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Microsystem for the single molecule analysis of membrane transport proteins
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Microsystem for the single molecule analysis of membrane transport proteins Rikiya Watanabe Molecular Physiology Laboratory, RIKEN, Saitama, Japan
A R T I C LE I N FO
A B S T R A C T
Keywords: Biomembrane Membrane protein Single molecule biophysics bioMEMS
Micro-chamber arrays enable highly sensitive and quantitative bioassays at the single-molecule level. Accordingly, they are widely used for ultra-sensitive biomedical applications, e.g., digital PCR and digital ELISA. However, the versatility of micro-chambers is generally limited to reactions in aqueous solutions, although various functions of membrane proteins are extremely important. To address this issue, microsystems using arrayed micro-sized chambers sealed with lipid bilayers, referred to here as a “biomembrane microsystems”, have been developed by many research groups for the analysis of membrane proteins. In this review, I would like to introduce recent progress on the single molecule analysis of membrane transport proteins using a biomembrane microsystem, and discuss the future prospects for its use in analytical and pharmacological applications.
1. Introduction The maintenance of an appropriate intracellular environment is a constant challenge for all living organisms, from prokaryotes to multicellular eukaryotes. Intracellular homeostasis is in general maintained by membrane proteins which mediate the translocation of various compounds across biomembranes [1]. Therefore, the analysis of transport mechanisms is crucial to understand cell physiology, as well as the action of drugs [2,3]. The transport of various substrates is mainly conducted by transport proteins embedded in the cell membrane. For several decades, the functional dynamics of transporters have been studied using a variety of single-molecule techniques [4,5], which offer key benefits over macroscopic assay methods, as they unlock the ability to quantify transport events [6]. One of the most robust systems used to analyze membrane transport at the single molecule level is patch clamp recording, which measures the flux of ions as an electric current under constant electrical voltage across a membrane. Recent developments have resulted in the automation of patch clamp recordings, thus enabling massively parallel analysis of transport activities, e.g. pores and channels [7,8]; however, transport rates of most transporters (< 102 molecules s−1) are much smaller than those of ion channels (> 107 molecules s−1), and therefore, it is difficult to detect their activities as an electric current. Moreover, patch clamp recording cannot detect the flux of electrically neutral substrates, and therefore, the development of a more versatile system has been long desired. Recently, microsystems arrayed with micro-sized chambers sealed
with lipid bilayers have been developed by many research groups for the analysis of membrane transport; in these systems, transport activity is measured optically based on the accumulation or consumption of a substrate present in the chambers [9–15]. Microsystems enhance both sensitivity and throughput, thereby allowing the highly sensitive analysis of various transporter proteins in a high throughput manner. Notably, our “biomembrane microsystem”, which contains > 100,000 arrayed atto- to femto-liter chambers sealed with stable lipid bilayers, has been used to perform the most sensitive and parallel analysis of membrane transport, enabling the single molecule analysis of transporters with extremely low transport activities (< 10 molecules s−1) [5,16]. Moreover, we have successfully demonstrated some physiological aspects of the membrane system, such as an asymmetric transbilayer phospholipid distribution [16,17], and the modulation of membrane potential across lipid bilayers [18]. Thus, our system paves the way for understanding the mechanism of membrane transport under semi-physiologic conditions, as well as allowing for further analytical and pharmacological applications. In this review, I would like to introduce our current microsystem, and discuss the future prospects for membrane transport analysis at the single molecule level. 2. Part I. Biomembrane microsystems Over the past few years, we have developed various “biomembrane microsystems” in order to carry out the single molecule analysis of membrane transport proteins under semi-physiological conditions [5,16–21]. The microsystem consists of a glass substrate containing the
E-mail address: [email protected]. https://doi.org/10.1016/j.bbagen.2019.03.016 Received 9 December 2018; Received in revised form 19 March 2019; Accepted 22 March 2019 0304-4165/ © 2019 Elsevier B.V. All rights reserved.
Please cite this article as: Rikiya Watanabe, BBA - General Subjects, https://doi.org/10.1016/j.bbagen.2019.03.016
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Fig. 1. Biomembrane microsystem. (a) Components of the system. Top, a glass block with a sample injection port (l: w: h = 20 mm: 20 mm: 5 mm); middle, a spacer sheet with one side open (Frame-Seal, BIO-RAD); bottom, a glass slide with > 10,000 microchambers printed. (b) Photograph (top) and illustration (bottom) of the assembled system. The access ports for sample injection are indicated using red circles. (c) Schematic illustration of the generation of phospholipid membranes. Membrane bilayers were formed over microchambers via sequential injection of liquids from the access port. Gray, blue, and yellow are the first aqueous solution, second aqueous solution, and chloroform solution containing lipids, respectively. (d) Fluorescent image of Alexa 647 encapsulated into microchambers sealed by lipid membranes.
chamber via the sequential injection of liquids from the access port (Fig. 1c). This technique is similar to the conventional technique known as the “painting technique,” which forms lipid bilayers through the deposition of a lipid solution across a small aperture in a hydrophobic material [12,23]. The experimental procedures used to form the lipid membrane are comprised of three steps. First, the micro-chambers are filled with an aqueous solution. A chloroform solution containing phospholipids, e.g., phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylserine (PS), is then infused via the access port. Due to the hydrophobicity of the top orifice, the interface between water and chloroform is formed over the orifice, where a lipid monolayer is then deposited. Finally, a second aqueous solution is infused to form another lipid monolayer at the interface with the chloroform solution, which is then deposited over the entire surface of the device. Thus, bilayers are formed over the micro-chambers, whereas monolayers are formed elsewhere. It is worth noting that we form lipid bilayers using chloroform as a solvent for the lipid, whereas decane or hexadecane have typically been used in conventional approaches [22].
microarray, a spacer sheet, and a glass block with an access port for sample injection (Fig. 1a). Microarrays with > 10,000 micro-chambers (ɸ = 2.4–8.2 μm and h = 0.03–0.5 μm: V = 0.2–71 fL) were fabricated using soft lithography on a hydrophilic glass substrate coated with CYTOP (Asahi-glass), a carbon‑fluorine hydrophobic polymer (Fig. 1b). Notably, in previous studies fluorine polymers have frequently been used as a support medium for lipid membranes [22]. The fabrication procedure is comprised of three steps. First, CYTOP is spin-coated onto a glass substrate of a specified thickness. Photolithography is then used to pattern mask the structures onto the photo-resist cover on the entire surface of the CYTOP layer. The resist-patterned substrate is then dryetched with O2 plasma using a reactive-ion etching system (RIE) to expose the glass substrate, resulting in the fabrication of the microchamber. Owing to the treatment with O2 plasma, the bottom and walls of the micro-chamber become hydrophilic, while the top orifice remains very hydrophobic. These different surfaces are crucial for forming lipid bilayers in a high throughput manner (described later). Lipid membranes are formed on the top orifice of the micro2
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A prominent feature of our approach is that the lipid membranes spontaneously thin down to form bilayers without mechanical perturbation, e.g., hydraulic pressure, which is required for bilayer formation using decane or hexadecane. This lack of mechanical perturbation is critically important, because typically more than half of the lipid membranes are broken by the mechanical perturbation, resulting in an efficiency of < 30% for bilayer formation using these approaches [22]. Thinning of the lipid membranes is driven by drainage of the organic solvent over the hydrophobic support. Because chloroform is more water soluble than decane or hexadecane, and so can be more easily dissolved in an aqueous solution, it is more easily drained over a hydrophobic support, and thus, our approach does not require any mechanical perturbation, which contributes to increasing the efficiency of bilayer formation to over 97% (Fig. 1d).
2.1.1. Membrane potential Membrane potential, one of the main driving forces for transport by membrane proteins, is indispensable in cell physiology. For example, numerous types of ion channels in neurons are controlled by membrane potential, allowing for signal transmission via intra- or inter-cellular interfaces. To mimic physiological conditions, a microsystem with a nano-sized electrode has been developed to modulate membrane potential in a highly quantitative manner [18] (Fig. 2). The system is fabricated using conventional vacuum metal deposition and photolithography. First, 500-nm-thick Au layers and 20-nmthick chromium adhesion layers are coated onto the glass substrate using a vacuum evaporator. Following this, CYTOP is spin-coated onto the Au layer at a thickness of 500 nm. Photolithography is then conducted to pattern-mask structures onto the photo-resist covering the entire surface of the CYTOP layer. The resist-patterned substrate is then dry-etched with O2 plasma using a reactive-ion etching system to expose the Au surface, and regions bare of Au and Cr are then etched using Au and Cr etchants, respectively. A schematic illustration of the system is shown in Fig. 2a. The Au and fluororesin layers are used as a nano-sized electrode to modulate membrane voltage, and to support the lipid bilayer membrane, respectively. The through-hole structures on these layers are used as micro-chambers to detect biological reactions with high sensitivity. To evaluate membrane voltage, the micro-chambers are filled with a buffer solution containing DiBAC4 [24], a fluorescent indicator which increases its fluorescence intensity in proportion to the amplitude of the membrane voltage, and the lipid bilayers are then formed over the top orifices as described above (Fig. 2b). As shown in Fig. 2c, the fluorescence intensity of DiBAC4 changes following voltage modulation through the nano-sized electrodes. In addition, the change in intensity of DiBAC4 is highly stable for > 10 min. Thus, a long-term quantitative modulation of membrane voltage using a nano-electrode has been demonstrated in this microsystem.
Fig. 2. Membrane protential. (a) Photograph (top) and illustration (bottom) of the fabricated microsystem to modulate membrane potential. Microchambers (ϕ = 3 μm) were fabricated on a double layer of fluororesin (h = 500 nm) and Au (h = 500 nm). (b) Membrane potential monitoring. A membrane potential indicator, DiBAC4, was encapsulated in the chamber. Membrane potential was modulated using electrodes on a chamber and a top glass block. (c) Fluorescence intensity of DiBAC4 (green) against applied membrane potential using nano-sized electrodes (black).
2.1.2. Asymmetric phospholipid distribution Most biological membranes possess an asymmetric transbilayer phospholipid distribution, the maintenance of which requires endogenous enzymes to expend energy by promoting phospholipid translocation [25]. In particular, the plasma membrane of most eukaryotes maintains a high degree of asymmetry. For example, PS and PE are strictly confined to the inner leaflet, which controls various cellular functions, such as signal transduction, membrane fusion, and cell apoptosis [26–28]. Two different methods have been developed to achieve asymmetric phospholipid distributions in our system in order to reproduce the physiological environment (Fig. 3) [16,17]. First, a method was developed whereby two lipid monolayers are independently prepared, and then assembled on the top orifice of the micro-chamber to create an asymmetric distribution (top, Fig. 3a) [17]. The experimental procedure used for this is comprised of three steps.
First, an aqueous solution is infused into the flow channel of the system. After this infusion, the micro-chamber is filled with an aqueous solution. Second, a lipid solution is then infused to flush away the first aqueous solution. The chloroform solution used for this process contains a small amount of lipid, and therefore, lipid monolayers, but not multilayers, are formed at the top orifice of chambers. Third, a second chloroform solution containing a different lipid composition, and a second aqueous solution are sequentially infused to form a second lipid monolayer at their interface, which is then deposited over the entire surface of the system. In this process, the hydrocarbon tails in the second lipid layer are zipped together with those of the first lipid layer 3
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Fig. 3. Assymtery of phospholipid distribution. (a) Asymmetric lipid bilayer formation using two compositions of phospholipids. First, an aqueous solution is injected into the microsystem through an access port. Second, the first lipid solution containing fluorescent-labeled lipids (fluorescein-DHPE) is injected. Third, another lipid solution without fluorescein-DHPE is injected. Finally, a second aqueous solution is injected to flush the second lipid solution. During this process, asymmetric lipid bilayers are formed on the orifices of the microchambers. The bottom panel is the fluorescence image of fluorescein-DHPE on the asymmetric lipid bilayers. (b) Asymmetric lipid bilayer formation by irradiating a high-energy laser. 1) Membrane bilayers formed over microchambers were exposed a laser to 2) photobleach fluorescently labeled phospholipids (red). 3) Fluorescent lipids diffused into the bilayer from the surrounding area, but were then restricted to the outer layer. (c) The fluorescence images of microchambers, TopFluor-TMR-PS (red) before and after photo-bleaching.
The experimental procedures for this approach are as follows: First, lipid membranes are formed in the microsystem using a fatty-acid labeled fluorescent phospholipid, e.g., 18:1–6:0 TopFluor-TMR-PC, -PS, -PE or 18:1-6:0 NBD-PS (Avanti, USA). The fluorescence of the bilayer membrane in the micro-chamber can be clearly seen to be higher than that of the surrounding monolayers (Fig. 3b, top Fig. 3c). Following
to form an asymmetric bilayer over the micro-chamber, whereas monolayers are formed elsewhere. Notably, this method allows for the formation of > 100,000 asymmetric lipid bilayer membranes with an efficiency of over 97% (bottom, Fig. 3a). A second method for creating an asymmetric distribution of phospholipids involves irradiation with a high-energy laser (Fig. 3b) [16]. 4
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this, the fluorescent lipids in the bilayer membranes over the microchambers are photobleached by exposure to a high-energy laser for a few seconds in order to establish an asymmetric lipid distribution As shown in the bottom panel of Fig. 3c, the fluorescence in the microchambers gradually recovers over a period of several hundred seconds to the same level as seen on the surrounding surfaces, confirming that the membrane over the micro-chambers is contiguous with the monolayer in other parts of the system. The recovery of the fluorescence over the micro-chambers do not exceed the fluorescence of surrounding surfaces (Fig. 3b), indicating that the vertical translocation of phospholipids between the two layers of the membrane does not occur during measurement or is very slow, as has been observed in plasma membranes [29]. In other words, this result shows that photobleaching generates an asymmetric membrane bilayer in which unbleached fluorescent-phospholipids are present only in the upper layer. Notably, the asymmetry is maintained over 2 h after photo-bleaching, and is moreover reproduced at the same bilayers for many times by laser irradiations. These technical advantages are useful for single molecule analysis of phospholipid transport proteins, such as flippase, floppase, and scramblase (see below).
3. Part II. Single molecule analysis of membrane transport proteins Over the past few years, single molecule analysis has been demonstrated for a variety of membrane transport proteins, e.g., pores, channels, ion-pumps, and phospholipid scramblases, using “biomembrane microsystems” [5,16–21]. In this section, I would like to introduce the achievements made for the single molecule analysis of transport proteins. 3.1. Pore: α-hemolysin A transmembrane pore is a membrane protein that passively transport substrates along their concentration gradient. The activities of pores are relatively higher than other transport proteins, e.g., ionpumps, because both sides of the permeation pathway are always open, and as a result, they have been well characterized mechanistically using patch clamp recordings. A single molecule transport assay was carried out on our microsystem using α-hemolysin (Fig. 5) [30], a toxic membrane protein that binds to lipid bilayer membranes and forms transmembrane nanopores (ϕ = 1–2 nm) that have been used as various biomedical sensors, e.g., in nanopore DNA sequencing [31]. While monomers of α-hemolysin are soluble in water, once bound to a lipid bilayer, α-hemolysin assembles into a heptameric ring and forms a nanopore at the center, resulting in the passive transport of small molecules as small as 1 nm through its pore. For the assay, a fluorescent dye, Alexa 488, was encapsulated in the chambers as the transport substrate, and a low concentration of α-hemolysin solution (i.e., 1 μg mL−1) was injected into the flow channel after lipid bilayer formation (Fig. 5a). In this setup, the passive transport activity of α-hemolysin was detectable by analyzing the fluorescent intensity of the chamber, because Alexa 488 passively diffuses out of the chamber through the α-hemolysin pores, resulting in a decrease in chamber fluorescent intensity. Notably, a simple physicochemical model has been established for the passive transport by αhemolysin pores in the microsystem, where diffusion of the dye molecule into the α-hemolysin pore is the kinetic bottleneck, and obeys Fick's law [5,19]. In this model, the fluorescence intensity, F(t), of Alexa 488 encapsulated in the chamber is expressed by Eq. (1):
2.1.3. Generation of concentration gradient Owing to the small volume of the chambers in our system, a minute quantity of product is detectable, allowing for the direct monitoring of the individual activities of isolated single membrane proteins in a highly sensitive and quantitative manner. However, even though the chambers are highly integrated in the system, it is difficult to conduct multiple bioassays under different conditions using integrated chambers in parallel, because the composition of the reaction solution encapsulated in the chambers is uniform over the entire area of the system. Recently, this technical issue was addressed by developing a novel system capable of forming a variety of concentration gradients of target molecules encapsulated in the chambers (Fig. 4) [21]. In this system, based on the advection diffusion model, a concentration gradient of target molecules is formed along the flow channel via the sequential injection of several liquids from the access port (Fig. 4a). First, an aqueous solution containing the target molecules is infused into the flow channel to fill the individual chambers, as described above. Second, a specified amount of a second aqueous solution lacking target molecules (7–12 μL) is infused at a defined flow rate (0.5–1.5 μL/s) using an electric pipette. In this step, the concentration gradients are generated based on the advection-diffusion process. The aqueous solution first filled in the flow channel is gradually diluted from the inlet as the second solution for dilution is infused. Then, the concentration gradients are generated along the flow direction, i.e., the first solution is not completely diluted because the volume of second solution is smaller than that of the flow channel. Finally, to encapsulate the target molecules into the chambers, a chloroform solution containing phospholipids and a third aqueous solution are successively infused. After infusion, the lipid-bilayer membrane is formed on the orifice of the individual chambers, as described above, resulting in the encapsulation of the target molecules. To examine the feasibility of this approach, a fluorescent dye (Alexa 488) was used as an indicator of the concentration gradient. The concentration gradient was formed using the first or second aqueous solutions, with or without 1 μM Alexa 488, respectively. As expected, the fluorescence intensity of Alexa 488 encapsulated in the chambers increased along the flow channel (Fig. 4b, c), confirming the formation of a concentration gradient of target molecules in our system. Furthermore, the concentration gradient of the target molecule, i.e., Alexa 488, became steeper as the flow rate increased, and/or the volume of the second aqueous solution decreased (Fig. 4b). Accordingly, the concentration gradient in the micro-chamber array can be quantitatively modulated in proportion to the distance from the access port, L (Fig. 4b, c). So far, using this system, multiple single-molecule bioassays have been performed on the system under various conditions.
N ∙D∙d 2∙π ⎞ F (t ) = Fo ∙exp ⎛− t + F1 4∙L∙V ⎠ ⎝ ⎜
⎟
(1)
where N is the number of α-hemolysin pores reconstituted in the lipid membrane, D is the diffusion coefficient of Alexa 488, d is the diameter of the α-haemolysin pore (~1 nm), L is the length of the α-hemolysin pore (~10 nm), and V is the volume of the chamber. According to Eq. (1), the fluorescence intensity of the chamber exponentially decreases due to the passive transport activity of α-hemolysin with a rate constant proportional to the number of α-hemolysin pores (N). Fig. 5b, c show typical time courses of the fluorescent signals of 1 μM Alexa 488 encapsulated in 7 fL chambers, where the decay of fluorescent intensity represents passive transport through the α-hemolysin pore. The response of each individual chamber is not homogeneous (Fig. 5b), representing the stochastic formation of α-hemolysin pores in lipid bilayers. The distribution of the rate constant for fluorescent decay, determined by fitting with an exponential function (black line in Fig. 5c), exhibits four distinctive peaks (Fig. 5d). The intervals between the peaks are essentially constant, a typical feature of a singlemolecule digital assay [32,33], and each peak corresponds to the activity of 0, 1, 2, or 3 α-hemolysin pores. The rate constant of the passive diffusion through a single α-hemolysin pore was determined from the peak interval to be 5.5 ± 1.5 × 10−4 s−1. The initial transport flux of a single α-hemolysin pore (v) was thus estimated as v = n · k, where n is the number of 5
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Fig. 4. Concentration gradient. (a) Schematic illustration of the concentration gradient of target molecules. The concentration gradients were formed on individual micro-chambers depending on the distance (L) from the access port for sample injection. The individual micro-chambers were sealed with lipid-bilayer membranes. (b) Concentrations of Alexa 488 in micro-chambers are plotted against the flow rate (top) or volume of the second buffer (bottom) used to generate the concentration gradients. The solid lines represent linear regressions. (c) Fluorescence image of Alexa 488 (green) encapsulated in micro-chambers (h = 0.5 μm) at each L. The flow rate of liquid was 1.5 μL/s and the volume of the second buffer solution was 7 μL.
fluorescent dye molecules encapsulated in each chamber, and n = 7 fL × 1 μM × NA = 4.2 × 103 molecules. The initial transport flux per α-hemolysin pore was 2.3 molecules s−1, which is much lower than the detection limit of conventional patch clump recordings (~107 ions s−1). Thus, through the use of the biomembrane microsystem the highly sensitive detection of passive transport by a transmembrane pore can be achieved down to the single molecule level.
extensive biochemical studies have been conducted for ion-pumps, a single-molecule analysis has not achieved due to the technical difficulties in detecting slow transport events in highly sensitive manner. For the assay, RhP-M, a fluorescent pH indicator which increases its fluorescent intensity under acidic conditions [35], is encapsulated in the chamber, and a buffer solution containing a low concentration of FoF1 is injected into the flow channel after lipid bilayer formation. Following this, ATP is injected to initiate proton pumping. In this setup, FoF1 molecules with an outward orientation of the catalytic core domain can hydrolyze ATP to pump protons into the chamber, decreasing the pH, and thereby increasing the fluorescent intensity of RhP-M (Fig. 6a). Fig. 6b, c show typical time courses of fluorescent images obtained after ATP injection. Notably, some RhP-M molecules bound nonspecifically to the chamber wall due to its hydrophobicity, exhibiting ring-like shapes in fluorescent images (Fig. 6b). Similarly to the fluorescence signals obtained from the single-molecule α-hemolysin assay, the fluorescent signals in this assay show heterogeneity. As expected, Fig. 6b (left) exhibited discrete signal levels, representing the stochastic reconstitution of the FoF1 molecules. The active chambers reached a plateau at approximately 1600, which corresponds to a pH of
3.1.1. Ion pump: F-type ATPase An ion pump is a membrane protein that actively transports ions by consuming electro-chemical energy, e.g., ATP hydrolysis or ion-motive force (imf). The activities of ion pumps are relatively low (> 100 molecules s−1) because one side of the ion permeation pathway is always closed, i.e., they have to induce conformational changes to open and close the outward or inward gate on the pathway, and therefore, it is technically challenging to measure the low activity of ion pumps at the single molecule level. A single molecule analysis of an ion pump was carried out on our microsystem using the F-type ATPase (FoF1) (Fig. 6a), which mediates proton transport by coupling with ATP hydrolysis [34]. Although 6
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Fig. 5. Single molecule analysis of pore, α-hemolysin. (a) Schematic illustration of passive transport of α-hemolysin using the biomembrane microsytem. The fluorescent dye (Alexa 488) diffused out from the chamber via the pore on the membrane formed by α-hemolysin. (b) Fluorescent images of the passive transport activity in the chambers. The images were recorded just after αhemolysin injection (left panel) and 3000 s later (middle panel). The right panel, diff., shows the intensity difference between these two images in the form of a color gradient. (c) Continuous recording of the passive transport activity of 1 μg mL−1 α-hemolysin. The chambers containing 0 α-hemolysin pores, 1 α-hemolysin pore, 2 α-hemolysin pores, and 3 α-hemolysin pores were plotted as gray, blue, red, and green, respectively. Solid lines represent the fittings with the single exponential decay: y = C1 · exp.(−k · t) + C2, where k is the rate constant of passive transport. (d) In the histogram, the number of chambers is plotted versus the rate constant of passive transport, k. The four peaks can be attributed to occupancies of 0, 1, 2, or 3 α-hemolysin pores per chamber. They were fitted to a sum of Gaussians.
3.1.2. Phospholipid scramblase: TMEM16F Phospholipid scramblase is a membrane protein which translocates phospholipids between the inner and outer leaflets of cell membranes in order to disrupt their asymmetric lipid-distribution [36]. The biochemical features of scramblase have not been well characterized because the purification of intact scramblase is difficult, and moreover, with a few exceptions, membrane bilayers containing asymmetrically distributed phospholipids are challenging to construct [17,37,38]. A single molecule analysis of phospholipid scramblase is carried out using our microsystem [16] and the transmembrane protein 16F (TMEM16F), a Ca2+-dependent phospholipid scramblase [39] that mediates PS exposure in activated platelets during blood clotting, and regulates hydroxyapatite release from osteoblasts during bone mineralization (Fig. 7a) [40,41]. Despite its physiological importance, a single-molecule analysis has not been achieved so far due to the technical difficulties in preparing asymmetric lipid bilayers and in detecting lipid translocation in a highly sensitive manner. For the assay, a buffer solution containing a low concentration of TMEM16F is injected into the flow channel after lipid bilayer formation. Following this, an asymmetric distribution of the fluorescent lipid is formed on membrane
approximately 5.4. The transmembrane proton gradient (ΔpH) generated was approximately 1.7, which is at a level of equilibrium with the free energy of ATP hydrolysis. At the end of the observation period, a H+ ionophore, nigericin (arrow in Fig. 6c), was injected into the flow channel. The active chambers uniformly recovered their fluorescence toward the original level. This nigericin-induced fluorescence recovery ensures that the fluorescence increase represents the ATP-driven proton-pumping activity of FoF1. To estimate the proton transport rate for FoF1 during the initial phase, the initial rate is measured from the linear portion of the timecourse (typically from 1500 s to 4000 s) where the fluorescence increases at a constant rate (Fig. 6c). Similarly to the α-hemolysin assay, the distribution of the proton-pumping rate exhibites three distinct, regularly spaced peaks, indicating that each peak represented chambers with 0, 1, or 2 molecules of FoF1 (Fig. 6d). The rate of proton pumping by FoF1 is estimated from the intervals between the peaks to be 27.5 s−1. Thus, the biomembrane microsystem is the first to achieve the highly sensitive detection of active transport by an ion pump at the single molecule level.
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Fig. 6. Single molecule analysis of ion-pump, F-type ATPase. (a) Schematic illustration of F-type ATPase (FoF1) active transport. FoF1 pumps the proton from outside to inside the microchambers by hydrolyzing ATP. The fluorescent pH indicator (RhP-M) is encapsulated in the microchamber for pH monitoring. (b) Fluorescence images of the proton pumping of FoF1 in the chambers. The images were recorded just after the injection of 200 μM ATP (left panel) and 6000 s later (middle panel). The right panel, diff., shows the intensity difference between the left and middle panels in the form of a color gradient. (c) Continuous recording of proton pumping. The chambers containing 0 molecules, 1 molecule, and 2 molecules of active FoF1 were plotted as gray, blue, and red, respectively. (d) In the histogram, the number of chambers is plotted versus the slope of fluorescence increments from 1500 to 4000 s. The three peaks can be attributed to occupancies of 0, 1, or 2 active FoF1 molecules per chamber. They were fitted with a sum of Gaussians.
fluorescence up to their original level in the presence of 100 μM Ca2+, but not in the presence of 100 μM Ca2+ plus 1.0 μM of the inhibitor EGCg [42] (Fig. 7c), demonstrating that phospholipid scrambling by TMEM16F can be triggered more than once in our microsystem. A kinetic analysis is carried out by conducting the scrambling assay under various experimental conditions, e.g., different membrane sizes or at different temperatures. The scrambling catalyzed by TMEM16F slowes down (Fig. 7d) as the area of the bilayer membrane increased. Because TMEM16F-mediated scrambling is reversible [39], the reaction should follow the reaction
bilayers by irradiation with a high-energy laser, as described above. The scrambling assay is initiated by injecting 100 μM Ca2+ into the channel to activate the phospholipid scrambling activity of TMEM16F. Using this setup, the phospholipid translocation of TMEM16F is detected by analyzing the fluorescent intensity of the bilayers, because TMEM16F can transports fluorescent lipids from the outer leaflet to the inner leaflet of bilayers, resulting in an increase in their fluorescent intensity. Fig. 7b, c display typical time courses of fluorescent images obtained after Ca2+ injection, where the fluorescent increase in the bilayers represents the transport of a fluorescent lipid by TMEM16F. The response of each individual bilayer is not homogeneous (Fig. 7b), showing the stochastic reconstitution of TMEM16F molecules. The fluorescent intensity of the bilayers gradually increases to reach the original intensity level before photobleaching (Fig. 7c), irrespective of the phospholipid composition of the membranes. Notably, when active bilayers are laser-photobleached for a second time to re-create an asymmetric distribution, the bilayers gradually regain their
k
[Lipidout ] ⇄ [Lipidin] k
(3)
where k is the rate constant, and [Lipidout] and [Lipidin] are the concentrations of fluorescent lipids at the outer and inner layers, respectively. Notably, [Lipidout] is considered to be constant because it is contiguous with a large lipid monolayer in the surrounding areas. Since 8
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Fig. 7. Single molecule analysis of phospholipid scramblase, TMEM16F. (a) Schematic illustration of the phospholipid scrambling by TMEM16F. 1) TMEM16F was incorporated into the asymmetric membrane bilayers with fluorescent phospholipids (red). 2) Addition of CaCl2 initiated phospholipid scrambling, followed by lateral diffusion of unbleached phospholipids from surrounding areas, increasing the fluorescence intensity in microchambers. (b) Purified TMEM16F was loaded at 3.4 μg mL−1, and scrambling was initiated with 100 μM CaCl2. Subsequently, the asymmetric membrane bilayers were again formed by laser irradiation with injection of 1 μM EGCg. Maximum intensity projections of confocal images of TopFluor-TMR-PS (red) obtained from the same microchambers before (top) and after addition of 100 μM CaCl2 (middle) or 100 μM CaCl2 + 1 μM EGCg (bottom). Scale bar: 5 μm. (c) Time course of TMEM16F-mediated phospholipid scrambling. Lipid bilayers with (red) or without (gray) TMEM16F were treated with 100 μM CaCl2 (black arrowheads), and fluorescence from TopFluor-TMR-PS was followed over time. Subsequently, the microchambers were re-exposed for a few seconds to a laser at time points marked in orange, with (lower panel) or without (upper panel) prior injection of 1 μM EGCg (red arrowhead). (d) Time course of TMEM16F-mediated phospholipid scrambling at different area (S) of the bilayer membrane, 8.6 or 71 μm2. Black curves are fits to the single-exponential function y = C1 + C2 × (1 – exp.[−k × (t – 1000)]). (e) The rate constant of fluorescence increase (k*) of TopFluor-TMR-PS was plotted against the inverse of membrane area (1/S). Data in blue, red, and orange were obtained using TopFluor-TMR-PS at 16 °C, 25 °C, and 31 °C, respectively. Black lines are linear regressions. (f) An Arrhenius plot for TMEM16F-mediated phospholipid scrambling, with k values from Fig. 7e. The solid line represents linear regression, and the inset contains thermodynamic parameters at 25 °C, as determined from the plot.
size of micro-chambers (ɸ < 100 μm) drastically affect the efficiency of bilayer formation. In addition, the experimental conditions for reconstituting membrane proteins into lipid bilayers are widely varied (not universal) among protein species and ways of purification. These points represent remaining technical challenges for single molecule analysis using the biomembrane microsystems. Single-molecule analysis using micro-chamber arrays is an emerging approach to detect various bio-reactions, e.g., hydrolysis, protein synthesis, and membrane transport, with a high sensitivity down to the single-molecule level, enabling biomedical applications, such as digital PCR and digital ELISA. The biomembrane microsystem described here has broadened the versatility of micro-chambers to a single molecule analysis for membrane proteins, which are promising drug targets for the treatment of various diseases. Thus, this microsystem will enable further analytical and pharmacological applications, such as use in high throughput drug screening, and as an early diagnosis tool.
the fluorescent intensity over the chamber (FI) is proportional to the concentration of fluorescent lipids and the membrane area, FI is thus determined as a function of time t;
FI = FIout + FIin⋅[1 − exp(−k ∗⋅t )] k∗ =
k 1 ⋅ η S
(4)
(5)
where FIout and FIin are the fluorescent intensities of the outer and inner layers, respectively, S is the area of the membrane over the chambers, and η is the number of lipid molecules (~2.0 × 106/μm2) per unit area of the monolayer, as determined previously [43]. Fitting the time course in Fig. 7d to Eq. (4) demonstrates k*, the rate constant of fluorescent increase at 25 °C. As expected from Eq. (5), k* is inversely proportional to S (Fig. 7e), demonstrating that k, the rate of TMEM16Fmediated lipid transport, is 4.5 × 104 lipids/s at 25 °C (Fig. 7f); this value is ~108-fold faster than the spontaneous vertical lipid flip-flop across membrane bilayers [29]. Importantly, the k value does not differ between fluorescence substrates, such as TopFluor-TMR-PS, -PC, and -PE, indicating that TMEM16F does not distinguish between the head group moieties of phospholipids, and confirming that TMEM16F nonspecifically scrambles phospholipids. The TMEM16F-mediated scrambling is temperature dependent (Fig. 7e), with k values of 1.4 × 104 and 7.1 × 104 lipids/s at 16 °C and 35 °C, respectively. The corresponding Arrhenius plot fitted well to a linear function (Fig. 7f), indicating that TMEM16F-mediated lipid transport is governed by a single rate-limiting step, at least from 16 °C to 31 °C. The thermodynamic parameters of this rate-limiting step are as follows: ΔG‡ = 47 kJ/mol, ΔH‡ = 82 kJ/mol, and TΔS‡ = 35 kJ/mol at 25 °C (inset, Fig. 7f). The ΔG‡ value for spontaneous vertical lipid flipflop is approximately 100 kJ/mol [29], showing that TMEM16F significantly reduces the activation free energy. Thus, this biomembrane microsystem is the first to achieve a highly sensitive detection of phospholipid transport by phospholipid scramblase at the single molecule level.
Acknowledgements This work was supported by a Grant-in-Aid for Scientific Research (JP17H03660, JP15H05591) from the Japan Society for the Promotion of Science, a PRESTO Grant (JPMJPR13LC) from the Japan Science and Technology Agency, and a PRIME Grant (JP17gm0910020) from the Japan Agency for Medical Research and Development (to R.W.). References [1] M.H. Saier Jr., A functional-phylogenetic classification system for transmembrane solute transporters, Microbiol. Mol. Biol. Rev. 64 (2000) 354–411. [2] C. International Transporter, et al., Membrane transporters in drug development, Nat. Rev. Drug Disc. 9 (2010) 215–236. [3] K. Sugano, et al., Coexistence of passive and carrier-mediated processes in drug transport, Nat. Rev. Drug Disc. 9 (2010) 597–614. [4] S. Veshaguri, et al., Direct observation of proton pumping by a eukaryotic P-type ATPase, Science 351 (2016) 1469–1473. [5] R. Watanabe, et al., Arrayed lipid bilayer chambers allow single-molecule analysis of membrane transporter activity, Nat. Commun. 5 (2014) 4519. [6] A.J. Garcia-Saez, P. Schwille, Single molecule techniques for the study of membrane proteins, App. Microbiol. Biotechnol. 76 (2007) 257–266. [7] S.B. Kodandaramaiah, et al., Automated whole-cell patch-clamp electrophysiology of neurons in vivo, Nat. Methods 9 (2012) 585–587. [8] J. Dunlop, et al., High-throughput electrophysiology: an emerging paradigm for ionchannel screening and physiology, Nat. Rev. Drug Discov. 7 (2008) 358–368. [9] O. Keminer, et al., Optical recording of signal-mediated protein transport through single nuclear pore complexes, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 11842–11847. [10] H. Bayley, P.S. Cremer, Stochastic sensors inspired by biology, Nature 413 (2001) 226–230. [11] K. Sumitomo, et al., Ca2+ ion transport through channels formed by alpha-hemolysin analyzed using a microwell array on a Si substrate, Biosens. Bioelectron. 31 (2012) 445–450. [12] S. Ota, et al., Microfluidic lipid membrane formation on microchamber arrays, Lab Chip 11 (2011) 2485–2487. [13] T. Tonooka, et al., Lipid bilayers on a picoliter microdroplet array for rapid fluorescence detection of membrane transport, Small 10 (2014) 3275–3282. [14] M. Urban, et al., Highly parallel transport recordings on a membrane-on-nanopore chip at single molecule resolution, Nano Lett. 14 (2014) 1674–1680. [15] A. Kleefen, et al., Multiplexed parallel single transport recordings on nanopore
3.1.3. Future prospects A high-throughput single-molecule analysis of the membrane transport proteins, α-hemolysin, F-type ATPase, and TMEM16F has been achieved using the “biomembrane microsystem”. The microsystem has the following advantages for single molecule analysis: i) a high sensitivity using ultra-small chambers, ii) high throughput using chambers in parallel, and iii) a high compatibility for membrane protein analysis. These features may extend the use of our microsystem for single molecule analysis toward other membrane transport proteins, e.g., ion-channels, secondary active transporters, and phospholipid flippases/floppases. Such studies will reveal the generality and uniqueness of the operating principles of membrane transport proteins, thereby promoting in vitro membrane protein studies. The currently developed microsystem has several limitations. First, the concentration of bivalent cations (< 10 mM Ca2+ or Mg2+) and the 10
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[29] T.C. Anglin, J.C. Conboy, Kinetics and thermodynamics of flip-flop in binary phospholipid membranes measured by sum-frequency vibrational spectroscopy, Biochemistry 48 (2009) 10220–10234. [30] L. Song, et al., Structure of staphylococcal alpha-hemolysin, a heptameric transmembrane pore, Science 274 (1996) 1859–1866. [31] D. Branton, et al., The potential and challenges of nanopore sequencing, Nat. Biotechnol. 26 (2008) 1146–1153. [32] Y. Zhang, H. Noji, Digital bioassays: theory, applications, and perspectives, Anal. Chem. 89 (2017) 92–101. [33] D. Kim, et al., Research highlights: digital assays on chip, Lab Chip 15 (2015) 17–22. [34] W. Junge, et al., Torque generation and elastic power transmission in the rotary F0F1-ATPase, Nature 459 (2009) 364–370. [35] D. Asanuma, et al., Acidic-pH-activatable fluorescence probes for visualizing exocytosis dynamics, Angew. Chem. Int. Ed. Engl. 53 (2014) 6085–6089. [36] K. Segawa, S. Nagata, An apoptotic 'eat me' signal: phosphatidylserine exposure, Trends Cell Biol. 25 (2015) 649–650. [37] S. Pautot, et al., Engineering asymmetric vesicles, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 10718–10721. [38] M. Iwamoto, S. Oiki, Contact bubble bilayers with flush drainage, Sci. Rep. 5 (2015) 9110. [39] J. Suzuki, et al., Calcium-dependent phospholipid scrambling by TMEM16F, Nature 468 (2010) 834–838. [40] N. Pedemonte, L.J.V. Galietta, Structure and function of TMEM16 proteins (anoctamins), Physiol. Rev. 94 (2014) 419–459. [41] S. Nagata, et al., Exposure of phosphatidylserine on the cell surface, Cell Death Differ. 23 (2016) 952–961. [42] T. Suzuki, et al., Functional swapping between transmembrane proteins TMEM16A and TMEM16F, J. Biol. Chem. 289 (2014) 7438–7447. [43] J.F. Nagle, S. Tristram-Nagle, Structure of lipid bilayers, Biochim. Biophys. Acta 1469 (2000) 159–195.
arrays, Nano Lett. 10 (2010) 5080–5087. [16] R. Watanabe, et al., Single-molecule analysis of phospholipid scrambling by TMEM16F, Proc. Natl. Acad. Sci. U. S. A. 115 (2018) 3066–3071. [17] R. Watanabe, et al., High-throughput formation of lipid bilayer membrane arrays with an asymmetric lipid composition, Sci. Rep. 4 (2014) 7076. [18] R. Watanabe, et al., Novel nano-device to measure voltage-driven membrane transporter activity, IEEE Trans. Nanotechnol. 15 (2016) 70–73. [19] N. Soga, et al., Attolitre-sized lipid bilayer chamber array for rapid detection of single transporters, Sci. Rep. 5 (2015) 11025. [20] R. Watanabe, et al., Arrayed water-in-oil droplet bilayers for membrane transport analysis, Lab Chip 16 (2016) 3043–3048. [21] R. Watanabe, et al., High-throughput single-molecule bioassay using microchamber arrays with a concentration gradient of target molecules, Lab Chip 18 (2018) 2849–2853. [22] M. Zagnoni, Miniaturised technologies for the development of artificial lipid bilayer systems, Lab Chip 12 (2012) 1026–1039. [23] R. Iino, et al., A single-cell drug efflux assay in bacteria by using a directly accessible femtoliter droplet array, Lab Chip 12 (2012) 3923–3929. [24] D.E. Epps, et al., Characterization of the steady-state and dynamic fluorescence properties of the potential-sensitive dye bis-(1,3-dibutylbarbituric acid)trimethine oxonol (Dibac4(3)) in model systems and cells, Chem. Phys. Lipids 69 (1994) 137–150. [25] G. van Meer, et al., Membrane lipids: where they are and how they behave, Nat. Rev. Mol. Cell. Biol. 9 (2008) 112–124. [26] J.M. Boon, B.D. Smith, Chemical control of phospholipid distribution across bilayer membranes, Med. Res. Rev. 22 (2002) 251–281. [27] D.L. Bratton, et al., Appearance of phosphatidylserine on apoptotic cells requires calcium-mediated nonspecific flip-flop and is enhanced by loss of the aminophospholipid translocase, J. Biol. Chem. 272 (1997) 26159–26165. [28] V.A. Fadok, et al., Loss of phospholipid asymmetry and surface exposure of phosphatidylserine is required for phagocytosis of apoptotic cells by macrophages and fibroblasts, J. Biol. Chem. 276 (2001) 1071–1077.
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Contents lists available at ScienceDirect
BBA - General Subjects journal homepage: www.elsevier.com/locate/bbagen
Microsystem for the single molecule analysis of membrane transport proteins Rikiya Watanabe Molecular Physiology Laboratory, RIKEN, Saitama, Japan
A R T I C LE I N FO
A B S T R A C T
Keywords: Biomembrane Membrane protein Single molecule biophysics bioMEMS
Micro-chamber arrays enable highly sensitive and quantitative bioassays at the single-molecule level. Accordingly, they are widely used for ultra-sensitive biomedical applications, e.g., digital PCR and digital ELISA. However, the versatility of micro-chambers is generally limited to reactions in aqueous solutions, although various functions of membrane proteins are extremely important. To address this issue, microsystems using arrayed micro-sized chambers sealed with lipid bilayers, referred to here as a “biomembrane microsystems”, have been developed by many research groups for the analysis of membrane proteins. In this review, I would like to introduce recent progress on the single molecule analysis of membrane transport proteins using a biomembrane microsystem, and discuss the future prospects for its use in analytical and pharmacological applications.
1. Introduction The maintenance of an appropriate intracellular environment is a constant challenge for all living organisms, from prokaryotes to multicellular eukaryotes. Intracellular homeostasis is in general maintained by membrane proteins which mediate the translocation of various compounds across biomembranes [1]. Therefore, the analysis of transport mechanisms is crucial to understand cell physiology, as well as the action of drugs [2,3]. The transport of various substrates is mainly conducted by transport proteins embedded in the cell membrane. For several decades, the functional dynamics of transporters have been studied using a variety of single-molecule techniques [4,5], which offer key benefits over macroscopic assay methods, as they unlock the ability to quantify transport events [6]. One of the most robust systems used to analyze membrane transport at the single molecule level is patch clamp recording, which measures the flux of ions as an electric current under constant electrical voltage across a membrane. Recent developments have resulted in the automation of patch clamp recordings, thus enabling massively parallel analysis of transport activities, e.g. pores and channels [7,8]; however, transport rates of most transporters (< 102 molecules s−1) are much smaller than those of ion channels (> 107 molecules s−1), and therefore, it is difficult to detect their activities as an electric current. Moreover, patch clamp recording cannot detect the flux of electrically neutral substrates, and therefore, the development of a more versatile system has been long desired. Recently, microsystems arrayed with micro-sized chambers sealed
with lipid bilayers have been developed by many research groups for the analysis of membrane transport; in these systems, transport activity is measured optically based on the accumulation or consumption of a substrate present in the chambers [9–15]. Microsystems enhance both sensitivity and throughput, thereby allowing the highly sensitive analysis of various transporter proteins in a high throughput manner. Notably, our “biomembrane microsystem”, which contains > 100,000 arrayed atto- to femto-liter chambers sealed with stable lipid bilayers, has been used to perform the most sensitive and parallel analysis of membrane transport, enabling the single molecule analysis of transporters with extremely low transport activities (< 10 molecules s−1) [5,16]. Moreover, we have successfully demonstrated some physiological aspects of the membrane system, such as an asymmetric transbilayer phospholipid distribution [16,17], and the modulation of membrane potential across lipid bilayers [18]. Thus, our system paves the way for understanding the mechanism of membrane transport under semi-physiologic conditions, as well as allowing for further analytical and pharmacological applications. In this review, I would like to introduce our current microsystem, and discuss the future prospects for membrane transport analysis at the single molecule level. 2. Part I. Biomembrane microsystems Over the past few years, we have developed various “biomembrane microsystems” in order to carry out the single molecule analysis of membrane transport proteins under semi-physiological conditions [5,16–21]. The microsystem consists of a glass substrate containing the
E-mail address: [email protected]. https://doi.org/10.1016/j.bbagen.2019.03.016 Received 9 December 2018; Received in revised form 19 March 2019; Accepted 22 March 2019 0304-4165/ © 2019 Elsevier B.V. All rights reserved.
Please cite this article as: Rikiya Watanabe, BBA - General Subjects, https://doi.org/10.1016/j.bbagen.2019.03.016
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Fig. 1. Biomembrane microsystem. (a) Components of the system. Top, a glass block with a sample injection port (l: w: h = 20 mm: 20 mm: 5 mm); middle, a spacer sheet with one side open (Frame-Seal, BIO-RAD); bottom, a glass slide with > 10,000 microchambers printed. (b) Photograph (top) and illustration (bottom) of the assembled system. The access ports for sample injection are indicated using red circles. (c) Schematic illustration of the generation of phospholipid membranes. Membrane bilayers were formed over microchambers via sequential injection of liquids from the access port. Gray, blue, and yellow are the first aqueous solution, second aqueous solution, and chloroform solution containing lipids, respectively. (d) Fluorescent image of Alexa 647 encapsulated into microchambers sealed by lipid membranes.
chamber via the sequential injection of liquids from the access port (Fig. 1c). This technique is similar to the conventional technique known as the “painting technique,” which forms lipid bilayers through the deposition of a lipid solution across a small aperture in a hydrophobic material [12,23]. The experimental procedures used to form the lipid membrane are comprised of three steps. First, the micro-chambers are filled with an aqueous solution. A chloroform solution containing phospholipids, e.g., phosphatidylcholine (PC), phosphatidylethanolamine (PE), and phosphatidylserine (PS), is then infused via the access port. Due to the hydrophobicity of the top orifice, the interface between water and chloroform is formed over the orifice, where a lipid monolayer is then deposited. Finally, a second aqueous solution is infused to form another lipid monolayer at the interface with the chloroform solution, which is then deposited over the entire surface of the device. Thus, bilayers are formed over the micro-chambers, whereas monolayers are formed elsewhere. It is worth noting that we form lipid bilayers using chloroform as a solvent for the lipid, whereas decane or hexadecane have typically been used in conventional approaches [22].
microarray, a spacer sheet, and a glass block with an access port for sample injection (Fig. 1a). Microarrays with > 10,000 micro-chambers (ɸ = 2.4–8.2 μm and h = 0.03–0.5 μm: V = 0.2–71 fL) were fabricated using soft lithography on a hydrophilic glass substrate coated with CYTOP (Asahi-glass), a carbon‑fluorine hydrophobic polymer (Fig. 1b). Notably, in previous studies fluorine polymers have frequently been used as a support medium for lipid membranes [22]. The fabrication procedure is comprised of three steps. First, CYTOP is spin-coated onto a glass substrate of a specified thickness. Photolithography is then used to pattern mask the structures onto the photo-resist cover on the entire surface of the CYTOP layer. The resist-patterned substrate is then dryetched with O2 plasma using a reactive-ion etching system (RIE) to expose the glass substrate, resulting in the fabrication of the microchamber. Owing to the treatment with O2 plasma, the bottom and walls of the micro-chamber become hydrophilic, while the top orifice remains very hydrophobic. These different surfaces are crucial for forming lipid bilayers in a high throughput manner (described later). Lipid membranes are formed on the top orifice of the micro2
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A prominent feature of our approach is that the lipid membranes spontaneously thin down to form bilayers without mechanical perturbation, e.g., hydraulic pressure, which is required for bilayer formation using decane or hexadecane. This lack of mechanical perturbation is critically important, because typically more than half of the lipid membranes are broken by the mechanical perturbation, resulting in an efficiency of < 30% for bilayer formation using these approaches [22]. Thinning of the lipid membranes is driven by drainage of the organic solvent over the hydrophobic support. Because chloroform is more water soluble than decane or hexadecane, and so can be more easily dissolved in an aqueous solution, it is more easily drained over a hydrophobic support, and thus, our approach does not require any mechanical perturbation, which contributes to increasing the efficiency of bilayer formation to over 97% (Fig. 1d).
2.1.1. Membrane potential Membrane potential, one of the main driving forces for transport by membrane proteins, is indispensable in cell physiology. For example, numerous types of ion channels in neurons are controlled by membrane potential, allowing for signal transmission via intra- or inter-cellular interfaces. To mimic physiological conditions, a microsystem with a nano-sized electrode has been developed to modulate membrane potential in a highly quantitative manner [18] (Fig. 2). The system is fabricated using conventional vacuum metal deposition and photolithography. First, 500-nm-thick Au layers and 20-nmthick chromium adhesion layers are coated onto the glass substrate using a vacuum evaporator. Following this, CYTOP is spin-coated onto the Au layer at a thickness of 500 nm. Photolithography is then conducted to pattern-mask structures onto the photo-resist covering the entire surface of the CYTOP layer. The resist-patterned substrate is then dry-etched with O2 plasma using a reactive-ion etching system to expose the Au surface, and regions bare of Au and Cr are then etched using Au and Cr etchants, respectively. A schematic illustration of the system is shown in Fig. 2a. The Au and fluororesin layers are used as a nano-sized electrode to modulate membrane voltage, and to support the lipid bilayer membrane, respectively. The through-hole structures on these layers are used as micro-chambers to detect biological reactions with high sensitivity. To evaluate membrane voltage, the micro-chambers are filled with a buffer solution containing DiBAC4 [24], a fluorescent indicator which increases its fluorescence intensity in proportion to the amplitude of the membrane voltage, and the lipid bilayers are then formed over the top orifices as described above (Fig. 2b). As shown in Fig. 2c, the fluorescence intensity of DiBAC4 changes following voltage modulation through the nano-sized electrodes. In addition, the change in intensity of DiBAC4 is highly stable for > 10 min. Thus, a long-term quantitative modulation of membrane voltage using a nano-electrode has been demonstrated in this microsystem.
Fig. 2. Membrane protential. (a) Photograph (top) and illustration (bottom) of the fabricated microsystem to modulate membrane potential. Microchambers (ϕ = 3 μm) were fabricated on a double layer of fluororesin (h = 500 nm) and Au (h = 500 nm). (b) Membrane potential monitoring. A membrane potential indicator, DiBAC4, was encapsulated in the chamber. Membrane potential was modulated using electrodes on a chamber and a top glass block. (c) Fluorescence intensity of DiBAC4 (green) against applied membrane potential using nano-sized electrodes (black).
2.1.2. Asymmetric phospholipid distribution Most biological membranes possess an asymmetric transbilayer phospholipid distribution, the maintenance of which requires endogenous enzymes to expend energy by promoting phospholipid translocation [25]. In particular, the plasma membrane of most eukaryotes maintains a high degree of asymmetry. For example, PS and PE are strictly confined to the inner leaflet, which controls various cellular functions, such as signal transduction, membrane fusion, and cell apoptosis [26–28]. Two different methods have been developed to achieve asymmetric phospholipid distributions in our system in order to reproduce the physiological environment (Fig. 3) [16,17]. First, a method was developed whereby two lipid monolayers are independently prepared, and then assembled on the top orifice of the micro-chamber to create an asymmetric distribution (top, Fig. 3a) [17]. The experimental procedure used for this is comprised of three steps.
First, an aqueous solution is infused into the flow channel of the system. After this infusion, the micro-chamber is filled with an aqueous solution. Second, a lipid solution is then infused to flush away the first aqueous solution. The chloroform solution used for this process contains a small amount of lipid, and therefore, lipid monolayers, but not multilayers, are formed at the top orifice of chambers. Third, a second chloroform solution containing a different lipid composition, and a second aqueous solution are sequentially infused to form a second lipid monolayer at their interface, which is then deposited over the entire surface of the system. In this process, the hydrocarbon tails in the second lipid layer are zipped together with those of the first lipid layer 3
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Fig. 3. Assymtery of phospholipid distribution. (a) Asymmetric lipid bilayer formation using two compositions of phospholipids. First, an aqueous solution is injected into the microsystem through an access port. Second, the first lipid solution containing fluorescent-labeled lipids (fluorescein-DHPE) is injected. Third, another lipid solution without fluorescein-DHPE is injected. Finally, a second aqueous solution is injected to flush the second lipid solution. During this process, asymmetric lipid bilayers are formed on the orifices of the microchambers. The bottom panel is the fluorescence image of fluorescein-DHPE on the asymmetric lipid bilayers. (b) Asymmetric lipid bilayer formation by irradiating a high-energy laser. 1) Membrane bilayers formed over microchambers were exposed a laser to 2) photobleach fluorescently labeled phospholipids (red). 3) Fluorescent lipids diffused into the bilayer from the surrounding area, but were then restricted to the outer layer. (c) The fluorescence images of microchambers, TopFluor-TMR-PS (red) before and after photo-bleaching.
The experimental procedures for this approach are as follows: First, lipid membranes are formed in the microsystem using a fatty-acid labeled fluorescent phospholipid, e.g., 18:1–6:0 TopFluor-TMR-PC, -PS, -PE or 18:1-6:0 NBD-PS (Avanti, USA). The fluorescence of the bilayer membrane in the micro-chamber can be clearly seen to be higher than that of the surrounding monolayers (Fig. 3b, top Fig. 3c). Following
to form an asymmetric bilayer over the micro-chamber, whereas monolayers are formed elsewhere. Notably, this method allows for the formation of > 100,000 asymmetric lipid bilayer membranes with an efficiency of over 97% (bottom, Fig. 3a). A second method for creating an asymmetric distribution of phospholipids involves irradiation with a high-energy laser (Fig. 3b) [16]. 4
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this, the fluorescent lipids in the bilayer membranes over the microchambers are photobleached by exposure to a high-energy laser for a few seconds in order to establish an asymmetric lipid distribution As shown in the bottom panel of Fig. 3c, the fluorescence in the microchambers gradually recovers over a period of several hundred seconds to the same level as seen on the surrounding surfaces, confirming that the membrane over the micro-chambers is contiguous with the monolayer in other parts of the system. The recovery of the fluorescence over the micro-chambers do not exceed the fluorescence of surrounding surfaces (Fig. 3b), indicating that the vertical translocation of phospholipids between the two layers of the membrane does not occur during measurement or is very slow, as has been observed in plasma membranes [29]. In other words, this result shows that photobleaching generates an asymmetric membrane bilayer in which unbleached fluorescent-phospholipids are present only in the upper layer. Notably, the asymmetry is maintained over 2 h after photo-bleaching, and is moreover reproduced at the same bilayers for many times by laser irradiations. These technical advantages are useful for single molecule analysis of phospholipid transport proteins, such as flippase, floppase, and scramblase (see below).
3. Part II. Single molecule analysis of membrane transport proteins Over the past few years, single molecule analysis has been demonstrated for a variety of membrane transport proteins, e.g., pores, channels, ion-pumps, and phospholipid scramblases, using “biomembrane microsystems” [5,16–21]. In this section, I would like to introduce the achievements made for the single molecule analysis of transport proteins. 3.1. Pore: α-hemolysin A transmembrane pore is a membrane protein that passively transport substrates along their concentration gradient. The activities of pores are relatively higher than other transport proteins, e.g., ionpumps, because both sides of the permeation pathway are always open, and as a result, they have been well characterized mechanistically using patch clamp recordings. A single molecule transport assay was carried out on our microsystem using α-hemolysin (Fig. 5) [30], a toxic membrane protein that binds to lipid bilayer membranes and forms transmembrane nanopores (ϕ = 1–2 nm) that have been used as various biomedical sensors, e.g., in nanopore DNA sequencing [31]. While monomers of α-hemolysin are soluble in water, once bound to a lipid bilayer, α-hemolysin assembles into a heptameric ring and forms a nanopore at the center, resulting in the passive transport of small molecules as small as 1 nm through its pore. For the assay, a fluorescent dye, Alexa 488, was encapsulated in the chambers as the transport substrate, and a low concentration of α-hemolysin solution (i.e., 1 μg mL−1) was injected into the flow channel after lipid bilayer formation (Fig. 5a). In this setup, the passive transport activity of α-hemolysin was detectable by analyzing the fluorescent intensity of the chamber, because Alexa 488 passively diffuses out of the chamber through the α-hemolysin pores, resulting in a decrease in chamber fluorescent intensity. Notably, a simple physicochemical model has been established for the passive transport by αhemolysin pores in the microsystem, where diffusion of the dye molecule into the α-hemolysin pore is the kinetic bottleneck, and obeys Fick's law [5,19]. In this model, the fluorescence intensity, F(t), of Alexa 488 encapsulated in the chamber is expressed by Eq. (1):
2.1.3. Generation of concentration gradient Owing to the small volume of the chambers in our system, a minute quantity of product is detectable, allowing for the direct monitoring of the individual activities of isolated single membrane proteins in a highly sensitive and quantitative manner. However, even though the chambers are highly integrated in the system, it is difficult to conduct multiple bioassays under different conditions using integrated chambers in parallel, because the composition of the reaction solution encapsulated in the chambers is uniform over the entire area of the system. Recently, this technical issue was addressed by developing a novel system capable of forming a variety of concentration gradients of target molecules encapsulated in the chambers (Fig. 4) [21]. In this system, based on the advection diffusion model, a concentration gradient of target molecules is formed along the flow channel via the sequential injection of several liquids from the access port (Fig. 4a). First, an aqueous solution containing the target molecules is infused into the flow channel to fill the individual chambers, as described above. Second, a specified amount of a second aqueous solution lacking target molecules (7–12 μL) is infused at a defined flow rate (0.5–1.5 μL/s) using an electric pipette. In this step, the concentration gradients are generated based on the advection-diffusion process. The aqueous solution first filled in the flow channel is gradually diluted from the inlet as the second solution for dilution is infused. Then, the concentration gradients are generated along the flow direction, i.e., the first solution is not completely diluted because the volume of second solution is smaller than that of the flow channel. Finally, to encapsulate the target molecules into the chambers, a chloroform solution containing phospholipids and a third aqueous solution are successively infused. After infusion, the lipid-bilayer membrane is formed on the orifice of the individual chambers, as described above, resulting in the encapsulation of the target molecules. To examine the feasibility of this approach, a fluorescent dye (Alexa 488) was used as an indicator of the concentration gradient. The concentration gradient was formed using the first or second aqueous solutions, with or without 1 μM Alexa 488, respectively. As expected, the fluorescence intensity of Alexa 488 encapsulated in the chambers increased along the flow channel (Fig. 4b, c), confirming the formation of a concentration gradient of target molecules in our system. Furthermore, the concentration gradient of the target molecule, i.e., Alexa 488, became steeper as the flow rate increased, and/or the volume of the second aqueous solution decreased (Fig. 4b). Accordingly, the concentration gradient in the micro-chamber array can be quantitatively modulated in proportion to the distance from the access port, L (Fig. 4b, c). So far, using this system, multiple single-molecule bioassays have been performed on the system under various conditions.
N ∙D∙d 2∙π ⎞ F (t ) = Fo ∙exp ⎛− t + F1 4∙L∙V ⎠ ⎝ ⎜
⎟
(1)
where N is the number of α-hemolysin pores reconstituted in the lipid membrane, D is the diffusion coefficient of Alexa 488, d is the diameter of the α-haemolysin pore (~1 nm), L is the length of the α-hemolysin pore (~10 nm), and V is the volume of the chamber. According to Eq. (1), the fluorescence intensity of the chamber exponentially decreases due to the passive transport activity of α-hemolysin with a rate constant proportional to the number of α-hemolysin pores (N). Fig. 5b, c show typical time courses of the fluorescent signals of 1 μM Alexa 488 encapsulated in 7 fL chambers, where the decay of fluorescent intensity represents passive transport through the α-hemolysin pore. The response of each individual chamber is not homogeneous (Fig. 5b), representing the stochastic formation of α-hemolysin pores in lipid bilayers. The distribution of the rate constant for fluorescent decay, determined by fitting with an exponential function (black line in Fig. 5c), exhibits four distinctive peaks (Fig. 5d). The intervals between the peaks are essentially constant, a typical feature of a singlemolecule digital assay [32,33], and each peak corresponds to the activity of 0, 1, 2, or 3 α-hemolysin pores. The rate constant of the passive diffusion through a single α-hemolysin pore was determined from the peak interval to be 5.5 ± 1.5 × 10−4 s−1. The initial transport flux of a single α-hemolysin pore (v) was thus estimated as v = n · k, where n is the number of 5
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Fig. 4. Concentration gradient. (a) Schematic illustration of the concentration gradient of target molecules. The concentration gradients were formed on individual micro-chambers depending on the distance (L) from the access port for sample injection. The individual micro-chambers were sealed with lipid-bilayer membranes. (b) Concentrations of Alexa 488 in micro-chambers are plotted against the flow rate (top) or volume of the second buffer (bottom) used to generate the concentration gradients. The solid lines represent linear regressions. (c) Fluorescence image of Alexa 488 (green) encapsulated in micro-chambers (h = 0.5 μm) at each L. The flow rate of liquid was 1.5 μL/s and the volume of the second buffer solution was 7 μL.
fluorescent dye molecules encapsulated in each chamber, and n = 7 fL × 1 μM × NA = 4.2 × 103 molecules. The initial transport flux per α-hemolysin pore was 2.3 molecules s−1, which is much lower than the detection limit of conventional patch clump recordings (~107 ions s−1). Thus, through the use of the biomembrane microsystem the highly sensitive detection of passive transport by a transmembrane pore can be achieved down to the single molecule level.
extensive biochemical studies have been conducted for ion-pumps, a single-molecule analysis has not achieved due to the technical difficulties in detecting slow transport events in highly sensitive manner. For the assay, RhP-M, a fluorescent pH indicator which increases its fluorescent intensity under acidic conditions [35], is encapsulated in the chamber, and a buffer solution containing a low concentration of FoF1 is injected into the flow channel after lipid bilayer formation. Following this, ATP is injected to initiate proton pumping. In this setup, FoF1 molecules with an outward orientation of the catalytic core domain can hydrolyze ATP to pump protons into the chamber, decreasing the pH, and thereby increasing the fluorescent intensity of RhP-M (Fig. 6a). Fig. 6b, c show typical time courses of fluorescent images obtained after ATP injection. Notably, some RhP-M molecules bound nonspecifically to the chamber wall due to its hydrophobicity, exhibiting ring-like shapes in fluorescent images (Fig. 6b). Similarly to the fluorescence signals obtained from the single-molecule α-hemolysin assay, the fluorescent signals in this assay show heterogeneity. As expected, Fig. 6b (left) exhibited discrete signal levels, representing the stochastic reconstitution of the FoF1 molecules. The active chambers reached a plateau at approximately 1600, which corresponds to a pH of
3.1.1. Ion pump: F-type ATPase An ion pump is a membrane protein that actively transports ions by consuming electro-chemical energy, e.g., ATP hydrolysis or ion-motive force (imf). The activities of ion pumps are relatively low (> 100 molecules s−1) because one side of the ion permeation pathway is always closed, i.e., they have to induce conformational changes to open and close the outward or inward gate on the pathway, and therefore, it is technically challenging to measure the low activity of ion pumps at the single molecule level. A single molecule analysis of an ion pump was carried out on our microsystem using the F-type ATPase (FoF1) (Fig. 6a), which mediates proton transport by coupling with ATP hydrolysis [34]. Although 6
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Fig. 5. Single molecule analysis of pore, α-hemolysin. (a) Schematic illustration of passive transport of α-hemolysin using the biomembrane microsytem. The fluorescent dye (Alexa 488) diffused out from the chamber via the pore on the membrane formed by α-hemolysin. (b) Fluorescent images of the passive transport activity in the chambers. The images were recorded just after αhemolysin injection (left panel) and 3000 s later (middle panel). The right panel, diff., shows the intensity difference between these two images in the form of a color gradient. (c) Continuous recording of the passive transport activity of 1 μg mL−1 α-hemolysin. The chambers containing 0 α-hemolysin pores, 1 α-hemolysin pore, 2 α-hemolysin pores, and 3 α-hemolysin pores were plotted as gray, blue, red, and green, respectively. Solid lines represent the fittings with the single exponential decay: y = C1 · exp.(−k · t) + C2, where k is the rate constant of passive transport. (d) In the histogram, the number of chambers is plotted versus the rate constant of passive transport, k. The four peaks can be attributed to occupancies of 0, 1, 2, or 3 α-hemolysin pores per chamber. They were fitted to a sum of Gaussians.
3.1.2. Phospholipid scramblase: TMEM16F Phospholipid scramblase is a membrane protein which translocates phospholipids between the inner and outer leaflets of cell membranes in order to disrupt their asymmetric lipid-distribution [36]. The biochemical features of scramblase have not been well characterized because the purification of intact scramblase is difficult, and moreover, with a few exceptions, membrane bilayers containing asymmetrically distributed phospholipids are challenging to construct [17,37,38]. A single molecule analysis of phospholipid scramblase is carried out using our microsystem [16] and the transmembrane protein 16F (TMEM16F), a Ca2+-dependent phospholipid scramblase [39] that mediates PS exposure in activated platelets during blood clotting, and regulates hydroxyapatite release from osteoblasts during bone mineralization (Fig. 7a) [40,41]. Despite its physiological importance, a single-molecule analysis has not been achieved so far due to the technical difficulties in preparing asymmetric lipid bilayers and in detecting lipid translocation in a highly sensitive manner. For the assay, a buffer solution containing a low concentration of TMEM16F is injected into the flow channel after lipid bilayer formation. Following this, an asymmetric distribution of the fluorescent lipid is formed on membrane
approximately 5.4. The transmembrane proton gradient (ΔpH) generated was approximately 1.7, which is at a level of equilibrium with the free energy of ATP hydrolysis. At the end of the observation period, a H+ ionophore, nigericin (arrow in Fig. 6c), was injected into the flow channel. The active chambers uniformly recovered their fluorescence toward the original level. This nigericin-induced fluorescence recovery ensures that the fluorescence increase represents the ATP-driven proton-pumping activity of FoF1. To estimate the proton transport rate for FoF1 during the initial phase, the initial rate is measured from the linear portion of the timecourse (typically from 1500 s to 4000 s) where the fluorescence increases at a constant rate (Fig. 6c). Similarly to the α-hemolysin assay, the distribution of the proton-pumping rate exhibites three distinct, regularly spaced peaks, indicating that each peak represented chambers with 0, 1, or 2 molecules of FoF1 (Fig. 6d). The rate of proton pumping by FoF1 is estimated from the intervals between the peaks to be 27.5 s−1. Thus, the biomembrane microsystem is the first to achieve the highly sensitive detection of active transport by an ion pump at the single molecule level.
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Fig. 6. Single molecule analysis of ion-pump, F-type ATPase. (a) Schematic illustration of F-type ATPase (FoF1) active transport. FoF1 pumps the proton from outside to inside the microchambers by hydrolyzing ATP. The fluorescent pH indicator (RhP-M) is encapsulated in the microchamber for pH monitoring. (b) Fluorescence images of the proton pumping of FoF1 in the chambers. The images were recorded just after the injection of 200 μM ATP (left panel) and 6000 s later (middle panel). The right panel, diff., shows the intensity difference between the left and middle panels in the form of a color gradient. (c) Continuous recording of proton pumping. The chambers containing 0 molecules, 1 molecule, and 2 molecules of active FoF1 were plotted as gray, blue, and red, respectively. (d) In the histogram, the number of chambers is plotted versus the slope of fluorescence increments from 1500 to 4000 s. The three peaks can be attributed to occupancies of 0, 1, or 2 active FoF1 molecules per chamber. They were fitted with a sum of Gaussians.
fluorescence up to their original level in the presence of 100 μM Ca2+, but not in the presence of 100 μM Ca2+ plus 1.0 μM of the inhibitor EGCg [42] (Fig. 7c), demonstrating that phospholipid scrambling by TMEM16F can be triggered more than once in our microsystem. A kinetic analysis is carried out by conducting the scrambling assay under various experimental conditions, e.g., different membrane sizes or at different temperatures. The scrambling catalyzed by TMEM16F slowes down (Fig. 7d) as the area of the bilayer membrane increased. Because TMEM16F-mediated scrambling is reversible [39], the reaction should follow the reaction
bilayers by irradiation with a high-energy laser, as described above. The scrambling assay is initiated by injecting 100 μM Ca2+ into the channel to activate the phospholipid scrambling activity of TMEM16F. Using this setup, the phospholipid translocation of TMEM16F is detected by analyzing the fluorescent intensity of the bilayers, because TMEM16F can transports fluorescent lipids from the outer leaflet to the inner leaflet of bilayers, resulting in an increase in their fluorescent intensity. Fig. 7b, c display typical time courses of fluorescent images obtained after Ca2+ injection, where the fluorescent increase in the bilayers represents the transport of a fluorescent lipid by TMEM16F. The response of each individual bilayer is not homogeneous (Fig. 7b), showing the stochastic reconstitution of TMEM16F molecules. The fluorescent intensity of the bilayers gradually increases to reach the original intensity level before photobleaching (Fig. 7c), irrespective of the phospholipid composition of the membranes. Notably, when active bilayers are laser-photobleached for a second time to re-create an asymmetric distribution, the bilayers gradually regain their
k
[Lipidout ] ⇄ [Lipidin] k
(3)
where k is the rate constant, and [Lipidout] and [Lipidin] are the concentrations of fluorescent lipids at the outer and inner layers, respectively. Notably, [Lipidout] is considered to be constant because it is contiguous with a large lipid monolayer in the surrounding areas. Since 8
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(caption on next page)
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Fig. 7. Single molecule analysis of phospholipid scramblase, TMEM16F. (a) Schematic illustration of the phospholipid scrambling by TMEM16F. 1) TMEM16F was incorporated into the asymmetric membrane bilayers with fluorescent phospholipids (red). 2) Addition of CaCl2 initiated phospholipid scrambling, followed by lateral diffusion of unbleached phospholipids from surrounding areas, increasing the fluorescence intensity in microchambers. (b) Purified TMEM16F was loaded at 3.4 μg mL−1, and scrambling was initiated with 100 μM CaCl2. Subsequently, the asymmetric membrane bilayers were again formed by laser irradiation with injection of 1 μM EGCg. Maximum intensity projections of confocal images of TopFluor-TMR-PS (red) obtained from the same microchambers before (top) and after addition of 100 μM CaCl2 (middle) or 100 μM CaCl2 + 1 μM EGCg (bottom). Scale bar: 5 μm. (c) Time course of TMEM16F-mediated phospholipid scrambling. Lipid bilayers with (red) or without (gray) TMEM16F were treated with 100 μM CaCl2 (black arrowheads), and fluorescence from TopFluor-TMR-PS was followed over time. Subsequently, the microchambers were re-exposed for a few seconds to a laser at time points marked in orange, with (lower panel) or without (upper panel) prior injection of 1 μM EGCg (red arrowhead). (d) Time course of TMEM16F-mediated phospholipid scrambling at different area (S) of the bilayer membrane, 8.6 or 71 μm2. Black curves are fits to the single-exponential function y = C1 + C2 × (1 – exp.[−k × (t – 1000)]). (e) The rate constant of fluorescence increase (k*) of TopFluor-TMR-PS was plotted against the inverse of membrane area (1/S). Data in blue, red, and orange were obtained using TopFluor-TMR-PS at 16 °C, 25 °C, and 31 °C, respectively. Black lines are linear regressions. (f) An Arrhenius plot for TMEM16F-mediated phospholipid scrambling, with k values from Fig. 7e. The solid line represents linear regression, and the inset contains thermodynamic parameters at 25 °C, as determined from the plot.
size of micro-chambers (ɸ < 100 μm) drastically affect the efficiency of bilayer formation. In addition, the experimental conditions for reconstituting membrane proteins into lipid bilayers are widely varied (not universal) among protein species and ways of purification. These points represent remaining technical challenges for single molecule analysis using the biomembrane microsystems. Single-molecule analysis using micro-chamber arrays is an emerging approach to detect various bio-reactions, e.g., hydrolysis, protein synthesis, and membrane transport, with a high sensitivity down to the single-molecule level, enabling biomedical applications, such as digital PCR and digital ELISA. The biomembrane microsystem described here has broadened the versatility of micro-chambers to a single molecule analysis for membrane proteins, which are promising drug targets for the treatment of various diseases. Thus, this microsystem will enable further analytical and pharmacological applications, such as use in high throughput drug screening, and as an early diagnosis tool.
the fluorescent intensity over the chamber (FI) is proportional to the concentration of fluorescent lipids and the membrane area, FI is thus determined as a function of time t;
FI = FIout + FIin⋅[1 − exp(−k ∗⋅t )] k∗ =
k 1 ⋅ η S
(4)
(5)
where FIout and FIin are the fluorescent intensities of the outer and inner layers, respectively, S is the area of the membrane over the chambers, and η is the number of lipid molecules (~2.0 × 106/μm2) per unit area of the monolayer, as determined previously [43]. Fitting the time course in Fig. 7d to Eq. (4) demonstrates k*, the rate constant of fluorescent increase at 25 °C. As expected from Eq. (5), k* is inversely proportional to S (Fig. 7e), demonstrating that k, the rate of TMEM16Fmediated lipid transport, is 4.5 × 104 lipids/s at 25 °C (Fig. 7f); this value is ~108-fold faster than the spontaneous vertical lipid flip-flop across membrane bilayers [29]. Importantly, the k value does not differ between fluorescence substrates, such as TopFluor-TMR-PS, -PC, and -PE, indicating that TMEM16F does not distinguish between the head group moieties of phospholipids, and confirming that TMEM16F nonspecifically scrambles phospholipids. The TMEM16F-mediated scrambling is temperature dependent (Fig. 7e), with k values of 1.4 × 104 and 7.1 × 104 lipids/s at 16 °C and 35 °C, respectively. The corresponding Arrhenius plot fitted well to a linear function (Fig. 7f), indicating that TMEM16F-mediated lipid transport is governed by a single rate-limiting step, at least from 16 °C to 31 °C. The thermodynamic parameters of this rate-limiting step are as follows: ΔG‡ = 47 kJ/mol, ΔH‡ = 82 kJ/mol, and TΔS‡ = 35 kJ/mol at 25 °C (inset, Fig. 7f). The ΔG‡ value for spontaneous vertical lipid flipflop is approximately 100 kJ/mol [29], showing that TMEM16F significantly reduces the activation free energy. Thus, this biomembrane microsystem is the first to achieve a highly sensitive detection of phospholipid transport by phospholipid scramblase at the single molecule level.
Acknowledgements This work was supported by a Grant-in-Aid for Scientific Research (JP17H03660, JP15H05591) from the Japan Society for the Promotion of Science, a PRESTO Grant (JPMJPR13LC) from the Japan Science and Technology Agency, and a PRIME Grant (JP17gm0910020) from the Japan Agency for Medical Research and Development (to R.W.). References [1] M.H. Saier Jr., A functional-phylogenetic classification system for transmembrane solute transporters, Microbiol. Mol. Biol. Rev. 64 (2000) 354–411. [2] C. International Transporter, et al., Membrane transporters in drug development, Nat. Rev. Drug Disc. 9 (2010) 215–236. [3] K. Sugano, et al., Coexistence of passive and carrier-mediated processes in drug transport, Nat. Rev. Drug Disc. 9 (2010) 597–614. [4] S. Veshaguri, et al., Direct observation of proton pumping by a eukaryotic P-type ATPase, Science 351 (2016) 1469–1473. [5] R. Watanabe, et al., Arrayed lipid bilayer chambers allow single-molecule analysis of membrane transporter activity, Nat. Commun. 5 (2014) 4519. [6] A.J. Garcia-Saez, P. Schwille, Single molecule techniques for the study of membrane proteins, App. Microbiol. Biotechnol. 76 (2007) 257–266. [7] S.B. Kodandaramaiah, et al., Automated whole-cell patch-clamp electrophysiology of neurons in vivo, Nat. Methods 9 (2012) 585–587. [8] J. Dunlop, et al., High-throughput electrophysiology: an emerging paradigm for ionchannel screening and physiology, Nat. Rev. Drug Discov. 7 (2008) 358–368. [9] O. Keminer, et al., Optical recording of signal-mediated protein transport through single nuclear pore complexes, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 11842–11847. [10] H. Bayley, P.S. Cremer, Stochastic sensors inspired by biology, Nature 413 (2001) 226–230. [11] K. Sumitomo, et al., Ca2+ ion transport through channels formed by alpha-hemolysin analyzed using a microwell array on a Si substrate, Biosens. Bioelectron. 31 (2012) 445–450. [12] S. Ota, et al., Microfluidic lipid membrane formation on microchamber arrays, Lab Chip 11 (2011) 2485–2487. [13] T. Tonooka, et al., Lipid bilayers on a picoliter microdroplet array for rapid fluorescence detection of membrane transport, Small 10 (2014) 3275–3282. [14] M. Urban, et al., Highly parallel transport recordings on a membrane-on-nanopore chip at single molecule resolution, Nano Lett. 14 (2014) 1674–1680. [15] A. Kleefen, et al., Multiplexed parallel single transport recordings on nanopore
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