Recording Electrical Currents across the Plasma Membrane of Mammalian Sperm Cells

All experiments were performed in accordance with NIH guidelines for animal research and approved by UC Berkeley Animal Care and Use Committee (AUP 2015-07-7742), with every effort made to minimize animal suffering. All described methods are consistent with the recommendations of the Panel on Euthanasia of the American Veterinary Medical Association and IACUC committee. All experimental procedures utilizing human derived samples were approved by the Committee on Human Research at the University of California, Berkeley, IRB protocol number 2013-06-5395. 1. Making glass micropipettes for whole-cell sperm patch-clamp recording. NOTE: The small size of the cytoplasmic droplet requires glass micropipettes with fine tips. For micropipette fabrication, start with borosilicate glass capillaries with an outer diameter 1.5 mm, inner diameter 0.86 mm and an internal filament. Pull the tip on a micropipette puller (Figure 3B). For shaping and polishing of the pipette tip, position a hot wire in close proximity of the tip for a moment. Perform this process of fire-polishing under 100x magnification using a pipette polisher (Figure 3C,D). Ensure that the inner diameter of the pipette tip is ~2 µm before fire polishing and is reduced to ~0.5 µm after proper polishing. NOTE: Only freshly pulled pipettes made on the day of recording are recommended for use. This avoids the accumulation of dust particles on the pipette tip. 2. Setting up the rig Assembly of the recording chamber Use a low-volume diamond-shaped perfusion chamber with laminar flow and relatively fast solution exchange rates (Figure 4). Connect the chamber via a manifold to a gravity-fed perfusion system (Figure 5). To perfuse a solution through the recording chamber, assemble a simple self-made gravity- fed perfusion system with a polytetrafluoroethylene manifold that connects the perfusion chamber to eight separate perfusion lines. NOTE: The tubing should not leach plasticizers into the solutions, since such chemicals can significantly impact the normal function of sperm cells60 and alter ion channel activity61,62 . A small diameter polytetrafluoroethylene tubing is highly recommended as the main perfusion tubing. As polytetrafluoroethylene tubing is rather stiff, make flexible connections with high purity silicon tubing fit over the polytetrafluoroethylene tubing. In addition, use syringes (e.g., Luer Lock syringe- type) that lack lubricant, since mineral oil or other lubricant additives can interfere with ion channel recording (Figure 5). Assembling an agar bridge To avoid liquid junction potential changes during recordings, keep the environment around the reference electrode stable by using an agar bridge (Figure 4). To make an agar bridge, make an L-shaped glass capillary by bending it under a small Bunsen burner fire and let it cool down. Make a solution of 1% agarose in 1 M KCl and heat it in a microwave until the agarose melts and the solution becomes transparent. Carefully fill the L- shaped glass capillary with the above solution to avoid air bubbles and let it cool to room temperature. The agar bridges can be stored in 1 M KCl for ~2 months at +4 °C. To use an agar bridge, fill an Ag/AgCl pellet reference electrode with 1 M KCl and insert the agar bridge carefully to avoid solidified agarose popping out of the glass capillary. Next, insert the gold pin (2 mm) of the amplifier headstage wire into the reference electrode 2 mm (gold) slot and insert the end of the L-shaped agar bridge in the perfusion chamber as shown in Figure 4. Solutions and recipes ​NOTE: All reagents and chemicals must be of the highest purity. All prepared solutions are filtered through a sterile 0.22 µm PES filter and stored at 4 °C for up to one month. It should be noted that the exact adjustment of osmolarity is a prerequisite for successful patch clamping. Prepare human tubular fluid (HTF) solution as described in Table 1. Adjust the pH to 7.4 with NaOH. Osmolarity should be 280 ± 5 mOsm, measured using a vapor pressure osmometer. Prepare high saline (HS) "bath" or "extracellular" solution as given in Table 2. Adjust the pH to 7.4 with NaOH. Osmolarity should be 320 ± 5 mOsm. Prepare the cesium methanesulfonate "bath" or "extracellular" solution as described in Table 3. Adjust the pH to 7.4 with CsOH and the osmolarity to 320 ± 5 mOsm. To prepare the cesium methanesulfonate (CsMeSO3) intracellular "pipette" solution follow Table 4 and adjust pH to 7.4 with CsOH. Osmolarity should be up to 10 mOsm higher than the HS solution. 3. Isolation and purification of mammalian spermatozoa NOTE: Euthanize C57BL/6 male mice aged 3-6 months by inhalation of CO2 followed by cervical dislocation. After cervical dislocation, perform tissue collection (cauda or corpus epididymis) from mice immediately. Isolation of epididymal mouse spermatozoa NOTE: Euthanize C57BL/6 male mice aged 3-6 months by inhalation of CO2 followed by cervical dislocation. After cervical dislocation, perform tissue collection (cauda or corpus epididymis) from mice immediately. Open the lower abdominal area of the mouse with scissors and extract both epididymides. Place them in a 35 mm cell culture dish filled with HS solution (high saline solution, see Table 2). Ensure that an aliquot of the HS solution is prewarmed to room temperature before contact with the sperm cells. Transfer the epididymides into a new cell culture dish containing HS solution and thoroughly remove all residual fat. Separate the epididymides into caput, corpus and cauda using a #15 scalpel blade (Figure 6). Transfer the corpus (or cauda if needed) of each epididymis into a new cell culture dish containing HS solution. Make multiple incisions in the isolated part of the epididymis using a pointed #11 scalpel blade. Transfer the parts of the epididymides with multiple incisions into a 1.5 mL microcentrifuge tube containing 1.5 mL HS solution. Briefly shake sperm cells from the epididymis into the solution using super-fine Dumont type 5a forceps. Discard epididymides and leave the tube at room temperature for 10 min. Wait until the solid matter (non-sperm cells) sediments to the bottom of the tube, and then transfer the supernatant into another 1.5 mL microcentrifuge tube. Store the sperm mixture at room temperature with access to air for up to 2 h, during which time the experiments can be performed. Isolated sperm lose their performance 3 h after isolation. Capacitation of mouse spermatozoa ​NOTE: All epididymal spermatozoa are suitable for patch-clamp, but only spermatozoa from the cauda epididymis are mature enough to become fertilization competent. Since caudal spermatozoa can undergo capacitation, perform experiments, particularly sperm capacitation, using caudal sperm cells. Perform sperm collection on caudal or ejaculated spermatozoa (see steps 3.1.1-4). Take caudae isolated as described in step 3.1.4. and apply multiple incisions as described in step 3.1.5. Transfer the caudae to a 10 mm cell culture dish with 2 mL of capacitation solution. Incubate the caudal tissue in this capacitation solution initially for 10 min at 37 °C and 5% CO2, then discard the tissue and incubate the released sperm cells for another 50-80 min. NOTE: In vitro capacitation of mouse caudal spermatozoa is achieved by 60-90 min incubation. Capacitated sperm cells are visually identified based on excessive asymmetric bending of their flagella known as hyperactivation. This type of motility allows spermatozoa to swim in a nonlinear manner. Collection and purification of human sperm NOTE: In this case, fresh semen samples were obtained from a total of 21 healthy volunteers aged 21-38 and spermatozoa purified by the room temperature swim- up technique as described in detail in48,63 . Briefly, the procedure is as follows: Leave the human ejaculate samples at room temperature for 60 min to allow for liquefaction before being purified (longer durations may negatively affect sperm quality). Preheat about 7 mL of HTF solution to 37 °C in a 50 mL centrifuge tube. Visually assess the volume of the ejaculate-if the volume is greater than 1 mL, prepare multiple tubes of HTF solution. Carefully underlay 1 mL of liquefied ejaculate to the bottom of each tube without lifting strands of semen to the surface of the buffer. The semen-to-HTF- interface should be as clear-cut as possible. Avoid air bubbles. Place the tubes in a 37 °C incubator for 1 h in a stand that holds them at a 45° angle for maximal surface area. If the incubator is a CO2 incubator, close the cap tightly to prevent acidification of the buffer (CO2 is not required for this procedure). During the incubation, motile sperm will gradually swim towards the surface of the HTF buffer. After 1 h collect the uppermost 1 mL fraction from all samples in a 15 mL centrifuge tube, thus combining the highest quality, highly motile sperm for patch clamping. Do not attempt to remove all the supernatant-leave several mL of HTF remaining above the seminal plasma to avoid contamination. The purified cell suspension can be stored at room temperature for several hours. NOTE: A normal ejaculate should contain at least 15 x 106 sperm cells per mL64, but sperm counts vary widely between samples. The obstacle of a low cell count may be overcome by letting the purified sperm suspension sit for 30-60 min at room temperature before patch clamping. Under this condition, the cells will gradually accumulate at the bottom of the centrifuge tube and form a visible cell cloud from which you can pipette cells into the recording chamber (see below). Capacitation of human spermatozoa NOTE: In-vitro capacitation of human spermatozoa can be accomplished by 4 h incubation in capacitation solution: 20% fetal bovine serum, 25 mM NaHCO3 in HTF or HS buffers48,51 . Capacitated sperm cells can be visually identified based on their vigorous motility and excessive asymmetric bending of the flagella that allows spermatozoa to swim in a nonlinear manner. Prepare a 2x capacitation solution and mix 1:1 with the sperm suspension. Perform incubation at 37 °C and 5% CO2 for a minimum of 4 h. To ensure a high percentage of capacitated cells, incubation time should be no shorter than 4 h. 4. Preparing coating solution (only needed for human sperm patch-clamp) NOTE: An essential step is to lift the attached spermatozoon from the coverslip before break-in. This step is only necessary for human sperm cells and requires coating of the glass coverslip to create a less adhesive glass surface. Coverslip coating reduces the sperm cells’ probability of sticking to the coverslip and allows human spermatozoa to be lifted from the glass coverslip after successful gigaseal formation. Dilute 200 µL of the ejaculate in 5 mL of HS solution in 15 mL centrifuge tube. Spin down at 300 x g for 5 min and remove supernatant. Resuspend the pellet in 1 mL of HS solution and transfer to a microcentrifuge tube. Sonicate for 10 min at 25 °C using a water-filled sonication bath. Spin down at 10,000 x g for 5 min. Transfer the supernatant to a fresh microcentrifuge tube and use it for coating (instructions below). 5. Recording ion conductance from the whole sperm plasma membrane. Attaching sperm to the coverslip. NOTE: Before the patch-clamp recording, spermatozoa are plated onto 5 mm round cover slips. If working with human cells, briefly dip the coverslips into the prepared coating solution using sharp forceps. Place four cover slips into a single well of a 4-well plate and cover with 300 µL of HS solution. Prevent the cover slips from floating by taping them to the bottom of the 4-well plate, pushing them with the plastic tip of a pipettor. Gently resuspend the concentrated suspensions of human (step 3.3.5) or mouse spermatozoa (step 3.1.9) from the storage tube (20-50 µL of concentrated sperm) in 300 µL of HS solution above the coverslip. Ensure that spermatozoa accumulate at the bottom of the well where their heads will attach to the coverslip and their flagella will vigorously beat in the solution just above the cover slip. It is important to allow spermatozoa to sediment onto the coverslip for about 10 min at room temperature before recording. NOTE: If human spermatozoa attach poorly, go back to step 5.1.1 and dilute the prepared coating solution with HS solution (1:20 to up to 1:2). This step must be adjusted to sperm samples from different human donors based on the variable sperm motility that is common in human sperm preparations. Placing the cover slip Take one coverslip with slightly adhered cells from the well and place into a diamond-shaped well of the recording chamber containing HS solution (Figure 4). Turn on the perfusion system (Figure 5) filled with HS solution to rinse the chamber and remove debris and excess detached spermatozoa from the chamber. This will increase the probability that the recording pipette stays clean when it reaches the cells. This step is helpful when working with non- capacitated cells that tend to stick to the coverslip stronger. NOTE: If recording from capacitated sperm, one must avoid turning on the perfusion immediately after a fresh cover slip is placed into the chamber. Capacitated spermatozoa have vigorous motility and only loosely attach to the coverslip. Therefore, to ensure that the largest number of capacitated sperm are available, it is important to wait ~ 10 min and let capacitated spermatozoa adhere to the coverslip before turning on perfusion. Once the appropriate capacitated cell is chosen and a gigaseal is formed, the perfusion can be turned back on. Selection of a sperm cell for patch-clamp recording. Find a suitable sperm cell with a cytoplasmic droplet using 600x magnification (see below). This is done using a 60x water immersion len as described in the methods with a 10x magnification eyepiece. In sperm cells isolated from corpus epididymis, the CD is usually located close to the center of the midpiece. In caudal cells, the CD can usually be found close to the annulus (Figure 2C). For human sperm, the CD is located in the neck region (Figure 2A,B). NOTE: Additionally, the pull-out 1.6x magnification option of the microscope can be used to closely examine sperm morphology (magnification 960x). Ensure that the cytoplasmic droplet is oval and has a slightly elongated (Figure 2G), spindle-like shape. CDs that appear very round and enlarged are often fragile and unsuitable for patching (Figure 2D,F). Select a spermatozoon that is motile with the head attached to the coverslip, such that the sperm cell is partly fixed, but the CD and the rest of the flagellum continues to move with flagellar beating. Ensure that the head of the sperm is loosely attached to the coverslip so that it rotates a little bit as the flagellum moves from side to side. NOTE: Loose attachment is important since after formation of the gigaohm seal and before the transition into the whole-cell mode, the spermatozoon must be lifted from the coverslip into the solution. Forming a gigaseal ​NOTE: To record monovalent CatSper currents in the whole-cell mode, pipettes are filled with Cs-methanesulfonate (also referred to as pipette or intracellular) solution (Table 4) – this yields a pipette resistance of 11-17 MΩ (Figure 7A), as automatically calculated by the amplifier upon applying a voltage step from 0 mV to 10 mV. Pipettes can also be filled with other solutions depending on the application. It is recommended that solutions are made with mobile anions such as methanesulfonate, Cl- , aspartate or similar ions to reduce pipette resistance. The osmolarity of the pipette solution should be up to 10 mOsm higher than the bath solution. Slightly higher tonicity of the pipette solution helps to keep the access resistance into the sperm cells as low as possible during patch-clamp recording. After visual selection of a sperm cell with a proper morphology (Figure 2G), fill the micropipette with a pipette solution (Table 4) and secure it into the pipette holder. In order to keep the pipette tip clean from debris, apply positive pressure to the pipette using the U-tube shaped assembly to ensure that the pipette solution flows out of the tip after it is immersed into the extracellular solution, also referred to below as the "bath" solution (see Table 2). NOTE: If conditions permit, the formation of the seal under constant perfusion is highly advised, as this condition ensures the cleanest pipette tip. Lower the pipette down and immerse its tip into the bath solution (Table 2). At this time, pipette resistance can be measured (Figure 7A). In order to clearly visualize the cell, position the tip of the pipette above the CD with the opening of the tip aligned diagonally (approximately 45°) toward the CD. Quickly lower the tip of the pipette toward the CD to be in the same focal plane, within a few µm of the CD. As soon as the tip of the pipette touches the CD, apply negative pressure to the pipette using the "mouthpiece" to move part of the droplet into the tip and form a gigaohm seal – also known in electrophysiology as a "suction event". Usually, after initial contact with the pipette tip, the negative pressure in the pipette can be maintained up to a minute to gradually achieve a gigaohm seal. After successful formation of the gigaohm seal (> 4-20 GΩ), lift the spermatozoon from the coverslip (Figure 7B). NOTE: This step is not required for rodent sperm cells due to their length; however, partial detachment of the sperm flagellum is beneficial to alleviate potential pipette drift. Human sperm detachment is an essential step, since the CD is small (1~3 µm); if the cell is left attached to the coverslip, even a slight drift of the pipette during the experiment can disrupt the CD. After the spermatozoon is lifted, its flagellum normally continues to beat, while the cell is attached only to the micropipette. Break-in and transition into the whole-cell mode. Compensate stray capacitance transients using the amplifier's compensatory mode before transitioning to the whole-cell mode (Figure 7B). To initially observe capacitance transients, hold the membrane potential at 0 mV and apply 10 mV pulses using the "Membrane Test" tool (Figure 7B). Perform a break-in and transition into the whole-cell mode by applying short (1 ms) gradually increasing (430-650 mV, ~50 mV increment) voltage pulses combined with a very light suction (Figure 7C). NOTE: To apply break-in pulses, use the previously programmed break-in protocol within the patch clamp software. The break-in voltage pulses are applied to the spermatozoon using the high voltage command input (rear-switched) on the back of the amplifier. After application of each break-in voltage pulse, launch the Membrane Test tool to check whether larger capacitance transients appear. NOTE: The presence of large capacitance transients (Figure 7C) indicates that the break-in has occurred, and the entire capacitance of the cell is being measured (~1 pF for human sperm and ~2.5 pF or more for murine sperm). Fit the large capacitance transients using the Membrane Test Tool to determine the capacitance of the whole cell as well as its access resistance. NOTE: The access resistance should be as low as possible for efficient perfusion of the interior of the sperm cell with the micropipette solution, as well as for reliable voltage clamp of the sperm plasma membrane. Access resistance is usually 25-30 MΩ and 50-70 MΩ for Cs+ /K+ – and NMDG- based intracellular solutions, respectively. After successful break-in, proceed with the planned whole cell patch-clamp experiments, such as applying various bath solutions (Figure 8) containing different compounds or measuring channel activities using voltage-step (Figure 8, Figure 9) or voltage-ramp protocols. If the break-in was not successful, return to step 6.4.1. by choosing a fresh micropipette and select another appropriate sperm cell.