The overall goal of this protocol is to characterize metabolites in live cells of developing embryos by in situ microsampling capillary electrophoresis-mass spectrometry. This method can help answer key questions in the cell and developmental biology field because it enables the analysis of metabolism in single cells directly in live developing embryos. The main advantage of this technique is that it is fast, scalable to different cell sizes and enables minimally invasive sampling of individual cells in live embryos.
Visual demonstration of this method is critical as some steps are difficult to learn because the embryo develops relatively fast and a custom built capillary electrophoresis electrospray ionization instrument is used. Begin this procedure with preparation of 2%agarose in one 1X Steinberg's solution. While still liquid, coat the bottom of 60 millimeter Petri dishes with the solution.
Once the agarose gel has cooled and solidified, flame the end of a six-inch Pasteur pipette until it forms a ball. Then, lightly touch the heated end to imprint five to 10 wells approximately one millimeter deep into the agarose. To fabricate tapered tip micropipettes, first, pull borosilicate capillaries in a Flaming/Brown type capillary puller using the settings found in the text protocol.
Next, break off the tip of the pulled micropipette using a pair of fine sharp forceps to obtain a capillary tip outer diameter of approximately 20 microns. Perform this step under a stereo microscope to aid precision and reproducibility. Obtain embryos through gonadotropin induced natural mating of adult xenopus laevis or via in vitro fertilization as referenced in the text protocol.
Remove the jelly coats surrounding the embryos as they begin to cleave into the two-cell stage by first letting the embryos rest in the dejellying solution for two minutes. Then, gently swirl them for an additional two minutes to prevent the embryos from adhering to the surface of the collection dish. Gently pour the dish contents into a clean beaker and quickly decant the dejellying solution from the beaker.
Immediately cover the eggs with 0.1X Steinberg's solution to rinse off the remaining dejellying solution. Gently swirl and then decant the solution. Repeat this step four times to thoroughly wash the embryos.
Transfer the dejellied embryos into 1X Steinberg's solution in a Petri dish. To minimize crowding within the plates, place approximately 100 embryos per 100 millimeter dish. Sort the cleaving embryos at the two-cell stage into a separate dish in which stereotypical pigmentation confidently marks the dorsal ventral access with reference to establish sulfate maps.
Identify correctly cleaving embryos by ensuring that the first cleavage furrow, which demarks the midsagittal plane, bisects the darkly pigmented animal pull and the lightly pigmented animal pull such that the two halves are mirror images. Next, mount a fabricated micropipette on a multi-axis micromanipulator. Connect the micropipette to a microinjector.
Use a plastic transfer pipette to aspirate approximately five of the eight-cell embryos and transfer them into the sampling dish containing 0.5X Steinberg's solution. Using a hair loop, position the embryo to be sampled into an individual well and ensure that the correctly identified cell of interest is facing the microprobe at an approximate angle of 90 degrees. While working under the stereo microscope, guide the tip of the micropipette into the identified single cell within the live embryo.
Withdraw a desired portion of the cell's contents by applying negative pressure pulses to the microcapillary using the microinjector. Gently retract the microprobe from the cell and transfer its tip into four microliters of the metabolite extraction solvent chilled in a micro-sized vial. Next, apply a pressure surge of positive 80 psi for one second to the capillary using the microinjector to expel the aspirate into the solvent.
Tightly close the vial to prevent evaporation and place the vial back on ice until sampling is complete. Once sampling is completed, vortex mix the sample containing microcentrifuge vials for approximately one minute to expedite extraction of metabolites. Then, centrifuge the vials at 8, 000 times g for five minutes at four degrees Celsius to pellet cellular debris and other particulates.
Store the cell extracts together with the cell debris and precipitate at Minus 20 degrees Celsius for a day or at minus 80 degrees Celsius for up to one month until measurement by CE-ESI-MS. Construct the CE injection platform capable of rapid vertical translation of a stage holding the background electrolyte vial and the sample loading microvial as referenced in the text protocol. Assemble the CE-ESI interface by first mounting the electrospray metal emitter into a three-port T-union.
Then, feed the separation CE capillary through the electrospray emitter, allowing it to protrude approximately 40 to 100 microns beyond the tip of the emitter. Work under a stereo microscope to aid accuracy. Connect the sheet solution capillary to the remaining port to supply the electrospray solution.
Use appropriate sleeves and finger-tightened connections for leak-free operation of the CE-ESI interface. Using a plateholder, mount the CE-ESI interface onto a three-axis translation stage and position the electrospray emitter tip approximately two millimeters from the mass spectrometer orifice. To clean the components of the interface, first supply the electrospray sheet solution through the electrospray emitter at one microliter per minute and the BGE through the CE separation capillary.
Use syringe pumps to feed the solvents at a steady rate. Then, flush the CE separation capillary before each measurement by connecting a syringe to the capillary inlet end. Use efficiently large syringes to minimize refilling and priming of solvent supply lines.
After rinsing the separation capillary for approximately five minutes, transfer its inlet into BGE solution located in the stainless steel vial. Position the electrospray emitter tip approximately two millimeters from the mass spectrometer orifice. Fine adjust this distance using a translation stage to generate electrospray in the stable cone jet regime while monitoring the spray using a stereo microscope.
Monitor the stability of the total ion current or TIC for approximately 30 to 45 minutes to ensure stable operation. Apply 21 plus or minus two kilovolts to the BGE vial by gradually ramping up the potential over approximately 15 seconds, typically generating approximately 7.5 microamps of current across the separation capillary using 1%percent formic acid as the BGE. Before each measurement, ensure system stability by monitoring the TIC profile for approximately five to 10 minutes and then stepwise lower the separation potential to zero volts.
Pipette one microliter of the acetyl cooling standard solution into the injection vial and transfer the separation capillary from the BGE vial into the injection vial. Lift the CE injection stage 15 centimeters in one second. After 60 seconds, hydrodynamically inject six nanoliters of the sample into the separation capillary.
Subsequently, translate the stage back to starting levels and gently move the capillary inlet end into the BGE. Immediately after, ramp up the CE voltage to start electrophoretic separation and EMA state acquisition. Once the standard has been detected, stop the data acquisition and lower the separation voltage stepwise to zero volts.
Then, retrieve the emitter to two centimeters from the orifice. Flush the separation capillary for five minutes before analyzing the cell extract. Measure 10 nanoliters of the single cell extract by repeating these steps using 90 seconds to hydrodynamically inject the sample.
A representative electropherogram of the cell extract is shown here, in which the separation of a series of identified metabolites is displayed. 70 different signals were identified with high confidence as small polar metabolites based on migration time and MS/MS fragmentation patterns of cell extract signals compared to those from a standard or from tandem mass spectral libraries. Metabolite identification is conducted by comparing the accurate mass, migration time and fragmentation behavior of the unknown signal from the sample against the metabolite standard or data available in a metabolite database.
This step is demonstrated for histidine. Here, the individual dorsal cells that were identified and sampled using microprobe CE-ESI-MS are shown at the eight-cell, 16-cell and 32-cell embryonic stages. Multi-varied analysis of identified metabolites in the three different cell types uncovered different metabolic profiles and complex metabolic trends across cell stages.
Principal component analysis of the quantitative metadata uncovered metabolic changes as an identified progenitor cell of the eight-cell embryo divided to form a cell clone in the 16-cell and 32-cell embryo. Representative trends are shown for select metabolites. Metabolites such as acetylcholine decreased in abundance with cell division, whereas trolamine and serine/arginine showed opposing trends in these cell types.
Once mastered, this technique can be done very fast, usually in under 10 seconds for each cell. We first had the idea for this method when we began the mapping of metabolic cell heterogeneity in the early xenopus laevis embryo. After watching this video, you should have a good understanding of how to use microprobe capillary electrophoresis-mass spectrometry to analyze metabolites in single embryonic cells.
While attempting this procedure, it's important to remember to work fast as the metabolome is dynamic and cells divide quickly in the developing embryo. Also, examine analytical details to ensure data of high qualitative fidelity are collected on cell metabolism. The implications of this technique extend toward diagnosis of live specimens because our microprobe mass spectrometry approach works fast, is minimally invasive and compatible with small samples.
Following this procedure, other methods like sulfate tracking can be performed in order to answer additional questions like the impact of metabolites on tissue specification. Don't forget that working with high voltage power supplies can be extremely hazardous and precautions such as electrically isolating electrified interfaces should always be taken while performing this procedure.
