Chemical Transformations in Individual Ultrasmall Biomimetic Containers

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Science  19 Mar 1999:
Vol. 283, Issue 5409, pp. 1892-1895
DOI: 10.1126/science.283.5409.1892


Individual phospholipid vesicles, 1 to 5 micrometers in diameter, containing a single reagent or a complete reaction system, were immobilized with an infrared laser optical trap or by adhesion to modified borosilicate glass surfaces. Chemical transformations were initiated either by electroporation or by electrofusion, in each case through application of a short (10-microsecond), intense (20 to 50 kilovolts per centimeter) electric pulse delivered across ultramicroelectrodes. Product formation was monitored by far-field laser fluorescence microscopy. The ultrasmall characteristic of this reaction volume led to rapid diffusional mixing that permits the study of fast chemical kinetics. This technique is also well suited for the study of reaction dynamics of biological molecules within lipid-enclosed nanoenvironments that mimic cell membranes.

Living systems usually carry out biochemical transformations within cellular compartments defined by a phospholipid bilayer boundary. At such small dimensions [zeptoliters (10−21 liters) to femtoliters (10−15liters)], the surface-to-volume ratio is very high and the contained molecules experience collisions with the phospholipid surface at high frequencies. A hard-sphere approximation and a simulation of Brownian motion indicate that, in a 170-nm-diameter vesicle, a single enzyme and a single substrate collide at a frequency of 300 kHz, which might be compared with the substrate-wall collisional frequency of 200 MHz (1). Thus, the biochemical reactivity of the contained molecules can be dominated by surface interactions, and such interactions can profoundly influence enzyme kinetics (2). A tool to study confined chemical reactions under biologically relevant conditions would offer valuable insights into in vivo reaction conditions.

Various approaches exist for carrying out chemical reactions in aqueous solutions at small dimensions (3–7). Most open volume methods involve micromachining techniques (4) by which nanoliter to femtoliter wells can be created in silicon-based substrates (5). For self-enclosed volume elements, microdroplets in an immiscible solvent have been used (6). However, these techniques do not create the biologically relevant nanoenvironment that can be achieved in lipid vesicles.

From the perspective of chemical kinetics (7, 8), this method also offers the ability to achieve rapid diffusional mixing (microsecond to millisecond time range), and it opens the opportunity to study fast chemical reaction kinetics that are inaccessible to traditional bulk turbulence mixing techniques. The diffusive mixing time of dye molecules is about 8 ms for a 2-μm vesicle and 20 μs for a 0.1-μm vesicle. The small volume element contained within individual vesicles also matches the probe volume dimension of many optical single-molecule detection and manipulation schemes (9).

Using a recently developed rotaevaporative technique, we prepared unilamellar vesicles between 50 nm and 50 μm in diameter from a wide range of different phospholipids (10). The vesicles can encapsulate one or more reagent molecules of choice. Purified vesicle preparations are transferred to a bare or poly-l-lysine–coated borosilicate cover slip mounted on the stage of an inverted fluorescence microscope. Attoliter to zeptoliter vesicles can be precisely positioned and manipulated in solution by optical trapping (11). We used micromanipulator-controlled ultramicroelectrodes for electroporation and electrofusion, an infrared laser for optical trapping, two lasers (Ar+ and HeNe) for excitation, and two independent detection channels (Fig. 1). Optical damage to the vesicles has been observed at high laser powers and short wavelengths (ultraviolet). The power used for laser-induced fluorescence, however, is not sufficient to induce membrane damage, especially in the visible region where the lipid bilayer lacks absorption features. The membrane integrity of vesicles is also very sensitive to buffer conditions, such as ionic strength, pH, and type of buffer used.

Figure 1

Schematic of the experimental setup. Two carbon-fiber electrodes (5 μm in diameter) controlled by micromanipulators (MM) were used for electroporation and electrofusion. Three colinear laser beams were sent into a ×100, 1.3 numerical aperture microscope objective (MO). The 488-nm output of an argon ion laser (blue arrows) causes excitation of carboxyrhodamine-6G, Ca2+- chelated fluo-3, and fluorescein. The 633-nm output of a HeNe laser (orange arrows) causes excitation of TOTO-3–intercalated DNA. The 992-nm output of a MOPA diode laser (black arrows) is used for optical trapping of vesicles. The resulting dye fluorescence (green and red arrows) is collected, separated by a dichroic beamsplitter (DC) into two different color channels, passed through a bandpass filter (F1 or F2), and then detected by one of two CCD cameras. This setup is also modified to operate in a confocal mode with a single-photon counting silicon avalanche diode detector. MI, mirror; PB, polychroic beamsplitter; SD, spinning disk.

Figure 2 illustrates the hydrolysis of fluorescein diphosphate catalyzed by alkaline phosphatase. Fluorescein dianion product inside an optically trapped 3-μm vesicle was monitored by confocal fluorescence microscopy (9) at 60-s intervals. The time zero of the reaction for each interval is set by bleaching any products present. A nondestructive alternative to photobleaching is to monitor the product buildup over time. The small dynamic range of our detection scheme, however, prevented the use of this approach. Fewer than 100 product chromophores were produced between the time of bleaching and detection. With further optimization, formation of single product chromophores could be followed in real time—for example, by using fluorescence detection.

Figure 2

Fluorescence intensity versus time as a measure of alkaline phosphatase (AP) catalytic activity inside an optically trapped 3-μm vesicle containing AP and fluorescein diphosphate (0.75 mM). At 60-s intervals, the amount of fluorescent product accumulated after bleaching at the same wavelength is probed at 488 nm. The bleaching resets the reaction clock for each run. The peak values for each run were fitted to an exponential function, y =A exp(−kx), with A = 1118.6,k = 0.198, and a correlation coefficient of 0.98.

Because of the small volume element in a vesicle, the initial number of substrate molecules is limited and typically becomes largely depleted during the time required to prepare the vesicle. One way to overcome this drawback is to use the lipid bilayer as a partition between the reactants. The reaction can then be initiated by breaking down the bilayer through electric field–mediated membrane pore formation or membrane fusion.

Pore formation in lipid bilayers typically occurs within 50 μs (12, 13) and is controlled by the applied electric field strength and pulse duration; resealing of the membrane occurs in microseconds and sometimes longer (12, 14). To electroporate reagents into individual reagent-encapsulated vesicles and to perform electrofusion on single vesicle partners, we developed a miniaturized version of electroporation and electrofusion for individual vesicles. From whole-vesicle patch clamp measurements, we found that pore formation after electroporation was rapid. The inward current was observed to reach its half-maximal value after 276 ± 31 μs. The shape of the curve (Fig. 3) shows striking similarities to studies on irreversible electrical breakdown of unilamellar planar lipid films (15). In addition, when phosphatidylcholine (PC) vesicles containing 50 μM fluorescein and 150 mM NaCl (pH 7.2) surrounded by a citrate buffer (pH 4.3) were electroporated, we found a similar temporal dependence for quenching of fluorescence from intravesicular fluorescein (Fig. 3). During the pore-opening time, H+ ions diffuse into the liposome and protonate the fluorescein dianions, causing fluorescence quenching. Control experiments, in which the pH was adjusted to 8 both inside and outside the vesicle, did not result in any decreased fluorescence during electroporation (Fig. 3, insets). To drive the selective influx of molecules into the vesicles, we can exploit both concentration gradients and size differences between the molecules inside and outside the vesicle. Much faster pore-opening kinetics has been achieved in some lipid systems, which could enable reactions to be studied on low-microsecond time scales (12–14). In addition, vesicles with dimensions of tens of nanometers can be prepared, which also opens the possibility of low-microsecond diffusive mixing times.

Figure 3

Electroporation of single vesicles. Electric field–induced (20 V, 10 μs) membrane breakdown is measured by using patch clamp recordings (open circles), which are fit to a function describing a statistical sigmoid. The time needed for the current to reach its half-maximum value is 276 ± 31 μs (N = 6). Vesicles (10 μm in diameter) made from 74% PC, 19% phosphatidic acid, 7% cholesterol are patched in the whole-vesicle configuration and clamped to a holding potential of −10 to −30 mV. Ultramicroelectrodes are placed on opposite sides of the vesicle at an interelectrode distance of about 35 μm. Bath and pipette solutions are 140 mM NaCl and 10 mM Hepes (pH 7.4). Patch clamp data are sampled and stored on videotape (digitized at 20 kHz, filter frequency 10 kHz) and analyzed without additional filtering of the signal. In the fluorescence measurements (solid squares), 1- to 10-μm-diameter PC vesicles containing 50 μM fluorescein (pK a6.5, where K a is the acid constant) in 150 mM NaCl (pH 7.2) are electroporated (25 V, 10 μs duration) in a citrate buffer (pH 4.3). The start pulse from the multichannel scalar electronically triggers and initiates electroporation, which helps us define time zero of the experiment. The signal was detected with a single photon-counting module (50-μs dwell time per channel), filtered by using a five-point Savitsky–Golay smoothing algorithm, and normalized. The time needed for the fluorescence to reach its half-maximum value was determined to be about 184 μs. The figure shows a sigmoidal fit of the decrease in fluorescence intensity from the electroporated vesicles (average of four experiments). The fluorescence and patch clamp curves were centered at t = 1.0 ms where the sigmoidal curves reached 50% of their maximal values. (Inset) Full-trace scans from electroporation (25 V, 10-μs duration) of a fluorescein-containing vesicle (pH 8 inside) in an acidic solution (upper) (600-μs dwell time per channel) and in a solution buffered at the same pH (lower) (800-ms dwell time per channel). In the upper trace the fluorescence decreases because of the inflow of protons from the surrounding solution after electroporation, as indicated by arrows. No such events can be detected in the lower trace after electroporation (arrows).

Electroporation is useful in initiating reactions with sharp time distributions, but it lacks the capability to control precisely the amount of reagents delivered. In contrast, electrofusion between two select vesicles can be used to mix precise amounts of reagents. Optical traps (11) were used to align two selected vesicles for fusion. Fusion, induced by the reversible dielectric breakdown of the bilayer membrane (15–18), is achieved by application of a short, intense, and highly directed electric field generated across a pair of carbon-fiber ultramicroelectrodes (1- to 5-μm tip diameter). Pulses with a duration of 10 to 30 μs and an electric field strength of 20 to 50 kV/cm provided a near unity fusion yield. Figure 4A is a frequency histogram showing the number of pulses required to achieve fusion in 28 separate trials. About 14% of the fusions occurred after only one pulse and >70% of fusions occurred within five pulses (19).

Figure 4

(A) Histogram of the number of pulses required to achieve fusion (28 trials). The electric-field pulse is rectangular with a field strength of 40 kV/cm and a duration of 30 μs. Pulses were applied to PC vesicles containing fluorescein (0.1 mM) in a 10 mM Hepes buffer with 140 mM NaCl (pH 8). (B) Fluorescence recovery versus number of pulses applied. Data with error bars are mean values ± standard deviations (n = 3 to 5) and were fitted to a single exponential decay plotted for the entire data set. Data without error bars are single measurements. The fluorescence recovery is calculated as fluorescence intensity ratio of the product liposome to the arithmetic sum of the vesicles before fusion. (Insets) Images before and after fusion taken in fluorescence (488-nm excitation).

The initial stages of electrofusion are similar to electroporation. Fusion pores are being generated in the process. To determine whether leakage limits the utility of this technique, we encapsulated fluorescent molecules into the vesicles and monitored the fluorescence of these molecules before and after fusion under low-intensity excitation to minimize photoinduced bleaching of the chromophores. The fluorescence recovery as a function of the number of pulses is plotted (Fig. 4B), and the fluorescence images of two vesicles before and after fusion are shown (Fig. 4B, insets). On average, more than 94% of the fluorescence remained for one-pulse fusions (see also Fig. 4B, insets). Even for fusions that required more than 10 pulses, the vesicle still retained 60% or more of its fluorescence. We conclude that pores are formed when the two vesicles are fused, but the pores open and close too rapidly for any substantial amount of dye molecules to escape.

To demonstrate mixing after fusion, we fused two vesicles, each containing a different fluorescent dye (Fig. 5, A to D). To perform two-color mixing in fluorescence, we simultaneously excited the contents of the fused vesicle with two different wavelengths. The emitted photons were collected and separated into their respective color channels and then detected by two charge-coupled device (CCD) cameras (see Fig 1). Figure 5 shows the fusion between a vesicle containing carboxyrhodamine-6G (green fluorescence) and one containing TOTO-3-intercalated 15-mer DNA (red fluorescence) to yield a fused vesicle that appears orange.

Figure 5

(A to D) Fluorescence color mixing. Fusion of two vesicles, each containing a dye whose fluorescence is at a different color: 20 μM carboxyrhodamine-6G (top vesicle) and 20 μM TOTO-3–intercalated 15-mer DNA (bottom vesicle). (A) and (C) are bright-field images taken before and after fusion; (B) and (D) are the corresponding fluorescence images, obtained as described in Fig. 1. The buffer and pulse conditions were similar to those in Fig. 4. Images before (E) and after (G) electrofusion (about 75 kV/cm; 10 μs) of a 10 μM fluo-3–containing vesicle (left) and a 10 μM Ca2+-containing vesicle (right) are shown under bright-field illumination in a buffer solution containing 10 mM Hepes and 140 mM NaCl (pH 7.4). Corresponding fluorescence images are shown in (F) and (H). The fluo-3 solution encapsulated in the liposomes and the extraliposomal solution were titrated with EGTA to reduce background fluorescence. The vesicles were initially immobilized on poly-l-lysine–coated borosilicate cover slips, followed by rinsing with Ca2+-free buffer solution.

Figure 5, E to H, shows a chemical reaction carried out inside these ultrasmall containers, one of which holds fluo-3 (10 μM) and the other Ca2+ (10 μM). Before fusion (Fig. 5G), no fluorescence was detected from the Ca2+-containing vesicle, but a small background fluorescence was observed in the vesicle with fluo-3. Binding of Ca2+ by fluo-3 increases the fluorescence quantum yield of this chelator by about 40-fold (20). This fluorescence enhancement was indeed observed, as demonstrated in Fig. 5H, which represents the image taken after the contents of the two vesicles were mixed and the chelating reaction was initiated. Although we focused on fluorogenic reactions and fluorescence quenching in our experiments, other optical techniques might also be used, including measurement of polarization, lifetime, and fluorescence energy transfer.

The technique we have described offers the special opportunity to probe the dynamics of chemical reactions in spatially confined biomimetic nanoenvironments. By systematically varying the membrane lipid, protein, and glycoprotein composition of a vesicle, the in vivo activity of multiple or single biological molecules might be inferred. In combination with single-molecule detection (9) and manipulation methodologies (11), single-molecule reactions can be studied within the lipid bilayer boundary of a vesicle. Because only one reaction is studied at a time, synchronization of the reactions is not necessary. Therefore, the observable chemical kinetics is limited only by the excited-state lifetime of the chromophore. The small volume characteristics of this technique should also find use as a general chemical or biochemical delivery system whose spatial and temporal locations can be precisely controlled.


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