Anal. Chem. 1997, 69, 4761-4767
Articles
Localized Sampling of Cytoplasm from Xenopus Oocytes for Capillary Electrophoresis Veronica Luzzi, Chao-Lin Lee, and Nancy L. Allbritton*
Department of Physiology and Biophysics, University of California, Irvine, California 92697-4560
Continued progress in cellular physiology requires new measurement strategies which can be applied to solitary cells. Since many cellular signaling pathways act on time scales of a few seconds, there is a critical need for singlecell techniques with subsecond time resolution. Capillary electrophoresis shows great promise as a tool for the analysis of individual cells. In the present work, we describe a technique to load a capillary with picoliter to nanoliter volumes of cytoplasm and initiate electrophoresis in less than 500 ms. When cytoplasm was sampled from a Xenopus laevis oocyte previously loaded with fluorescein, calcium green, or a mixture of the two fluorophores, their fluorescent peaks were readily identifiable on the electropherogram. Since the volume of cytoplasm (e30 nL) loaded into the capillary was much smaller than the l µL oocyte volume, spatially localized biochemical measurements were also possible. To demonstrate the utility of this new technique, the activity of the enzyme β-galactosidase was measured in small regions of the Xenopus oocyte. Subcellular, subsecond sampling of oocyte cytoplasm will enable biochemical measurements with the resolution required to understand many cellular signal transduction pathways. Since cells in a group respond asynchronously to external stimuli, single-cell measurements are becoming increasingly important in the study of biochemical pathways.1-6 The ability to perform rapid, subcellular measurements is opening new avenues of investigation into the dynamics of biological processes. Enhancing the spatial and temporal resolution of such measurements will improve our ability to understand the physiology of cells. The Xenopus laevis oocyte is an important model system in biology. Its large 1 mm diameter permits many single-cell biochemical measurements not possible with other cell types.7-10 Since the endogenous biochemical pathways of the oocyte are (1) Tsien, R. Y. Am. J. Physiol. 1992, 263, C723-C728. (2) Giuliano, K. A.; Post, P. L.; Hahn, K. M.; Taylor, D. L. Annu. Rev. Biophys. Biomol. Struct. 1995, 24, 405-434. (3) Kennedy, R. T.; Oates, M. D.; Cooper, B. R.; Nickerson, B.; Jorgenson, J. W. Science 1989, 246, 57-63. (4) Yeung, E. S. Acc. Chem. Res. 1994, 27, 409-414. (5) Ewing, A. G. J. Neurosci. Methods 1993, 48, 215-224. (6) Jankowski, J. A.; Tracht, S.; Sweedler, J. V. Trends Anal. Chem. 1995, 14, 170-176. (7) Nussberger, S.; Foret, F.; Hebert, S. C.; Karger, B. L.; Hediger, M. A. Biophys. J. 1996, 70, 998-1005. (8) Stith, B. J.; Goalstone, M.; Silva, S.; Jaynes, C. Mol. Biol. Cell. 1993, 4, 435-443. S0003-2700(97)00550-7 CCC: $14.00
© 1997 American Chemical Society
similar to those found in mammalian cells, the oocyte is utilized to study a multitude of signaling pathways, including those involving ion channels, the cell cycle, transporter proteins, and Ca2+ waves. The Xenopus oocyte is also a valuable tool to study the functions of foreign proteins, since the oocyte synthesizes large quantities of proteins from microinjected mRNA. Development of new strategies to perform biochemical measurements on the Xenopus oocyte and other single cells has the potential to contribute to the understanding of many different biochemical signaling pathways. Most biochemical measurements require the cytoplasm of an entire oocyte or several oocytes, so measurements are made without spatial resolution.7,8 Since biochemical reactions do not occur simultaneously throughout the oocyte, a spatially averaged measurement is obtained. These averaged determinations frequently do not accurately represent true events. Large spatial gradients in the concentrations of molecules and presumably the activities of enzymes can span distances of less than 1 µm to hundreds of micrometers in the Xenopus oocyte.9,10 To investigate the formation and function of these biochemical gradients, measurements must be performed on cytoplasmic volumes of nanoliters and less. These required small sample volumes suggest that capillary electrophoresis (CE) may be well suited for localized measurements in Xenopus oocytes. Olefirowicz and Ewing have pioneered the use of subcellular sampling techniques for CE.11,12 They measured the intracellular concentration of electroactive species, such as dopamine, in a subcellular region of the giant neuron of the snail Planorbis corneus. The tip of a capillary etched with hydrofluoric acid to a 6-10 µm outer diameter was inserted into a neuron, and ∼50100 pL of cytoplasm was electromigrated into the capillary for 5 s. This strategy is suitable for the measurement of biologic entities whose intracellular concentration or activity is constant during the 5 s of electromigration following penetration of the plasma membrane. More recently, Sweedler and colleagues have performed subcellular measurements on neurons from the sea slug Aplysia californica.6 An etched capillary tip was used to mechanically disrupt the cell body from its neuronal processes, after which the individual pieces of the cell could be sampled. This technique is appropriate for measurement of biologic analytes whose properties are not altered by fragmentation of the cell and (9) Lechleiter, J.; Girard, S.; Peralta, E.; Clapham, D. Science 1991, 252, 123126. (10) Busa, W. B.; Nuccitelli, R. J. Cell. Biol. 1985, 100, 1325-1329. (11) Olefirowicz, T. M.; Ewing, A. G. J. Neurosci. Methods 1990, 34, 11-15. (12) Olefirowicz, T. M.; Ewing, A. G. Anal. Chem. 1990, 62, 1872-1876.
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which remain unchanged during the time before final lysis of the cellular fragments. While the above strategies are applicable to the analysis of many cellular properties, the techniques do not have sufficient temporal resolution to measure the activity of proteins and the concentration of metabolites that change on time scales of a few seconds or less. During normal cellular functions, the activities of a wide range of enzymes and the concentrations of many important metabolites can be substantially altered within several seconds.13 Additionally, the properties of these analytes can be dramatically modified within 5 s after damage to the plasma membrane by insertion of a capillary or fragmentation of a cell. A breach in the plasma membrane creates a rapid influx of extracellular ions, such as Ca2+, activating many enzymes including kinases and proteases.13 To broaden the range of cellular proteins and metabolites measurable by CE, we sought to develop a subsecond sampling technique for spatially localized measurements in Xenopus oocytes. In the present work, we describe a technique to sample picoliter to nanoliter volumes of cytoplasm from an intact living Xenopus oocyte in less than 500 ms. Biochemical measurements can also be performed on cytoplasm obtained at varying locations and from different depths within the oocyte. With this methodology, we also demonstrate measurement of the activity of an enzyme, β-galactosidase, in Xenopus oocytes. Decreasing the time required to obtain cytoplasm will improve the temporal resolution of most cellular biochemical measurements employing CE. EXPERIMENTAL SECTION Materials. Calcium green 2 and fluorescein di-β-D-galactopyranoside were purchased from Molecular Probes, Eugene, OR. [R-32P]ATP (800 Ci/mmol, 10 mCi/mL) was obtained from New England Nuclear, Boston, MA. Agarose II, a low-melting-temperature agarose, was from Amresco, Solon, OH. All other reagents were purchased from Fisher Scientific, Pittsburgh, PA. Oocyte Isolation and Preparation. Stage V and VI oocytes were isolated from X. laevis frogs as described previously.14 The oocytes were cultured at room temperature in ND96 buffer solution [96 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES (pH 7.6), 2.5 mM sodium pyruvate] until used. The oocytes were visualized with a dissecting microscope (Olympus or Applied Fiberoptics, Inc.) for all manipulations. The membraneimpermeant reagents [R-32P]ATP (56 nM), calcium green 2 (150 µM), fluorescein (10-50 µM), and β-galactosidase (0.5 mg/mL) were microinjected into oocytes in 20-50 nL of buffer A [135 mM NaCl, 5 mM KCl, 2 mM MgCl2, 1 mM CaCl2, 10 mM HEPES (pH 7.4)]. After microinjection, the oocytes recovered at room temperature for a minimum of 1 h, which allowed the radioactive ATP, calcium green 2, or fluorescein to diffuse throughout the oocyte. For aspiration of oocyte cytoplasm into a capillary, a single oocyte was placed into a “V”-shaped well drilled into a block of Teflon or polycarbonate. The total volume of this chamber was variable, ranging from 50 µL to 5 mL. In some instances, a small amount of low-melting-temperature agarose (1.5%) was poured around the edges of the oocyte to hold it in position. Healthy oocytes can leak or extrude small amounts of the microinjected reagents into the surrounding media. These buffer contaminants can enter the capillary prior to insertion of the capillary into the (13) Barritt, G. J. Communication Within Animal Cells; Oxford University Press: New York, 1992. (14) Zagotta, W. N.; Hoshi, T.; Aldrich, R. W. Proc. Natl. Acad. Sci. U.S.A. 1989, 86, 7243-7.
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Figure 1. Schematic of the cytoplasmic sampling apparatus for X. laevis oocytes.
oocyte, confounding subsequent measurements.16 To eliminate this possibility, the oocytes were continuously washed in a slowly flowing buffer stream. Alternatively, the buffer surrounding the oocytes was removed just prior to cytoplasmic sampling. All experiments were performed on several different days. Expression of β-Galactosidase in Oocytes. A linearized SinRep/lac Z RNA template was used to produce capped mRNA encoding β-galactosidase with the InvitroScript CAP Kit (Invitrogen Corp., San Diego, CA). Oocytes were microinjected with β-galactosidase RNA (184 ng in 46 nL of buffer A) and incubated in the ND96 buffer solution for a minimum of 24 h. The oocytes were loaded with fluorescein di-β-D-galactopyranoside by incubation with 2 mM for 2 min.15 The oocytes were then prepared for cytoplasmic sampling as described above. Aspiration of [r-32P]ATP from a Buffer Solution or Oocyte. A capillary (50 µm i.d., 360 µm o.d., 57-90 cm long) was mounted on the edge of a 1.3 cm × 3.8 cm × 0.05 cm piezoelectric motor element (Piezo Systems, Cambridge, MA) with a custom-built clamp. A micromanipulator, which held the piezoelectric element, was used to position the capillary adjacent to an oocyte or a bead of a buffer solution. The tip of the capillary had previously been etched to a sharp tip with hydrofluoric acid.12 A “T” junction (Valco Instruments, Houston, TX) was spliced into the capillary 12 cm from the tip (Figure 1). The outlet of the capillary 45 cm from the T junction was placed in a buffer solution held at ground potential. A second capillary (50 µm i.d., 360 µm o.d., 5 cm long) was attached to the third outlet of the T junction and to a 2 cm length of Teflon tubing (0.38 mm i.d.). The tubing was cemented to the common outlet of a three-way valve (The Lee Co., Westbrook, CT). The normally open inlet of the valve was at atmospheric pressure, while the normally closed inlet was connected to a vacuum (709 or 284 mmHg). The capillary tip moved vertically downward (0-600 µm) following application of a voltage (-150 to +150 V) across the piezoelectric motor. The capillary tip moved 300 µm in less than 20 ms. The power supply was controlled remotely using the digital to analog channel of a data acquisition board (DAS-1802ST-DA or DAS-1802HR-DA, Keithly Metrabyte, Taunton, MA) in a personal computer (Gateway, North Sioux City, SD). Opening the valve between the capillary and the vacuum line applied a vacuum to the third port of the T junction. To remotely open the valve, a relay switch (ODC-05 output module, Metrabyte) interposed between the valve and a 12 V power supply was closed via a digital to analog output channel on the Metrabyte board. The response times of the relay switch (15) Lin, S.; Yang, S.; Hopkins, N. Dev. Biol. 1994, 161, 77-83. (16) Lee, C. L.; Allbritton, N. L. unpublished work, Univiversity of CaliforniasIrvine, 1997.
and valve were 750 µs and 10 ms, respectively. A digital to analog output channel was also used to turn on the electrophoretic voltage across the inlet and outlet ends of the capillary. To sample cytoplasm or buffer solution, a customized program (in Viewdac, Keithley Metrabyte) controlled the piezoelectric motor, the valve, and the time delays between piezoelectric motor movement and valve opening and closing. After positioning the capillary tip adjacent to the desired sampling location, a series of softwarecontrolled events were triggered: (i) movement of the piezoelectric motor downward driving the capillary into the oocyte or buffer, (ii) followed by a variable time delay, (iii) opening of the vacuum valve causing aspiration of cytoplasm or buffer into the capillary, (iv) closing of the vacuum valve, (v) another time delay, (vi) upward movement of the piezoelectric element withdrawing the capillary tip from the oocyte or buffer solution, (vii) a third time delay, and (viii) initiation of electrophoresis by application of a voltage across the capillary. The three time delays were each 100 ms unless stated otherwise. When electrophoresis was not performed, steps vii and viii were eliminated. Sampling of Cytoplasm from Oocytes Microinjected with Fluorescein, Calcium Green 2, or β-Galactosidase RNA. Cytoplasmic sampling from oocytes microinjected with fluorescein, calcium green 2, or β-galactosidase RNA was performed similarly to that of the oocytes loaded with [R-32P]ATP with two exceptions. The vacuum was not used to aspirate cytoplasm into the capillary, and the software was modified to reflect the absence of the vacuum. An etched capillary tip (50 or 20 µm i.d., 3-15 cm long) was attached to a second capillary (50 µm i.d., 360 µm o.d., 3380 cm long), utilizing the T junction or a union connector (Valco Instruments). The inner diameter of these connectors was 150 µm. For the experiments in Figure 4 only, a continuous capillary (50 µm i.d., 360 µm o.d., 57-90 cm long) with an etched tip was used. The software-controlled steps were as follows: (i) downward movement of the piezoelectric motor driving the capillary a variable distance into the oocyte, (ii) a 100 ms delay in which the capillary remained in the oocyte, (iii) upward movement of the piezoelectric motor, removing the capillary from the oocyte, (iv) another 100 ms delay, and (v) application of a voltage across the ends of the capillary initiating electrophoresis. Capillary Electrophoresis. Fused-silica capillaries were uncoated (Polymicro Technologies, Phoenix, AZ) or possessed a proprietary neutral coating (Supelco, Bellafonte, PA). With the exception of [R-32P]ATP, analytes contained in buffer solutions were loaded into capillaries by gravitational fluid flow, and the loaded volumes calculated from Poiseulle’s equation.17 Electrophoresis was performed in buffer A. For electrophoresis, the outlet reservoir was maintained at ground potential while the inlet reservoir was held between -6 and -15 kV with currents ranging from 50 to 100 µA. After electrophoresis of cytoplasm, uncoated capillaries were washed extensively with water or 0.1 N NaOH/ water/0.1 N HCl, respectively. The number of theoretical plates (N) was calculated as described previously.17 Since substantial peak tailing occurred, the width of the peak at 10% of its height was used rather than the peak width at the baseline. Fluorescence Detection. The fluorescence of analytes electrophoresed through the capillary was detected in one of two ways. In the first, an optical window in the capillary was created by burning off a short length of the polyimide coating 50 cm from
the inlet. The capillary was interrogated at the optical window by the focused laser beam of an argon ion laser (488 nm, Coherent, Santa Clara, CA). Fluorescence was collected at a right angle to the capillary and laser beam with a collecting lens, and the light was measured with a photomultiplier tube (PMT, R928, Hamamatsu, Bridgewater, NJ) after spectral filtering (533DF56, Omega Optical, Brattleboro, VT). Alternatively, the fluorescence of analytes was measured using a fluorescent microscope. The outlet end of a capillary was positioned at a right angle to a cover slip mounted on the stage of an inverted microscope equipped with a xenon lamp (Diaphot 200, Nikon, Melville, NY). A Teflon ring cemented to the coverslip held the electrophoretic buffer solution and ground electrode. The fluorescence of analytes was measured with a 100× oil immersion lens as the compounds eluted from the capillary. A standard fluorescein filter set (Chroma Technology Corp., Brattleboro, VT and Omega Optical) was used for the excitation and emission of fluorescein and calcium green 2. Fluorescence was collected with a photomicrographic attachment (PFX, Nikon) equipped with a PMT (R928, Hamamatsu, San Jose, CA). For both measurement strategies, the PMT current was amplified and converted to a voltage with a preamplifier (Model 1212, DL Instruments, Dryden, NY) or a custom-built preamplifier based on an operational amplifier (LF355N, Motorola, Tempe, AZ).18 The signal was digitized by a data acquisition board (DAS-1802ST-DA or DAS-1802HR-DA, Keithley Metrabyte, Taunton, MA) in a personal computer (Gateway, Sioux City, SD). The data were plotted and peak areas calculated using Origin (Microcal, Northhampton, MA). Calibration of Volumes Sampled from Oocytes Loaded with Fluorescein. To estimate the volume of cytoplasm obtained from oocytes containing fluorescein, cytoplasm was sampled from oocytes loaded with [R-32P]ATP or fluorescein without application of a vacuum. A capillary was inserted 270 µm for 100 ms into an oocyte containing [R-32P]ATP and the volume of cytoplasm introduced into the capillary was measured. Cytoplasm was then obtained from fluorescein-loaded oocytes in the same manner, and the peak area of the fluorescein measured during electrophoresis. The average volume obtained from the measurements with the [R-32P]ATP (n ) 4) was equated with the average area of the fluorescein peaks (n ) 4). Since the cytoplasmic samples were obtained with identical technique, their average volume was the same. The volume of subsequent cytoplasmic samples was calculated by comparing the area of the fluorescein peak to that of the calibration point.
(17) Weinberger, R. Practical Capillary Electrophoresis; Academic Press: San Diego, 1993.
(18) Horowitz, P.; Hill, W. The Art of Electronics; Cambridge University Press: New York, 1990.
Table 1. Typical Time Scales for Cellular Events cellular event DNA replication protein synthesis protein phosphorylation ion channel opening
time scale13,19a hours minutes seconds milliseconds
a Defined as the time required for the biochemical event to occur to a degree significant enough to alter cellular physiology or behavior. As an example, the time scale for protein phosphorylation is the typical time needed to phosphorylate a substantial percentage of a protein in vivo, not the time required to place a phosphate on a single residue of a protein.
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RESULTS AND DISCUSSION In the course of normal biologic responses, the properties of a cell change on many different time scales (Table 1). A biological value such as a concentration or an activity may change substantially during the time required to measure it. To accurately measure many biochemical processes, the temporal resolution of the measurement must be less than the time it takes for the measured value to change significantly. For biological entities that change on a time scale of a second, the temporal resolution of the measurement technique must be subsecond. For analysis of intracellular species by CE, the temporal resolution of the biologic measurement is determined by the time between the initiation of cytoplasmic sampling and the termination of cellular reactions with the molecule of interest. Termination of the cellular reactions is achieved by separation of the reactants, usually by diffusion and electrophoresis, or by inactivation of reactants, typically by denaturation of proteins.11,12,20-26 Decreasing the time required to obtain cellular cytoplasm and diminishing the delay between sampling and the initiation of electrophoresis can, therefore, enhance the time resolution of subcellular measurements by CE. To improve the time resolution of biochemical analyses on oocytes, we automated each step of the sampling process and used a vacuum rather than electromigration to load the capillary with cytoplasm. Aspiration of Nanoliter-Sized Volumes from a Buffer Solution. To determine the range of sample volumes and the reproducibility of the sampling apparatus, a buffer solution containing [R-32P]ATP was aspirated into the capillary. Application of the vacuum to the side port of the T junction was regulated by a computer-controlled valve (Figure 1). The sharpened tip of a capillary was placed just above, but not in contact with, the radioactive solution. The small bead of radioactive solution and the fluid level in the capillary outlet reservoir were at the same height. To introduce the sample, the capillary was lowered 300 µm into the buffer by the piezoelectric motor. After a 100 ms delay, the valve to the vacuum line was opened for varying lengths of time. After closure of the valve and a subsequent 100 ms delay, the capillary was removed from the radioactive solution by the piezoelectric motor. The radioactivity in the tip was measured to determine the volume of fluid sampled. Increasing the applied vacuum time from 125-1000 ms increased the volume aspirated into the capillary from 6 to 108 nL (Figure 2). Decreasing the vacuum strength decreased the aspirated volume (data not shown). When a vacuum was not applied to the side port of the T junction, the volume of radioactive solution introduced into the capillary was too small to quantitate. Aspiration of Nanoliter-Sized Volumes from Oocytes. To determine whether cytoplasm could be obtained from intact living X. laevis oocytes, the aspiration apparatus was used to withdraw cytoplasm from oocytes previously microinjected with [R-32P]ATP. The etched tip of the capillary was placed in contact with, but did (19) Lodish, H.,; Baltimore, D.; Berk, A.; Zipursky, S. L.; Matsudaira, P.; Darnell, J. Molecular Cell Biology; W. H. Freeman and Co.: New York, 1995. (20) Orwar, O.; Fishman, H. A.; Ziv, N. E; Scheller, R. H.; Zare, R. N. Anal. Chem. 1995, 67, 4261-4268. (21) Xue, Q.; Yeung, E. S. Anal. Chem. 1994, 66, 1175-1178. (22) Oates, M. D.; Cooper, B. R.; Jorgenson, J. W. Anal. Chem. 1990, 62, 15731577. (23) Gilman, S. D.; Ewing, A. G. J. Capillary Electrophor. 1995, 2, 1-13. (24) Lee, T. T.; Yeung, E. S. Anal. Chem. 1992, 64, 3045-3051. (25) Zhang, R.; Cooper, C. L.; Ma, Y. Anal. Chem. 1993, 65, 704-706. (26) Rosser, B. G.; Powers, S. P.; Gores, G. J. J. Biol. Chem. 1993, 268, 2359323600.
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Figure 2. Aspiration from a buffer solution into a capillary. The valve to the vacuum line was opened for the times displayed on the logarithmic x axis. The volume of buffer drawn into the capillary is displayed on the y axis. The error bars represent one standard deviation. The number of data points at each time is 9 (125 ms), 11 (250 ms), 5 (500 ms), and 4 (1000 ms).
Figure 3. Aspiration of cytoplasm from a Xenopus oocyte. The time that the valve to the vacuum line was open and the volume of cytoplasm that was drawn into the capillary are displayed on a semilogarithmic plot. The vacuum strength was 709 (triangles) or 354 mmHg (circles). The error bars represent one standard deviation. The number of data points at each time is 3 (100 ms), 5 (125 ms), 9 (250 ms), and 4 (500 ms) for 709 mmHg and 3 (250 ms), and 3 (500 ms) for 354 mmHg.
not breach, the oocyte’s plasma membrane. To obtain cytoplasm, the capillary was driven 150 µm into the oocyte followed by aspiration for varying lengths of time. Other parameters of the experiment were identical to those utilizing the [R-32P]ATP in a buffer solution. Aspiration for 100 ms loaded 3 nL of cytoplasm into the capillary, while 500 ms of applied vacuum aspirated 27 nL (Figure 3). Due to the greater viscosity of cytoplasm compared to buffer solution, longer times were required to aspirate equal volumes of cytoplasm. The volume of cytoplasm loaded into the capillary plateaued at ∼30 nL as the applied vacuum time was increased beyond 500 ms. At a 30 nL volume, the column of cytoplasm was ∼1.5 cm in length. At this plug length, the increased force required to move the viscous cytoplasm through the column exceeded that of the vacuum and little additional cytoplasm could be loaded into the capillary. A stronger vacuum is expected to load larger volumes of cytoplasm. A weaker vacuum loaded smaller volumes of cytoplasm (Figure 3). Application of the reduced vacuum force for 100 ms aspirated 0.7 nL into the capillary. In this range of sample volumes, decreasing
Figure 5. Illustration of the distances that determine the spatial resolution in the z (A) and x-y (B) dimensions. In (A), cytoplasm enters the capillary as the capillary moves parallel to the z axis into the oocyte. When a vacuum is applied to the capillary, additional cytoplasm enters the capillary from an approximately spherical region in the oocyte. The final depth of the capillary in the oocyte, d, and the diameter of the sampled spherical region, s, determine the spatial resolution along the z axis. As shown in (B) the x-y spatial resolution is determined from s and the inner diameter of the capillary, c.
Figure 4. Volume of cytoplasm loaded into the capillary dependent on its depth in the oocyte. (A) Electropherograms of cytoplasm obtained after inserting the capillary 270, 150, or 60 µm deep into oocytes previously microinjected with fluorescein. Electrophoresis was performed in a 90 cm long capillary with a neutral surface coating. The fluorescence detection window was 50 cm from the inlet end and the electrophoretic voltage was -15 kV (current, ∼60 µA). (B) Capillaries were inserted varying depths into oocytes, the sample electrophoresed, and the volume of cytoplasm calculated from the area of the fluorescein peak. The error bars represent one standard deviation. The number of data points at each depth is 5 (60 µm), 5 (150 µm), and 4 (270 µm).
the initial 100 ms delay to 1 ms did not alter the volume sampled. The initial time delay occurred between insertion of the capillary tip into the oocyte and opening of the vacuum valve. In contrast, decreasing the second time delay to 10 ms or less, decreased the volume of aspirated solution. This delay time, which occurred between closure of the vacuum valve and removal of the capillary from the oocyte, was required to fully exhaust the vacuum contained in the capillary. During delay times of 10 ms or less, the capillary was withdrawn from the oocyte prior to full dissipation of the vacuum, decreasing the volume aspirated into the capillary. Procurement of Picoliter-Sized Volumes from Oocytes. Fluorescein instead of [R-32P]ATP was microinjected into oocytes for aspiration of volumes less than 500 pL since these volumes contained insufficient amounts of radioactivity for measurement. After introduction of cytoplasm into the capillary and removal of the capillary from the oocyte, electrophoresis was initiated. A
single fluorescent peak was present on the electropherogram, and its migration time was similar to that of fluorescein in a buffer solution (Figure 4A). In this subnanoliter range, the quantity of material introduced into the capillary was influenced not only by the duration and strength of the applied vacuum but also by the depth that the capillary was inserted into the oocyte. In the absence of a vacuum, inserting the capillary 270 µm into the oocyte loaded 500 pL of cytoplasm into the capillary (Figure 4). Decreasing the capillary depth to 60 µm decreased the volume to 22 pL. Decreasing the inner diameter of the capillary tip from 50 to 20 µm also decreased the volume of cytoplasm sampled (data not shown). These results are consistent with cytoplasmic sampling by coring of the oocyte with the capillary: the sample represents a cylinder of cytoplasm extending from the plasma membrane to the deepest point of the capillary in the oocyte. The capillary could not be inserted less than 60 µm into a normal oocyte due to the elasticity of the outer cell membrane. However, shallower capillary depths and smaller sampling volumes were achieved by removing the oocyte’s vitelline membrane, which eliminates the pliability of the outer membrane (data not shown). Estimated Spatial Resolution of the Sampling Technique. The spatial resolution of the sampling technique can be estimated from the volume of cytoplasm sampled, the inner diameter of the capillary, and the depth of insertion of the capillary into the oocyte. The volume of sampled cytoplasm is determined by the coring action of the capillary as it enters the oocyte and by the vacuummediated suction of cytoplasm into the capillary (Figure 5). For the 22 pL volume obtained by inserting the capillary (50 µm i.d.) a depth of 60 µm into the oocyte (see Figure 4B), the estimated spatial resolutions are 60 (z axis) and 50 µm (x and y axes) (Figure 5). The small size of the lumen and shallow depth of the capillary relative to the dimensions of the Xenopus oocyte, and the ability of CE to efficiently transport and separate small sample volumes, enable measurements with subcellular resolution. Separation of Fluorescein and Calcium Green 2. To determine whether molecules contained in the cytoplasm were separated during electrophoresis, we loaded oocytes with fluorescein, calcium green 2 (a fluorophore derived from fluorescein), or a mixture of fluorescein and calcium green 2. The capillary was inserted 60 µm into the oocyte for 100 ms without application Analytical Chemistry, Vol. 69, No. 23, December 1, 1997
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Figure 6. Separation of fluorescein and calcium green 2 dissolved in a buffer solution (A) or oocyte cytoplasm (B, C). Electrophoresis was performed in a 41 cm long capillary with a neutral surface coating and the T junction placed 3 cm from the etched tip of the capillary. Fluorescence was measured at the outlet of the capillary with a microscope as described in the Experimental Section. The electrophoretic voltage was -8 kV (current, ∼90 µA). (A) Shown are electropherograms of solutions of fluorescein (1 nL, 10 nM, solid line) or calcium green 2 (1 nL, 30 nM, dashed line) in buffer A. (B, C) Shown are electropherograms of fluorescein (B, solid line), calcium green 2 (B, dashed line), or a mixture of fluorescein and calcium green 2 (C) obtained from oocytes. A 50 nL volume of fluorescein (50 µM), calcium green 2 (150 µM), or a mixture of fluorescein and calcium green 2 (50 and 150 µM, respectively) was microinjected into an oocyte. The capillary was loaded with ∼30 pL of cytoplasm from the oocyte and electrophoresis initiated as described in (A). A different oocyte was used for each trace in (B) and (C). The difference in height and area of the fluorophore peaks is most likely due to variations in the oocytes’ ability to transport the dyes from the cytosol to the extracellular space. 4766
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of a vacuum. When cytoplasm containing only fluorescein or calcium green 2 was electrophoresed (Figure 6B) a single peak was present on the electropherogram with a migration time similar to that of the corresponding fluorophore loaded from a buffer solution (Figure 6A). The mobility of fluorescein and calcium green 2 was altered only slightly when the two fluorophores were introduced into the capillary in the viscous and highly structured cytoplasm. Electropherograms of cytoplasm containing a mixture of fluorescein and calcium green 2 revealed two fluorescent peaks with migration times identical to that of fluorescein alone and calcium green 2 alone in cytoplasm (Figure 6 B,C). These results also suggest that the fluorophores are rapidly separated from the cytoplasmic plug after the onset of electrophoresis. The high molecular weight, cytosolic proteins, membranes, and organelles are expected to migrate much slower than either fluorescein or calcium green 2. Molecules that bind tightly to cytoplasmic proteins or structures may exhibit migration times in the presence of cytoplasm that are substantially different from that in buffer alone. This was the case for some molecules, particularly those with a net positive charge (data not shown). Fluorescein and calcium green 2 migrated closer together when loaded in cytoplasm rather than buffer, demonstrating that the presence of cytoplasm decreased the resolution. The calculated number of theoretical plates (N)17 decreased in the presence of cytoplasm. When the T junction was not spliced into the capillary, N was 26 900 + 800 (n ) 7) for fluorescein loaded from a buffer solution and 19 700 + 800 (n ) 7) for fluorescein sampled from an oocyte. The effect of the T junction on the separation efficiency was much greater than that of cytoplasm. When the T junction was spliced into the capillary, N was 1920 + 660 (n ) 6) for fluorescein loaded from a buffer solution and 2040 + 300 (n ) 7) for fluorescein sampled from an oocyte (Figure 6 B,C). The T junction had a much larger inner diameter (150 µm) than that of the capillary (50 µm); consequently, the dead volume in the connector was quite large. Use of a smaller inner diameter connector with zero dead volume might greatly increase N when the T junction is present. Measurement of β-Galactosidase Activity in Oocytes. Measurement of β-galactosidase activity is used throughout the biologic sciences to determine how gene transcription and RNA translation are regulated. To illustrate the utility of the new sampling technique, the activity of the enzyme β-galactosidase was measured in oocytes. RNA encoding the β-galactosidase protein was microinjected into oocytes. After 24 h allotted for protein expression, the oocytes were loaded with the nonfluorescent, fluorescein di-β-D-galactopyranoside, which is cleaved by β-galactosidase to yield fluorescein. After varying times, cytoplasm from an oocyte was introduced into the capillary and electrophoresed as described for the fluorescein/calcium green 2 experiments (Figure 7). The concentration of fluorescein in the oocytes increased linearly over time at a rate of ∼3.8 fmol/min when 184 ng of RNA was injected into each oocyte. Capillary electrophoresis in conjunction with fluorescence detection can be used to measure the change in β-galactosidase activity over time in a single oocyte or the relative β-galactosidase activity of a set of oocytes. CONCLUSION We have developed an apparatus to rapidly obtain cytoplasm from small regions of Xenopus oocytes. Picoliter to nanoliter quantities of cytoplasm can be obtained from an intact, living
Figure 7. Measurement of β-galactosidase activity in oocytes microinjected with β-galactosidase RNA. At 3, 6, 12, and 24 min after loading fluorescein di-β-D-galactopyranoside into oocytes, ∼100 pL of cytoplasm was sampled and electrophoresed. Shown is the concentration of fluorescein in the oocyte at the different times. The number of moles of fluorescein in the sampled cytoplasm was determined by comparing the peak area of the sample from the oocyte to the peak area of electrophoresed, fluorescein standards. The concentration of fluorescein in the oocyte was then calculated from the number of moles of fluorescein in the sample and the sample volume. Cytoplasm was obtained from five different oocytes at 3 and at 6 min while four different oocytes were used at 12 and at 24 min. The error bars represent one standard deviation. Electrophoresis was performed in a 36 cm long capillary with a neutral surface coating. The electrophoretic voltage was -6.5 kV (current, ∼60 µA).
oocyte in less than 1 s. Furthermore, by combining this sampling strategy with capillary electrophoresis, analytes from within the oocyte can be separated and detected. The time elapsing between the initiation of cytoplasmic sampling and the termination of cellular reactions with the molecule of interest is the temporal resolution of the biologic measurement. While the temporal resolution is different for each analyte, diminishing the sampling time enhances the temporal resolution for all measurements,
which broadens the range of biologically relevant analyses. With this technique it is also possible to define a spatial resolution to the measurement. The spatial resolution depends on the volume sampled, the internal diameter of the capillary, and the insertion depth of the capillary into the oocyte. For the data shown, the smallest spatial resolution was estimated to be 50 µm. Finer spatial resolutions may be achievable with capillaries possessing an inner diameter less than 50 µm. This micrometer-sized spatial resolution enables subcellular biochemical measurements to be performed on the 3 mm circumference oocyte. With this sampling strategy, gradients of metabolites across the oocyte can now be identified. Endogenous metabolites or microinjected reporter molecules as demonstrated with the β-galactosidase substrate can be sampled from the oocyte and quantitated. The metabolites or analytes can then be identified by their electrophoretic migration times. Moreover, reporter molecules need not undergo alterations in their absorbance or fluorescence properties when acted upon by cellular proteins. It is far easier to engineer indicator molecules that shift their electrophoretic rather than optical properties in response to activation of a cellular enzyme. Examples are proteases, phosphatases, nucleases, and many others. With an enhanced spatial and temporal resolution for cytosolic acquisition, capillary electrophoresis will become widely applicable to the study of cellular physiology. ACKNOWLEDGMENT The authors thank Chris Sims for technical assistance and Harvey Fishman for stimulating discussions. This work was supported by the Arnold and Mabel Beckman Foundation and the Searle Scholars Program. C.-L.L. was supported by a fellowship from the National Defense Medical Center, Taiwan R.O.C. Received for review May 29, 1997. Accepted September 12, 1997.X AC970550O X
Abstract published in Advance ACS Abstracts, October 15, 1997.
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