Fast and Efficient Expression of Multiple Proteins in Avian Embryos Using mRNA Electroporation

Summary

We report messenger RNA (mRNA) electroporation as a method that permits fast and efficient expression of multiple proteins in the quail embryo model system. This method can be used to fluorescently label cells and record their in vivo movements by time-lapse microscopy shortly after electroporation.

Abstract

We report that mRNA electroporation permits fluorescent proteins to label cells in living quail embryos more quickly and broadly than DNA electroporation. The high transfection efficiency permits at least 4 distinct mRNAs to be co-transfected with ~87% efficiency. Most of the electroporated mRNAs are degraded during the first 2 h post-electroporation, permitting time-sensitive experiments to be carried out in the developing embryo. Finally, we describe how to dynamically image live embryos electroporated with mRNAs that encode various subcellular targeted fluorescent proteins.

Introduction

Electroporation is a physical transfection method that uses an electrical pulse to create transient pores in the plasma membrane, allowing substances like nucleic acids or chemicals to pass into the cytosol. Electroporation is widely used to deliver DNA into bacteria, yeast, plants, and mammalian cells^1,2,3. It is routinely used to introduce genetic payloads into target cells and tissues within the developing avian embryo to study the genetic control of development or label migrating populations of cells^4,5,6,7. However, several experimental limitations also exist with DNA electroporation⁸. For instance, DNA electroporation often introduces highly variable numbers of expression vectors per cell and subsequently the mRNAs and proteins they encode. This variability can lead to considerable cell-cell heterogeneity that complicates both image analysis and data interpretation^9,10. Additionally, proteins from DNA electroporation only begin to express ~3 h post-electroporation and do not reach the maximum efficiency in cell number and fluorescence intensity until 12 h, likely due to the time required to transfer into the nucleus and complete both transcription and translation in vivo¹¹.

In contrast, mRNA transfection has been effectively used in a variety of model systems, including Xenopus laevis oocytes by microinjection^12,13, reprogramming human stem cells by mRNA lipofectamine transfection¹⁴, and electroporating recalcitrant neural stem cells in adult mice¹⁵. We tested the ability of mRNA electroporation to efficiently label cells during early avian embryonic development using in vitro synthesized mRNAs that encode distinct fluorescent proteins (FPs). For our studies, we used the pCS2+ vector, a multipurpose expression vector that is commonly used for expressing proteins in Xenopus and zebrafish embryos. The SP6 and T7 RNA polymerase promoters in the pCS2+ permit the synthesis of mRNA and protein from any cloned gene when used in an in vitro transcription/translation system.

Here, we demonstrate that mRNA electroporation allows fast and efficient expression of fluorescent proteins (FPs) in gastrulating quail embryos. We designed and generated many of the expression vectors used in these studies. For example, we subcloned the LifeAct-eGFP gene¹⁶ into the pCS2+ vector¹⁷ to express from the CMV promoter and SP6 promoter. The inserted gene lies downstream of the SP6 promoter and upstream of the SV40 poly(A) tail¹⁸. In embryos co-electroporated with mRNA and DNA, FPs encoded from in vitro transcribed mRNAs were first detected within 20 min of electroporation, whereas FPs from DNA expression vectors were detected only after 3 h. Multiple mRNAs encoding for nuclear, Golgi, and membrane proteins can be electroporated into an embryo simultaneously, resulting in the quick and efficient expression of multiple proteins in individual cells. Finally, using an in vivo fluorescence recovery after photobleaching (FRAP) assay, we show that a majority of the electroporated mRNAs decay within 2 h. Thus, fast initial protein production combined with limited new protein translation makes mRNA electroporation a valuable technique when temporal control of expression is necessary.

Protocol

All animal procedures were carried out in accordance with approved guidelines from the Children’s Hospital Los Angeles and the University of Southern California Institutional Animal Care and Use Committees.

1. Generation pCS2-based Expression Vectors

1. To clone pCS2.Lifeact-eGFP, prepare the vector backbone by digesting 2 µg of pCS2.CycB1-GFP (a construct containing a different insert) with BamHI (10 U) and BsrGI (10 U) in appropriate digestion buffer (see Table of Materials) diluted to 1x in a total reaction volume of 50 µL for 1 h at 37 °C (see Figure 1 for the schematic of cloning procedure).
   1. Dephosphorylate the free 3’OH ends of the vector backbone from the restriction enzyme reaction by adding Shrimp Alkaline Phosphatase (1 U). Incubate for 30 min at 37 °C.
   2. Run the whole mixture in a 1% agarose gel/1x Tris Acetate EDTA (TAE) buffer at 90 V for ~50 min and then stain the gel in a 0.5 µg/mL solution of ethidium bromide in 1x TAE buffer for 15 min with gentle rocking. Also, make sure to load molecular weight markers in a free lane to help determine DNA sizes.
   3. Using a DNA safe UV Transilluminator to avoid nicking DNAs, cut out the vector backbone (expected band size of 4kb) from the agarose gel quickly and isolate the DNA fragment from the gel piece using a gel purification kit using manufacturer’s instructions.
2. Simultaneously with step 1.1, prepare the insert by digesting 2 µg of pEGFP-N1-Lifeact with BglII (10 U) and BsrGI (10 U) in appropriate digestion buffer diluted to 1x in a total reaction volume of 50 µL for 1 h at 37 °C. Run the whole mixture in a 1% agarose gel to isolate the 800 bp band as previously explained in step 1.1.2-1.1.3.
3. After both the vector and insert are purified and quantified on the spectrophotometer, combine 50 ng of the vector and 38 ng of the insert (1:3 molar ratio of vector to insert) in T4 DNA ligase buffer prior to adding T4 DNA ligase to catalyze the ligation reaction. Additionally, set up a no DNA control, a vector only control, and an insert only control. Incubate all reactions at room temperature for 30 min.
   1. Transform 1 µL of the ligation mixture(s) into competent E. coli DH10. Also, transform water only negative control and 20 pg of pUC19 positive control. Spread bacteria onto Luria Broth (LB) agar plates containing 50 µg/mL ampicillin and incubate overnight at 37 °C.
   2. On the following morning, count the colonies on each plate.
      NOTE: Ideally, there should be no colonies on the negative control plates, >100 colonies on the vector+insert ligation plates, and fewer colonies on the vector-only plate.
   3. Pick at least 8 bacterial colonies from the agar plate transformed with the vector+insert ligation mixture and inoculate them using sterile toothpicks or pipet tips into 2 mL of liquid LB broth containing 50 µg/mL Ampicillin.
   4. Extract DNA from each of the clones using commercial plasmid miniprep kits.
   5. Run a diagnostic restriction digest using 500 ng of DNA from each clone using NotI (10 U) and BamHI (10 U) in the appropriate digestion buffer diluted to 1x for 1 h at 37 °C and run the digested DNA on a 1% agarose gel in 1x TAE buffer. Expected band sizes for pCS2.Lifeact-eGFP positive clones are 3.9 kb and 978 bp.
   6. Transform 1 pg-100 ng of DNA from the positive clone into competent E. coli DH10, and prepare 3-4 miniprep reactions as detailed in previous steps to obtain a sufficient amount of DNA for in vitro transcription reaction (10 µg minimum).

2. Preparation of mRNA by In Vitro Transcription

1. Linearize 10 µg of pCS2.LifeAct-eGFP on the 3’ end of the insert with NotI (10 U) in an appropriate digestion buffer diluted to 1x in a total reaction volume of 50 µL at 37 °C overnight.
   NOTE: To prevent RNase contamination and reduce mRNA degradation, wear gloves while handling mRNA samples.
2. Purify the DNA using a mixture of phenol: chloroform: isoamyl alcohol (25:24:1, v/v).
   1. Add 150 µL of RNase-free water to make the total volume of the digestion reaction equal to 200 µL. Then, add 200 µL of phenol: chloroform: isoamyl alcohol and vortex the mixture for 20 s.
   2. Centrifuge in a microfuge at max speed (18,400 x g) for 2 min. Carefully remove the top aqueous phase that contains the linearized DNA. Repeat this step to remove additional impurities and be careful not to disrupt the white precipitate that may form between the bottom and top liquid phases.
3. Precipitate linearized DNA by adding 1/10 volume 3M sodium acetate (RNase-free) and 2.5 volumes of 100% ethanol. Leave the mixture at -20 °C for >30 min, then pellet the DNA by centrifuging at max speed (18,400 x g).
   1. Wash the DNA pellet with 70% ethanol, then air-dry the pellet for >5 min.
   2. Dissolve the DNA pellet in 5 µL of RNase-free water. Quantify the DNA by spectrophotometer.
      NOTE: The expected DNA concentration is ~0.5-1 µg/µL with an A260/A280 ratio between 1.7-2.0.
4. Use 1 µg of the linearized pCS2.LifeAct-eGFP DNA for in vitro transcription (IVT). Follow the manufacturer’s instructions in the commercial kit (see Table of Materials) to include 10 µL of cap analog (final concentration 0.8 mM) and NTPs (final concentration 1 mM for ATP, CTP, and TTP; 0.2 mM for GTP), 2 µL of 10x reaction buffer (final concentration 1x), 2 µL of SP6 RNA polymerase, and RNase-free water up to 20 µL.
   1. Incubate the IVT mixture for about 2 h or longer, depending on transcript size.
      NOTE: 2 h incubation works well for a 3 kb transcript, but overnight incubation works better for transcripts that are greater than 5 kb.
   2. To remove free nucleotides from the transcription reaction, add 30 µL of LiCl RNA precipitation solution (7.5 M lithium chloride, 50 mM EDTA) to precipitate the mRNA. Vortex the mixture briefly and store at -20 °C for 30 min. Spin the mRNA down for 15 min in a microfuge at max speed (18,400 x g) and rinse with 70% ethanol (RNase-free).
5. Dissolve the synthesized mRNA in 15 µL of RNase-free water. Dispense the synthesized mRNA at 5 µL/tube (3 tubes total) and place at -80 °C for long-term storage. Quantitate the mRNA with a spectrophotometer.
   NOTE: Expected yield is 15-20 µg of mRNA (~1 µg/µL). 1 µL (1 µg/µL) is sufficient for an experiment with ~10 embryos, assuming that the mRNA is diluted from 1 µg/µL to 500 ng/µL and that each embryo is injected with ~200 nL. Each tube should, therefore, contain enough mRNA for ~5 experiments.
6. Run ~300 ng of mRNA on a 1% agarose gel in 1x TAE buffer after cleaning all gel equipment with the RNase decontamination solution (e.g., RNase Away) and ensure that the mRNA appears as one band (multiple bands may be present if secondary structures form) and that no smear appears in the gel lane, which indicates RNA degradation.

3. Preparation of mRNA Electroporation Mix

1. Thaw and dilute mRNA at the desired concentration (250-500 ng/µL in RNase-free water works well for all mRNAs tested in this work). Add 1/10 volume of colored dye (see Table of Materials, RNAse-free, 0.1% final concentration) to help visualize the mRNA injection site and to properly place the electrodes.
   NOTE: If DNA and RNA are combined to perform a co-electroporation, ensure that DNA is free from contaminating RNases by using phenol-chloroform extraction and ethanol precipitation prior to mRNA and DNA mixture.
   1. Be sure to prepare a negative control using a mock (no RNA added) electroporation solution. If possible, prepare a positive control using pre-validated mRNA (mRNA that has been shown to produce FP successfully in previous experiments).
      NOTE: The negative control is essential for all imaging experiments to establish background fluorescence levels for normalization of data. The positive control is especially important when working with newly transcribed mRNA by helping to confirm that the electroporation settings worked.
2. Store the mRNA electroporation solution on an ice slurry to prevent degradation until ready to proceed with electroporation.

4. Electroporate mRNAs into Living Quail Embryos

1. Collect freshly laid fertilized quail eggs daily and store at 13 °C in a humidified refrigerator for no longer than 1 week. Incubate the quail eggs at 38 °C until the desired embryonic developmental stages^19,20,21.
   NOTE: HH3 to HH5 were used in this study for both static and dynamic imaging. For HH3 embryos, leaving the eggs at room temperature for 2 h prior to harvesting makes the isolation process much easier as the embryos are generally more resistant to physical manipulations when cooled down.
2. Isolate and prepare embryos according to the EC culture system²². Collect at least 5 embryos per condition, including at least one to serve as a negative control (not electroporated).
   1. Gently break and pour the egg into a 10 cm Petri dish. Remove the majority of the thick albumin with a transfer pipette and remove the remaining thick albumin around the embryo by gently wiping the surface of the yolk with a tissue wipe to ensure that the embryo sticks tightly to the paper ring.
   2. Lay precut filter paper (see Table of Materials) upon the embryo and use scissors to smoothly cut around the perimeter of the embryo.
   3. Use a Pasteur pipette to underlie the embryo with PBS, using gentle streams to vacate any yolk sticking to the embryo.
      NOTE: This step is critical when working with younger (<HH3) embryos because these embryos tend to stick more to the yolk and will often detach from the vitelline membrane during subsequent washing steps.
   4. Slowly pull the embryo/paper ring at an oblique angle up and off the yolk into a Petri dish filled with PBS for further cleaning. Once most of the yolk has been removed, place the embryo ventral side up on a 35 mm Petri dish covered with a semi-solid mixture of agar/albumen.
3. Prepare six to eight 10 cm-long glass microcapillaries (O.D. = 1.2 mm) by using a glass micropipette puller instrument.
4. Place the embryo ventral side up in the electroporation chamber filled with PBS. Using a glass microcapillary, inject a bolus of 200 nL of the mRNA or DNA/mRNA electroporation mix into the cavity between the epiblast and vitelline membrane covering the desired region.
   NOTE: The entire anterior area for pellucida and some area for opaca was electroporated in most of the experiments shown in this manuscript.
5. Place the positive and negative electrodes (platinum flat square electrode; side length of 5 mm) on top and bottom of the embryo respectively and electroporate using the following pulse sequence: five square electric pulses of 5 V, 50 ms duration with 100 ms intervals using an in vivo electroporator. Ensure that the distance between the electrodes is ~5 mm.
   NOTE: Optimizing the electroporation parameters is crucial to avoid conditions that can kill the fragile embryonic cells. Parameters of voltage, pulse length, pulse intervals, and the number of pulses for DNA and mRNA should be considered for various electroporation devices.
6. Incubate the electroporated embryos at 38 °C to the desired developmental stage.
   NOTE: A fluorescent dissecting stereoscope (see Table of Materials) helps to screen transfected vs. non-transfected embryos.
7. If the embryos are to be statically imaged, fix them in 4% paraformaldehyde in PBS for 1 h at room temperature or overnight at 4 °C.
   1. Remove the embryos from the vitelline membrane by smoothly cutting around the perimeter of the filter paper with scissors and peeling off the shiny membrane on the dorsal surface with sharp forceps gently.
   2. Wash the fixed embryos in PBS/Triton (0.1%) 2x for 5 min and continue with in situ hybridization or immunostaining if desired.
   3. Finally, stain the embryo in 0.5 µg/µL DAPI in PBS/Triton (0.1%) for at least 30 min at room temperature. Wash embryos in PBS/Triton 2x for 5 min and clear the embryo in SCALE-U2 solution²³ overnight.
8. To analyze the efficiency of electroporation (see Figure 2), use the binary and particle analysis tool and DAPI channel on ImageJ to obtain nuclear outlines from all cells in the image.
   1. To use the binary tool on ImageJ, use a single z-slice in the DAPI channel containing a majority of cells and click Process > Binary > Make Binary. To separate nearby cells, click Process > Binary > Watershed. Obtain cell outlines by clicking Analyze > Analyze Particles, size set to 100-500 (µm²).
   2. Ensure that a majority of the cells are outlined in the DAPI channel and save the cell outlines by clicking More > Save on the ROI manager pop-up window.
   3. Use these outlines to obtain fluorescence intensity values for the mRNA and DNA channel by opening the previously saved file on a single z-slice in the mRNA or DNA channel and then click Measure on the ROI manager.
   4. Finally, filter these intensity values, counting cells with <6,000 fluorescence intensity as non-transfected and cells with >6,000 fluorescence intensity as transfected.

5. Image FPs Encoded by Electroporated mRNAs

1. Choose the healthiest and best electroporated embryo for the dynamic imaging experiments after looking at all electroporated embryos under the fluorescent dissecting stereoscope.
   1. Continue to incubate the other electroporated embryos and the non-electroporated embryo (the negative control) in a separate incubator while imaging the chosen embryo in case this embryo dies during the experiment.
2. For the dynamic imaging, use the whole-mount ex ovo avian embryo culture as previously described^24,25,26 with an inverted confocal microscope.
   NOTE: The microscope is equipped with an onstage incubator (see Table of Materials) that maintains the temperature at 36 °C during imaging. It was observed during the microscope set-up that embryos incubated at 36 °C survive longer than at higher temperatures, possibly because the laser may cause local heating on the embryo. Readers should determine the optimum on-stage incubation temperature for their own microscope set-up.
   1. To dynamically image and visualize embryogenesis, clean the electroporated embryo briefly with PBS to remove any bubbles that may form on the dorsal side of the embryo during the electroporation process by moving the embryo around using forceps in a PBS clean solution.
   2. Place the clean embryo directly onto an imaging dish containing a thin layer of albumen-agar (~150 µL) making sure not to generate any bubbles on the dorsal surface of the embryo²².
   3. To ensure survival for long-term imaging, add a small moist rolled-up piece of tissue paper at the inside edges of the imaging dish and seal the dish using paraffin film to minimize evaporation during the imaging and incubation.
   4. Move this dish quickly to the pre-warmed stage of a confocal microscope and locate the colored dye in the embryo using the brightfield channel (PMT laser 20%), which identifies the injected and electroporated region.
3. Set imaging software to the desired objective (10 or 20x), dichroic mirror (488 nm for GFP nm, 561 for RFP), emission spectra (499-562 nm for GFP, 570-695 nm for RFP), and turn on an appropriate laser (488 nm for GFP, 561 nm for RFP).
   NOTE: Electroporated mRNA was translated into proteins that were seen within 20 min (see Figure 3). The imaging metadata used for most of the images in this paper were: an inverted confocal microscope with a 20x objective (see Table of Materials); pixel dwell time, ~1.5 μs; mean of 4-line scans.
   1. Click Live on the imaging software and adjust the laser power to a setting that is appropriate for the fluorescence intensity depending on each microscope laser power. Start by imaging the embryo using 1% laser power, 800 gain, and increase the laser power slowly by 1% increments until saturated pixels are seen.
   2. Follow this by decreasing the laser power slightly until no saturated pixels are seen anymore.
      NOTE: The laser power chosen for the beginning of an imaging session of an embryo may be good for earlier time points but not ideal at later time points if the fluorescence of the cells becomes much brighter or dimmer over time. To address this, image the embryo at slightly lower power settings initially since the electroporated cells generally get brighter over the first 6 hours after electroporation (see Figure 3A-E for quantification of signal increase). If the later images are saturated, continue to image with the original imaging settings, but take an additional image immediately after with weaker imaging settings (smaller pinhole or weaker laser power).
   3. Image the embryo every 3-5 min in order to track individual cell migration across different time points. For this work, the images were z-stacks (~50 µm thick) of the entire electroporated area, plus some extra room towards the bottom of the z-stack in case the embryo sinks into the agarose bed throughout the imaging session.
   4. Observe how fast the cells are moving by checking the first few time points of the first movie. If the cells are moving at a fast rate (meaning that they will exit the area of the imaged region within a couple more time points), consider expanding the zoom of the imaged area (1x à 0.8x) or imaging a different region.
      NOTE: Embryonic regions in the area pellucida move much more rapidly than those in the area opaca. Additionally, younger embryos (HH3, HH5) often contain cells that are undergoing more rapid movement compared to older embryos (>HH7).
   5. After imaging the electroporated region of the embryo, image an un-electroporated region of the same embryo to determine autofluorescence levels (should be minimal if low laser power <10% is used to image the embryo).

6. Fluorescence Recovery After Photobleaching (FRAP) to Assay mRNA Integrity

NOTE: An in vivo fluorescence recovery after photobleaching (FRAP) assay can be used to determine how long transfected mRNA could be translated into FPs. The following protocol outlines a FRAP experiment to detect the half-life of H2B.Citrine mRNA in an electroporated embryo.

1. Perform FRAP experiments on the inverted confocal microscope using a 20x 0.8 NA objective and a completely open pinhole.
   1. After confirming electroporation of H2B-Citrine on the stereomicroscope and setting the embryo on the pre-heated stage on the inverted confocal microscope (see step 5.2.4), photobleach most of the cellular fluorescence from H2B-Citrine at a variety of time points (45 min, 2 h, and 5 h post-electroporation) by using the 405 nm laser with 70% laser power, 100 iterations, scan speed 4, which leaves only 5% of fluorescence remaining.
      NOTE: This process should take a couple of minutes.
   2. Continue to incubate the embryos on the stage at 36 °C after photobleaching.
2. Be sure to pay attention to actively dividing cells within the photobleached region, which indicate that the photobleached cells have not completely died post-treatment.
3. Acquire post-bleach images (z-stacks of the electroporated region) for up to 30 min at regular time intervals (3 or 5 min) using the confocal microscope.
   NOTE: If available, use a transgenic H2B-XFP line as a positive control for ensuring cell survival in the photobleached region. The rate of fluorescence recovery for the FP encoded by the electroporated mRNA should decrease, but that for the FP encoded via transgene should remain consistent throughout the entire movie.
4. To ensure that the imaging conditions do not deleteriously affect embryo survival, concurrently incubate electroporated embryos that are not photobleached, which may serve as a control for imaging.
5. To quantitate the photobleaching results for mRNA decay post-electroporation, track the cell fluorescence over time (3 or 5 min) by measuring the fluorescence intensity of the center (a 7.5 µm circle) of each cell using ImageJ. Measure this for all cells within the photobleached region that are not undergoing mitosis and have been completely photobleached.
   NOTE: Consider omitting the mitotic cells from the quantification since mitotic nuclei have stronger fluorescence than interphase nuclei due to chromatin condensation.
6. Plot the fluorescence intensity over time post-bleach at a variety of time points (45 min, 2 h, and 5 h).

Results

mRNA electroporation is more efficient than DNA electroporation

We used pCS2+.H2B-Citrine to prepare in vitro transcribed mRNA. Since DNA electroporation is usually performed at 1-2 µg/µL, we used an equimolar concentration of mRNA (calculated to be around 0.25-0.5 µg/µL for H2B-Citrine) for mRNA electroporation. We first tested the electroporation efficiency of pCS2+.H2B-Citrine DNA compared to H2B-Citrine mRNA...

Discussion

In this protocol, we provided step by step instructions on how to precisely microinject and electroporate mRNA into the cells of gastrulating quail embryos. We demonstrated that in vitro synthesized mRNA electroporation allows fast and efficient expression of fluorescent proteins (FPs) in gastrulating quail embryos (Figure 2 and 3). Fluorescence from H2B-citrine protein translated from electroporated mRNAs could be detected by confocal microscopy within ~20 min and increased...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
BamHI-HF	New England Biolabs	R3136L
BglII	New England Biolabs	R0144S
BsrG1-HF	New England Biolabs	R3575S
NotI-HF	New England Biolabs	R3189L
SalI-HF	New England Biolabs	R3138L
Phenol:Chloroform:Isoamyl Alcohol	Thermo Fisher	15593031
SP6 mMessage Machine in vitro transcription kit	Thermo Fisher	AM1340
Fast Green FCF	Sigma Aldrich	F7252
Triton X-100	Sigma Aldrich	93443	4-(1,1,3,3-Tetramethylbutyl)phenyl-polyethylene glycol, t-Octylphenoxypolyethoxyethanol, Polyethylene glycol tert-octylphenyl ether
DAPI	Sigma Aldrich	D9542	2-(4-Amidinophenyl)-6-indolecarbamidine dihydrochloride, 4′,6-Diamidino-2-phenylindole dihydrochloride, DAPI dihydrochloride
Whatman No.1 filter paper	Sigma Aldrich	WHA1001125
glycerol	Sigma Aldrich	G9012
Urea	Sigma Aldrich	51457
pmTurquoise2-Golgi	Addgene	36205	pmTurquoise2-Golgi was a gift from Dorus Gadella (Addgene plasmid # 36205 ; http://n2t.net/addgene:36205 ; RRID:Addgene_36205)
pmEGFP-N1-LifeAct			Nat. Methods 2008;5:605-7. PubMed ID: 18536722
pCS2.Lifeact-mGFP	Addgene		This paper
pCS.H2B-citrine	Addgene	53752	pCS-H2B-citrine was a gift from Sean Megason (Addgene plasmid # 53752 ; http://n2t.net/addgene:53752 ; RRID:Addgene_53752)
pCS.memb-mCherry	Addgene	#53750	pCS-memb-mCherry was a gift from Sean Megason (Addgene plasmid # 53750 ; http://n2t.net/addgene:53750 ; RRID:Addgene_53750)
Zeiss LSM-780 inverted microscope	Carl Zeiss Microscopy GmbH		The LSM-780 is a confocal and multi-photon microscope that offers the sensitivity required for vital imaging work. Equipped with a motorized stage, an autofocus device, and a full stage-top blackout incubator, the 780 is an excellent microscope for high-end live cell/embryo imaging. The high-sensitivity 32-channel Quasar detector allows for spectral imaging, linear unmixing, and high color count (>4) image acquisition. Excitation can be performed with 6 lines single photon lasers (405, 458, 488, 514, 564 and 633 nm), Chameleon (Coherent) 2-photon laser (range from 690nm to 1000nm), and run with ZEN 2011 SP7 (Black) system software.
CUY-21 EDIT in vivo electroporator	Bex Co., Ltd.
Platinum flat square electrode, side length 5 mm	Bex Co., Ltd.	LF701P5E
Olympus MVX10 FL Stereo Microscope	Olympus LifeScience
XM10 Monochrome camera	Olympus LifeScience
Phosphate-Buffered Saline (PBS) for HCR (10×, pH 7.4)			To prepare 1 L of a 10× stock solution, combine 80 g of NaCl (Sigma-Aldrich S3014), 2 g of KCl (Sigma-Aldrich P9541), 11.4 g of Na2HPO4 (anhydrous; Sigma-Aldrich S3264), and 2.7 g of KH2PO4 (anhydrous; Sigma-Aldrich P9791). Adjust the pH to 7.4 with HCl, and bring the final volume to 1 L with ultrapure H2O. Avoid using CaCl2 and MgCl2 in PBS for HCR. It is important that the PBS for HCR is prepared as an RNase-free solution (e.g., via diethylpyrocarbonate [DEPC] treatment).
1.37 M NaCl
27 mM KCl
80 mM Na2HPO4 20 mM KH2PO4
PBS/Triton			Add 1 mL of Triton X-100 (Sigma Aldrich 93443) and 100 mL of 10× PBS to 890 mL of ultrapure distilled H2O. Filter the solution through a 0.2 μm filter and store it at 4 ?C until use.
1× phosphate-buffered saline (PBS) (DEPC-treated; pH 7.4)
0.1% Triton X-100