(2) Select the expression host and optimize growth conditions to
induce the expression of the target transport protein. When using a eukaryotic
expression system, researchers need to identify the membrane localization of
target transporters. Green fluorescent protein fusions are useful to visualize
whether transporters are expressed in plasma membranes or internal membranes.
Using green fluorescent protein-tagged transporters, Bai et al showed
that human sodium-dependent dicarboxylate co-transporter protein 1
predominantly locates on the plasma membrane, consistent with the results
predicted by a bioinformatics approach [17]. Proteins localized to different
membranes might also require different assays for activity. If protein
expression is detected in plasma membranes, the transport assay can be carried
out in intact cells. However, for proteins located at internal membranes,
vesicles or purified protein/proteoliposomes would need to be used. In a
different expression strategy, the Xenopus oocyte expression system is
used and the researcher needs to inject mRNA or DNA into the eggs to initiate
protein expression [18].
(3) Following the expression of target proteins, membrane vesicles
containing expressed transport proteins can be isolated. Different types of
membrane vesicles can be obtained at this stage. For plasma membranes,
researchers can use the French press technique to obtain reversed membrane
vesicles [inside out membrane vesicles (IOVs)] [19]. Or researchers can use the
method described by Hugenholtz et al [20] to isolate right side out
membrane vesicles (RSOVs). Internal membrane vesicles such as mitochondria and
chloroplasts can be isolated by disrupting cells and subsequent centrifugation
[21,22].
(4) With proper detergents, membrane proteins can be solubilized and
affinity purified. Detergents have two major roles in membrane protein
purification: disrupting lipid membranes and releasing lipid-bound membrane
proteins; and maintaining the native and functional structure of lipid-free
membrane proteins [23]. Detergents vary in their chemical structures and
physical properties. Researchers need to screen and find the most appropriate
detergent for the protein of interest. Purification of lipid-free membrane
proteins is similar to that of soluble ones. Detergents have to be included at
a concentration above critical micelle concentration in buffers to maintain the
native structure of membrane proteins. Purified membrane transporters can be
assayed for interactions with substrates. Alternatively, these purified
transporters could be reconstituted into liposomes for transport assay [13].
The remainder of this paper will give more information regarding
activity assays for transport proteins. To monitor substrate transport across
lipid membranes, transporters should be bound in vesicular membranes. In the
assay, researchers need to examine changes of substrates in the vesicle lumen.
To monitor interaction between ligands and transport proteins, the protein can be
in purified form or bound in lipid membranes. Various techniques can be applied
to investigate ligand-protein interactions.
Assaying Transport Activity of
Membrane Proteins
Assay strategies and
principles
The basic procedure for transport assay (uptake assay) is as
follows: (1) prepare membrane vesicles bearing transport protein of interest;
(2) initiate uptake by supplying with substrate and/or energy; and (3)
terminate uptake at the desired time and monitor the intake of substrate [24].
The conventional method to index the intake of substrate is by using
isotopically-labeled substrates [24]. Alternatively, movement of fluorescent
substrates or fluorescence change in indicators related to a transport process
can be measured [25,26]. Changes in pH and the volume of vesicles can also be
used as transport assay indexes [25,27].
In transport assays, the intake of labeled substrate mediated by
transport proteins is described by Equation 1:
Eq. 1
where Vin refers to the substrate
intake rate, Vmax refers to the maximal substrate intake
rate, [S]out refers to the environmental substrate
concentration, and Km refers to the Michaelis-Menten
constant.
There are three destinations of intake substrate.
(1) The substrate can be metabolized and will result in the decrease
of substrate concentration within intact cells. If this happens, researchers
can eliminate the metabolism system by either using enzyme-deficient mutants or
using membrane vesicles instead of intact cells. Or researchers could select a
non-metabolizable analog. For example, 3-O-methylglucose is a non-metabolizable
glucose analog and can be used as a marker to assess transport by evaluating
its uptake within various cells [27–29].
(2) If the substrate is not metabolized and does not exit, it will
remain in the cell. This is the ideal situation and the change of substrate
within the cell or vesicles can be monitored. For determination of
Michaelis-Menten kinetics, researchers have to measure the linear initial
transport rate. This is impossible because the reaction occurs too quickly.
Practically, a short transport period is used to approximate initial velocities
of transport. For example, researchers monitored the 15 s or 1 min uptake for
estimation of the apparent Vmax and Km values for nucleoside transport in E. coli or putrescine
transport in the cyanobacterium Synechocystis sp. PCC 6803 [15,30].
(3) The substrate can exit from the cell or closed membrane vesicle.
One example is the transport mediated by transport systems such as pores or
channels. Another example is artificial proteoliposomes where the same amounts
of reconstituted proteins are facing both sides of the lipid membranes. In both
cases the passage of substrates takes place in two directions. The intake rate
(Vin) is determined by Equation 1 and the outflow rate (Vout) is given by Equation 2:
Eq. 2
where Vmax refers to the maximal
substrate transport rate, [S]in refers to the substrate
concentration in the lumen, and Km refers
to the Michaelis-Menten constant.
If intake and outflow rates are similar and the environmental
substrate concentration is low, there is a difficulty with this technique in
monitoring the change of substrate concentration between the lumen and the
environment. To solve this problem, the entrance counterflow assay was
developed [13] that relies on two types of substrates, isotope-labeled and
unlabeled. Membrane vesicles are preloaded with unlabeled substrate at a high
concentration. Transport assay is initiated by dilution of preloaded vesicles
into large volumes of buffers containing isotope-labeled substrate at a low
concentration. Both labeled and unlabeled substrates can be transported in dual
directions (towards the outside or the inside of membrane vesicles). It can be
defined that [SU]in and [SR]in are the concentrations of unlabeled and radiolabeled substrates,
respectively, in vesicle lumen. [SU]out and [SR]out are
environmental concentrations of unlabeled and radiolabeled substrates, respectively.
The intake and outflow rates of radiolabeled substrate (VRin and VRout) and unlabeled substrate (VUin and VUout) are determined by the
following Equations:
Eq. 3
Eq. 4
Eq. 5
Eq. 6
The different stages of the entrance counterflow assay are shown in Fig.
1. This assay is extremely useful in assaying different types of
substrates. Practically, it is difficult and expensive to radiolabel all types
of substrates. However, with the entrance counterflow method, researchers can
assay different substrates by changing the types of loaded unlabeled substrate
in the vesicle lumen and comparing resultant changes in radiolabeled substrate
uptake. Using the entrance counterflow assay, transporters such as galactose
transporter GalP and nucleoside transporter NupG were investigated for their
activity in proteoliposomes [13,15].
Transport assay in intact
cells
There is at least one disadvantage of the uptake assay in intact cells,
in that live cells have a metabolic mechanism to modify or degrade substrates
and will affect the subsequent measurement. However, there are some advantages,
including the fact that the transport protein is in its natural status, and
intact cells have a complete system to support transport. To monitor substrate
transport in intact cells, researchers must first mix intact cells with labeled
substrates, and initiate uptake with energy and oxygen at the appropriate
temperature. Then for different expression systems, researchers will choose
different methods, such as centrifugation or filtration, to stop the uptake
interval by separating cells from the environmental labeled substrate [24].
Other methods can also be used, such as reducing the temperature or adding
inhibitors [24]. Table 2 compares different procedures for ending the
uptake interval. A combination of the above methods can also be considered. The
amount of proteins or cells involved in assays can be determined by a variety
of methods, such as the bicinchoninic acid protein assay or spectrometry. More
detailed protocols for transport assay in intact cells have been described by
Jarvis [24].
Transport assay in RSOVs and
IOVs
RSOVs are obtained by removing cell plasma and keeping the original
orientation of plasma membranes [20]. These vesicles can be assayed for uptake
by supplying an appropriate substrate and establishing a driving force, such as
proton motive force, for secondary transporters. The advantage of using RSOVs
for transport assay is significant: the intrinsic metabolic system is removed
so that the intake substrate will not be degraded or metabolized. Sometimes
researchers might also obtain reversed membrane vesicles (IOVs) [19]. The IOVs
are broadly used for assaying substrate transport in primary transport systems.
Researchers can easily supply ATP to the transport system, which is exofacial
in IOVs, and establish the driving force for substrate transport.
Lots of transport proteins have been assayed using RSOVs and IOVs.
For example, Xie et al [15] observed and assayed nucleoside transport in
RSOVs of E. coli. Ames et al [41] examined histidine transport in
IOVs of E. coli and found that ATP induced efflux of histidine from
IOVs. Thanassi et al [42] investigated bile salts transport in IOVs of E.
coli and found that everted membrane vesicles accumulated bile salts in an
energy-dependent manner. Soksawatmaekhin et al [43,44] also compared
transport properties of transporter CadB in RSOVs and IOVs.
However, there are still disadvantages for activity assay in RSOVs
or IOVs. As well as target transport proteins, membrane vesicles still contain
other irrelevant membrane proteins, such as pores or enzymes that might affect
transport assays.
Transport assay in artificial
proteoliposomes
Purified transporters can be reconstituted into liposomes by
dilution of a ternary mixture containing proteins, lipids, and detergents
[13,45]. Or researchers can use BioBeads (Bio-Rad, Hercules, USA) to absorb and
remove detergents [13]. Once the free detergent concentration in the mixture is
lower than the critical micellar concentration, detergent is recruited from the
bound detergent pool, and the association of proteins and lipids is initiated.
The reconstituted proteoliposomes contain a single type of transporter facing
both sides of the lipid membrane and can be assayed for transport activity. For
example, Xie et al [15] purified and reconstituted nucleoside
transporter NupG into liposomes and observed nucleoside transport in
proteoliposomes. Juge et al [46] co-reconstituted vesicular glutamate
transporter VGLUT1 and bacterial F-ATPase into liposomes and found that ATP
induced L-glutamate uptake in proteoliposomes. Bowsher et al
reconstituted amyloplast envelope membrane proteins from spring wheat and
assayed ADP, AMP, and ADP-glucose transport in these proteoliposomes [47].
Eytan et al reconstituted P-glycoprotein from cultured
multidrug-resistant Chinese hamster ovary cells and observed ATP-driven,
valinomycin-dependent uptake of rubidium in these proteoliposomes [48,49].
Assaying Interactions between
Substrate and Transport Proteins
Sometimes transport efficiency is low (for example, use of
inhibitors) and it results in technique difficulties for activity assay. An alternative
choice is to measure the interaction between substrates (inhibitors) and
transport proteins. Both purified transport proteins and membrane vesicles can
be used for ligand-protein interaction assays. Although lipid membranes still
play important roles in maintaining the functional structure of transport
proteins, they are not essential for ligand-protein interaction assay. Most
assaying techniques for ligand-protein interactions can be used for transport
proteins. The concentration change of substrate after ligand-protein
interaction can be directly measured. Research examples include investigation
into the interaction between galactose or glucose transporters and inhibitors
such as forskolin and cytochalasin B using the equilibrium dialysis method [50–52].
The interaction between ligands and proteins can change the
conformation and energy status of proteins. Scientists take advantage of these
changes and have developed techniques such as isothermal titration calorimetry
(ITC) and nuclear magnetic resonance (NMR) spectroscopy to monitor interaction
between ligands and proteins. ITC is a biophysical quantitative technique used
to determine the thermodynamic parameters (binding affinity, enthalpy changes,
and binding stoichiometry) of biochemical interactions. Using ITC, tungstate
transport protein A was observed to bind tungstate and molybdate and the
dissociation constant for binding was also determined [53]. Wei and Fu also
observed selective metal binding to a membrane-embedded aspartate in the E.
coli metal transporter YiiP [54].
NMR spectroscopy is a powerful technique used to obtain physical,
chemical, electronic, and structural information about molecules. NMR
spectroscopy depends on the splitting of nuclear energy levels in a magnetic
field and the transition induced between the levels. Interaction between many
transporters and substrates has been assayed using this technique [6]. For
example, Patching et al [55] assayed the interaction between methyl-b–D-glucuronide
with glucuronide transporter GusB and revealed the dissociation constant KD is higher than the Michaelis-Menten constant Km for energized transport.
Other Techniques for
Functional Assay of Transport Proteins
Fluorometry
Fluorometry is an analytical technique for identifying and
characterizing minute amounts of a substance by excitation of the substance
with a beam of ultraviolet light and detection and measurement of the
characteristic wavelength of fluorescent light emitted. Fluorometry is often a
technique used to monitor substrate transport by transporters. For example,
Woebking et al [26] assayed ethidium transport mediated by ATP-binding
cassette transporter MsbA expressed in intact Lactococcus lactis cells.
In transport assay, cells were preloaded with the fluorescent substrate until a
steady-state level was reached. Then glucose was added to the cells as a source
of metabolic energy, after which the ethidium fluorescence was monitored. With
this method, Woebking et al [26] showed that MsbA-mediated efflux of
ethidium is affected by the protein expression level. They also investigated
the Michaelis-Menten kinetic of MsbA-mediated ethidium transport.
Scintillation proximity assay (SPA)
SPA is a technique for carrying out binding assays without
separation of bound and unbound radiotracers. In SPA, the scintillant is
incorporated into small fluomicrospheres (beads). If a radioactive molecule is
bound to the bead, it can stimulate the scintillant to emit light. The unbound
radioactivity is too distant from the scintillant and the energy released is
dissipated before reaching the bead, therefore these beads do not produce a
signal. Quick and Javitch [56] described a direct scintillation proximity-based
isotope-binding assay for determining transport protein functions in crude cell
extracts and in purified form. The copper chelate SPA scintillation beads were
used to immobilize His-tagged Na+/tyrosine transporter Tyt1,
which binds radiolabeled tyrosine. The bound radiolabeled tyrosine stimulated
SPA beads and produced a signal as an index for the binding assay. With SPA,
the activity of the Na+/tyrosine transporter Tyt1 has
been investigated and confirmed.
Surface plasmon resonance (SPR)
SPR is a technique to measure biomolecular interactions in real time
in a label-free environment. SPR-based instruments use an optical method to
measure the refractive index near the sensor surface. Using SPR, Benabdelhak et
al [57] characterized a specific interaction between the nucleotide-binding
domain of the ATP-binding cassette transporter HlyB and a C-terminal fragment
of its transport substrate haemolysin A. The C-terminal fragment of haemolysin
A was expressed and immobilized on the sensor surface. The specific interaction
between this peptide and the nucleotide-binding domain of HlyB results in the
change of reflected light and is used as an index for binding assays.
Electrophysiological assays
For any transporter that transports a net charge, there are
electrophysiological assays, including patch clamping and two-electrode voltage
clamping, for recording membrane potential [58]. The assays can be applied to
both cultured cells and Xenopus oocytes. For example, Glaaser et al
[59] and Romanenko et al [60] assayed activity of sodium channels and
potassium channels expressed in cultured cells, and Reinders et al [39]
and Vicente et al [61] investigated the transport activity of Arabidopsis
sugar alcohol permease homolog AtPLT5 and potassium channels expressed in Xenopus
oocytes.
Spectrometric assays
Heuberger et al [62,63] developed a spectroscopic
carbohydrate-transport assay that does not require isotopically-labeled
substrates. They constructed a membrane system (hybrid membranes or
proteoliposomes) bearing the transport system of interest, and soluble glucose
dehydrogenase and the electron acceptor 2,6-dichloroindophenol enclosed in the
vesicle lumen. After transport across the vesicular membrane, the sugar is
oxidized by soluble glucose dehydrogenase. The accompanying reduction of
2,6-dichloroindophenol results in a decrease in A600 and is used as the index for sugar transport. With this method both
solute/H+ symport and exchange types of transport can be measured with high
sensitivity in crude membranes as well as in proteoliposomes.
Stopped-flow assays
Stopped-flow is one of the most frequently used rapid kinetics
techniques. Small volumes of solutions are driven through a mixer and pass
through a measurement flow cell. Using appropriate techniques such as
fluorescence spectrometry, the kinetics of the reaction in the solutions can be
measured in the cell. For example, for activity measurement of metal
transporters YiiP and ZitB of E. coli [25,54,64], researchers preloaded
proteoliposomes with metal-sensitive fluorescence indicator fluozin-1 mixed
with a buffer containing Zn2+ or Cd2+. The
uptake of metal into proteoliposomes resulted in fluorescence change of the
indicator and was recorded in a stopped-flow apparatus. Water permeabilities in
liposomes can also be measured by detecting the light scattering.
Proteoliposomes are preloaded with sorbitol, sucrose, or mannitol at low
concentrations, and mixed with assay buffers containing a high concentration of
these chemicals in an assay. Because these chemicals are impermeant for
proteoliposomes, the osmotic gradient drives the water efflux, and the
consequent reduction in liposome volume is measured as an increase in the
intensity of scattered light at 600 nm. The rate constant of the normalized light
intensity increase indicates the rate constant of water efflux, which is
proportional to the water permeability coefficient [65,66]. Using this method,
transport activity of water transporters like aquaporin-8 from rat, human, and
mouse were assayed in proteoliposomes [40]. Mallo and Ashby showed that this
method can also be used for assaying water permeability mediated by water
transporter AqpZ in intact E. coli cells [67].
Conclusions
The unique properties and functions of membrane
transport proteins make it possible to take advantage of various assay
strategies and techniques. Although the conventional isotope-based technique is
still the most reliable method, novel techniques such as ITC and SPA provide
more options in assaying activity. Sometimes a combination of these methods
will be more applicable for a specific transport protein. Researchers need to
consider the physiological status of proteins (bound in lipid membranes or
purified) and the purpose of the transport assay (screening, identifying, or
comparing substrates and inhibitors) before choosing the most suitable assay
strategies and techniques.
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