Programmed Vesicle Fusion Triggers Gene Expression - American

Sep 16, 2011 - Center for Fundamental Living Technology (FLinT), Institute of Physics and ... Graduate School of Frontier Biosciences, Osaka Universit...
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Programmed Vesicle Fusion Triggers Gene Expression Filippo Caschera,† Takeshi Sunami,‡ Tomoaki Matsuura,‡,§ Hiroaki Suzuki,‡,|| Martin M. Hanczyc,† and Tetsuya Yomo*,‡,||,^ †

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Center for Fundamental Living Technology (FLinT), Institute of Physics and Chemistry, University of Southern Denmark, Campusvej 55, DK-5230 Odense M, Denmark ‡ Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Agency, Yamadaoka 1-5, Suita, Osaka 565-0871 § Department of Biotechnology, Graduate School of Engineering, Osaka University, Yamadaoka 2-1, Suita, Osaka 565-0871, Japan Department of Bioinformatic Engineering, Graduate School of Information Science and Technology, and Department of Frontier Biosciences, ^Graduate School of Frontier Biosciences, Osaka University, Yamadaoka 1-5, Suita, Osaka 565-0871, Japan

bS Supporting Information ABSTRACT: The membrane properties of phospholipid vesicles can be manipulated to both regulate and initiate encapsulated biochemical reactions and networks. We present evidence for the inhibition and activation of reactions encapsulated in vesicles by the exogenous addition of charged amphiphiles. While the incorporation of cationic amphiphile exerts an inhibitory effect, complementation of additional anionic amphiphiles revitalize the reaction. We demonstrated both the simple hydrolysis reaction of β-glucuronidase and the in vitro gene expression of this enzyme from a DNA template. Furthermore, we show that two vesicle populations decorated separately with positive and negative amphiphiles can fuse selectively to supply feeding components to initiate encapsulated reactions. This mechanism could be one of the rudimentary but effective means to regulate and maintain metabolism in dynamic artificial cell models.

’ INTRODUCTION Increasing technical competency has allowed scientists to scrutinize the living cell down to the angstrom and nanosecond, with terms such as artificial, synthetic, and in vitro becoming more applicable in understanding and recapitulating biology. For example, scientists have synthesized over a half billion base pairs of the Mycoplasma genome from scratch and introduced the synthetic genetic material into a living cell,1 thereby producing a semisynthetic cell. Several researchers worldwide are currently developing methods to reconstitute or synthesize a living cell in the laboratory.2 15 To produce an artificial cell, a certain level of complexity will be needed to support all of the necessary characteristics of life, achievable by the stepwise integration of functional subsystems.9,11,12 An artificial cell based on lipid vesicles encapsulating basic biochemical metabolism, e.g., an in vitro transcription/translation (gene expression) system of specified components or the PURE system,16 is one current approach.2 8,15,17 It is believed that such a system can support the self-replication of basic genomes.17 The next step is to sustain the encapsulated metabolism by supplying fresh components over time. As nutrients are essential for an artificial cell, Noireaux and Libchaber demonstrated that a pore protein creates a selective passage for nutrients into a vesicle and that the encapsulated cell-free expression system is active for more than 4 days.2 r 2011 American Chemical Society

In this paper, we use another mechanism, the controlled addition of components using vesicle fusion, in which the internal aqueous contents in vesicles are mixed. This method is appropriate for the introduction of large biomolecules that cannot pass through the membrane. The fusion of membranes is used by nature for the transfer of material.18 20 We intend to use vesicle fusion21 24 as a primitive way to supply our artificial cell with components with low permeability. We previously used the concept of electrostatic charge charge interaction23 by vesicle membrane decoration to induce the selective fusion of vesicles.25,26 The detection of vesicle association and fusion (internal contents mixing) was demonstrated using Co2+, EDTA, and calcein, which are small chemical compounds. However, it is not clear whether the same vesicle fusion strategy can be applied to mixing metabolic components and complexes (proteins and RNAs). For example, these molecules may not be active inside the vesicles, or the reaction machineries may not work in concert due to unfavorable molecular interactions with the lipids inside of the vesicles. Here, we demonstrate certain key functionalities needed to regulate the encapsulated metabolism by vesicle fusion. Received: March 10, 2011 Revised: September 5, 2011 Published: September 16, 2011 13082

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Langmuir First, we show that the exogenous addition of charged amphiphiles can exert suppression and the subsequent restoration of enzymatic activity and gene expression. Second, a mechanism to introduce metabolic components is demonstrated using the selective fusion of oppositely charged vesicles to initiate gene expression from a DNA template using encapsulated T7 RNA polymerase and the PURE system.

’ EXPERIMENTAL SECTION Materials. 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) was purchased from Avanti Polar Lipids (Alabaster, AL). Oleic acid, didodecyldimethylammonium bromide (DDAB), β-glucuronidase enzyme from Escherichia coli (GUS), protease from Streptomyces griseus, and Hepes buffer were purchased from Sigma-Aldrich Corporation (St. Louis, MO). R-phycoerythrin (R-PE) and transferrin from human serum, Alexa Fluor 647 conjugate (TA647), APC (Allophycocyanin), and the substrate 5-(pentafluorobenzoylamino) fluorescein di-β-D-glucuronide (PFB-FDGlcU) were purchased from Invitrogen (Carlsbad, CA). TokyoGreen-βGlcU was purchased from Sekisui Medical Co., Ltd., Japan. The PURE system is an optimized laboratory-made system that features increased translation efficiency; the components of the laboratory-made system and the DNA construct used are detailed in our previous article.27 The DNA used in the experiments was a PCR product amplified from the plasmid pET-uidA using two primers (Primer 1: CCCGCGAAATTAATACGACTCACTATAGGG, Primer 2: CTCCTTTCAGCAAAAAACCCCTCAAGACCC). The β-glucuronidase gene was placed after the T7 promoter. The T7 RNA polymerase was purchased from Takara (Japan). The dialysis experiment was performed with a Biodialyser system Z367923, Sigma-Aldrich. The membranes for dialysis (polycarbonate filter with a pore size of 0.4 μm) were purchased from Nuclepore Track-Etch Membranes; Whatman, Maidstone, Kent, UK. Preparation of Freeze Dried Empty Liposome Membranes. Giant vesicles were prepared by the freeze dried empty liposome (FDEL) method using previously described procedures.25,27 Briefly, POPC was dissolved in dichloromethane/diethyl ether (1:1, v/v), dried using a rotary evaporator, and then hydrated with Milli-Q water (12 mM lipid suspension). After vortexing and sonication, the vesicles were extruded through a polycarbonate filter with a pore size of 0.4 μm (Nuclepore Track-Etch Membranes; Whatman, Maidstone, Kent, UK). The suspension was dispensed into small aliquots (40 μL each) and freeze dried overnight (Labconco Corp., Kansas City, MO). After filling with argon gas, the freeze dried membranes were stored in a freezer at 20 C. POPC giant vesicles were obtained from the lyophilized membranes by adding 10 μL of the buffer (50 mM Hepes-KOH, pH 7.6) or the reaction mixture and further diluting 20-fold with the appropriate buffer. This protocol produces a vesicle suspension with a lipid concentration of 2.4 mM, with the most frequent vesicle volume of ∼1 fL distributed over 1 order of magnitude.25 Decoration of Pre-Existing POPC Vesicles with Positive or Negative Amphiphiles. The stock solutions of positive (DDAB) and negative (oleic acid) amphiphiles were prepared in ethanol and were injected into the POPC vesicle populations to modify (decorate) the electric charge of the vesicle membranes as previously described.25,26 We defined the molar percentage of the added amphiphiles to the total amount of POPC and amphiphiles as MP = Mamp/(Mamp + MPOPC), where Mamp and MPOPC are the moles of added amphiphiles and pre-existing POPC in the final vesicle suspension, respectively. After each injection, the final amount of ethanol in solution was always 1% v/v. In our previous study, we confirmed that the vesicle size was maintained after the decoration step.25 The zeta potential of the vesicles before and after decoration was measured using a zeta potential meter (Zetasizer Nano, Malvern Instruments).

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Enzyme and Gene Expression Activity in the Presence of Charged Amphiphiles and Charged Vesicles. Charged vesicles at MP = 25% were obtained when 0.6 μL of 80 mM DDAB or 80 mM oleic acid dissolved in ethanol was added to 59.4 μL of POPC vesicle suspensions. The vesicle suspensions were mixed at a volume ratio of 1:2 v/v (30 μL total), which were separately decorated by DDAB and oleic acid at MP = 25%, respectively. Four vesicle populations with different lipid compositions were prepared: POPC, POPC/DDAB, POPC/oleic acid, and POPC/ DDAB/oleic acid. To 30 μL of each resulting vesicle solution, 0.5 μL of 730 nM β-glucuronidase and 0.5 μL of 250 μM PFB-FDGlcU were dispensed while the tubes were kept on ice. The time courses of the substrate hydrolysis at 37 C were measured using a real-time PCR machine (Mx3005P QPCR system; Agilent, La Jolla, San Diego, CA) with optical filters for excitation and emission wavelengths of 492 and 516 nm, respectively. Similarly, gene expression was examined in the presence of vesicles. Vesicle suspensions decorated, respectively, by DDAB or oleic acid at MP = 20% were prepared. The mixed membrane composition was prepared by mixing the DDAB and oleic acid vesicle populations at a 1:2 v/v ratio. Afterward, the PURE system reaction mixture27,28 containing 50 μM PFB-FDGlcU and 20 nM DNA encoding β-glucuronidase was aliquoted into four different test tubes and kept on ice. Finally, 1 μL of the vesicle suspensions composed of POPC, POPC/ DDAB, POPC/oleic, or POPC/DDAB/oleic acid was added to each aliquot. The signal of the fluorogenic product was measured with the real-time PCR machine during incubation at 37 C using the same optical filter. The enzymatic activity in the presence of a cationic and a anionic mixture of amphiphiles was assayed further as follows. To the 20 μL aliquot of the 1 nM β-glucuronidase and 50 μM TokyoGreen-βGlcU suspension in 50 mM Hepes KOH buffer (pH 7.6), ethanol containing DDAB was injected at various final concentrations. To examine the restoration of enzymatic activity, a solution containing 1 nM β-glucuronidase and 50 μM TokyoGreen-βGlcU, supplemented with 600 μM DDAB in 50 mM Hepes-KOH buffer (pH 7.6), was prepared. To this solution, ethanol containing oleic acid was injected at various final concentrations. In both cases, the time courses of substrate hydrolysis at 37 C were measured as described above. Preparation of Vesicles for Fusion: β-Glucuronidase Assay. Vesicles encapsulating reaction components were obtained by adding 10 μL of each mixture to an aliquot of freeze dried membranes. One mixture contained 50 mM Hepes-KOH (pH 7.6), 290 nM β-glucuronidase and 1 μM TA647. The other contained 50 mM Hepes-KOH (pH 7.6), 50 μM PFB-FDGlcU, and 0.4 μM R-PE. Then, the vesicle suspension was dialyzed to remove the enzyme to inhibit the reaction from occurring outside of the vesicles. The resulting vesicle suspension was diluted to a ratio of 1:20 with 50 mM Hepes KOH (pH 7.6) and dialyzed using a microbiodialyzer. Precisely 100 μL of each vesicle solution encapsulating the reaction mixture was dispensed in the chamber with a polycarbonate membrane featuring a 0.4 μm pore size and dialyzed in 200 mL of buffer with stirring to remove the nonencapsulated molecules in the suspension.29 The buffer was changed every 60 min for 5 h, and then the final dialysis round was continued overnight. The removal of enzyme was confirmed indirectly by monitoring the intensity of fluorescent protein APC (80 kDa) in the dialysis buffer (Supporting Information Figure S1). When the dialysis was completed, 100 μL of vesicle suspension was collected from each chamber. The retrieved vesicle suspensions were decorated either by DDAB or oleic acid at MP = 25%. For fusion experiments, the prepared vesicle populations were mixed in various combinations and incubated at 37 C (Cool Stat 5000, Anatech). In addition to dialysis, 0.7 mg/mL protease was added after mixing and equilibration (10 min) to ensure that the enzyme activity in the outside solution was completely inhibited. The state of vesicles (association, fusion, and subsequent reaction) was analyzed with FACS (FACSAria cytofluorometer, BD Biosciences).25 For each 13083

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Langmuir specified time during FACS measurement, 5 μL of the vesicle suspension was sampled and diluted with 45 μL buffer. Vesicle Fusion to Trigger Gene Expression. Two POPC vesicle populations with complementary components were constructed. In addition, one vesicle population without the T7 RNA polymerase was used as negative control. The vesicle populations were made by adding 10 μL of each premix to an aliquot of freeze dried membranes. Three different inner solutions were prepared on ice: (i) 0.4 μM R-PE and 19 nM DNA in the PURE system solution;27 (ii) 1 μM TA647, 100 μM PFB-FDGlcU, and T7 RNA polymerase in the PURE system solution; and (iii) (negative control) 1 μM TA647 and 100 μM PFB-FDGlcU in the PURE system solution. After encapsulation, 9 μL of each vesicle suspension was diluted with 121 μL of 50 mM Hepes-KOH (pH 7.6) and used for large-pore dialysis in a cold room at 4.5 C to remove the nonencapsulated molecules, as was described above. After dialysis, the population encapsulating the DNA (mixture i) was decorated by adding 0.1 μL of 83 mM DDAB in ethanol to 9.9 μL of vesicles suspension (MPDDAB = 20%). Instead, the population with T7 RNA polymerase (mixture ii) and the negative control without T7 RNA polymerase (mixture iii) were decorated by adding 0.2 μL of 83 mM oleic acid in ethanol to 19.8 μL of each POPC vesicle suspension. After charging, the samples were left to equilibrate for 20 min at room temperature. Vesicle association and fusion was induced by mixing the positive and negative vesicle populations in a volume ratio of 1:2 v/v (30 μL total volume). After mixing and equilibration for 10 min at room temperature, 270 μL of the outer feeding solution mixture, which is the PURE system solution without proteins and tRNA, was used to dilute the mixed vesicle suspension. This was done to maintain an equilibrium of the specific factors across the lipid membrane (i.e., amino acids and energy sources) to allow efficient gene expression inside vesicles. The feeding solution was composed of 0.27 mM of each 20 amino acid mixture, 3.4 mM ATP, 2.26 mM GTP, 1.13 mM CTP, 1.13 mM UTP, 90.2 mM Hepes (pH 7.6), 252.2 mM K-Glu, 1.35 mM Spermidine, 18 mM Mg(OAc)2, 23 mM CP, 1.35 mM DTT, and 9.18 ng/μL FD. The reaction was incubated at 37 C for 240 min. The fluorescence intensities of the vesicles sampled at time 0 (after fusion and equilibration), 60, 120, 180, and 240 min were measured with the FACS machine to obtain the time course of the reaction in vesicles. Measurement of Giant Vesicles by FACS. Three fluorescent signals, from PFB-fluorescein, R-PE, and TA647, respectively, were measured by FACS. We obtained 100 000 data samples for each measurement. Briefly, PFB-fluorescein was excited with a 488 nm semiconductor laser, and the emission was detected through a 530 ( 15 nm bandpass filter. R-PE was excited with a 488 nm semiconductor laser, and the emission was detected through a 575 ( 13 nm bandpass filter. TA647 was excited with a HeNe laser (633 nm), and the emission was detected through a 660 ( 10 nm bandpass filter. Fluorescence Microscopy. The vesicle populations were observed using an inverted light microscope (IX81; Olympus, Japan) with a 60 oil-immersion objective lens and a digital color charge-coupled-device camera (VB-7000, Keyence, Japan). Bright-field images were obtained through differential interference contrast (DIC) observation. Images of the substrate PFB-FDGlcU processed inside the fused vesicles were obtained through the corresponding filter and dichroic mirror unit (NIBA, Olympus, Japan; excitation 470 490 nm, emission 510 550 nm).

’ RESULTS Schema of Gene Expression through Vesicle Fusion. In previous studies, we demonstrated that the fusion of vesicles was induced by modifying (decorating) the zwitterionic lipid membrane of vesicles with amphiphiles of opposite charges (oleic acid and DDAB as negative and positive amphiphiles, respectively).25,26 Here, we utilized this strategy to initiate biochemical reactions

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Figure 1. Schema of gene expression by vesicle fusion. Separate vesicle populations encapsulating DNA and T7 RNA polymerase/substrate were prepared. After decoration with opposite electrostatic charges by EtOH injection, the vesicles were allowed to associate and fuse to initiate gene expression.

Figure 2. Zeta potential of vesicle populations after decoration with charged amphiphiles at various concentrations (MP: molar percentage of charged amphiphiles). Error bars represent standard deviations of points measured in triplicate.

encapsulated in lipid vesicles via content mixing. The strategy used to regulate the encapsulated gene expression reaction based on vesicle vesicle fusion is outlined in Figure 1. One vesicle population contains DNA coding for the enzyme (e.g., β-glucuronidase) and the in vitro transcription/translation system (the PURE system). The complementary vesicle population contains T7 RNA polymerase, the PURE system, and the fluorogenic substrate that can be cleaved by the enzyme. When these contents are mixed via vesicle fusion, gene expression is initiated by T7 RNA polymerase, and the synthesized enzyme hydrolyzes the substrate, thus producing a detectable green fluorescent signal. The vesicle population was analyzed by FACS and fluorescence microscopy to evaluate the reaction initiated by vesicle fusion. As different fluorescence markers are encapsulated in each vesicle population, associated or fused vesicles emit signals from both markers simultaneously. Thus, fused vesicles in which the reactions proceeded were detected as the events that emitted all three fluorescence signals. Decoration of Neutral Vesicles by Charged Amphiphiles. To complete the schema in Figure 1, several intermediate tests were performed. The zeta potential of vesicle populations after charging with DDAB or oleic acid was measured to confirm that an electric charge was added to pre-existing zwitterionic POPC vesicles (Figure 2A). The graph shows that the POPC vesicle was electrically charged by the insertion of exogenous charged amphiphiles. The zeta potential increased with the amount of added amphiphiles until the phase of charge saturation was reached, as was also observed in other reports.26,30 When the 13084

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Figure 3. (A,B) Activity assays in the presence of vesicles. (A) Initial reaction velocity Vini of the hydrolysis reaction of β-glucuronidase in the presence of neutral or charged vesicles, plotted on a logarithmic scale, with no vesicles (buffer) as a control. (B) Time course of the synthesis of β-glucuronidase enzyme with the PURE system and the subsequent hydrolysis of the substrate in the presence of neutral or charged vesicles. (C,D) Activity assays of β-glucuronidase in the presence of charged amphiphiles. (C) Vini of β-glucuronidase with DDAB (dotted line) and oleic acid (solid line) at various concentrations in solution. (D) Vini of β-glucuronidase, which was inactivated by 600 μM DDAB, after the addition of oleic acid at various concentrations. In (C) and (D), Vini was normalized by the Vini without the presence of charged amphiphiles.

oppositely charged vesicle suspensions prepared at MP = 50% were mixed at 1:1 v/v, the net zeta potential of mixed vesicles was ∼ 10 mV, which is in between the values of the positive and negative populations. This result explains that vesicles associate to become nearly neutral in terms of the surface charge. For fusion experiments, we previously found that mixing the excess oleic acid vesicle suspension produces a higher vesicle fusion yield,26 likely because the absolute value of the zeta potential of vesicles with DDAB was twice that of oleic acid at the equivalent molar percentage. We note that, at pH 7.6, we expect more than half of the oleic acid molecules to be protonated with no charge since the pKa of the acid in the vesicle structure is near pH 8.5. Therefore, an excess of oleic acid is necessary to impart enough charge on the vesicle membrane. Exogenous Amphiphiles Affect Activities of Enzyme Reaction and Gene Expression. Introducing exogenously charged amphiphiles may affect the reactions encapsulated in vesicles. We examined the extent of this effect for the hydrolysis reaction of the enzyme and for gene expression of the enzyme by performing the reactions in the presence of the vesicles in solution. First, the reaction time course of 12 nM β-glucuronidase enzyme was tested by adding POPC only vesicles (GV), POPC vesicles charged with DDAB (MPDDAB = 25%, GV+), POPC vesicles charged with oleic acid (MPOA = 25%, GV ), and a mixture of these populations (1:2 v/v, thus MPDDAB = 8.3% and MPOA = 16.6% in the final mixture, GV+/ ) to the reaction mixture. As shown in the plot of initial reaction velocity (Vini) in Figure 3A, vesicles decorated with DDAB suppressed the enzymatic activity ∼1/1000-fold, whereas it was unaffected in the presence of vesicles decorated with oleic acid. It should be noted that the enzymatic activity was also unaffected by the mixture of positive

and negative vesicles (GV +/ ). In this case, the positive charge of DDAB was balanced with the negative charge of oleic acid, and the enzymatic activity was restored. Next, we tested the effect of charged vesicles on the activity of the gene expression reaction. A PURE system, complete with T7 RNA polymerase, DNA template encoding β-glucuronidase, and fluorogenic substrate, was prepared. The in vitro synthesis of active β-glucuronidase was monitored through quantification of the fluorescent product. To this system, the same set of vesicle populations was added, and reaction time courses were measured. Figure 3B shows the fluorescence signal of the PFBFDGlcU hydrolysis product over time. This experiment qualitatively confirmed the same effects with the enzyme reaction (Figure 3A). The enzyme was synthesized in vitro and was active in the presence of POPC and POPC/oleic acid vesicles (MPOA = 20%), whereas no activity was detected in the presence of POPC/DDAB vesicles (MPDDAB = 20%). However, in the mixture of positive and negative vesicles (1:2 v/v), the reaction condition was compatible with the expression and activity of β-glucuronidase. Figure 3A,B indicates that DDAB is a strong inhibitor of the enzyme and the gene expression system, the effect of which is canceled out by the addition of oleic acid. To clarify this point, we measured the activity of the β-glucuronidase enzyme in the presence of different concentrations of amphiphiles (Figure 3C). We found that the enzymatic activity is lost when DDAB is added at a concentration above 60 μM, whereas the addition of oleic acid has no significant effect. In addition, when oleic acid was added to a DDAB-inactivated enzyme solution, the enzymatic activity was restored (Figure 3D). DDAB has been reported to be a cationic amphiphile that can affect the conformation of 13085

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Figure 4. Detection of enzymatic activity through vesicle vesicle fusion by FACS. (A) Two-dimensional density maps showing the fluorescence intensities from the mixture of vesicle populations containing an enzyme and fluorogenic substrate, both decorated by oleic acid. The plot of two volume markers (A1) is used to visualize the extent of vesicle association, whereas the plot of the green reaction product and one of the volume markers (A2) is used to evaluate the extent of fusion and the reaction in vesicles. (B) The same density maps showing the mixture of vesicle populations that were decorated by positive or negative amphiphiles and then mixed together. (C1) Temporal change in the ratio of associated vesicles Ra. (C2) Temporal change in the ratio of fused vesicles Rf in which the β-glucuronidase-catalyzed reaction proceeded.

proteins.31 It is thus reasonable to assume that β-glucuronidase has lost its activity due to the conformational change induced by the addition of DDAB. It should be noted that the loss of activity caused by DDAB is not specific to β-glucuronidase. The activity of another protein (fluorescent protein APC) was also lost at a DDAB concentration of above 60 μM (Supporting Information Figure S2). The recovery of the activity by the addition of oleic acid can thus be explained by the restoration of the conformation of the enzyme by neutralizing the charge of DDAB. We conjecture that the same inhibition and restoration mechanism also took place in the gene expression system (Figure 3B). Initiating an Enzyme Substrate Reaction via Vesicle Fusion. Because β-glucuronidase was active when the positive charge from DDAB was neutralized by oleic acid, we tested whether the enzyme was also active after the fusion of vesicles that were decorated with opposite charges. Vesicle populations of POPC, containing either the fluorogenic substrate or β-glucuronidase enzyme, were prepared and decorated by DDAB or oleic

acid. The vesicle populations were then mixed to allow association and fusion. The resulting vesicle population was evaluated by the fluorescence signal of the encapsulated reaction before and after incubation at 37 C for 3 h. To ensure the exclusion of artifacts, three control samples were measured in parallel. Figure 4A, B shows the 2-D density maps plotted for three fluorescence intensities from vesicles after 3 h of incubation (plots at t = 0 are shown in the Supporting Information Figure S3). As an example of the control experiments, a mixture of negative negative vesicle populations is shown in Figure 3A; that is, two populations each containing enzyme and substrate, both decorated by oleic acid, were mixed. In this case, as shown in the plot of two internal volume fluorescence markers (R-PE and TA647, Figure 4A1), an obvious distinction between the two populations was observed. This pattern indicates that two vesicle populations were present in the suspension without physically interacting with each other because of the electrostatic repulsion between the vesicles carrying the same charge. The same measurement 13086

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Figure 5. Expression of β-glucuronidase enzymatic activity from a DNA template triggered by selective vesicle fusion. (A) Two-dimensional density maps showing the fluorescence intensities from the mixture of vesicle populations containing DNA and substrate (without T7 RNA polymerase), decorated by DDAB and oleic acid. (B) The same density maps for the mixture of vesicle populations, in which T7 RNA polymerase was included to complement the reaction. (C1) Temporal change in the ratio of associated vesicles Ra. (C2) Temporal change in the ratio of fused vesicles, Rf, in which the synthesis of β-glucuronidase and its substrate hydrolysis proceeded.

data were plotted for the green fluorescence from the hydrolyzed substrate along the x-axis and for one of the volume markers (R-PE) along the y-axis, respectively (Figure 4A2). This combination illustrates the extent of the enzymatic reaction in the vesicles. In this plot, the majority of the vesicle population remained in the region with low green intensity, even after incubation, reflecting the fact that the mixing of vesicle internal contents did not take place. None of the other controls tested (combinations of positive positive and neutral neutral vesicles) showed an appreciable green fluorescence signal compared to the following fusion condition (Supporting Information Figure S3). Figure 4B shows the measurement results for the mixture of positive-negative vesicle populations (i.e., DDAB-decorated vesicles with PFB-FDGlcU and oleic-acid-decorated vesicles with β-glucuronidase). The plot of two volume markers (Figure 4B1) shows that two vesicle populations are associated (physically attached) because most of the vesicles exhibit R-PE and TA647 fluorescence simultaneously. In Figure 4B2, it is obvious that the

intense green signal, which is proportional to the vesicle volume, appeared in the majority of vesicles, indicating that the enzyme reaction took place in vesicles as a result of the content mixing in associated vesicles. From the time-sampled data sets, we evaluated the variation of vesicle association and fusion according to the definition used in our previous report.25 Here, we focused on vesicles larger than ∼2 fL (i.e., vesicles that had an R-PE intensity greater than a certain threshold), because we could not resolve the association and the reaction of subfemtoliter vesicles.25 The association ratio Ra was defined as the fraction of vesicles that had both volume markers (region R1) among all vesicles that had an R-PE intensity above the threshold. The fusion ratio Rf was defined as the fraction of fused vesicles that had high green fluorescent intensity (events that fell in both regions R1 and R2) among all vesicles that had an R-PE intensity above the threshold. The threshold for green fluorescence was determined by the background signal at t = 0 min (Supporting Information Figure S3). The time courses 13087

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Langmuir of Ra and Rf, measured for various combinatorial mixtures of charged vesicles are plotted in Figure 4C1 and 4C2, respectively. In the case of oppositely charged vesicles (positive negative mixture, GV( in Figure 4C1), the association ratio Ra reached ∼60% immediately after mixing. The Ra of the neutral neutral (GV) and positive positive (GV+) mixtures showed a comparable extent of association with a slight delay (∼20 min), whereas the negative negative mixture exhibited practically no association (GV ). Meanwhile, only the positive negative mixture exhibited a considerable increase in the reacted vesicles, which reached a plateau of ∼50% in less than 20 min (Figure 4C2). These results confirm the following features in the vesicle fusion system. First, although relatively slow association occurs with neutral neutral and positive positive vesicle mixtures, the enzymatic reaction associated with internal content mixing occurs only in the positive negative mixture with rapid association. Second, the reaction in the positive negative mixture shows that the lipid mixture of POPC, DDAB, and oleic acid that results from the vesicle fusion is compatible with the encapsulated enzymatic activity. Third, the association and fusion (content mixing) mostly completed within 20 min. To visually confirm the third proposition, we observed vesicles after the fusion process with a fluorescent microscope. In the mixture of two vesicle populations dyed with different fluorescent colors, most of the vesicle aggregates exhibited overlapping colors with respect to both lipids and internal contents within 30 min (Supporting Information Figures S4 and S5). This observation is in accordance with the measurements shown in Figure 4C1,C2. Triggering a Gene Expression Reaction via Vesicle Fusion. Finally, we demonstrated the expression and activity of the β-glucuronidase enzyme27,28 inside vesicles triggered by vesicle fusion, as depicted in Figure 1. Two vesicle populations, each containing DNA and T7 RNA polymerase/substrate decorated, respectively, with DDAB and oleic acid at MP = 20%, were mixed in a 1:2 volume ratio. During incubation at 37 C, the vesicle suspension was sampled at various times, and the intensities of three fluorescence signals from the vesicles in the mixed suspension were measured. The plot in Figure 5A1 represents vesicle association after 240 min of incubation for the condition in which the T7 RNA polymerase was excluded. Association was detected by the simultaneous emission of the volume markers (TA 647 and R-PE) along the x y axes. Figure 5A2 shows that the intensity of the green signal remained almost unchanged compared to the background level at t = 0 (Supporting Information Figure S6). This result also shows that the activity of the reporter gene (β-glucuronidase) was not observed, which indicated that the lack of gene expression due to the absence of transcription produced no green signal. However, when T7 RNA polymerase was included in the negative vesicle population, we observed an apparent increase in green intensity after 240 min of incubation (Figure 5B2), whereas the pattern of vesicle association appeared to be similar as expected due to the oppositely charged vesicles used in both cases. The time courses of Ra and Rf were evaluated and are plotted in Figure 5C. It is clear that the association immediately occurred and remained constant under both conditions (Figure 5C1). However, the values of Rf showed a completely different trend (Figure 5C2). Without T7 RNA polymerase, the fraction of vesicles with a green signal did not increase over time. In contrast, when T7 RNA polymerase was included, ∼10% of the vesicles among associated vesicles expressed β-glucuronidase activity synthesized in the PURE system after 240 min of incubation. Moreover, the time course exhibited

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Figure 6. Micrographs of fused vesicles showing the product of gene expression. Images of vesicles are presented in pairs with DIC on the left and green fluorescence on the right.

a concave curve, which is interpreted as a result of a multistep reaction.28 Thus, the increase in green signal intensity was the result of the expression of the β-glucuronidase enzyme that then cleaved the fluorogenic substrate and was not due to other confounding effects of the complex system, as evident in the control (Figure 5A2). In addition, the vesicle populations after fusion and the gene expression reaction were examined by microscopy. In Figure 6, DIC and epi-fluorescence micrographs of POPC/DDAB MP = 20% and POPC/oleic acid MP = 20% mixed in a ratio of 1:2 (v/v) and incubated for 240 min at 37 C are shown. The activity of the enzyme inside the mixed volume of the fused compartments was detected by the presence of the green fluorescent product. This provides visual confirmation of the production of green fluorescent signal inside the fused vesicles.

’ DISCUSSION Several steps toward the creation of an artificial cell are presented. It is known that the membrane composition can affect the activity of enzymes and protein expression systems.15,32,33 We have shown that, for our system, the excess positive charge produced by decorating POPC vesicles with DDAB almost completely inhibited the ability of β-glucuronidase to process its fluorogenic substrate (Figure 3A). The same inhibition was observed when the PURE system was mixed with POPC/DDAB vesicles (Figure 3B). The inhibitory effects were again overcome when the net positive charge in the membrane was neutralized by oleic acid. Remarkably, the same inhibition and restoration effects were also observed for reactions encapsulated within the minute volume of the vesicles. The reaction activity was restored through a fusion event between positively and negatively charged vesicles because the net charge of the membrane was neutralized via mixing with amphiphiles. This may serve as a useful tool for either regulating encapsulated reactions by the external manipulation of the charge ratios on the vesicles or self-regulation through which the vesicles can support such an enzymatic reaction only if they undergo a successful fusion event to neutralize the membrane charge. We argue that not only is functionality necessary for an artificial cell, but rudimentary regulation of activity is also required to produce the desired level of sophistication for an artificial cell. The present mechanism demonstrates the utility of membrane composition in creating regulation and material transfer pathways, essential mechanisms in cells and cell models. Successful regulation and initiation of a complete gene expression network via vesicle fusion is highlighted here as this brings us one 13088

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Langmuir step closer to making an artificial cell capable of controlled and sustained protein synthesis, a process ubiquitous in natural cells. We are currently developing the system further to allow successive decoration steps to control successive fusion to continuously feed the encapsulated metabolism as well as to introduce variation and complexity to the system. Such controlled steps are necessary for the dynamic construction and evolution of an artificial cell.3,11,14 Not only can the internal contents of the vesicles be changed and renewed over time, but the membrane components can also be renewed, allowing for perpetuation, perturbation, and maintenance of the entire system over time. In an effort to synthesize an artificial cell, important steps have already been made. For instance, biosynthesis of phospholipids that exploit cells’ metabolic pathways inside vesicles,4 dynamic compartmentalization in synthetic cells,34 reconstitution of a cytoskeleton actin network within GUV,35 and RNA replication by selfencoded RNA-dependent RNA replicase within giant vesicles17 have been reported. Here, we show for the first time that genetic expression from DNA can be activated after controlled vesicle fusion with internal content mixing. Although a universal definition for life might be controversial, helpful guidelines that focus on experimentation in the laboratory are proposed.5,14,36 38 A Turing test for artificial cells is possibly a good strategy10 to argue that artificial cells are truly alive. We would like to create a basic artificial cell based on modern biological functionalities. The artificial cell would have a basic genome and metabolism encapsulated by a membrane. With the membrane acting as a semipermeable barrier, many of the resources required to sustain minimal metabolism (such as ribosomes) will not be taken up through permeation. Other mechanisms, such as controlled vesicle vesicle fusion, are necessary to sustain and, eventually, with selection imposed, evolve artificial cells. Our goal to create such an evolving artificial cell stems from our study of the processes of evolution in living systems. Such living systems are too complex to understand how evolution changes in an organism at all levels of description over time. It is our goal to create a simplified artificial cell capable of evolution for which every component is specified and can be characterized over time. In this way, we may gain valuable insight into the processes of evolution, especially in regard to more simplified forms of life that may have existed during Earth’s early history.

’ ASSOCIATED CONTENT

bS

Supporting Information. Graph of the fluorescence intensity of APC fluorescent protein measured in the dialysis buffer at different time intervals during the dialysis process. Inhibition and restoration of the activity of fluorescent protein APC by addition of DDAB and oleic acid. Density maps for the mixture of vesicle populations, each containing an enzyme and a substrate, which were decorated by different combinations of amphiphiles. Micrographs show the extent of lipid and content mixing. Density maps for the mixture of vesicle populations, each containing complementary mixtures for gene expression, were decorated by DDAB and oleic acid. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*Tel: +81-6-6879-4171. Fax: +81-6-6879-7433. E-mail: yomo@ ist.osaka-u.ac.jp.

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’ ACKNOWLEDGMENT This research was supported in part by “Special Coordination Funds for Promoting Science and Technology: Yuragi Project”, the Global COE (Centers of Excellence) Program of the Japanese Ministry of Education, Culture, Sports, Science, and Technology and the JASSO organization (Japan student services organization), which provides scholarships for exchange programs at Osaka University. Dr. Kazufumi Hosoda is gratefully acknowledged for helpful discussion on protein synthesis inside vesicles. We also thank Dr. Yasuaki Kazuta, Ms. Hitomi Komai, and Tomomi Sakamoto for producing the PURE system. ’ REFERENCES (1) Gibson, D. C.; et al. Science 2010, 5987, 52. (2) Noireaux, V.; Libchaber, A. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 17669. (3) Ichihashi, N.; Matsuura, T.; Kita, H.; Sunami, T.; Suzuki, H.; Yomo, T. Cold Spring Harbor Perspect. Biol. 2010, 2 (6), a004945. (4) Kuruma, Y.; Stano, P.; Ueda, T.; Lusi, P. L. Biochim. Biophys. Acta 2009, 1788, 567. (5) Murtas, G.; Kuruma, Y.; Bianchini, P.; Diaspro, A.; Luisi, P. L. Biochim. Biophys. Res. Commun. 2007, 363, 12. (6) Yu, W.; Sato, K.; Wakabayashi, M.; Nakaishi, T.; Ko-mitamura, E. P.; Shima, Y.; Urabe, I.; Yomo, T. J. Biosci. Bioeng. 2001, 92, 590. (7) Nomura, S-i. M.; Tsumoto, K.; Hamada, T.; Akiyoshi, K.; Nakatani, Y.; Yoshikawa, K. ChemBioChem 2003, 4, 1172. (8) de Souza, T. P.; Stano, P.; Luisi, P. L. ChemBioChem 2009, 10, 1056. (9) Rasmussen, S.; Chen, L.; Deamer, D.; Krakauer, D.; Packard, N.; Stadler, P.; Bedau, M. Science 2004, 303, 963. (10) Cronin, L.; Krasnogor, N.; Davis, B. G.; Alexander, C.; Robertson, N.; Steinke, J. H. G.; Schroeder, S. L. M.; Khlobystov, A. N.; Cooper, G.; Gardner, P. M.; Siepmann, P.; Whitaker, B. J.; Marsh, D. Nat. Biotechnol. 2006, 24, 1203. (11) Szostak, J. W.; Bartel, D. P.; Luisi, P. L. Nature 2001, 409, 387. (12) Forster, A. C.; Church, G. M. Mol. Syst. Biol. 2006, 2, 45. (13) Jewett, M. C.; Forster, A. C. Curr. Opin. Biotechnol. 2010, 21, 697. (14) Luisi, P. L.; Ferri, F.; Stano, P. Naturwissenschaften 2006, 93, 1. (15) Sunami, T.; Hosoda, K.; Suzuki, H.; Matsuura, T.; Yomo, T. Langmuir 2010, 26, 8544. (16) Shimizu, Y.; Inove, A.; Tomari, Y.; Suzuki, T.; Yokogawa, T.; Nishikawa, K.; Ueda, T. Nat. Biotechnol. 2001, 19, 751. (17) Kita, H.; Matsuura, T.; Sunami, T.; Hosoda, K.; Ichinashi, N.; Tsukada, K.; Urabe, I.; Yomo, T. ChemBioChem 2008, 9, 2403. (18) Prestegard, J. H.; O’Brien, M. P. Annu. Rev. Phys. Chem. 1987, 38, 383. (19) Grafmuller, A.; Shillocock, J.; Lipowsky, R. Biophys. J. 2009, 96, 2658. (20) Tamm, L. K.; Crane, J.; Kiessling, V. Curr. Opin. Struct. Biol. 2003, 13, 453. (21) Haluska, C. K.; Riske, K. A.; Marchi-Artzner, V.; Lehn, J. M.; Lipowsky, R.; Dimova, R. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 15841. (22) Franzin, C. M.; Macdonald, P. M. Biochemistry 1997, 36, 2360. (23) Pantazatos, D. P.; MacDonald, R. C. J. Membr. Biol. 1999, 170, 27. (24) Tsumoto, K.; Kamiya, K.; Yoshimura, T. Proc. Int. Symp. MicroNanomechatronics. Human Sci. 2006, 7. (25) Sunami, T.; Caschera, F.; Morita, Y.; Toyota, T.; Nishimura, K.; Matsuura, T.; Suzuki, H.; Hanczyc, M. M.; Yomo, T. Langmuir 2010, 26, 15098. (26) Caschera, F.; Stano, P.; Luisi, P. L. J. Colloid Interface Sci. 2010, 345, 561. (27) Hosoda, K.; Sunami, T.; Kazuta, Y.; Matsuura, T.; Suzuki, H.; Yomo, T. Langmuir 2008, 24, 13540. 13089

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