Method and Protocol
A low-cost electric micromanipulator and its application to single-cell electroporation
2025 Volume 22 Issue 2 Article ID: e220010
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2025 Volume 22 Issue 2 Article ID: e220010
Micromanipulation techniques are essential in studies of cell function, both for single cells and for cell collectives. Various types of micromanipulators are now commercially available. Hydraulic micromanipulators have the advantage of analogue operation, allowing the user to move the glass microneedle in direct response to their own hand movements. However, they require regular maintenance to maintain their performance. On the other hand, some electric micromanipulators can operate in minute steps of several hundred nanometers, but they are expensive. This paper describes our assembly of a low-cost electric micromanipulator. The device consists of three commercially available stages, three linear DC motors to drive them, and a lab-made control circuit. Using this device, we were able to direct a glass microneedle to cut an MDCK cell sheet. We also manipulated an aspiration pipette to aspirate a portion of a Dictyostelium cell. In addition, we were able to gently touch the tip of an electroporation pipette to the surface of a single target cell in a sheet of fish epidermal keratocytes and load FITC into the cell. Our device can be assembled at one-fourth the cost of commercially available hydraulic micromanipulators. This could make it easier, both economically and technically, to add micromanipulators to all of a laboratory’s microscopes.
We present here a low-cost electric micromanipulator that anyone can easily assemble. It has the advantages of costing about one-fourth the price, in Japanese yen, of a commercially available hydraulic micromanipulator, and does not require regular maintenance to maintain performance. The device can be used for cutting a cell collective sheet by manipulation of a glass microneedle, for micropipette aspiration of a single adherent cell, and single-cell electroporation in a cell collective by gentle contact of an electroporation pipette tip with the cell surface.
Micromanipulation techniques [1–3] are essential in studies of cell function. These techniques have long been used to transplant intracellular organelles into single cells, such as mitotic apparatus from sea urchin eggs [4,5]; in electrophysiological experiments using glass microelectrodes on single muscle fibers [6], neurons [7] and Paramecium cells [8,9] and for microinjection [10] of various substances such as calcium buffer into Paramecium cells [11] or aequorin, a calcium-sensitive photoprotein, into the cytoplasm of the green alga Nitella [12,13]. They are still widely used for microinjection of newly developed fluorescent dyes such as Fura-2 and Indo-1 [14,15] and genes such as chameleon [16] and pericam [17], as well as for microsurgery, such as the removal of lamellipodia from migrating cells [18]. Micromanipulation techniques are also used in cell collectives. For example, the role of front-row cells in the collective migration of epithelial cells was demonstrated to be that of “leaders” by cutting these cells from the epithelial cell sheets [19].
Various types of micromanipulators are now available for purchase. A hydraulic micromanipulator [20] consists of an operator part and a moving part, connected by a thin rigid tube filled with oil or water. Movement of the cylinder in the operator part is transmitted by hydraulic pressure to the moving part. This type of micromanipulator has the advantage of analog operation, allowing the user to move the glass microneedle in direct response to their own hand movements. It is suitable for chasing fast-moving cells such as Paramecium cells with a pipette [21–23]. However, hydraulic micromanipulators require maintenance every few years, as their range of motion decreases with continuous evaporation of the hydraulic fluid. In addition, while they are not too expensive to be out of reach of general research funds, they are not cheap enough to allow them to be attached to all the microscopes in a laboratory. Electric micromanipulators have the advantage of not requiring regular maintenance. Operationally, they can move continuously over long distances at a constant speed. This appears ideal for microsurgery on large cell collectives. High-precision versions using piezoelectric elements are also available for purchase. However, commercially available electric micromanipulators are even more expensive than hydraulic types.
Most laboratories would benefit from access to a micromanipulator that was as precise as a hydraulic micromanipulator, but required no maintenance. Here we show a low-cost electric micromanipulator that we have assembled and currently use. The cost in Japanese yen to assemble it is about a quarter of what needed to purchase a commercial hydraulic micromanipulator. We then give examples of the use of our micromanipulator to cut a collective cell sheet with a microneedle, to aspirate a portion of a cell, and to load a fluorescent dye into single cells by single-cell electroporation. The techniques of which our micromanipulator is capable may not be cutting-edge, but the device is very low-cost and versatile and is thus potentially useful to a wide range of researchers.
The three-dimensional (3D) stage was constructed by combining three single-axis stages (x- and y-axis: TADC-401S; z-axis: TADC-401SZ; Sigma Koki, Tokyo, Japan) with an L-shaped bracket (LBR-4053; Sigma Koki) (Figure 1). The micrometer heads were removed from all the stages and replaced with linear DC motors (SDGC10-13; Sigma Koki), following the manufacturer’s instructions. The 3D stage was mounted on a 15 mm square rectangular aluminum bar attached to the post of an inverted microscope (Ti; Nikon, Tokyo, Japan). A ball joint (B-8B; Narishige, Tokyo, Japan) was connected to the 3D stage to hold the glass microneedle, aspiration pipette, or electroporation pipette as described below. The linear DC motors were operated from a lab-made operation box (Figure 2A). The electric circuit for motor control in the box is shown in Figure 2B. To keep the motor torque constant, we controlled the motors by pulse width modulated (PWM) signals using DC motor driver ICs (TB6612; Toshiba, Tokyo, Japan). A simplified schematic diagram is also shown in Figure 2C. The pulse generator, using a timer IC (NE555; Texas Instruments, Dallas, TX), outputs sequential pulses whose period (t0 in Figure 2C) is fixed at 3.9 ms and whose duty ratio (t1/t0 in Figure 2C) is continuously adjustable from 0.012 to 0.49 on the dial, or can be set at 0.98 by a toggle switch (Figure 2A and B, Speed controller). The 0.98 duty ratio is designed for rapid prepositioning of the glass microneedle or pipettes prior to micromanipulation. When one of the buttons in Figure 2A is pressed, a pulse is sent to the corresponding motor, which is then activated. The motion speed of the motor is a function of the duty ratio. Approximate production cost is shown in Table S1.
The new micromanipulator was tested for operation as follows. A square chamber (18 mm length and width, 2 mm depth), the bottom of which was made of a 24×24 mm coverslip (No. 1; Matsunami, Osaka, Japan), was filled with distilled water (DW). The chamber was then placed on the stage of an inverted microscope (TE300; Nikon). An aspiration pipette was held by the ball joint of the micromanipulator (Figure 1A and B), and the tip of the pipette was immersed into DW in the chamber, as in the case of manipulating cells in a culture medium. Then, the pipette was moved by PWM control pulses, and its tip movement was recorded by a high-speed CCD camera (HAS-220; DITECT, Tokyo, Japan) at 1 ms time intervals through a 40×objective.
Preparation of the single-cell electroporatorThe electroporation pipette and the single-cell electroporator were prepared according to the method of Rae and Levis (2002) [24,25], with small modifications. Briefly, to construct the pipettes, glass capillary (O.D. 1 mm, I.D. 0.6 mm) with a fused filament on the inner wall (GDC-1; Narishige) was used. A capillary was drawn using a vertical puller (PP-830; Narishige) to make two pipettes. One of the pipettes was then bent using a microforge (PG-1; Narishige) approximately 2 mm from the tip to allow vertical contact of the pipette tip with the cell surface. The rear end of the pipette was inserted into the tube containing the fluorescein isothiocyanate (FITC) medium (see below) and left for about 5 hours with the tip facing downwards. The medium filled the tube up to about 10 mm from the tip. The rest of the pipette was back-filled with distilled water (DW). Finally, a 0.2 mm-diameter Ag-AgCl electrode was inserted from the rear end and fixed such that the tip of the electrode was in contact with the FITC medium.
A lab-made electric circuit following Rae and Levis (2002) [24,25] and its photo are shown in Figure 3A and B. A low-bias-current operational amplifier (OPA129; Texas Instruments) is used in the current-to-voltage conversion circuit in the first stage to detect low currents. Circuits in all the following stages, including the subtraction circuit in the second stage, use a low-offset, high-precision operational amplifier (OP07; Texas Instruments). To better understand the complicated real circuit, a simplified schematic diagram is also shown in Figure 3C. The circuit can be viewed as a simple series circuit consisting of two power supplies, a pipette resistor, a gap resistor between the pipette tip and the cell surface, and an oscilloscope to measure current (Figure 3C, Left). As the pipette is lowered to make contact with the cell surface (Figure 3C, Middle, white arrow), the gap resistance increases and the current through the circuit decreases (Figure 3C, Middle, blue arrows). After checking the decreasing current on the oscilloscope, the second power supply is connected via a switch (Figure 3C, Right, white arrow). The current value then increases, releasing the substances by a combination of electrophoresis and electroosmosis.
FITC (F7250; Sigma-Aldrich, St Louis, MO) was dissolved at 5 mg/ml in DMSO and then diluted 50 times with Dulbecco’s phosphate-buffered saline without Ca2+ or Mg2+ (07269-84; Nacalai Tesque, Kyoto, Japan).
Cell cultureMadin-Darby canine kidney (MDCK) cells were provided by Riken BRC (No. RCB0995) through the National BioResource Project of the MEXT, Japan. The cells were cultured using Eagle’s minimum essential medium (MEM, M4655; Sigma-Aldrich) containing 10% fetal calf serum (Nichirei, Tokyo, Japan), 1% MEM-nonessential amino acids (06344-56; Nacalai Tesque) and antibiotic/antimycotic solution (09366-44; Nacalai Tesque).
A Dictyostelium discoideum cell line (myosin II heavy-chain-null cells expressing GFP-myosin II) was used. Cells were developed in Bonner’s standard saline (BSS, 10 mM NaCl, 10 mM KCl, and 3 mM CaCl2) until they became aggregation-competent, as described previously [26].
Keratocyte sheets from the scales of African cichlids (Maylandia lombardoi) were cultured as previously described [27]. Briefly, without sacrificing the fish, a few of their scales were removed and washed in a culture medium (Leibovitz’s medium: L-15, L5520; Sigma-Aldrich) supplemented with 10% fetal calf serum (Nichirei) and antibiotic/antimycotic solution (09366-44; Nacalai Tesque). The scales were placed external side up on the floor of the aforementioned square glass bottom chamber (18×18 mm and 2 mm in depth). They were then covered with another small coverslip and allowed to adhere to the bottom coverslip for 30 min at 23°C. Then, after removal of the upper coverslip, culture medium was added to the chamber and the scales were kept at 23°C again for about 6 h to allow the cells to spread out from the scale. All the experiments were carried out in accordance with national guidelines and were approved by Yamaguchi University’s Animal Use Committee.
Confocal microscopyThe migrating cells were observed using an inverted microscope (Ti; Nikon) equipped with a laser confocal scanner unit (CSU-X1; Yokogawa, Tokyo, Japan). The fluorescence images were detected using an EM CCD camera (DU897; Andor, Belfast, UK).
Micropipette aspirationA portion of a migrating cell was aspirated into a pipette as previously described [28]. Briefly, a suction pipette with an inner diameter of 3 μm was drawn from a glass capillary (G-1; Narishige) using a horizontal pipette puller (PG-1; Narishige) and a microforge (MF-830; Narishige). The pipette was then connected to a vertical open-ended glass tube and a 3-ml syringe via a silicone tube, and all three were filled with BSS. The syringe was then used to adjust the height of the water surface in the glass tube.
We first tested the operation of the new electric micromanipulator. An aspiration pipette was held by the micromanipulator. The tip of the pipette was immersed into a chamber (18×18×2 mm) filled with DW. We sent PWM control pulses with different duty ratios (Figure 4A, inset, t1/t0) at 3.9 ms period (Figure 4A, inset, t0) to the x- or y-axis motors and recorded the left/right (Figure 4A, inset, –x/+x) or front/back (Figure 4B, inset, –y/+y) movement of the pipette. The speed of the pipette was then calculated from the images. Note that the speeds here are averaged over 30 ms to match the time resolution of the human eye. The speed of the pipette was consistent in all directions (+x, –x, +y, –y) at all the duty ratios tested (Movie S1 at a duty ratio of 0.25). Although we did not compare the speeds in the vertical directions (±z), they are likely to be similar to those in the ±x- and ±y-direction. The fact that the speed remains the same in all directions is important for ease of use.
Next, to determine the limits of the device, we detected pipette movement using high-speed recordings at 1 ms intervals (Figure 4C–L). PWM control pulses with duty ratios of 0.13, 0.25, or 0.49 at 3.9 ms cycles were applied (Figure 4C–E). In all tested directions (+x, –x, +y, –y), the speed fluctuations were synchronized with the PWM control pulses (Figure 4F–K). They were more pronounced in the ±x directions (Figure 4F–H) than in the ±y directions (Figure 4I–K), probably because the force of viscous resistance of DW on the pipette is stronger in the y direction. For all duty ratios, there was no significant difference between the mean speeds in all directions (Figure 4L and Movie S2 at a duty ratio of 0.25).
Micromanipulation using the new micromanipulatorCutting of cell sheets is a useful experimental technique. For example, scratch wound assays are traditionally performed, in which a portion of an epithelial cell sheet is cut out to create a simulated wound and the repair process is examined [29]. In another case, leader cells were cut out of cell sheets to clarify their exact role in driving the sheets [19]. We therefore experimented with cutting an MDCK cell sheet by manipulating a glass microneedle with our micromanipulator (Figure 5A and Movie S3). The cell sheet could be accurately cut into squares. Some of the cells at the edge of the square fragment began to move as leaders, with the other cells following (Figure 5B and Movie S4). This is consistent with the typical behavior of MDCK cells in scratch wound assays [30,31].
Micropipette aspiration is a good way to study the mechanical properties of cells [32–34]. This technique requires carefully touching the tip of an aspiration pipette to the cell surface using a micromanipulator. A portion of the cell is then aspirated using the negative pressure of the pipette. Mechanosensitive accumulation of myosin II was observed by micropipette aspiration of several cell types, including Dictyostelium cells [28,35–39], neutrophils [40] and Drosophila embryos [41]. Using our micromanipulator, we brought the tip of an aspiration pipette into contact with the surface of a Dictyostelium amoeba. In response to the negative pressure applied, a portion of the cell was sucked into the pipette. Myosin II was confirmed to accumulate at the tip of the aspirated cytoplasm (Figure 5C, Movie S5). When the negative pressure was released, the portion stretched by aspiration returned to normal, and the cells moved away from the pipette tip and began to migrate on the substrate, indicating that the cell had not been damaged. These results indicate that our micromanipulator is sufficiently practical, at least for MDCK cells and Dictyostelium amoebae.
Combination of the new micromanipulator with single-cell electroporationPressure microinjection is a suitable method for loading substances into single cells several hundred μm in size, such as Paramecium cells and eggs, but is difficult with cells about 10 μm in size. It would be useful if dyes and genes could be easily loaded into single cells as small as 10 μm. To achieve this goal, various single-cell electroporation methods have been developed [24,42–49]. For example, Rae and Levis (2002) reported a sophisticated single-cell electroporation method using modified patch-clamp techniques [24,25]. Their elegant device uses a simple voltage-clamp circuit to detect contact between the pipette tip and the surface of small cell types such as CHO cells, HEK293 cells and α-TN4 cells, and can also load substances into them. Commercially-available devices such as the SU10 (Yokogawa), which uses the electroporation method, can also load substances into cells very easily and efficiently. Combining our electric micromanipulators with the single-cell electroporator of Rae and Levis (2002) [24,25] provides a low-cost system for loading substances into cells.
To perform single-cell electroporation, the tip of a pipette containing the substance must be lowered almost vertically to contact the surface of an adherent cell that is itself only a few μm thick. Here, we made the electroporator used by Rae and Levis (2002) [24,25] and experimented with manipulating the electroporation pipette using our electric micromanipulator. In collective cell studies, loading substances into each cell is a useful technique for studying the role of individual cells in the collective. Fish epidermal keratocytes show collective migration to affect wound repair [50]. All the cells at the leading edge of their sheet extend their lamellipodia and act as leader cells, pulling forward the follower cells behind them [27,51,52]. We tested the feasibility of loading a fluorescent dye, FITC, into one leader cell in the keratocyte sheet (Figure 6A–D). The power supply voltage of the single-cell electroporator (Figure 3C, Left, V1) was set to –1 V and a pipette containing FITC was moved to just above the target cell, using our micromanipulator, and lowered the pipette vertically (Figure 6A, Movie S6). As the pipette was lowered, the current through the pipette decreased from 70 to 60 nA (Figure 6B and C, Movie S7), indicating that the pipette tip had made contact with the cell surface. The resistance of the pipette (Figure 3C, Left, R1) and the contact resistance between the pipette tip and the cell surface (Figure 3C, Left, R2) can be calculated as 1 V/70 nA=14.3 MΩ and 1 V/(70–60) nA=2.3 MΩ, respectively. These values are similar to those reported by Rae and Levis (2002) [24,25]. With the pipette in contact with the cell surface, an additional –4 V supply (Figure 3C, Left, V2) was applied for 500 ms (Figure 3C, Right) to load FITC into the cell. The FITC-loaded cells continued to migrate as leader cells without any apparent damage (Figure 6D, Movie S8). These results indicate that our micromanipulator is capable of fine manipulation.
Finally, we tested the applicability of this device to cells other than keratocytes by performing single-cell electroporation on MDCK cells. We confirmed that by performing electroporation for 500 ms at different voltages, the FITC fluorescence intensity, i.e., the amount of FITC loaded into the MDCK cells, was greater at higher negative voltages (Figure 6E and F), but was somewhat variable at a voltage of –10 V. These results indicate that our electric micromanipulator is capable of precise manipulation, and that the combination of our micromanipulator and the single-cell electroporator of Rae and Levis (2002) [24,25] can serve as a low-cost device for loading substances into cells.
Many commercially-available experimental devices are very advanced, thanks to the efforts of many companies, and contribute to revealing entirely new research results. These cutting-edge devices are inevitably expensive. Devices that are not the most advanced, but can be easily used by many users for routine work, may not be worth developing commercially because they are not very cost effective. In 2013 we invented a low-cost device for loading substances directly into adherent cells using an autopipette [53] and have been using it in our studies [18,54,55]. Although several researchers have expressed a wish to use a device of this type, few manufacturers have expressed any interest in commercializing it. The purpose of this report is not to develop cutting-edge technology, but rather to generate, introduce, and disseminate ideas for low-cost experimental methods.
In this report, we were able to make a micromanipulator at a low cost. One of the contributions to the cost reduction is that we made the electric circuits, etc. by hand, but another important one is that we drastically reduced the performance of our device to just what is necessary and sufficient. For example, as shown in Figure 4, this device, which is driven by a PWM pulse-controlled DC motor, has a slight vibration. However, it is negligible in the general micromanipulations routinely performed by many researchers, and these micromanipulations do not require a precise and expensive mechanism such as a piezoelectric drive. The electric micromanipulator we made enabled us to perform cutting of MDCK cell sheets (Figure 5A and B), aspiration of Dictyostelium amoebae (Figure 5C), and single-cell electroporation of fish epidermal keratocytes and MDCK cells (Figure 6). When we brought the tip of a glass microneedle, an aspiration pipette, or an electroporation pipette into contact with the cell surface, we operated the micromanipulator with a duty ratio of 0.25 or less for PWM control pulses. At least in this speed range, the speed fluctuation synchronized with the PWM control pulses (Figure 4C–L) did not affect the cells. These facts indicate that our device is suitable for manipulating at least stationary or slowly moving cells. Commercially-available hydraulic micromanipulators would, however, be easier to use for pipetting fast-moving cells such as swimming ciliates [56,57]. A hydraulic micromanipulator’s 3D control knob is easy to operate with one hand than our push-button devices, and once accustomed to its use, it can be operated in direct response to the user’s hand movements.
Selecting one cell from a cell collective and analyzing its function while keeping it within the collective would provide new findings that would not be possible using an isolated cell. Staining or manipulating only the targeted individual cells in a cell collective has hitherto required sophisticated techniques or expensive equipment. By combining our device with the single-cell electroporator of Rae and Levis (2002) [24,25] as described in this report, it should be easy for any investigator to load substances into specifically-targeted cells in a cell collective. In future, due to their low cost, multiple micromanipulators could be mounted onto a single microscope and developed into a system to load different substances into each cell in a cell collective. Also, since our device is pulse-controlled, it could easily be developed for computer-controlled movement of pipettes to provide a constant trajectory or repeated movements.
We have assembled a push-button, motorized 3D electric micromanipulator that works with the same level of precision as commercially-available hydraulic micromanipulators, while costing about a quarter of the manufacturing cost of the latter. In addition to its low cost, it does not require regular maintenance, and can be installed on all microscopes in a laboratory for general use.
The authors declare no competing interests.
C.O. and Y.I. designed the research and wrote the manuscript. K.S. and Y.I. designed and made the devices. C.O. supervised all the experiments. N.N. performed the cutting experiments on MDCK cell sheets. M.O. performed micropipette aspiration experiments on Dictyostelium cells. K.S. and N.N. performed the single-cell electroporation experiments. All the authors edited and approved the manuscript.
The evidence data generated and/or analyzed during the current study is available from the corresponding author on reasonable request.
We thank M. Kikuyama (Niigata University) for helpful discussions and Y. Nishigami (Hokkaido University) for inspiring us to write and submit. This study was supported by MEXT/JSPS Kakenhi Grants Nos. 24H01483, 24K21936, 23K27144, 22H05683, 21K19228, 20H03227 to Y.I., and 24K18086 and 21K15055 to C.O.
