Static and Dynamic Permeability Assay for Hydrophilic Small

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A Static and Dynamic Permeability Assay for Hydrophilic Small Molecules Using a Planar Droplet Interface Bilayer Yohan Lee, Hyun-Ro Lee, KyuHan Kim, and Siyoung Q. Choi Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b03004 • Publication Date (Web): 08 Jan 2018 Downloaded from http://pubs.acs.org on January 8, 2018

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Analytical Chemistry

A Static and Dynamic Permeability Assay for Hydrophilic Small Molecules Using a Planar Droplet Interface Bilayer

Yohan Lee, Hyun-Ro Lee, KyuHan Kim*, Siyoung Q. Choi* Department of Chemical and Biomolecular engineering, Korea Advanced Institute of Science and Technology (KAIST), Daejeon 34141, Korea *Corresponding author, E-mail: [email protected], [email protected]

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Abstract Because numerous drugs are administered through an oral route and primarily absorbed at the intestine, the prediction of drug permeability across an intestinal epithelial cell membrane has been a crucial issue in drug discovery. Thus, various in vitro permeability assays have been developed such as the Caco-2 assay, the parallel artificial membrane permeability assay (PAMPA), the phospholipid vesicle-based permeation assays (PVPA) and Permeapad®. However, due to the time-consuming and quite expensive process for culturing cells in the Caco-2 assay and the unknown microscopic membrane structures of the other assays, a simpler yet more accurate and versatile technique is still required. Accordingly, we developed a new platform to measure the permeability of small molecules across a planar freestanding lipid bilayer with a well-defined area and structure. The lipid bilayer was constructed within a conventional UV spectrometer cell, and the transport of drug molecules across the bilayer was recorded by UV absorbance over time. We then computed the permeability from the time-dependent diffusion equation. We tested this assay for five exemplary hydrophilic drugs and compared their values with previously reported ones. We found that our assay has a much higher permeability compared to the other techniques, and this higher permeability is related to the thickness of the lipid bilayer. Also we were able to measure the dynamic permeability upon the addition of a membrane-disrupting surfactant demonstrating that our assay has the capability to detect real-time changes in permeability across the lipid bilayer.


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Biological cells are encapsulated by a cell membrane separating their internal contents from external environments.1,2 Mass transport across the cell membrane is ubiquitous and is closely related to numerous physiological phenomena3–10 such as the uptake of essential nutrients into cells,3,4 the absorption of small molecules through the cell membrane,5,6 cell-to-cell communications by releasing and accepting signaling molecules,7,8 and the maintenance of the ion concentration gradient to control the proper membrane potential.9,10 Mass transport through cell membranes is also important for new drug discovery because drug absorption through a cell membrane is crucial in determining its future efficiency. Drug administration through an oral route is the most frequently used method because of its convenience. These drug molecules are mostly absorbed at the intestine in the human body.11,12 It thus has been emphasized that estimating the permeability across membranes of an intestinal epithelial cell for screening effective drug candidates is essential. Hence, various in vitro assays have been developed to estimate the drug permeability.13–17 In the Caco-2 assay,18–24 cells derived from human colon carcinoma cells are cultivated on permeable filters and form monolayers that are similar to intestinal epithelial cells structurally and functionally. By mimicking the human intestinal environment, this technique could provide a more accurate prediction of the absorption of drug candidates at the human intestine. However, the Caco-2 assay contains time-consuming and quite expensive processes for culturing cells, and its reproducibility for measuring drug permeability is still poor as well.25–28 Meanwhile, a passive diffusion process that is driven by a concentration gradient of drug molecules across the cell membrane is a very important transport mechanism in the intestine.29,30 Therefore, a precise quantification of the passive diffusion across the cell



membranes in vitro would be extremely helpful for the first screening of newly developed drugs in the pharmaceutical industry, so various much simpler in vitro assays considering only the passive diffusion, have been developed as well. For example, parallel artificial membrane permeability assay (PAMPA), first introduced by Kansy et al.,31 (ref 31) uses an artificial membrane and this artificial membrane is conventionally prepared by immersing a porous filter into a lipid solution.14,31–36 Even though PAMPA is a much simpler and costeffective method, compared to the Caco-2 assay, it has been known that these PAMPAs have oil residues within the membrane structure, and how this membrane forms is still not clear yet.36,37 Furthermore, because drug permeation occurs through thousands of micron-sized pores in PAMPA,14,32,35,37 drug transport might be affected significantly by the edges of the small pores, too. Additionally, two different permeation assays for assessing passive transport, phospholipid vesicle-based permeation assays (PVPA)38 and Permeapad®,39 have been developed too. In these two assays, thin layers of phospholipid vesicles or lipid solution are deposited on the filter support as well, thus possibly containing similar problems that PAMPA has. To overcome these issues of artificial membranes, the development of a new artificial membrane, whose structure is defined better as a model cell membrane, is needed. For this reason, a large freestanding lipid bilayer could be an alternative solution. Previously, various methods for obtaining stable lipid bilayers40–48 have been developed, and the droplet interface bilayer (DIB) technique is one of the promising candidates that contains a freestanding lipid bilayer between two aqueous phases, once the geometry of the DIB is modified.42–44,47 In this paper, we report a new in vitro drug permeability assay that is easy to set up and simple to use. The freestanding lipid bilayer is formed easily by attaching a droplet lipid monolayer formed at the surface of a water droplet containing drug molecules inside, to a planar lipid monolayer at an oil-water interface within a typical UV spectrometer cell. Once


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the lipid bilayer is created, a significant concentration gradient across the lipid bilayer is applied, and then, the molecules move across the lipid bilayer by passive diffusion (Figure 1a). Exploiting the automated UV-Vis spectroscopy system, the concentration change of the drug molecules could be recorded over time, and the static permeability of the specific drug could be calculated from its concentration-time curve (Figure 1b). Using our technique, we investigate the permeability of five different hydrophilic drugs (caffeine, theophylline, antipyrine, acetaminophen and cimetidine), whose transport rates are an order of magnitude different. We then compare the permeability with previous results from various in vitro assays and summarize the advantages of our technique over other assays. Finally, we also show that the dynamic permeability (a change of the drug permeability over time) could be measured if we impair the membrane integrity in situ while the passive diffusion occurs.

Experimental Section Materials 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) was purchased from Avanti Polar Lipids. Squalane (99%), silicone oil (Silicone Oil AR 20), Triton X-100 (TX100) and all drug compounds including caffeine, theophylline, antipyrine, acetaminophen and cimetidine were purchased from Sigma-Aldrich and used without further purification. Ultrapure deionized water of pH 7.4, prepared with a Milli-Q system (Merck-Millipore, resistivity of 18.2 MΩ·cm at 25 °C) was used throughout all the experiments. Lipid Solution Preparation To prepare the lipid solution, 1,2-dioleoyl-sn-glycero-3-phosphatidylcholine (DOPC) was dissolved in chloroform initially at a concentration of 25 mg/mL. This stock solution was then transferred to a glass vial and evaporated under a nitrogen gas stream to remove the



chloroform completely resulting in a dried lipid film. This dried lipid film was dissolved again in a 1:1 (v/v) mixture of squalane and silicone oil at a concentration of 4 mg/mL, and the lipid solution was bath-sonicated for 15 minutes at 20 ºC right before use (Ultrasonic Cleaner Set, DAIHAN Scientific). Formation of the Lipid Bilayer at the Oil-Water Interface Prior to the real-time measurement of the drug transport using UV-Vis spectroscopy, a stable freestanding lipid bilayer was formed at the oil-water interface. To do so, 1 mL of deionized water was transferred first into a quartz UV cuvette (1.8 mL stirring cell, Starna Cells), and then, 250 µL of the DOPC solution dissolved in the 1:1 (v/v) mixture of squalane and silicone oil with a 4 mg/mL concentration was added carefully to the top of the water phase. After a few minutes, a self-assembled lipid monolayer (denoted as a planar monolayer) formed at the oil-water interface. Next, using a micropipette (10 µL, Eppendorf), 1 µL of an aqueous droplet containing the drug molecules inside at various concentrations (30 mM for caffeine and theophylline, 25 mM for acetaminophen, 15 mM for cimetidine, and 5 mM for antipyrine) was produced carefully at the end of the tip in the upper oil phase near the oilwater interface, also leading to the formation of a lipid monolayer (denoted as a droplet monolayer) at the surface of the aqueous droplet. When both the planar monolayer and the droplet monolayer were gently brought into contact, the adhesion of the two monolayers started to occur along with oil drainage thus resulting in the formation of a lipid bilayer. These adhesion processes were observed at similar time scale after bringing two monolayers into contact whether the drug molecules are presented or not. The gravity effect could be negligible here due to the small size of the droplet (1.2mm of diameter), and the stable bilayer could be formed with over 80% of success rate. Schematic illustration for an overall experimental procedure is shown in Figure S1. Observation of the DOPC Bilayer Formation by Fluorescence Microscopy


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As a fluorescent dye, Texas Red DHPE (Texas Red 1,2-dihexadecanoyl-snglycero-3-phosphoethanolamine, Thermo Fisher Scientific) was used, and the final concentration of the dye was set as 0.3 µg/mL in the DOPC solution. All processes of the bilayer formation were observed and recorded by an inverted-typed optical microscope (Olympus) equipped with an EMCCD camera (iXon3, Andor). Real-Time Measurement of Drug Transport Across the Lipid Bilayer After the lipid bilayer formation, drug molecules began to transfer from the aqueous droplet to the bottom water phase due to the concentration gradient across the lipid bilayer. During the transfer process of the drug molecules, UV light with a fixed wavelength was applied to the bottom water phase in the UV cuvette, and the UV absorbance of the bottom water phase was measured over time using a UV-Vis spectrometer (UV-2600, Shimadzu). To correlate the UV absorbance at a specific wavelength and the concentration of the drug in the bottom water phase, each UV spectrum (from 220 nm to 300 nm) was first obtained for the drug solutions with several concentrations that varied from 10 to 100 µM. Then, UV absorbance-concentration curves were extracted at each wavelength and linearly fitted. Finally, the wavelength where the R2 value for the fitting process is higher than 0.99 was selected for converting the UV absorbance, I, to the drug concentration, C, in the bottom water phase with the relation, C = αI, where α is a converting constant (see Supporting Information Figure S2 and Table S1). During the experiments, we assumed that the drug molecules are uniformly distributed over the whole region of the bottom water phase because the bottom water phase was continuously mixed using a magnetic cell stirrer (Spinette, Starna cells with stirring bar of 5 mm x 2 mm, at 1000 rpm). We confirmed that the stirring process did not affect the UV absorbance as well. The Introduction of Triton X-100 during Drug Transport To check if our technique is able to track a sudden change in the membrane



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permeability during drug transport, we inserted Triton X-100 (TX100), a nonionic surfactant, into the DOPC bilayer in the middle of the drug transport. To do so, 1-2 µL of an aqueous TX100 solution was injected carefully into the bottom water phase with a syringe (10 µL Gastight, Hamilton), resulting in the final TX100 concentrations in the bottom water phase from 0.1 CMC (critical micelle concentration ~ 0.20 mM) to 2.0 CMC, in the middle of the acetaminophen transport whose process was recorded by a UV spectrometer at a wavelength of 245 nm. Each UV absorbance of TX100 at 245 nm with various concentrations was measured independently, and these values were subtracted from the UV absorbance measured during the drug transport to achieve the UV absorbance for only the drug molecules. Permeability Calculations When molecules were transferred through the membrane by a concentration gradient, the permeability of the molecules was calculated by eq 123,32,36,49,50:

ܲ=

ܸ஺ 1 ݀‫ܥ‬஺ ሺ‫ݐ‬ሻ ‫ܥ[ ܣ‬஽ ሺ‫ݐ‬ሻ − ‫ܥ‬஺ ሺ‫ݐ‬ሻ] ݀‫ݐ‬

(1)

where P is the permeability; A is the membrane area through which molecules are transferred (bilayer area); VA is the volume of the acceptor (1 mL, bottom water phase); CA is the concentration of the acceptor, and CD is the concentration of the donor (upper water droplet). The bilayer area was calculated as A = πR2, where R is the radius of the bilayer estimated from the fluorescence microscopic images. Assuming that there is no loss of drugs into the membrane or the oil phase, eq 2 was established by mass conservation:

‫ܥ‬஽ ሺ0ሻܸ஽ = ‫ܥ‬஺ ሺ‫ݐ‬ሻܸ஺ + ‫ܥ‬஽ ሺ‫ݐ‬ሻܸ஽ .

(2)

1) Static permeability In contrast to the previous conventional model whose transport occurs from the very beginning, our measurements included the formation process of the lipid bilayer, thus leading


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to the transport across the lipid bilayer after some period of time. As shown in Figure 1b, each graph of CA(t) vs time for the various drug compounds contains the inflection point where the second time derivative of CA(t) is zero, and it turns out that t* is the time when transport across the lipid bilayer begins (A detailed explanation is in Results and Discussion). Therefore, to estimate the static permeability that remains constant even over time, we solved eq 1 analytically using two different conditions simultaneously, such as the mass conservation of eq 2 and the initial condition of CA(t*)=M, where M is the concentration at t=t* thus resulting in the successful establishment of eq 3.

‫ܥ‬஺ ሺ‫ݐ‬ሻ ‫ܯ‬ 1 1 = 1 − ቆ1 − ቇ ݁‫ ݌ݔ‬൤−ܲ‫ ܣ‬൬ + ൰ ሺ‫ ݐ‬− ‫ ∗ ݐ‬ሻ൨, ‫ܥ‬௘௤ ‫ܥ‬௘௤ ܸ஽ ܸ஺

(3)

where Ceq is the equilibrium concentration, defined as Ceq = CD(0)VD/(VD+VA). Then, the static permeability was calculated by a nonlinear fitting of eq 3 to the CA(t) curve whose process started at t=t*. For each drug, 3 independent sets of CA(t) data were measured and their averaged value at each time was used to estimate the static permeability. 2) Dynamic permeability In contrast to the estimation of the static permeability, the dynamic permeability could be calculated numerically. From the fact that Ceq = CD(0)VD/(VD+VA), eq 2 was modified into eq 4,

‫ܥ‬஽ ሺ‫ݐ‬ሻ − ‫ܥ‬஺ ሺ‫ݐ‬ሻ = ‫ܥ‬஽ ሺ0ሻ ቆ1 −

‫ܥ‬஺ ሺ‫ݐ‬ሻ ቇ. ‫ܥ‬௘௤

4)

Then, eq 4 was applied to eq 1 to get rid of CD(t) term, thus resulting in eq 5,

ܲ ሺ‫ ݐ‬ሻ =

‫ܥ‬௘௤ ܸ஺ ݀‫ܥ‬஺ ሺ‫ݐ‬ሻ . ‫ܥܣ‬஽ ሺ0ሻ ሺ‫ܥ‬௘௤ − ‫ܥ‬஺ ሺ‫ݐ‬ሻሻ ݀‫ݐ‬

5)

The time derivative of the acceptor concentration dCA(t)/dt was obtained by calculating the slope between two adjacent points, (t, CA(t)) and (t+∆t, CA(t+∆t)), where ∆t is the time interval (for cimetidine, ∆t =300 s; for the rest of the drugs, ∆t =50 s), and then, the dynamic



permeability at each time was calculated numerically from eq 5. Error Analysis for the Dynamic Permeability When measuring the dynamic permeability, their values started to fluctuate significantly as the concentrations of each drug, CA(t), became closer to the equilibrium value, Ceq, shown in Figure S3. It is because the concentration change is so small that the sensitivity of the instrument is insufficient to provide reliable permeability values. Thus, we simply took out the experimental data when ∆(Absorbance) for ∆t was smaller than 10 times of the instrumental accuracy. We provide additional data for this analysis in Figure S3.

Results and Discussion Visualization of the Lipid Bilayer Formation Exploiting the geometry of the planar freestanding lipid bilayer that was recently developed,47 we succeeded in forming a stable freestanding lipid bilayer with a large area (> 1 mm2) within the UV cell. Using a DOPC solution that is dissolved in a 1:1 (v/v) mixture of squalane and silicone oil at a concentration of 4 mg/ml, a DOPC bilayer is achieved by the adhesion of the droplet DOPC monolayer and the planar DOPC monolayer. We specifically use a 1:1 (v/v) mixture of squalane and silicone oil as an oil phase to obtain oil-free lipid bilayer. Squalene has been frequently used for lipid bilayer formation because it is known that squalene cannot be located within the lipid bilayer due to its bulky molecular structure44 thus resulting in a solvent-free lipid bilayer. However, its highly unsaturated hydrocarbon structure is vulnerable to oxidation, and thus, a fully saturated version of squalene is used instead. Furthermore, silicone oil, a bad solvent for phospholipid molecules,45 is added to squalane, to facilitate the adhesion of the two lipid monolayers.51,52 To further support the formation of the lipid bilayer, we visualized the adhesion


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process of the monolayers using fluorescence microscopy (Video 1, Supporting Information). At 120-180 seconds after placing the lipid coated droplet on the planar lipid monolayer, oil drainage was clearly observed between the two lipid monolayers implying that the adhesion of the two lipid monolayers occurs successfully as they become closer shown in Figure 2. This adhesion process was completed within another 120-180 seconds, and thus, it takes 240360 s in total to form the freestanding lipid bilayer from the time when the droplet is introduced to the planar monolayer. Our lipid bilayer was stable for at least a day, which is sufficient for the permeability measurement of various drugs. Permeability Assay We selected five different hydrophilic drug molecules, caffeine, theophylline, antipyrine, acetaminophen, and cimetidine whose physicochemical properties as well as permeability are well-known shown in Table 1. All of these drugs are water-soluble and known to be absorbed by passive transport at the intestine.14,35 The concentration change of the drugs over time in the acceptor region (the bottom water phase) can be measured by the UV absorbance over time, and these are then normalized by the equilibrium concentration, Ceq = CD(0)VD/(VD+VA), when the transport is finished so that the concentrations of the two regions are matched (See experimental section). During the measurement, the incident UV light can only cover the part that is several millimeters below where the lipid bilayer is located. Therefore, it takes some time for the drug molecules to diffuse into the part where the UV light can detect them after they pass through the lipid bilayer. Given that the diffusivity of a drug in water is generally on the order of 10-10 m2/s, it would take 103 - 104 seconds for the drug molecules to diffuse a distance of a few millimeters, and the membrane permeability cannot be measured because of the diffusion-dominant process. To solve this problem, the bottom water phase is continuously stirred by a magnetic cell stirrer to fully mix the bottom water phase so that the concentration



in the water phase is spatially uniform during the measurement. Because this mixing is completed within a few seconds, compared with the shortest time interval for the UV measurement (50 seconds), we were able to detect all the transported molecules without considering the effect of diffusion for the drug molecules. As shown in Figure 3, the concentration of the five different drugs in the acceptor region increases over time and eventually reaches their equilibrium concentrations. This strongly suggests that almost all of the drug molecules are transferred to the bottom water phase, rather than partitioned into the oil phase or trapped in the bilayer. In addition, at the early stage of transport, the CA(t)/Ceq curves for each drug increases sharply with a positive curvature, which is not expected from the simple diffusion model. After this region, the slope changes to a negative curvature with an inflection point. This appears at ~ 250, 350, 300, 450 and 600 s for caffeine, theophylline, antipyrine, acetaminophen and cimetidine, respectively. After this inflection point, the transport of the drugs exhibits a simple exponential behavior which can be explained by the diffusion model. These time scales where the inflection point occurs agree quite well with the time required to finish the bilayer formation based on our visualization results. The initial increase in concentration before the inflection point is likely to be a result of the transport through the two lipid monolayers and an additional thin oil film when the top droplet approaches the planar monolayer. As the oil drainage and bilayer formation finish by the inflection point, it appears to be the starting point for the transport of a drug across the oil-free lipid bilayer. We set the inflection point as a starting point of the fitting by the diffusion model, eq 3, and confirmed that each CA(t)/Ceq curve is fitted quite well with an R2 > 0.97 (dotted lines in Figure 3). In contrast, if this is fitted for the entire time range, then R2 varies from 0.92 to 0.96. This fitting provides a “static” permeability, which is a specific bilayer membrane property for each drug. The permeability obtained by our technique is summarized in the last column of Table 1.


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As easily seen in Figure 3, the transport time through the lipid bilayer is significantly different for each drug. Quantitatively, the characteristic time scale, τ = [PA(1/VD + 1/VA)]-1, varies two orders of magnitude from 340 s to 11,600 s showing that our technique could measure orders of magnitude different permeability values. In fact, the maximum permeability that could be reasonably measured by our technique can be estimated as follows: the change in concentration for a time interval of the UV absorbance measurement, ∆t, is ஼ಲ ሺ∆௧ሻ/஼೐೜ ∆௧

. Because CA(t)/Ceq becomes one for the fastest transport during this time period, the

shortest characteristic time for the transport is simply ∆t. The maximum permeability is 8500 × 10-6 cm/s with ∆t = 10 s, which is more than 30 times larger than the largest permeability (caffeine) from our measurements. When a concentration gradient of a solution set up between two compartments is blocked by a membrane, the system can also be equilibrated by the transport of solvent which also depends on the transport rate. Because a lipid bilayer membrane is permeable to both drug and water molecules, the water molecules can move from the bottom water phase to the top droplet to decrease the concentration of drugs in the droplet. To determine whether this water transport affects our measurements, we take the water flux in a reverse direction into account in eq 2 and eq 4 by considering the volume change of the donor region (water droplet) at each time interval, and the volume change is given as

∆ܸ஽ ܸ஽ ሺ0ሻ ‫ܥ‬஺ ሺ‫ݐ‬ሻ = ܲ௪ ‫ݒܣ‬௠ ‫ܥ‬஽ ሺ0ሻ ቆ1 − ቇ, ∆‫ݐ‬ ܸ஽ ሺ‫ݐ‬ሻ ‫ܥ‬௘௤

(6)

where Pw is the water permeability of the DOPC bilayer, 0.0056 cm/s;53 A is the bilayer area, 0.0121 cm2, and vm is the molar volume of water, 18 × 10-6 mM-1. For cimetidine with the slowest permeability, which could have the strongest effect, the maximum volume change in the water droplet during a single time interval could be simply estimated by ∆VD ~ PwAvmCD(0)∆t ~ 0.0055 µL, using CD(0) = 15 mM and ∆t = 300 s at t = 0. This volume



change is only 0.55%, and thus, this strongly suggests that the concentration change by the upward water flux during our time interval is negligible. A more detailed investigation for elucidating the effect of the water flow on the permeability is presented in Figure S4. Comparison of the Permeability with Other Assays As presented in Table 1 and Figure 4, the permeability of the five drugs measured by our technique are quite larger than those by the other in vitro assays, even though the rank of the permeability remains the same. For example, our technique has a 1.5-7 times larger permeability than Caco-2, 25-60 times larger values than PAMPA, 8-20 times lager values than PVPA and 12-33 times larger values than Permeapad®. Even the permeability of cimetidine is reported as 0 for PAMPA whereas our technique could measure it. Such a large difference is likely to originate mainly from the thickness of the membrane. Specifically, PAMPA uses the artificial membrane whose microstructure has not been known precisely. The PAMPA membrane is conventionally prepared by dipping a porous filter into lipid solutions whose solvents are linear hydrocarbons such as decane, dodecane, and hexadecane.14,31,32,35,36 It is known that the use of these kinds of solvents leaves a residual oil layer between the lipid layers.36,37 This oil layer results in a relatively thicker barrier for drugs compared to the lipid bilayer thus leading to the slow transport. The membranes of PVPA and Permeapad® could also have the thicker barriers due to the similar procedure for membrane preparation. In addition, for the Caco-2 assay, drug molecules are required to pass through the interior of the cell as well as the outer bilayer membranes thus resulting in a longer permeation path, too. Therefore, these thicker membranes of the other in vitro assays result in a lower permeability compared to our assay in which the drugs transport only through the oil-free phospholipid bilayer because the permeability is inversely proportional to the membrane thickness.36 Furthermore, the larger permeability values from our platform could also be explained by the simple component of our membrane; our lipid bilayer consists


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of only DOPC, which has a liquid disordered phase at our experimental temperature. In the Caco-2 assay, the outer layer of the cells consists of a lipid bilayer embedded with various proteins, polysaccharides, glycolipids, etc., and these possibly hinder the passive transport of drug molecules as well. Besides the fact that our assay provides a more accurate permeability of small molecules across a lipid bilayer, it could also provide several advantages over other in vitro assays. For example, to obtain the drug permeability properly using the previous assays, it is necessary to make additional efforts to find the appropriate time for sampling the transported drugs because the time scale for how fast drugs are transported is not known prior to the experiments.36,50 If the sampling is done when the concentration of the transported drug is quite close to the equilibrium concentration, this might give imprecise permeability.50 In contrast, our platform keeps track of the concentration change every few seconds; thus, highly permeable drugs could be measured without any additional efforts. Dynamic Permeability In addition to the estimation of the static permeability of a drug, the dynamic permeability (time-dependent permeability) could also be obtained from the experimental data using eq 5 (See experimental section). Figure 5 shows the dynamic permeability within the reliable range, and the details of this error analysis to determine the reliable range are presented in the experimental section. The dynamic permeability of all our tested drugs indicates two distinct regimes: the early stage where the permeability gradually increases from zero and the equilibrium stage where permeability remains constant over time. For caffeine, the permeability slightly decreases after reaching the equilibrium value, and we do not understand this behavior. In the early stage before the lipid bilayer is formed, the two lipid monolayers become closer as the top droplet monolayer approaches the bottom planar monolayer. Drug molecules



could be transferred through the two lipid monolayers and the additional oil film, as we mentioned before, and the permeability could increase as the oil in between the monolayers drains out. Meanwhile, the lipid bilayer formation is completed before the equilibrium stage, and the membrane thickness would be maintained during this regime thus resulting in constant permeability over time. This transition from the early stage to the equilibrium stage agrees well with the time required for the bilayer formation as well as with the time when the inflection point occurs in the graph of CA(t)/Ceq (Figure 3). To further demonstrate the dynamic permeability, we used Triton X-100 (TX100), a nonionic detergent often used for cell lysis,54 for an in situ modification of the permeability of the lipid bilayer. We added TX100 to the bottom water phase in the middle of the transport measurement of acetaminophen (600 s after starting measurement) shown in Figure 6, possibly inducing a more permeable lipid bilayer by the incorporation of the TX100 molecules into the bilayer. Before the addition of TX100 to the bottom water phase, the dynamic permeability in Figure 6a is identical as we expected. However, the addition of TX100 results in ~ 1.5 times increase in the permeability (from ~100 to ~150 × 10-6 cm/s) at 600 seconds after injecting the TX100 at 0.9 CMC of concentration. Furthermore, the change of static permeability after injecting TX100 with concentration from 0.1 CMC to 2 CMC is also indicated in Figure 6b, and 1.35 times and 1.25 times larger permeability is observed with the addition of 0.9 CMC and 0.5 CMC of TX100, respectively, whereas little change is observed with 0.1 CMC of TX100. However, over 1.0 CMC of TX100, the bilayer rupture always occurs possibly due to the disruption of the membranes, and these results are well agreed with the previous research.54 Thus, this demonstrates that our assay provides a solid platform to measure any dynamic permeability measurements and has a great advantage for detecting unexpected realtime behaviors during the transport across the lipid bilayer such as a coalescence of the top


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droplet to the bottom water phase (Figure S5). Further systematic investigation on this dynamic permeability upon the addition of TX100 is in progress.

Conclusions In conclusion, we developed a new technique to measure the permeability of various drug compounds across a freestanding lipid bilayer. By placing a DOPC coated water droplet that contains drug molecules inside onto the planar DOPC monolayer, a freestanding DOPC bilayer with a large interfacial area was formed successfully in the UV-Vis spectrometer cell. In addition, based on this platform, we succeeded in estimating the drug permeability for five representative drug compounds by obtaining reproducible concentration-time curves even though their transport time scales are quite different from several hundreds to tens of thousands of seconds. We confirmed that the rank order of the permeability from our assay agrees well with those from the other in vitro assays, even though transport through our thin and oil-free lipid bilayer is 1.5-60 times faster than that of the other assays. Furthermore, we also showed that the dynamic permeability over time could be measured. By adding TX100 to the bottom water phase in the middle of a transport, the increase in the drug permeability presumably due to the disruption of the lipid bilayer was successfully observed. This thus strongly suggests that our technique could be used to investigate important physiological phenomena such as cell apoptosis, a well-programmed cell death entailing a sudden increase in the mitochondrial outer membrane permeability.55,56 Although our technique might not be suitable for rapidly screening hundreds of thousands of drug candidates at the present stage, we believe that further improvements would make our platform a more appropriate tool for high-throughput screening eventually.

Supporting Information.



Schematic illustrations for an overall experimental procedure, converting the UV absorbance to the concentration, cut-off process for achieving reliable dynamic permeability, dynamic permeability with consideration of the water flux in a reverse direction, the result with the coalescence of the top droplet and one video clip for visualization of the lipid bilayer formation.

References (1)

Stein, W. The movement of molecules across cell membranes; Academic Press: New York, 1967.

(2)

Saier, M. H. Microbiol. Mol. Biol. Rev. 2000, 64, 354–411.

(3)

Conner, S. D.; Schmid, S. L. Nature 2003, 422, 37–44.

(4)

Palm, W.; Thompson, C. B. Nature 2017, 546, 234–242.

(5)

Artursson, P.; Palm, K.; Luthman, K. Adv. Drug Deliv. Rev. 2001, 46, 27–43.

(6)

Bareford, L. M.; Swaan, P. W. Adv. Drug Deliv. Rev. 2007, 59, 748–758.

(7)

Ratajczak, J.; Wysoczynski, M.; Hayek, F.; Janowska-Wieczorek, A.; Ratajczak, M. Z. Leukemia 2006, 20, 1487–1495.

(8)

Engelberg-Kulka, H.; Amitai, S.; Kolodkin-Gal, I.; Hazan, R. PLoS Genet. 2006, 2, 1518–1526.

(9)

Blaustein, M. P.; Lederer, W. J. Physiol. Rev. 1999, 79, 763–854.

(10)

Gadsby, D. C. Nat. Rev. Mol. Cell Biol. 2009, 10, 344–352.

(11)

Zhang, Y.; Benet, L. Z. Clin. Pharmacokinet. 2001, 40, 159–168.

(12)

Pang, K. S. Drug Metab. Dispos. 2003, 31, 1507–1519.

(13)

Bohets, H.; Annaert, P.; Mannens, G.; Van Beijsterveldt, L.; Anciaux, K.; Verboven, P.; Meuldermans, W.; Lavrijsen, K. Curr. Top. Med. Chem. 2001, 1, 367–383.


Page 18 of 28

Page 19 of 28 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(14)

Zhu, C.; Jiang, L.; Chen, T.-M.; Hwang, K.-K. Eur. J. Med. Chem. 2002, 37, 399–407.

(15)

Yeon, J. H.; Park, J. K. Anal. Chem. 2009, 81, 1944–1951.

(16)

Li, S.; Hu, P.; Malmstadt, N. Anal. Chem. 2010, 82, 7766–7771.

(17)

Kuhn, P.; Eyer, K.; Allner, S.; Lombardi, D.; Dittrich, P. S. Anal. Chem. 2011, 83, 8877–8885.

(18)

Hilgers, A. R.; Conradi, R. A.; Burton, P. S. Pharm. Res. 1990, 7, 902–910.

(19)

Karlsson, J.; Artursson, P. Int. J. Pharm. 1991, 71, 55–64.

(20)

Yazdanian, M.; Glynn, S. L.; Wright, J. L.; Hawi, A. Pharm. Res. 1998, 15, 1490– 1494.

(21)

Yamashita, S.; Furubayashi, T.; Kataoka, M.; Sakane, T.; Sezaki, H.; Tokuda, H. Eur. J. Pharm. Sci. 2000, 10, 195–204.

(22)

Marino, A. M.; Yarde, M.; Patel, H.; Chong, S.; Balimane, P. V. Int. J. Pharm. 2005, 297, 235–241.

(23)

Hubatsch, I.; Ragnarsson, E. G. E.; Artursson, P. Nat. Protoc. 2007, 2, 2111–2119.

(24)

Sun, H.; Pang, K. S. Drug Metab. Dispos. 2010, 38, 536–537.

(25)

Bergström, C. A. S.; Strafford, M.; Lazorova, L.; Avdeef, A.; Luthman, K.; Artursson, P. J. Med. Chem. 2003, 46, 558–570.

(26)

Ungell, A. L. B. Drug Discov. Today Technol. 2004, 1, 423–430.

(27)

Balimane, P.; Chong, S. Drug Discov. Today 2005, 10, 335–343.

(28)

Balimane, P. V; Han, Y.-H.; Chong, S. AAPS J. 2006, 8, E1–E13.

(29)

Stenberg, P.; Luthman, K.; Artursson, P. J Control. Release 2000, 65, 231–243.

(30)

Sugano, K.; Kansy, M.; Artursson, P.; Avdeef, A.; Bendels, S.; Di, L.; Ecker, G. F.; Faller, B.; Fischer, H.; Gerebtzoff, G.; Lennernaes, H.; Senner, F. Nat. Rev. Drug Discov. 2010, 9, 597–614.

(31)

Kansy, M.; Senner, F.; Gubernator, K. J. Med. Chem. 1998, 41, 1007–1010.



(32)

Avdeef, A.; Strafford, M.; Block, E.; Balogh, M. P.; Chambliss, W.; Khan, I. Eur. J. Pharm. Sci. 2001, 14, 271–280.

(33)

Wohnsland, F.; Faller, B. J. Med. Chem. 2001, 44, 923–930.

(34)

Sugano, K.; Takata, N.; Machida, M.; Saitoh, K.; Terada, K. Int. J. Pharm. 2002, 241, 241–251.

(35)

Chen, X.; Murawski, A.; Patel, K.; Crespi, C. L.; Balimane, P. V. Pharm. Res. 2008, 25, 1511–1520.

(36)

Avdeef, A. Absorption and Drug Development: Solubility, Permeability, and Charge State, 2nd ed.; Wiley: New York, 2012.

(37)

Sugano, K. In Comprehensive Medicinal Chemistry Ⅱ: Volume 5: ADME-Tox Approaches; Testa, B., Van de Waterbeemd, H., Eds.; Elsevier, 2007; pp 453–487.

(38)

Flaten, G. E.; Dhanikula, A. B.; Luthman, K.; Brandl, M. Eur. J. Pharm. Sci. 2006, 27, 80–90.

(39)

Di Cagno, M.; Bibi, H. A.; Bauer-Brandl, A. Eur. J. Pharm. Sci. 2015, 73, 29–34.

(40)

Funakoshi, K.; Suzuki, H.; Takeuchi, S. Anal. Chem. 2006, 78, 8169–8174.

(41) Stachowiak, J. C.; Richmond, D. L.; Li, T. H.; Liu, A. P.; Parekh, S. H.; Fletcher, D. A. Proc. Natl. Acad. Sci. 2008, 105, 4697–4702. (42)

Hwang, W. L.; Chen, M.; Cronin, B.; Holden, M. A.; Bayley, H. J. Am. Chem. Soc. 2008, 130, 5878–5879.

(43)

Bayley, H.; Cronin, B.; Heron, A. J.; Holden, M. A.; Hwang, W. L.; Syeda, R.; Thompson, J. R.; Wallace, M. I. Mol. Biosyst. 2008, 4, 1191–1208.

(44)

Leptihn, S.; Castell, O. K.; Cronin, B.; Lee, E.-H.; Gross, L. C. M.; Marshall, D. P.; Thompson, J. R.; Holden, M.; Wallace, M. I. Nat. Protoc. 2013, 8, 1048–1057.

(45)

Thiam, A. R.; Bremond, N.; Bibette, J. Langmuir 2012, 28, 6291–6298.

(46)

Beltramo, P. J.; Van Hooghten, R.; Vermant, J. Soft Matter 2016, 12, 4324–4331.


Page 20 of 28

Page 21 of 28 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(47)

Jeong, D.-W.; Jang, H.; Choi, S. Q.; Choi, M. C. Sci. Rep. 2016, 6, 38158.

(48)

Osaki, T.; Takeuchi, S. Anal. Chem. 2017, 89, 216–231.

(49)

Palm, K.; Luthman, K.; Ros, J.; Grasjo, J.; Artursson, P. J. Pharmacol. Exp. Ther. 1999, 291, 435–443.

(50)

Tavelin, S.; Gråsjö, J.; Taipalensuu, J.; Ocklind, G.; Artursson, P. In Epithelial Cell Culture Protocols; Wise, C., Ed.; Humana Press: New Jersey, 2002; pp 233–272.

(51) Wauer, T.; Gerlach, H.; Mantri, S.; Hill, J.; Bayley, H.; Sapra, K. T. ACS Nano 2014, 8, 771–779. (52)

Villar, G.; Heron, A. J.; Bayley, H. Nat. Nanotechnol. 2011, 6, 803–808.

(53)

Rawicz, W.; Smith, B. A.; McIntosh, T. J.; Simon, S. A.; Evans, E. Biophys. J. 2008, 94, 4725–4736.

(54)

Koley, D.; Bard, A. J. Proc. Natl. Acad. Sci. 2010, 107, 16783–16787.

(55)

Green, D. R.; Kroemer, G. Science. 2004, 305, 626–629.

(56)

Tsujimoto, Y.; Shimizu, S. Apoptosis 2007, 12, 835–840.

Acknowledgments This work was supported by the Basic Science Research Program through the National Research

Foundation

of

Korea

(NRF-2012R1A6A3A04040395,

NRF-

2015R1C1A1A01054180, NRF-2016R1A6A3A11931836) and by the Agency for Defense Development of Korea (ADD 14-70-06-10).

For TOC only



Figure 1. A platform to analyze the real-time transport of drug molecules across the lipid bilayer. (a) Schematic illustration of the permeability assay for drug molecules constructed in a UV cuvette. Lipid molecules that are dissolved initially in an oil phase form self-assembled monolayers at the oil-water interface and the droplet surface first. Then, when two different monolayers are brought into contact each other, the lipid bilayer is formed, and the drug molecules start to transfer through it due to a concentration gradient. During the transport process, UV light with a fixed wavelength is applied to the bottom water phase, and the UV absorbance is measured over time using a UV-Vis spectrometer. (b) A representative


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time series of UV absorbance during drug transport, which contains an inflection point (blue circle) at the early stage that corresponds to the starting point of the drug transport across the lipid bilayer.

Figure 2. Visualization of the lipid bilayer formation by fluorescence microscopy. (a) Schematic illustration of the visualization process using an inverted-typed fluorescence microscope. The adhesion between two lipid monolayers that are formed at the droplet surface and the planar oil-water interface, respectively, occurs along with oil drainage resulting in the lipid bilayer formation. (b) Fluorescence microscopic images over time. The elapsed time since the oil drainage started is indicated in each image. The boundary between the two regions where oil remains and is absent is indicated by white dotted lines arrows. The entire adhesion process is completed within 180 seconds.



Figure 3. The graph of CA(t)/Ceq. Each concentration change of the 5 different drug compounds (caffeine, theophylline, antipyrine, acetaminophen, and cimetidine) over time at the acceptor region (bottom water phase) is normalized by their equilibrium concentrations, Ceq, respectively. Each dataset indicates averaged experimental values by repeating three independent trials, and each dotted line corresponds to the fitted results from eq 3, and this fitting process starts at t=t*, not t=0. t* indicates the inflection points of each graph, whose values vary depending on the drug compound, but they stay within the shaded region (250 s – 600 s). The inset represents a full curve of cimetidine with a time over 30,000 s. Error bars represent standard deviations.


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Table 1. Physicochemical properties of the drugs and their permeability values from the Caco-2 assay, PAMPA, Permeapad®, PVPA and this work.



Figure 4. Comparison of the permeability for various drug compounds depending on the different permeability assays. Error bars in this work correspond to the standard deviations of the permeability values by fitting eq 3 to the CA(t)/Ceq curves, shown in Figure 3.


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Figure 5. Dynamic permeability for the five different drug compounds. Dynamic permeability of the drug is calculated by calculating eq 5 numerically from the CA(t)/Ceq curve, shown in Figure 3. Red dots indicate each dynamic permeability for the various drugs, caffeine, theophylline, antipyrine, acetaminophen, and cimetidine. Solid blue line corresponds to the static permeability of each drug, indicated in Figure 4.



Figure 6. Permeability change of acetaminophen with the addition of Triton X-100. (a) Dynamic permeability of acetaminophen over time in the presence/absence of TX100 during the drug transport. With the addition of TX100 (0.9 CMC) ~ 1.5 times larger dynamic permeability of acetaminophen, compared to the reference (solid blue triangles), is observed at ~ 600 seconds after introducing TX100 (solid red dots). The arrow indicates the time when Triton X-100 is introduced. (b) Static permeability of acetaminophen with varying concentrations of TX100 from 0.1 to 0.9 CMC. Error bars represent standard deviations.


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Static and Dynamic Permeability Assay for Hydrophilic Small

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