May 2020



Measuring Extracellular Proton and Anionic Fluxes in Arabidopsis Pollen Tubes

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The ion-selective vibrating probe has been used to detect and quantify the magnitude and direction of transmembrane fluxes of several ions in a wide range of biological systems. Inherently non-invasive, vibrating probes have been essential to access relevant electrophysiological parameters related to apical growth and morphogenesis in pollen tubes, a highly specialized cell where spatiotemporal tuning of ion dynamics is fundamental. Of relevance, crucial processes to the cell physiology of pollen tubes associated with protons and anions have been elucidated using vibrating probes, allowing the identification of diverse molecular players underlying and regulating their extracellular fluxes. The use of Arabidopsis thaliana as a genetic model system posed new challenges given their relatively small dimensions and difficult manipulation in vitro. Here, we describe protocol optimizations that made the use of the ion-selective vibrating probe in Arabidopsis pollen tubes feasible, ensuring consistent and reproducible data. Quantitative methods like this enabled characterizing phenotypes of ion transporter mutants, which are not directly detectable by evident morphological and reproductive defects, providing valuable insights into molecular and cellular mechanisms. The protocol for quantifying extracellular proton and anionic fluxes detailed here can be adjusted to other systems and species, while the sample preparation can be applied to correlated techniques, facilitating the research of pollen tube growth and development.

Keywords: Ionic fluxes (离子通量), Ion-selective vibrating probe (非损伤微测技术), Non-invasive measurements (非侵入式测量), Phenotyping (表型), Plant ion dynamics (植物离子动力学), Pollen tubes (花粉管)


The relevance of bioelectricity and ion exchange for living cells is unquestionable, having a functional impact in a range of phenomena, from pattern formation, signaling and development to cancer and other diseases (Levin, 2014). Diverse techniques can be employed to detect action potentials, electric fields, extracellular electric currents, and ion fluxes. However, the assessment of their function in vivo requires non-invasive methods. Ideally, any biological system of interest should be studied with the minimum interference and under the most physiological condition possible. Such criteria are attained by the non-invasive ion-selective vibrating probe, which has been used for measuring multiple transmembrane ion fluxes in a wide variety of experimental systems, including Drosophila (Browne and O’Donnell, 2016), zebrafish (Guh et al., 2016), mouse skin (Sun et al., 2015), roots (He et al., 2015), Daphnia (Stensberg et al., 2014), C. elegans (Adlimoghaddam et al., 2014), etc. In pollen tubes, quantitative measurements of extracellular ion fluxes using the ion-selective vibrating probe have been fundamental in establishing the role of major ions (especially Ca2+, H+, K+, and anions) in apical growth. When used in association with reverse genetics and pharmacology targeting specific molecular players, these methods also allowed the identification of mutation effects and subtle phenotypes, such as through quantitative analysis of aberrant oscillatory behavior (Certal et al., 2008; Michard et al., 2011 and 2017; Portes et al., 2015; Wudick et al., 2018; Hoffmann et al., 2020).

The ion-selective vibrating probe is a technique designed to reduce the electric noise related with ionophore loaded, glass microelectrode probes, bringing the signal/noise ratio to levels compatible with the measurement of physiological extracellular electric/ionic currents, associated with single cells and other biological systems (Shipley and Feijó, 1999; Kunkel et al., 2006). The experimental setup comprises a customized microelectrode, front-loaded with ion-selective ionophore cocktails (LIX) that measures the voltage at two locations at close vicinity of the plasma membrane (with µm precision), so that most noise and drift induced bias is subtracted from the final output. In the described protocol/setup, two routines allow for extra quantitative precision: on one hand, the background reference is measured away from any cell and also subtracted from the final voltage difference for convection, thermal or ionic gradient compensation; on the other hand, background concentration of the measured ion is continuously monitored, allowing the quantitative normalization of the fluxes in real time using Fick’s laws of diffusion.

The entire setup consists on an inverted microscope, nanometer 3-D positioners, and electrode impedance/capacitance correcting amplifier head stages, all placed inside a Faraday’s cage to reduce environmental electrical noise. Individual probes are calibrated at the start and end of experiments by measuring their potential in 3 appropriate solutions with concentrations ranging 3 orders of magnitude, deemed usable if closer than 95% to the Nernstian potential over the reference solutions. All experimental output is generated by an external, variable-gain amplifier with an analog read-out, and fed through an A/D board into a dedicated computer. All data processing including the calibration procedure, probe quality control, 3-D stepper motor-driven probe positioning and movement (vibration) system is performed by the ASET software (Automated Scanning Electrode Techniques - Applicable Electronics). The spatial resolution is limited by the dimension of the probe tip, usually at 1-3 μm of diameter, allowing sampling of small and specific patches at the cell surface. Depending on the ion, flux resolution goes into the pmol cm-2 s-1 range (Shipley and Feijó, 1999; Kunkel et al., 2006).

The use of a first-generation wire/voltage detection vibrating probes led to the discovery of an electric field around pollen tubes, which have been proposed to be cells behaving as electrical dipoles (Weisenseel et al., 1975). While highlighting the importance of ion dynamics for pollen tube growth and development, these early measurements and interpretative models were affected by technical artefacts of these early wire electrode probes, namely the stirring induced by the hundreds of Hz vibration needed for the noise subtraction by lock-in amplifiers. Discussing these limitations (e.g., Shipley and Feijó, 1999) allowed to assess ionic derived currents, and the subsequent development of ion-specific glass microelectrodes methods. Besides measuring only a specific ion, instead of the electric current stemming from the sum of all ion transport over a given surface, it also vibrates at very low frequencies (< 0.5 Hz) thus respecting the formation and stability of ion gradients over the cell surface during data acquisition. The resulting extensive effort led to the description of ion dynamics of pollen tubes in species as lily and tobacco (Feijó et al., 1999; Certal et al., 2008, Michard et al., 2008). The adoption of such species as study models was presumably related to the convenience of appropriated cell dimensions and high pollen germination and pollen tube growth rates, making them easy to obtain and experimentally manipulate in vitro. Despite the progress made using these species, the search for molecular mechanisms related to apical cell growth and sexual plant reproduction required the adoption of Arabidopsis thaliana as a genetic model. The application of these techniques to study ion dynamics and morphogenesis in Arabidopsis pollen is challenging due to the reduced dimensions of the flowers and difficulties in pollen manipulation in vitro. Besides these complications, the development of sophisticated molecular and genetic tools available for Arabidopsis required substantial protocol adaptations and optimizations, which were crucial for obtaining consistent data. Efforts to develop these protocols and approaches allowed the molecular identification and cellular localization of many ion transporters underlying the ion fluxes using ion-specific vibrating probes, together with the identification of key players involved in regulating ion homeostasis and morphogenesis in pollen tubes (reviewed in Michard et al., 2017).

The importance of protons and anions for pollen tube growth has been reported in diverse studies, where the presence of an intracellular gradient of these ions and the identification of the ion transporters promoting their movement was determined (Zonia et al., 2002; Michard et al., 2008; Gutermuth et al., 2013; Domingos et al., 2019). Of relevance, protons have been reported to enter mainly at the tip and generalized efflux occurring along the tube shank and pollen grain (Feijó et al., 1999; Certal et al., 2008; Hoffman et al., 2020). Anions, deemed as mostly chloride (Cl-) for being the only anion in the germination medium, have been shown to have an opposite flux direction, with influx along the pollen tube and grain and massive efflux at the tip (Zonia et al., 2002; Gutermuth et al., 2013; Domingos et al., 2019). Herein, a detailed description of the sample preparation and experimental protocol for measuring proton and anionic fluxes in pollen tubes with the ion-selective vibrating probe is presented.

Using quantitative techniques able to access measurable variables and parameters is complementary to genetics and imaging tools, offering a complete tool kit for characterizing mutant phenotypes. In this context, the use of ion-selective vibrating probes allowed the identification of phenotypes being otherwise undetectable, once diverse mutations do not display evident morphological and reproduction defects due to the high genetic redundancy and other compensatory mechanisms present in pollen tubes (Gutermuth et al., 2013; Wudick et al. 2018; Domingos et al., 2019). Experimental approaches coupling biophysical with molecular data are fundamental for an integrated and multidisciplinary investigation, contributing to better understand the mechanisms regulating intracellular cell physiology. The detailed protocol described below can be easily reproduced and adjusted to other systems, also enabling interested researchers to study pollen tube growth and development.

Materials and Reagents

  1. Reference electrode – Dri-Ref (World Precision Instruments, catalog number: DRIREF-2 )

  2. Glass microfiber filters (Sigma-Aldrich, catalog number: WHA1820047 )

  3. Safe-Lock tubes, 1.5 ml (Eppendorf, catalog number: 0030120086)

  4. Acrylic rectangular dish with a round chamber of 10 mm (custom-made) being the commercial equivalent the glass bottom dish (Cellvis, catalog number: D35-10-1-N )

  5. Cover slip (Fisher Scientific, catalog number: 12-545 )

  6. Glass syringe (Hamilton, catalog number: 80600 )

  7. Silica gel (Sigma-Aldrich, catalog number: 13767 )

  8. Silver/silver chloride wire (Fisher Scientific, catalog number: AA41390G2 )

  9. Borosilicate glass capillaries (World Precision Instruments, catalog number: TW150-4 )

  10. N,N-Dimethyltrimethylsilylamine (Sigma-Aldrich, catalog number: 41716 )

  11. Hydrogen ionophore II-cocktail A (Sigma-Aldrich, catalog number: 95297 )

  12. Chloride ionophore I-cocktail A (Sigma-Aldrich, catalog number: 99408 )

  13. Sucrose (Sigma-Aldrich, catalog number: S7903 )

  14. Potassium chloride (Sigma-Aldrich, catalog number: P9333 )

  15. Magnesium sulfate (Sigma-Aldrich, catalog number: 746452 )

  16. Boric acid (Sigma-Aldrich, catalog number: B6768 )

  17. Calcium chloride (Sigma-Aldrich, catalog number: C1016 )

  18. 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) (Sigma-Aldrich, catalog number: H3375 )

  19. Poly-L-lysine solution (Sigma-Aldrich, catalog number: P4707 )

  20. Agarose low gelling temperature (Sigma-Aldrich, catalog number: A9045 )

  21. Germination medium (see Recipes)

  22. Stock solutions (see Recipes)


  1. pH meter

  2. Vortex

  3. Microwave

  4. Oven

  5. Desiccator

  6. Fine needle-sharp tweezers (Sigma-Aldrich, catalog number: T4537 )

  7. Benchtop microcentrifuge (Eppendorf, model: 5415D )

  8. Micropipette puller (Sutter Instrument Company, model: P-97 )

  9. Ion-Selective Vibrating Probe (System Applicable Electronics, model: SIET-Scanning Ion-selective Electrode Technique )

  10. Inverted microscope (Nikon, model: Eclipse TE300 )

  11. Microscope light power supply (Nikon, model: TE-PS100 )

  12. Camera (Andor, model: iXon3 )

  13. 60x/1.40 Objective lens oil immersion (Nikon, model: Plan Apo )


The procedure description includes the detailed preparation of germination medium, experimental dishes, pollen collection, ion-selective microelectrodes, and saline bridges (only for anionic fluxes). The germination protocol was optimized from Boavida and McCormick (2007). Critically, the ionic background concentrations must be lowered as much as possible to minimize background noise when using the ion-selective vibrating probe technique. Thus, the concentration of all components of the germination medium was reduced aiming to reach a minimal, but functional, culture medium sufficient for pollen germination while increasing the signal-to-noise ratio.

  1. Dishes preparation


    1. Keep stock solutions under -20 °C.

    2. Prepare fresh germination medium on the experimental day.

    3. Due to the reduced dimensions of Arabidopsis pollen grains, the glass microelectrode can easily touch the glass bottom and break. To solve this problem, a thin layer of liquid agarose is placed over the glass, just enough to cover it. After drying, the agarose pad is coated with poly-L-lysine to increase the pollen adherence avoiding pollen tube floating. The ideal condition is having the pollen tube growing attached to the agarose pad facilitating the probe positioning. Pollen grains remain attached to the agar for few hours without dish agitation.

    4. The poly-L-lysine should be properly removed to avoid toxicity.

    1. Prepare fresh liquid germination medium from stock solutions.

    2. Fix a cover slip at the bottom of the round chamber with 10 mm of diameter of the acrylic rectangular dish (not needed when using the commercial equivalent).

    3. Melt 0.01% of low melting agarose in 1 ml of germination medium in a safe-lock tube (1.5 ml) for 30 s in the microwave.

    4. Add 50 μl to the bottom of the dish and remove the excessive liquid with a glass microfibre filter keeping only the volume sufficient to cover the glass surface but creating an agarose pad.

    5. Add 50 μl of poly-L-lysine on top of the agarose pad.

    6. Wait 5 min.

    7. Remove poly-L-lysine.

    8. Wash twice with liquid medium to remove the excessive poly-L-lysine but still keeping the adhesive property.

  2. Pollen collection and germination


    1. Growing Arabidopsis plants preferentially under short-day photoperiod conditions increase pollen quality and stability, achieving high germination rates (> 90%). Although, flowers grown under other photoperiods conditions can also be used.

    2. Start to collect flowers from the 10th silique. The first siliques have a low density of pollen and the germination rate is reduced.

    3. Collect flowers until 1:00 pm, otherwise, the flowers start to close and pollen viability decreases;

    4. Density is very important for Arabidopsis pollen germination. Collect 20-25 flowers per dish of 10 mm diameter well.

    5. Resuspend pollen precipitate with a 200 μl tip with the first millimeters cut to enlarge its aperture, as to avoid pollen grains damage.

    6. To prepare the wet chamber use a Petri dish with a paper filter soaked in distilled water, to avoid medium evaporation during pollen germination.

    1. Grow Arabidopsis plants until flowering (ecotype Columbia - Col-0) in growth chambers under short-day photoperiod conditions (12 h light/12 h dark cycle) at 22 °C with 70% humidity and light intensity of ~100 μmol m-2 s−1 to improve pollen integrity and density.

    2. Collect flowers immediately after anthesis (stage 13 – Smyth et al., 1990) using a thin tweezer and transfer to a 1.5 ml tube (no more than 100 flowers per tube - use 20-25 flowers per dish).

    3. Add 1 ml of germination liquid medium.

    4. Vortex at high speed (~2,500 rpm) for 30-40 s.

    5. Centrifuge at 1,600 x g for 3 min.

    6. Remove flowers and supernatant.

    7. Resuspend gently the pollen precipitate in 100 μl of liquid medium.

    8. Add one single drop (~25 μl) of the pollen precipitate in the experimental dish.

    9. Add 200 μl of liquid medium very slowly to keep pollen grains attached to the agarose pad.

    10. Incubate the dishes in a wet chamber at 22 °C, preferentially in the dark.

    11. After 2-3 h growing pollen tubes with ≥ 200 μm of length can be assayed.

    The procedure is illustrated in Figure 1.

    Figure 1. Schematic representation of flower collection and sample preparation steps. The illustration shows how to obtain pollen from flowers, followed by resuspension and incubation of pollen grains to promote pollen germination. Finally, extracellular ionic measurements can be performed in sufficiently long Arabidopsis pollen tubes.

  3. Glass micropipette silanization and ion-selective probe preparation


    1. Optimizing the puller parameters to obtain ion-selective probes according to the desired pipette configuration. The tip aperture should have ~1-3 μm of diameter for measuring both protons and anions, while the taper is slightly longer for anions to decrease ionophore leakage (see Shipley and Feijó, 1999 for reference).

    2. Ionophores are commercialized by different brands, being purity and quality important factors that can drastically vary according to the manufacturer, having impact on the Nernst values measured by the electrode. Furthermore, discrepant Nernst values could be an indication that the ionophore is old or contaminated.

    1. Pulling proton- and anionic-specific micropipettes.

    2. Place the micropipettes in a wire net or Teflon chamber that enables air circulation (Figure 2).

      Figure 2. Glass micropipettes silanization using a wire net or a Teflon chamber. A. Micropipettes placed on a Teflon chamber for silanization. Syringe indicates the aperture where the N, N-Dimethyltrimethylsilylamine is applied. B. Closed chamber ready for the silanization procedure. C. Micropipettes placed on a wire net for silanization. D. Closed chamber ready for the silanization procedure.

    3. Keep the micropipettes overnight in the oven at 210 °C.

    4. Add 90 μl of N,N-dimethyltrimethylethylsilamine with a glass syringe.

    5. Wait 30 min.

    6. Let the micropipettes cool down inside the oven.

    7. Store the silanized micropipettes in a desiccator with silica gel.

    8. Using a syringe, backfill the microelectrode with a 20-25 mm column of 40 mM KH2PO4/15 mM KCl, pH 7.5, and front-loaded with a ~25 μm column of the hydrogen or chloride ionophore cocktail.

  4. Ion-selective probe calibration


    1. Calibration solutions used for H+ flux measurements are pH 5, 6, and 7 and 0.1, 1, and 10 mM KCl for anionic measurements.

    2. As the chloride ionophore detects other anions in addition to Cl, the estimates are referred to anionic fluxes instead of chloride fluxes.

    3. A stable H+-electrode can be used for 4-5 h of measurements; it is recommended to check its stability by measuring one of the calibration solutions every 2 h.

    4. The anionic-electrode should be prepared and stabilized for ~2 h before calibration. Electrode stabilization consists of keeping the electrode in the 0.1 mM KCl calibration solution. Moreover, after stabilization and calibration, the anionic-electrode can be used only for 1 h for ensuring reliable measurements.

    5. The reference electrode (Dri-Ref) should be kept in 3 M KCl and properly rinsed with water before use.

    6. The silver/silver chloride wire connected to the pipette holder has to go through the electro-chloridizing process before use. The process consists of removing any coating residue from the wire with a small piece of thin sandpaper. After that, a 9 V battery is connected to a dual crocodile clip cable, where in one end has a thick silver wire attached and the other end has the metal pin of the pipette holder. Both wires ends must be immersed at the same time in a solution of 1 M KCl for a few seconds. When the silver wire of the pipette holder becomes darker, it is ready to use.

    1. Connect the microelectrode at the holder ensuring the contact of silver/silver chloride wire with the filling solution.

    2. Place the microelectrode in the 3 calibration solutions recording the voltage values individually, estimating the adequate Nernst potential fitting.

    3. After calibration, place the microelectrode in the experimental dish.

    4. Close the circuit by inserting the dry reference electrode also into the dish (Figure 3).

      Figure 3. Experimental setup and sample dish. Ion-selective vibrating probe setup consisting of the reference electrode (left), experimental dish and the ion-selective microelectrode connected to pipette holder.

    5. Position the microelectrode close enough to the plasma membrane of the pollen tube but avoid touching it (Figure 4, Videos 1 and 2).

    6. Measure the background reference in a region > 500 μm away from any pollen grain or pollen tube.

      Figure 4. Proton- and anionic-electrodes. Examples of a proton-specific (left) and anionic-specific electrode (right) placed in front of the tip of Arabidopsis pollen tubes. Anionic-specific electrodes have a longer taper and slightly smaller tip aperture.

      Video 1. Timelapse from extracellular proton measurement at the tip of Arabidopsis pollen tube. Acquisition of 100 frames with 4 s interval.

      Video 2. Timelapse from extracellular anionic measurements at the tip of Arabidopsis pollen tube. Acquisition of 100 frames with 4 s interval.

  5. Data processing

    1. Save the output data from the ASET software.

    2. Use data from ASET output to calculate fluxes as set up on the Excel file provided. The spreadsheet has as input the following constants: Electrode excursion, ionophore efficiency, diffusion coefficient, slope, and intercept of the calibration curve. In the cells below the constants one should input the ASET ‘Background’ and ‘Voltage difference’ measurements that will produce an output in the columns to the right of the calculation spreadsheet. One should then select all rows of the output and drag it down until it matches the end of the input rows.

          The spreadsheet subtracts the background reference values from the voltage differential recordings and calculates ion fluxes using Fick’s law for protons using diffusion coefficient 9.37 x 10−5 cm−2 s−1 efficiency 1; and for chloride diffusion coefficient -2.03 x 10−5 cm−2 s−1 and efficiency 0.5.

  6. Salt bridges (used only for anionic flux measurements)

    Note: Salt bridges are used for isolating the biological sample from the reference electrode, avoiding the germination medium contamination with high concentrations of KCl, leaking from the reference electrode. Otherwise, the chloride concentration would substantially increase in the culture medium altering the signal-to-noise ratio, generating a source of error in these estimates. The salt bridge establishes the electrical contact between the two dishes but keeps the reference electrode isolated from the biological sample avoiding contamination and inaccuracies related to the ionic concentration fluctuation during the measurement.

    1. Create a bridge by heating the center of a glass capillary and folding it using two forceps to create a bridge with a “V” format.

    2. Fill the bridge with a jellified solution of germination liquid medium with 0.01% of low melting point agarose.

    3. Place one part of the bridge in the dish where the pollen tubes are growing (where the ion-selective probe is placed) and the other part of the bridge is inserted in the other dish filled only with liquid medium (where the reference electrode is placed) (Figure 5).

      Figure 5. Salt bridge for anionic fluxes measurements. The anion-specific electrode is placed into the experimental dish while the reference electrode is placed in another dish filled with germination medium. The salt bridge connects both dishes closing the measuring circuit. The setup is zoomed in on the right.


  1. Germination medium

    500 μM KCl

    500 μM CaCl2

    125 μM MgSO4

    0.005% (w/v) H3BO3

    125 μM HEPES

    16% (w/v) sucrose

    Adjust pH to 7.5 with NaOH

  2. Stock solutions

    Stock KCl      100 mM

    Stock CaCl2     100 mM

    Stock MgSO4  100 mM

    Stock H3BO3   1%

    Stock HEPES   100 mM


J.F. lab was supported by the National Science Foundation grants (MCB 1616437/2016 and 1930165/2019) and the University of Maryland. We thank Custódio de Oliveira Nunes and Michael A. Lizzio for the photos.

Competing interests

There are no conflicts of interest or competing interest.


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[摘要]离子选择性振动探针已被用于检测和量化各种生物系统中几种离子的跨膜通量的大小和方向。固有Ñ上侵入性,振动探针已经必需访问有关在花粉管,与心尖生长和形态发生电生理参数高度专业化的细胞,其中离子动力学的时空调谐是根本。与此相关的是,已使用振动探针阐明了与质子和阴离子相关的花粉管细胞生理学的关键过程。 ,可以识别潜在分子并调节其细胞外通量。利用拟南芥作为遗传模型系统所带来的相对赋予了新的挑战LY尺寸小,不易操纵体外。在这里,我们描述了协议优化,该优化使在拟南芥花粉管中使用离子选择性振动探针成为可能,从而确保了一致且可重复的数据。Q这样的定量的方法启用表征离子转运蛋白的突变体表现型,这是不被明显形态学和生殖缺陷直接可检测的,提供了有价值的见解分子和细胞机制。可以将此处详述的用于量化细胞外质子和阴离子通量的方案调整为其他系统和物种,同时将样品制备方法应用于相关技术,从而促进对花粉管生长和发育的研究。

[背景]生物电和离子交换的对活体细胞的相关性是毋庸置疑的,具有一个功能影响的范围内的现象,从图案形成,信令和发展癌症和其他疾病(莱,2014)。多样的技术可以被用于检测动作电位,电场,胞外电流,和离子通量。然而,它们的功能评估在体内需要小号的非侵入性的方法。理想情况下,任何感兴趣的生物系统都应在尽可能少的干扰下和最生理的条件下进行研究。这些标准是由非侵入性的离子选择性振动探头,它已被用于在很宽的测量多个跨膜离子通量达到各种实验系统,包括果蝇(布朗和O'Donnell的,2016),斑马鱼(GUH的等。,2016),小鼠皮肤(孙等人,2015年),根(他等人,2015年),水蚤(Stensberg等,2014),C.线虫(Adlimoghaddam等,2014),等等。在花粉管中,使用离子选择性振动探针定量测量细胞外离子通量对于确定主要离子(尤其是Ca 2 + ,H + ,K +和阴离子)在根尖生长中的作用至关重要。当与相关联地使用反向遗传学和pharmacolog ÿ靶向特定的分子的玩家,这些方法也允许编的识别突变效果和细微的表型,如叔hrough的异常定量分析振荡行为(Certal等人,2008; Michard等人。,2011和2017;波特斯等人,2015; Wudick 。等人,2018;霍夫曼。等人,2020)。

离子选择性振动探针是一种旨在减少与装有离子载体的玻璃微电极探针相关的电噪声的技术,使信噪比达到与测量与单个细胞和其他细胞相关的生理性细胞外电流/离子电流兼容的水平生物系统(Shipley和Feijó,1999; Kunkel等,2006)。实验设置包括定制的微电极,前装载用离子选择性离子载体鸡尾酒(59) ,其测量在两个位置上的电压在接近附近的质膜(与μ米精度),因此,大部分噪声和DRIF吨诱导偏压从最终输出中减去。在所描述的协议/设置中,有两个例程可实现更高的定量精度:一方面,在远离任何单元的位置测量背景参考,并从对流,热或离子梯度补偿的最终电压差中减去背景参考;另一方面,可以连续监控被测离子的背景浓度,从而可以使用Fick扩散定律实时对通量进行定量归一化。

的整个装置由一个Ñ倒置显微镜,纳米3- d定位器,和电极阻抗/电容校正放大器头阶段,所有放置在法拉第笼内部减少环境的电噪声。个别探针在通过用浓度范围3个数量级的测量在3个适当的解决办法它们潜在的开始和结束实验校准,认为可用如果距离小于95% ,以在能斯特电势的Reference olutions。所有实验输出均由具有模拟读数的外部可变增益放大器生成,并通过A / D板馈入专用计算机。所有数据处理包括在校准程序,探针的质量控制,3-d步进电机驱动探头定位和移动(振动)系统是由ASET软件( -适用的电子自动扫描电极技术)来进行。空间分辨率受探针尖端尺寸的限制,通常直径为1-3μm,从而可以在细胞表面采样小而特定的斑块。取决于离子,通量分辨率进入pmol cm -2 s -1范围(Shipley和Feijó,1999; Kunkel等人,2006)。

在使用的第一代代线/电压检测振动导致探针发现一个的电场围绕花粉管,这已被建议为细胞behav荷兰国际集团作为电偶极子(Weisenseel等人,1975) 。而突出离子动力学的花粉管生长和发育的重要性,这些早期的测量和interpre吨ative模型受这些早期的线电极探针的技术假象,即搅拌诱导所需噪声数百赫兹振动的减法通过锁相放大器。讨论这些局限性(例如希普利和费若,1999 )允许吨ö评估离子衍生的电流小号,以及特定离子的玻璃microele后续发展Ç TRODE小号方法。乙eside小号仅测量一个特定的离子,而不是ë LECTRIC电流所产生从所有的总和离子,我在给定表面输送吨也振动小号在很低的频率(< 0.5赫兹)从而尊重离子的形成和稳定性数据采集过程中细胞表面的梯度。导致花粉管离子动力学的物种如律的描述将得到广泛的努力ÿ和烟草(费若。等人,1999; Certal 。等人,2008年,Michard等人,2008。 )。这样的通过物种作为研究模型中推测有关到拨的便利性细胞的尺寸和高花粉萌发和花粉管生长速率,使它们易于获得和实验操作在体外。尽管使用这些物种取得了进展,但寻找与根尖细胞生长和有性植物繁殖相关的分子机制仍需要采用拟南芥作为遗传模型。由于花朵的大小减少和体外花粉处理的困难,应用这些技术研究拟南芥花粉的动力学和形态发生具有挑战性。除了这些并发症之外,拟南芥可利用的先进分子和遗传工具的开发还需要进行大量的方案调整和优化,这对于获得一致的数据至关重要。È ffort s到开发这些协议和方法允许分子识别和细胞定位许多离子转运底层的离子通量使用离子特异性振动探针,连同参与调节花粉管离子稳态和形态发生的主要参与者的识别(综述在Michard等人,2017年)。

质子和一个的重要性离子为花粉管的生长已被报道在不同的研究中,其中这些离子的胞内梯度的存在,并且所述离子转运促进它们的移动的标识确定(Zonia等人,2002; Michard等等人,2008; Gutermuth等人,2013; Domingos等人,2019)。的相关性,质子已经被报道主要进入在尖端和广义流出发生环沿管柄和花粉粒(费若等人,1999; Certal 。等人,2008;霍夫曼。等人,2020)。阴离子,视为大多氯(氯- )为是在发芽培养基中的唯一的阴离子,已经显示出具有相反的通量DIRECTI上,沿着花粉管和谷物和大量流出在尖端(Zonia涌入等。,2002; Gutermuth等,2013; Domingos等,2019)。本文中,将详细介绍使用离子选择振动探针测量花粉管中质子和阴离子通量的样品制备和实验方案。


关键字:离子通量, 非损伤微测技术, 非侵入式测量, 表型, 植物离子动力学, 花粉管

参比电极– Dri-Ref(世界精密仪器,目录号:DRIREF-2)
玻璃微纤维过滤器(Sigma-Aldrich,目录号:WHA1820047 )
盖玻片(Fisher Scientific,目录号:12-545)
银/氯化银丝(Fisher Scientific,目录号:AA41390G2)
琼脂糖低胶凝温度(Sigma-Aldrich,目录号:A9045 )


倒置显微镜(尼康,型号:Eclipse TE300)
60x / 1.40物镜浸油(尼康,型号:Plan Apo)




保持库存溶液在-20以下 ℃。

将0.01%的低熔点琼脂糖在安全锁定管(1.5 ml)中的1 ml发芽培养基中融化,在微波炉中加热30 s 。
加50 μ升到培养皿的底部,并除去过量的液体,具有一玻璃微纤维过滤器仅保留体积足以覆盖玻璃表面,而且产生的琼脂糖垫。
添加50 μ聚-L-赖氨酸的升琼脂上的顶部OSE垫。


增长荷兰国际集团优先在短日照条件下光照的拟南芥植物的花粉提高质量和稳定性,实现高发芽率(> 90%)。虽然,也可以使用在其他光周期条件下生长的花朵。
要准备湿室,请使用带滤纸的P etri皿,将滤纸浸入蒸馏水中,以避免在花粉萌发过程中介质蒸发。

在短时光周期条件下(12小时光照/ 12小时黑暗周期)于22 °C ,湿度为70%,光强度为〜100μmolm -2的条件下,在生长室中种植拟南芥植物直至开花(生态型Columbia - Col-0)。s -1改善花粉的完整性和密度。
收集花后立即nthesis(阶段13 -史密斯等人,1990),使用薄的镊子和转移到1.5ml管(每管不超过100鲜花-使用每皿20-25花)。
以高速(〜2500 rpm)涡旋30-40 s 。
以1600 x g离心3分钟。
在实验皿中加入一滴(〜25μl )花粉沉淀。
加入200 μ液体介质的升得非常缓慢,以保持polle Ñ附着到琼脂晶粒OSE垫。
将碟子在22 °C的潮湿箱中孵育,最好在黑暗中孵育。
后2-3小时生长花粉管与≥200 μ长度的米可用于检测。




根据所需的移液器配置优化拉拔器参数,以获得离子选择性探针。末端孔应具有〜1-3 μ直径用于测量两个质子和阴离子的米,而锥形稍长为阴离子降低离子载体泄漏(见希普利和费JO,1999为参考ENCE) 。



将微量移液器在210 °C的烤箱中过夜。
添加90 μ N个升,N -dimethyltrimethylethylsilamine用玻璃注射器。
使用注射器,回填用40mM KH的20-25毫米柱的微小电极2 PO 4 / 15mm以上的KCl,pH 7.5和前加载有〜25 μ氢或氯化物离子载体鸡尾酒的m个列。


Ç使用用于h alibration溶液+通量测量为pH 5,6 ,和7及0.1,1 ,和10mM KCl的阴离子测量。
由于氯化物离子载体检测除了氯以外的阴离子- ,估计被称为阴离子通量代替氯化物通量。
稳定的H +电极可用于4-5小时的测量;建议通过每2小时测量一种校准溶液来检查其稳定性。
校准前应准备好阴离子电极并使其稳定约2 h。电极稳定化包括在0.1 mM KCl校准溶液中对电极进行密封。此外,经过稳定和校准后,阴离子电极只能使用1小时,以确保可靠的测量。
参比电极(Dri-Ref)应保存在3 M KCl中,并在使用前用水正确冲洗。
连接到移液器支架的银/氯化银导线在使用前必须经过电氯化处理。该过程包括用一小片薄砂纸去除导线上的任何涂层残留物。之后,将9 V电池连接到双鳄鱼夹电缆,其中一端连接有粗银线,另一端具有移液器支架的金属销。两条导线两端必须在1米氯化钾的溶液的同时被浸入一个几秒钟。当移液器支架的银线变深时,就可以使用了。



测量的区域中的背景参考> 500 μ m距离任何花粉粒或花粉管程。


视频1.缩时从尖端细胞外质子测量拟南芥花粉管。以4 s的间隔采集100帧。

视频2.拟南芥花粉管尖端细胞外阴离子测量的时移。以4 s的间隔采集100帧。

根据提供的Exce l文件中的设置,使用ASET输出中的数据来计算通量。电子表格具有作为输入下列常数:ë lectrode偏移,离子载体效率,扩散系数,斜率,以及校准曲线的截距。在常数下方的单元格中,应输入ASET“背景”和“电压差”测量值,这将在计算电子表格右侧的列中产生输出。然后,应选择输出的所有行并将其向下拖动,直到与输入行的末尾匹配为止。
电子表格从电压差记录中减去背景参考值,并使用菲克定律针对质子使用扩散系数9.37 x 10 -5 cm -2 s -1效率1来计算离子通量;对于氯化物扩散系数为-2.03×10 -5 cm -2 s -1和效率为0.5。

注意:盐桥用于隔离从参比电极的生物样品,避免了与高浓度的萌发培养基污染的KCl ,泄漏荷兰国际集团从参比电极。否则,培养基中的氯化物浓度将大大增加,从而改变信噪比,从而在这些估计中产生错误的来源。盐桥在两个培养皿之间建立了电接触,但使参比电极与生物样品保持隔离,避免了污染以及与测量过程中离子浓度波动有关的不准确性。

通过加热玻璃毛细管的中心并使用两个镊子将其折叠以创建“ V”格式的桥来创建桥。



0.005%(w / v )H 3 BO 3

125μM HEPES   
16%(w / v)蔗糖 

股票KCl 100毫米                         
库存CaCl 2 100 mM           
股票硫酸镁4 100mM的           
股票H 3 BO 3 1%           
库存HEPES 100 mM           


JF实验室获得了美国国家科学基金会(MCB 1616437/2016和1930165/2019)和马里兰大学的资助。我们感谢Custódiode Oliveira Nunes和Michael A. Lizzio的照片。




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引用:Portes, M. T. and Feijó, J. A. (2021). Measuring Extracellular Proton and Anionic Fluxes in Arabidopsis Pollen Tubes. Bio-protocol 11(3): e3908. DOI: 10.21769/BioProtoc.3908.

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