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Vesicle Encapsulation of a Non-biological Photochemical System Capable of Reducing NAD+ to NADH David P. Summers* and David Rodoni†
*Carl Sagan Center, SETI Institute, c/o NASA Ames Research Center, Mail Stop 239-4, Moffett Field, CA 94035 † Foothill College, Los Altos, CA KEYWORDS. Vesicles, Liposomes, Artificial Cell, Origin of Life, Origin of Photosynthesis, Energy Transduction, Minerals, Semiconductors, Particles
ABSTRACT. One of the fundamental structures of a cell is the membrane. Self-assembling lipid bilayer vesicles can form the membrane of an artificial cell and could also have plausibly assembled prebiotically for the origin of life. Such cell like structures that encapsulate some basic subset of the functions of living cells are important for research to infer the minimum chemistry necessary for a cell, to help understand the origin of life, and to allow the production of useful species in microscopic containers. We show that encapsulation of TiO2 particles has the potential to provide the basis for an energy transduction system inside vesicles which can be used to drive subsequent chemistry. TiO2 encapsulated inside vesicles can be used to produce biochemical species such as NADH. The NADH is formed from NAD+ reduction and is produced in a form that is able to drive
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further enzymatic chemistry. This allows us to link a mineral-based, non-biological, photosystem to biochemical reactions. This is a fundamental step toward being able to use this mineral photosystem in a protocell/artificial cell.
INTRODUCTION. There is interest in methods of creating cell like structures, “artificial cells”, that contain some basic subset of the functions of living cells. There are a number of motivations for such research. For example, to be able to infer the minimum chemistry necessary for a cell1-4, to help understand the origin of life1, 5-8, and to allow the production of useful species in microscopic containers.3, 9 One of the fundamental structures of a cell is the membrane. Self-assembling lipid bilayer vesicles can form the basis for the membrane of an artificial cell into which a chemical system can be incorporated.1, 2, 4, 9 Such structures could also have plausibly assembled prebiotically to play a role in the origin of life.1, 5, 8, 10-12 Another important cellular function is photosynthesis, providing the basis for most of life today. Incorporation of a photochemical system into a vesicle can allow it to capture and transduce light energy, enabling the vesicle to grow in size, reproduce, or produce species of technological or scientific interest.1, 3 How, and when, photosynthesis developed is also a central issue in the origin and early evolution of life, as is the role vesicles may have played in that development.1, 11, 13-18 Minerals can introduce a number of useful properties into vesicle systems. The incorporation of clay particles into vesicles has been demonstrated to be a way to introduce catalytic properties and to template the formation of macromolecules.19-21 We have shown that semiconducting particles, such as titanium dioxide, can use light energy to drive photoelectrochemical reactions inside vesicles.22 Here, we wish to report on a chemical system where the photocatalytic activity of titanium dioxide particles can be used to produce species that have the potential to carry out subsequent
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biochemical reactions inside vesicles, specifically the production of NADH (Nicotinamide adenine dinucleotide hydride) from NAD+ (Nicotinamide adenine dinucleotide). NADH (and the very similar molecule, NADPH) are important biochemical carriers in living systems and drive a number of enzyme catalyzed reactions. The results described here show that a titanium dioxide photocatalytic system can, while encapsulated inside of a vesicle, photochemically produce enzymatically active NADH. For the first time, this provides a way to link a mineral-based, non-biological, photosystem to biochemical reactions. This is a fundamental step toward being able to use this mineral photosystem in a protocell/artificial cell. EXPERIMENTAL. Colloidal suspensions of TiO2 (10%, < 50nm BET, 23% rutile & 77% anatase) were purchased from Sigma-Aldrich (cat. No. 700347). For all experiments here, suspensions were centrifuged, for 60-75s at 13750 rpm, before use in order to remove heavier particles, which may be less photoactive and less likely to remain suspended for a long period of time. The remaining concentration was determined by the weight if TiO2 remaining after removal of the water. Centrifugation can limit the concentration of TiO2 and was eventually found not to be required. Suspensions made up from a nanopowder (Sigma-Aldrich) were also active. No surfactant was added in any experiment. Typical experiments have used 0.025 M serine or 3% methanol as an electron donor. Concentrations are as noted in results and discussion. Rh(bipyridine)3+3, Rhodium tris(2,2’-bipyridine) chloride, was prepared by the method of Kirch et al.23 It is present as an electron mediator, becoming reduced at the particle surface by two electrons23, which are then passed on, as a pair, to the NAD+. (See Figure 1.) It has been previously established that Rh(bipyridine)3+3 is necessary for the formation of enzymatically active products such as NADH.24, 25 In current experiments, we also do not observe enzymatically active product without it. TiO2 is also required. Before encapsulation experiments, we observed that, in the
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absence of TiO2, NAD+ degradation products were observed instead of the formation of NADH or NAD dimer (see Supplementary Information SI1). To encapsulate species in vesicles, in a typical experiment, chicken egg L-α-phosphatidylcholine in chloroform (Avanti Polar Lipids), in sufficient volume to constitute the desired amount of lipid (as indicated below), was dried down under a stream of nitrogen and exposed in vacuo for ½-1 hour. An aqueous suspension of TiO2 particles, dispersed in the desired solution, was added with vortex stirring and vigorous shaking until the lipid was all suspended. Fatty acid vesicles were prepared by using the method of Hargreaves & Deamer.26 A solution of oleic acid was dissolved in a minimum amount of 0.4 M NaOH. The pH was then lowered by the addition of water and 0.4 M boric acid to create a pH 9, 0.1 M borate, buffer. Dodecanol, which strengthens the membrane, was then added to the suspension and allowed to dissolve in the vesicle walls. Species to be encapsulated were present in solution before the pH was lowered. Species can also be added after vesicle formation by sonication of the suspension to break open and reform vesicles, allowing them to encapsulate the added species. External species were removed by size exclusion, as described below. Unless otherwise indicated, the method of Deamer and Barchfield was used to concentrate species inside the vesicles.27 A suspension was produced as described above and evaporated to dryness under a stream of nitrogen. A moist kimwipe™ was placed in the top of the tube and the residue was hydrated overnight. The residue was resuspended in buffer by vigorous vortex stirring and passed through a micropore filter with a 1 µm pore size. This procedure was not applied to fatty acid vesicles, which will encapsulate the solution without concentration.26 The suspension was, after passage through a micropore filter, passed down a Sephadex™ G-50 size exclusion column (2.2 cm x 17.5 cm) from which the turbid fractions were collected.27 This removed NAD+/NADH, and smaller species, external to the vesicles, allowing us to observe just the activity encapsulated inside.4, 27, 28 (TiO2 is held up, and needs to be periodically cleaned from the
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column. It is not known how efficiently this occurs. In any case, TiO2 external to the vesicle will have no NAD+ with which to react.) Typically, the last turbid fraction, ~10% of the total, was discarded as a precaution. The separation of vesicles, from externally dissolved species, was confirmed by taking the UV-Vis spectrum of aliquots as they came off the column. This showed clean separation of the vesicles from the external NAD+. Species which can be lost during drying, or during size exclusion chromatography (such as methanol), were re-added prior to experiments. Experiments had ~1015 vesicles/liter. Samples were purged with nitrogen in a septum sealed quartz cuvette prior to irradiation. The light source was a medium pressure mercury arc lamp (Hanovia, 450 watt) in quartz, water cooled, well. A glass bandpass filter was used to isolate the 365 nm peak. Sample were placed ~1 cm outside of the filter with a resultant intensity of ~7.5 mW/cm2. Except when noted, NADH was detected by fluorescence (see below), after the titanium dioxide was removed, since TiO2 was found to quench NADH fluorescence. Removal of the TiO2 was accomplished by extracting twice with butanol (to remove vesicle materials and open up the vesicles), extraction with hexane/dichloromethane (to remove the butanol), and centrifugation for 15 min at 13750 rpm (to remove titanium dioxide particles). An excitation wavelength of 340 nm was used to obtain emission spectra. No emission from TiO2 or Rh(bipy)3+3 was observed over the wavelengths studied, ~390 nm and up. Rh(bipy)3+3, which was present in a concentration 80x lower than that of NAD+, can emit in this region23 but the emission was either too faint, the complex couldn’t compete with other species for the absorption of excitation light,23, 29 and/or the emission was quenched by NAD+, and/or NADH. Fluorescent Microscopy was done using a Nikon Microphot – FXA instrument with a FXA/SA Epifluorescent attachment using UV excitation. The excitation was 330 to 380nm in wavelength, peaking at 365nm. The camera sold with the instrument, FX-35DX, was used with 800 ASA film and 2 second exposure. Suspensions used for fluorescent microscopy were not passed through a 1 µm filter to
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allow for larger, easier to view, vesicles. We tested for NADH enzymatic activity with lactate dehydrogenase (Sigma Aldrich) using pyruvate as a substrate.24, 25, 30 NADH was not consumed unless both enzyme and pyruvate were present. RESULTS AND DISCUSSION. A suspension (8.5 mM NAD+, 2% TiO2, and 3% methanol in a 0.5 M pH 9 carbonate buffer with 0.1 mM Rh(bipy)3+3 as a mediator, see above) was encapsulated inside vesicles formed from 22 mM phosphatidylcholine. A schematic of the reaction system is shown in figure 1. After dehydrate/rehydrate cycling and removal of external species, as described in the experimental section, samples were irradiated with 365 nm light. After irradiation, the NADH formed was detected by fluorescence. Since the emission of NADH is quenched by TiO2, the vesicles were opened and the TiO2 particles were removed as described in the experimental sections. In figure 2 is shown the emission spectrum, using 340 nm excitation, of the NADH produced. The peak at 383 nm varied with the excitation wavelength and is due to inelastic scattering. The spectrum observed has the same shape and wavelength maximum as an authentic sample of NADH. The excitation profile matches the lowest energy absorption band for NADH and the excitation profile of authentic NADH samples under the same conditions. In experiments where the vesicles had not been concentrated by dehydrate/rehydrate cycling, we observe similar results, but with a lower intensity emission. We have also seen similar results if NADP+ is used instead of NAD+. Experiments were only run until a low conversion to NADH was achieved, since NADH self-quenching limits emissions at higher concentrations. Based on the total amount of NADH produced (from the intensity of the NADH emission, after removal of the vesicles and particles), and the estimated encapsulation volume using an area of 0.25 nm2 for a lipid pair, an estimate of the conversion of NAD+ to NADH was ~1%. As illustrated in Figure 1, vesicles can be unilamellar or multilamellar (single walled and multiwalled). Under the conditions used, to allow larger vesicles (and easier detection of product), the vesicles were a mix of unilamellar and multilamellar vesicles. With 1 µm pore size extrusion,
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about a third of the vesicles are multilamellar.31 This is not expected to affect the results. The photosystem has been demonstrated in unilamellar vesicles.22 Since the emission spectrum of NADH is similar to that of the enzymatically inactive NAD dimer, the enzymatic activity of the product was tested. It was verified that the product will, with the enzyme lactate dehydrogenase, reduce pyruvate to lactate (equation 1). NADH + pyruvate + H+ -> NAD+ + Lactate
(1)
It was also necessary to test for enzymatic activity because the importance in producing NADH lies in it ability to drive enzymatic reactions. A sample (similar to that described above) was created, the lipid and titanium dioxide were removed, the sample was adjusted to pH 7.5, and enzyme and pyruvate were added. In figure 3, one can see that the NADH was consumed by the presence of enzyme and pyruvate. Clearly NADH was being produced and the NADH was capable of being a substrate in enzymatic reactions. In figure 4 is a micrograph showing emission after a solution (15 mM NAD+, 0.2% TiO2, 0.1 mM Rh(bipy)3+3, and 3% methanol in a 0.5 M pH 9 carbonate buffer) that was encapsulated inside vesicles (22 mM phosphatidylcholine) was irradiated for two hours. The field of view is about 175 µm across. The source of emission is associated with the vesicles and the color is consistent with the wavelengths of NADH emission. The emission from these vesicles is more intense than most in the sample (likely having enough NADH to be observed well, but not having too much NADH, or TiO2, to quench or self-quench). The reader may be able to make out some dim spots in the right center of the image which correspond to isolated and dimmer vesicles. The bright spots in the leftcenter of the picture are from clumps of vesicles. See also Supplementary Information SI2 and SI3. In these experiments, one can typically see a large number of vesicles which glow very faintly to the eye, but which are too dim to be captured on film. Experiments were also conducted inside fatty acid/fatty alcohol vesicles, instead of phospholipid vesicles. While fatty acids and fatty alcohols are considered more plausible prebiotic species, for
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the origin of life, this result also shows that the encapsulated chemistry is not dependent on the type of vesicles, or how they were made (see experimental). In figure 5, we see that NADH is formed from irradiation of a solution (8 mM NAD+, 2% TiO2, 0.1 mM Rh(bipy)3+3, and 3% methanol in a 0.1 M borate buffer, pH 9) encapsulated in vesicles made with oleic acid/dodecanol (46 mM oleic acid, 32 mM dodecanol). Testing with pyruvate and lactate dehydrogenase proved this product to be enzymatically active, figure 6. The intensity of the emission show in figure 5 is higher than in figure 2. It is not known if the dehydrate/rehydrate cycles used on the previous experiment equally concentrate all species, which may affect the results. However, one likely explanation is simply that this experiment used a larger amount of lipid, resulting in a larger encapsulated volume. CONCLUSIONS. Mineral particle photocatalysis can be used to drive photochemical reactions inside vesicles. Such energy transduction can produce biochemical species that are associated with cellular processes, in this case a species responsible for biochemical redox transfer. The species produced here, NADH, is enzymatically active and so is able to drive subsequent enzymatic chemistry and feed into more complex biochemistry. This provides a fundamental and necessary link between a non-biological, mineral-based, photosystem and enzymatic reactions that can then carry out biochemistry in a protocell/artificial cell. Thus, such a system can provide the basis for a simple photosynthetic system that can provide energy transduction for an artificial cell. The ability to photochemically drive enzymatic reactions can allow a wide range of possible reactions that can be useful for technological applications. In a prebiotic setting, a source of energy transduction could provide a basis for a “protobiochemistry” for protocells and the origin of life. Such a system can also represent a model of a simple photochemical system that preceded the current complex system in the origin of photosynthesis. Similarly, a simple system that doesn’t require complex machinery would make an origin of life involving photosynthetic organisms more plausible.
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ACKNOWLEDGEMENTS. The Authors would like to thank NASA’s Astrobiology: Exobiology and Evolutionary Biology program for funding. SUPPORTING INFORMATION AVAILABLE. SI1: Emission Spectrum of Product Generated without TiO2. SI2: Micrograph of the vesicles using both UV fluorescence and visible light transmittance. SI3: Micrograph of a large clump of vesicles using; visible light transmittance microscopy, visible transmittance and UV fluorescence microscopy, and UV fluorescence microscopy. This information is available free of charge via the Internet at http://pubs.acs.org/.
*
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REFERENCES. (1) Pohorille, A.; Deamer, D. Artificial Cell: Prospects for Biotechnology. Trends Biotech 2002, 20, 123-128. (2) Walde, P. Buidling Artificial Calls and Protocell Models: Experiments Approaches with Lipid Vesicles. BioEssays 2010, 32, 296-303. (3) Zhang, Y.; Ruder, W. C.; LeDuc, P. R. Artificial Cells: Building Bioinspired Systems Using Small-Scale Biology. Trends. Biotech. 2008, 26, 14-20. (4) Oberholzer, T.; Wick, R.; Luisi, P. L.; Biebricher, C. K. Enzymatic RNA replication in selfreproducing vesicles: an approach to a minimal cell. Biochemical and biophysical research communications 1995, 207, 250-257. (5) Luisi, P. L.; Stano, P.; Rasi, S.; Mavelli, F. A possible route to prebiotic vesicle reproduction. Artificial Life 2004, 10, 297-308. (6) Morigaki, K.; Dallavalle, S.; Walde, P.; Colonna, S.; Luisi, P. L. Autopoietic self-reproduction of chiral fatty acid vesicles. Journal of the American Chemical Society 1997, 119, 292-301. (7) Szostak, J. W.; Bartel, D. P.; Luisi, P. L. Synthesizing life. Nature 2001, 409, 387-390. (8) Walde, P.; Wick, R.; Fresta, M.; Mangone, A.; Luisi, P. L. Autopoietic self-reproduction of fatty acid vesicles. Journal of the American Chemical Society 1994, 116, 11649-11654. (9) Chandrawati, R.; Caruso, F. Biomimetic Liposome- and Polymersome-Based Multicompartmentalized Assemblies. Langmuir 2012, 28, 13798-13807. (10) Hargreaves, W. R.; Mulvihill, S.; Deamer, D. W. Synthesis of Phospholipids and Membranes in Prebiotic Conditions. Nature 1977, 266, 78-80. (11) Koch, A. L. Primeval Cells: Possible Energy-Generating and Cell-Division Mechanisms. Journal of Molecular Evolution 1985, 21, 270-277. (12) Pohorille, A.; Wilson, M. A. Molecular Dynamics Studies of Simple Membrane-Water Interfaces: Structure and Functions in the Beginnings of Cellular Life. Orig. Life Evol. Biosphere 1995, 25, 21-46. (13) Blankenship, R. E. Chapter 11: Origin and Evolution of Photosynthesis. In Molecular Mechanisms of Photosynthesis, Blackwell Science: London, 2002; pp 220-257. (14) Willner, I.; Mandler, D.; Maidan, R. Bio-Models and Artificial Models for Photosynthesis. New Journal of Chemistry 1987, 11, 109-121. (15) King, C. C. Did Membrane Electrochemistry Precede Translation? Origins Life Evol. Biosphere 1990, 20, 15-25. (16) Deamer, D. W.; Harang, E. Light-dependent pH gradients are generated in liposomes containing ferrocyanide. Biosystems 1990, 24, 1-4. (17) Sun, K.; Mauzerall, D. A Simple Light-Driven Transmembrane Proton Pump. Proceedings of the National Academy of Sciences, USA 1996, 93, 10758-10762. (18) Deamer, D.; Dworkin, J. P.; Sandford, S. A.; Bernstein, M. P.; Allamandola, L. J. The First Cell Membranes. Astrobiology 2002, 2, 371-381. (19) Li, Z.; Guo, Y.; Scriven, L. E.; Davis, H. T. Stabilization of Aqeous Clay Suspensions with AOT Vesicular Solutions. Colloids and Surfaces A 1996, 106, 149-159. (20) Hanczyc, M. M.; Fujikawa, S. M.; Szostak, J. W. Experimental Models of Primitive Cellular Compartments: Encapsulation, Growth, and Division. Science 2003, 302, 618-622. (21) Hanczyc, M. M.; Mansy, S. S.; Szostak, J. W. Mineral Surface Directed Membrane Assembly. Orig. Life Evol. Biosphere 2007, 37, 67-82. (22) Summers, D. P.; Noveron, J.; Basa, R. C. B. Energy Transduction Inside of Amphiphilic Vesicles: Encapsulation of Photochemically Active Semiconducting Particles. Orig. Life Evol. Biosphere 2009, 39, 127–140.
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(23) Kirch, M.; Lehn, J. M.; Sauvage, J. P. Hydrogen generation by visible light irradiation of aqueous solutions of metal complexes. An approach to the photochemical conversion and storage of solar energy. Helvetica Chimica Acta 1979, 62, 1345-1384. (24) Bojarska, E.; Pawlicki, K.; Czochralska, B. Photocatalytic Reduction of Nicotinamide Coenzymes in the presence of Titanium Dioxide: The Influence of Aliphatic Amino acids. Journal of Photochemistry and Photobiology A 1997, 108, 207-213. (25) Cuendet, P.; Grätzel, M. Photoreduction of NAD+ to NADH in Semiconductor Dispersions. Photochem. Photobiol. 1984, 39, 609-612. (26) Hargreaves, W. R.; Deamer, D. W. Liposomes from ionic, single-chain amphiphiles. Biochemistry 1978, 17, 3759-3768. (27) Deamer, D. W.; Barchfield, G. L. Encapsulation of Macromolecules by Lipid Vesicles under Simulated Prebiotic Conditions. Journal of Molecular Evolution 1982, 18, 203-206. (28) Paula, S.; Volkov, A.; Van Hoek, A.; Haines, T.; Deamer, D. W. Permeation of protons, potassium ions, and small polar molecules through phospholipid bilayers as a function of membrane thickness. Biophysical Journal 1996, 70, 339-348. (29) Nishizawa, M.; Suzuki, T.; Sprouse, S.; Watts, R.; Ford, P. Ligand steric effects on the photophysics of bis-and tris (2, 2'-bipyridine) complexes of rhodium (III). Inorganic Chemistry 1984, 23, 1837-1841. (30) Cantet, J.; Bergel, A.; Comtat, M. Bioelectrocatalysis of NAD+ reduction. Bioelectrochemisry and Bioenergetics 1992, 27, 475-486. (31) Berger, N.; Sachse, A.; Bender, J.; Schubert, R.; Brandl, M. Filter extrusion of liposomes using different devices: comparison of liposome size, encapsulation efficiency, and process characteristics. International journal of pharmaceutics 2001, 223, 55-68.
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Figure 1. Schematic outline of chemistry encapsulated in unilamellar and multilamellar vesicles. 104x132mm (300 x 300 DPI)
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Figure 2. Emission spectra of NADH generated by the Rh(bipyridine)3+3 mediated photoreduction of NAD+ by TiO2 particles encapsulated in phosphatidylcholine vesicles. 83x84mm (300 x 300 DPI)
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Figure 3. Enzymatic consumption of NADH by Lactate Dehydrogenase and pyruvate after removal from phosphatidylcholine vesicles by lipid extraction. Pyruvate (0.2 M) and enzyme (25 units) were added and the fluorescence was taken as a function of time. 82x82mm (300 x 300 DPI)
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Figure 4. Micrograph of the fluorescence of vesicles generated by the Rh(bipyridine)3+3 mediated photoreduction of NAD+ by TiO2 particles encapsulated in phosphatidylcholine vesicles. 96x111mm (300 x 300 DPI)
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Figure 5. Emission spectra of NADH generated by the Rh(bipyridine)3+3 mediate photoreduction of NAD+ by TiO2 particles encapsulated in oleic acid/dodecanol vesicles after removal from vesicles by sonication. 77x73mm (300 x 300 DPI)
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Figure 6. Enzymatic consumption of NADH by Lactate Dehydrogenase and pyruvate after removal from fatty acid vesicles by sonication. Pyruvate (0.2 M) and enzyme (20 units) were added and the fluorescence was taken after 1 hr. 81x80mm (300 x 300 DPI)
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299x140mm (180 x 180 DPI)
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