Use of chick neural tube for optimizing the PSM and epithelial somites electroporation parameters: A detailed protocol

Somite myogenesis is one of the crucial early embryonic events that lead to the formation of muscular tissue. A complex of dynamic gene regulatory networks masters this event. To understand and analyze these networks, there remains a genuine need for the use of a reproducible and highly efficient gene transfer technique. In vivo electroporation has proven to be amongst the best approaches in achieving a high level of gene transfer. However, unoptimized electroporation conditions can directly cause varying degrees of cellular damage which may induce abnormal embryonic development as well as changes in the endogenous gene expression. Presegmented mesoderm and epithelial somites are not easy to electroporate. Chick neural tube has served in many functional studies as an ideal experimental model organ which is both robust and easily manipulated. In the current detailed protocol, the neural tube was used as a tool to optimize the electroporation conditions which were subsequently applied in the electroporation of the presegmented mesoderm and epithelial somites. The protocol highlights important notes and hints that enable reproducible results and could be applied in the in vivo electroporation of other chick embryo tissues.


INTRODUCTION
Somite formation is a complex of developmentally orchestrated cell signaling that leads to skeletal muscle and vertebral column formation. Somite myogenesis participates in the formation of ribs, cartilage, tendons, ligaments and limb buds. This process begins by segmenting a solid rod of mesoderm, the presegmented mesoderm (PSM) from which a new epithelial somite (ES) is produced every 90 min. Once a somite is formed, it starts to differentiate from the caudal to the rostral direction to form dorsal (epaxial) and ventral (hypaxial) domains [1]. The dorsal domain gives rise to the dermis and skeletal muscles while the sclerotome migrates to give rise to the vertebrae, smooth muscle cells, fibrocytes, chondrocytes, osteocytes and endothelial cells [2].
A considerable number of studies investigating gene expression and the governing regulatory networks during somite myogenesis have been carried out (reviewed in [3]). However, these networks are yet to be completely understood [4]. Gain-and loss-of-gene expression using either in vivo and/or in vitro electroporation is a very powerful approach for manipulating gene function [5]. This includes electroporating single (aptamer) [6] or double [7] strand DNA, microRNA inhibitor (antagomir) [8], siRNA [9], morpholino (MO) [10], and Crispr-Cas9 [11] into the whole embryo, the neural tube (NT), neural crest, somites, retinal explants, and brain. However, gene transfer by electroporation has few main limitations which include direct damage to the cells, difficulty in controlling the number of electroporated cells, and possible off-target gene transfer. Some of these disadvantages could be controlled through a suitable optimization of the electroporation conditions [12].
Amongst animal model systems, chick embryo offers many advantages as a simple, experimental, and manipulative embryonic model for the study of different aspects of development. Advantages of the chick embryo include a high level of similarity with the human genome, and the low cost and ease by which it can be obtained and developed in the laboratory. In vivo chick embryo electroporation approach has been used to study gene function and regulation during somite myogenesis [13]. However, from our own experience and as has been reported by Wang and his colleagues, targeting muscle cells is not easy at any stage of Tungsten needles Sharpen the Tungsten wire using a micro-torch flame then insert the sharpened-wire into a glass rod by slightly melting the glass using the flame. The Tungsten needles should be kept in a dissecting tools' box to avoid damage to the needles' fine tips. Tungsten needles are used for embryo microdissection and removal of the inner vitelline membrane.

Egg incubation
Incubate chicken eggs at 38°C and 75% humidity to obtain either stage HH12 or HH16 embryos according to Hamburger and Hamilton timetable [17].

HINTS:
(1) Fertilized eggs should be pathogen free; (2) Upon arrival to the lab, store eggs in the BOD incubator at 15°C for 24 h prior incubation to allow the eggs to settle; (3) Eggs stored for more than one-week at 15°C should not be used for electroporation since they usually develop poor quality embryos; (4) Temperature and humidity of the eggs' incubator should be monitored; and (5) Rotation of the eggs during incubation is strongly recommended (to prevent adhe-sion of the egg membranes) which usually leads to producing high quality embryos.

Electroporation station setting up
For setting up the electroporation station ( Fig. 1A): (1) Obtain a commercially available galvanized iron sheet with approximate measurements of 80 cm length, 40 cm width, and 3 mm thickness, and then place the iron sheet in the middle of the electroporation station; (2) Place the stereomicroscope with its associated reflected light units in the middle of the iron sheet; (3) Place a manual micromanipulator (loaded on a mounting magnetic base) on each side of the stereomicroscope, one micromanipulator should hold the injection microcapillary holder and another should carry the electrodes' holder; switch the magnetic controls of the mounting bases "on" to fix them firmly in place on the iron sheet; (4) Place the Eppendorf microinjector (Fig. 1B) on one side of the electroporation station and the electroporator (connected to the current amplifier) (Fig. 1C) on the other side; and (5) Connect the injection microcapillary to the microinjector using the provided tubing and the electrodes to the electroporator using the provided electric wires.

Micropipette pulling
CRITICAL STEP: Using the P97 micropipette puller (Fig. 1D), insert the microcapillary in position (see the P97 manual for illustration) and carry out a ramp test according to the manu-facturer's instructions (see P97 manual). This test determines at which temperature the glass microcapillary should start to melt. At the end of the ramp test, a heating value will be displayed on the puller's screen. Use this heating value to set up a new microcapillary pulling program in which the heat POL Scientific protocol value is set up within the range of +15 to −15 of the ramp test heating value to achieve the optimal microcapillary pulling.
The following program can be used as a guide to pull out the injection microcapillary: Heat = ramp test heat value ± 15, pull: 40, velocity: 200, time: 50 NOTE: Pulling a very fine microcapillary tip is essential for carrying out neat and precise injections.
Electrodes assembly CRITICAL STEP: Assemble two lengths of Platinum wire in an L-shape ( Fig. 2A) with one length serving as a negative (−ve) and another as a positive (+ve) electrode. The current carrying section of the electrodes should be 3 mm long (Fig. 2B) for precise tissue targeting and this can be made by coating the remaining bent part of the electrodes with a nail varnish thus allowing a small window to permit the passing of current between the −ve and +ve electrodes. The distance from the edge to the holding point should be 7 mm long. Solder the assembled electrodes to a black (−ve electrode) and red (+ve electrode) electric wires. Check the conductivity and resistance after soldering. Mount both the −ve and +ve electrodes on the electrodes' holder by which the gap between the electrodes is approximately 3-4 mm. Mount the electrodes' holder onto the left-handed side micromanipulator and fix it using its magnet switch on the left-hand side of the stereomicroscope.

NOTE:
(1) Coating of the electrodes prevents damage to the embryos and increases the precision of the tissue targeting; and (2) Platinum electrodes with Platinum coating can be bought ready-made (BEX, fixed Platinum needle electrode, LF613 series, LF611P7-3).

DNA preparation
Expression vectors carrying GFP or RFP such as pCMV-IRES-GFP ( Fig. 3A) or pCMV-IRES-RFP (Fig. 3B) can serve as a control electroporated DNA. Extract DNA using Qiagen EndoFree Plasmid Maxi kit according to the manufacturer's instructions. Carefully measure the DNA concentration using NanoDrop based on which DNA can be further concentrated by EtOH precipitation to obtain a concentration ranging from 1 mg/ml to a maximum of 3 mg/ml. Wash DNA pellet twice with 70% EtOH to eliminate any salts interference which could lower the electroporation efficiency. DNA at 1-2 mg/ml should give good fluorescence intensity of GFP or RFP after electroporation. A DNA concentration above 3 mg/ml will be too viscous and difficult to inject due to clogging of the microcapillary.

HINTS:
(1) In case of the expression vectors which do not have GFP (such as pCDNA3 vector, Thermo Fisher cat. # V79020), mix with 2-3 µl of 1 mg/ml IRES-GFP vector to enable tracing the injected DNA (for example: add 15 µl of 2 mg/ml pCDNA3 + 3 µl 1 mg/ml pCMV-IRES-GFP); and (2) Centrifuge DNA mix at 12000 rpm for 2 min at room temperature before using it for microinjection.

Morpholino preparation
Dissolve control morpholino (CMO) according to the manufacturer's instructions in double distilled autoclaved water at a concentration of 300 mM.
CRITICAL STEP: MO should be heated up to 60°C for 5 min to denature before injection and then keep on ice during injection. Store after injection at 4°C.

HINT:
If fluorescently-labelled MO (yellow/greenish in color) is used, there is no need to mix it with Fast Green since it will be clearly visible during injection.

Microcapillary assembly
Fill in the microcapillary with DNA/Fast Green mix or fluorescent CMO using Eppendorf microloader fine tip and then mount the microcapillary onto the microcapillary holder. Mount the holder on the right-handed-side micromanipulator and fix it on the right-hand side of the stereomicroscope. Under the stereomicroscope, snip off the tip of the microcapillary using the Watchmaker's forceps No. 5. Test the flow of the DNA or MO out of the microcapillary by continuously pressing the inject button of the Eppendorf FemtoJet microinjector until a bolus of the DNA or MO starts to come out of the needle's tip. If this does not happen, then carefully snip off the tip of the microcapillary again until the DNA or MO bolus can be seen coming out of the needle's tip. Try to keep the needle's tip as fine as possible.

1.1.
Take an egg out of the incubator, and then wipe out the blunt end (air space side) with 70% EtOH, place the egg under the stereomicroscope with its blunt end facing up, and stick a small strip of tape on the eggshell blunt end.

1.2.
Using a pair of small dissecting scissors, cut through the tape and the underneath eggshell to make a circle of 1-1.5 cm diameter egg window.
HINT: Care should be taken while making the egg window not to damage the underneath embryo. This can be achieved by keeping the dissecting scissors in a horizontal position while cutting through the eggshell.
1.3. Add 2-3 drops of PBS/Penicillin-Streptomycin (10 μl/ml) on top of the outer vitelline membrane and then gently pierce the membrane using the Watchmaker's forceps No. 5. PBS should infiltrate through the membrane which then swells and can be easily ruptured and removed by the forceps. The embryo should be immediately exposed after the membrane removal.
HINT: (1) Contamination could be a major problem and hence Penicillin-Streptomycin should be added to the POL Scientific protocol PBS in all steps of the embryo manipulations; and (2) Dissecting tools should be sterilized either with 70% EtOH or by autoclaving before use.

1.4.
Inject Indian ink buffer using 1 ml syringe with 26-gauge needle underneath the blastoderm disc to visualize the embryo.

1.5.
Carefully remove the inner vitelline membrane on top of the embryonic disc at the site where the tissue of interest to be electroporated using a very fine Tungsten needle.

2.
Neural tube electroporation and optimization 2.1. At stage HH16 embryos, make a small egg window as described in step 1, remove the outer and inner vitelline membranes on top of the NT, and then manoeuvre and gently insert the microcapillary containing DNA mix (1-2 mg/ml IRES-GFP/Fast Green) into the NT lumen.

2.2.
Fill in the NT with the DNA, gently take out the microcapillary, and then carefully place the electrodes to sandwich the NT. Add 2-3 drops of PBS/Penicillin-Streptomycin and electroporate the DNA. Add immediately 2-3 drops of PBS to cool down the electroporation site and then gently elevate the electrodes using the micromanipulator.

NOTE:
It is important to take in consideration that careful assembly of the electrodes combined with the critical optimization of the electroporation parameters are essential in producing a highly efficient electroporation.

2.4.
Aspirate some of the egg albumen using 10 ml syringe with 18-gauge needle until the embryo is slightly brought down. Seal the egg with tape and re-incubate until the desired embryo stage is reached. Embryos can be checked at any time during incubation.

NOTE:
(1) Once the embryos are electroporated and re-incubated, rotating the incubator's shelves should be stopped to avoid adhesion of the embryos to the sealing tape; and (2) The egg window should be well-taped to avoid gaps which could lead to leakage of hot air into the embryos which will affect their survival.

2.5.
Check GFP signals after 4 to 24 h of DNA electroporation. IRES-GFP can be detected by Alexa Fluor 488 ηm filter and IRES-RFP by Alexa Fluor 565 ηm filter. After incubation is complete, dissect out the electroporated embryos.

2.6.
Assess the electroporation efficiency based on the following parameters (see also Table S1).

2.6.1.
The overall embryo survival: should be at least 90%.

2.6.2.
The embryo morphology: at least 90% of normal development should be achieved.

2.6.4.
Electroporation can be considered as successful when it results in a high survival rate, and normal morphology but associated with strong or very strong GFP fluorescence intensity.

HINTS:
(1) If the electroporation is producing kinky embryos, then this suggests that the electroporation conditions are not properly optimized. The voltage (v) could be lowered as well as the number of pulses until normal embryo development is achieved; and (2) Take images before fixation since PFA fixative could cause auto-fluorescence and thus gives false signals.

3.1.
Manipulate stage HH12 embryos as described in step 1, and then manoeuvre the microcapillary containing DNA mix or MO to approach the PSM. Carefully insert the microcapillary fine tip into the rostral POL Scientific protocol domain of the PSM. Inject the molecule to be electroporated which should spread in a rostrocaudal direction of the PSM until it is filled ( Fig. 4A and 4B).

3.2.
Place the electrodes gently to sandwich the PSM and then add 2-3 drops of PBS/Penicillin-Streptomycin before DNA is electroporated. Apply the optimized parameters obtained from the NT electroporation to electroporate the PSM (Fig. 4B and 4C).

4.1.
At stage HH16, manipulate the embryos as in step 1, manoeuvre the microcapillary to approach the first ES and gently insert the tip to pierce the somite epithelium until it reaches the somitocoel and then inject the DNA. Similarly, inject the rest of the ESs to be electroporated. Gently place the electrodes to sandwich the somites, add 2-3 drops of PBS and electroporate the epaxial domain (Fig. 5A-5C).
If the hypaxial somite domain is to be electroporated, then similarly inject the somite but reverse the current as shown in Fig. 5D-5F.
HINT: Keep the tip of the microcapillary as fine as possible during the injection which not only helps in reducing any possible damage to the somite but will also minimize any leakage of the injected DNA.

4.2.
After incubation is completed, dissect out the electroporated embryos and then assess the embryos' survival, normal morphology and GFP signals intensity as described in step 2.6.

5.1.
Using a pair of dissecting scissors, carefully remove the electroporated embryos in a Petri-dish containing DEPC-PBS. Carefully transfer the embryos using a 5 ml plastic Pasteur pipette with its tip cut off.

5.2.
Remove the embryonic membranes using Watchmaker's forceps No. 5 before capturing images.

5.5.
mRNA probe synthesis for ISH can be carried out as described in Geisha protocols [16].

ANTICIPATED RESULTS AND DISCUSSION
The current protocol consists of two stages (see the graphical abstract): In the first stage, the NT of HH16 chick embryo (Fig. 6A) was used as a tool to optimize the electroporation conditions. To achieve this, we initially tested a number of voltages (15, 20, 25, 30, and 35). Each voltage was combined with 5 pulses (p), 30 ms pulse space (ms/ps) and 100 ms pulse width (ms/pw). After each experiment, the efficiency of the electroporation was assessed as described in step 2.6 by which strong to highly strong GFP or RFP signals (level 3) constituted efficient or successful electroporation. For each voltage we tested, 50 embryos were electroporated (total n = 250 embryos for all voltage trials). The results revealed that only 25 v combined with 5 p, 30 ms/ps and 100 ms/pw led to achieving level 3 of GFP expression. After this initial electroporation parameters testing, we applied 25 v, 5 p, 30 ms/ps, and 100 ms/pw to electroporate the NT in three separate experiments to ensure reproducibility (total n = 147 embryos). Each experiment resulted in a survival rate and normal development of at least 96% of the electroporated embryos (total n = 141). This was combined with level 3 of GFP signals intensity in 95% of the embryos (n = 139) (Fig.  6B). According to the criteria established in this protocol, we considered these results to represent efficient electroporation. showing strong GFP signals in the NT, note the overall normal morphology of the embryo. e: eye, h: heart, fl: fore-limb, hl: hind-limb, mb: mid-brain.
Number of pulses was another important parameter to test during the electroporation optimization stage. A varying number of pulses (3, 4, 5, and 6) in a combination of 25 v, 30 ms/ps, and 100 ms/pw were applied to electroporate the NT. Three (n = 46 embryos) and four (n = 41 embryos) pulses resulted in either absence (level 0) or weak (level 1) GFP signals. Five pulses were tested as mentioned above. Six pulses (n = 43) resulted in very strong GFP signals but this was associated with either a low survival rate and/or kinky embryos. At the end of this optimization stage, we concluded that 25 v, 5 p, 30 ms/ps, and 100 ms/pw were the best electroporation parameters to apply in the electroporation of the PSM and ES.
In stage 2, we applied the optimal NT electroporation conditions to electroporate MO into the PSM. This showed efficient electroporation results similar to those produced by NT electroporation using 25 v, 5 p, 30 ms/ps, and 100 ms/pw program. This was indicated by strong GFP signals (level 3) after 3-4 h of electroporation (Fig. 7A) (total n = 121, 94%, three experiments). Green fluorescent MO was easily detectable later in the ES (Fig. 7B). PSM electroporated embryos also showed normal morphology in at least 93% (total n = 112). In addition, the effects of the electroporation on the endogenous gene expression were checked by ISH which showed normal Myog expression (Fig. 7C as an example, 100%). The non-injected but electroporated left side of the embryo served as the internal control.

POL Scientific protocol
The electroporation program (25 v, 5 p, 30 ms/ps, and 100 ms/ pw) applied to the NT and PSM was used to electroporate the epaxial domain of the ES (total n = 152, three experiments) (Fig. 8A). These conditions showed reproducible and efficient results indicated by strong (level 3) GFP signals ( Fig. 8B and 8C) (94%, total n = 142). When the electroporated embryos were incubated for up to 3 d, they showed strong GFP expression with no signs of abnormal development (Fig.  8D). Similarly, efficient electroporation results were obtained when IRES-RFP was electroporated (Fig. 8E) (total n = 88, 92%, three experiments). The effects of IRES-RFP electroporation on the endogenous MyoD expression was checked by ISH which showed normal expression pattern in both electroporated and injected somites as well as the electroporated non-injected control side (Fig. 8F).
In vivo electroporation is based mainly on applying current to the tissue of interest at a low voltage to cause a temporary perforation of the cell membrane to allow uptake of exogenous molecules. Such method proved to be a powerful approach for gene transfer and functional studies. Our results showed that the NT is highly suitable for electroporation due to its inherent structural advantages, including a narrow lumen that can retain the DNA after injection, as well as a dorsoventral structural depth-making it highly receptive to the current. These advantages allowed us to utilize this robust system to easily optimize the electroporation parameters and to apply these parameters to electroporate the PSM and ES. However, additional parameters including proper electrodes' assembly, microcapillary assembly, concentration of the injected molecule(s), and purity and quality of the injected DNA should be all considered to achieve efficient electroporation.
The current approach to optimize the electroporation parameters using the NT is not limited to electroporating the PSM and ES. Our previous work has showed that the same approach for optimizing the electroporation conditions can be applied to electroporate other tissues such as the epibranchial placodes [18] and surface ectoderm [20]. Furthermore, we previously used the same approach to successfully electroporate more than one molecule such as two morpholinos, and a mix of double strand DNA and MO [13]. In conclusion, the current protocol may be of use in the electroporation of embryonic tissues which are considered to be "difficult to electroporate" such as the somites, optic and auditory vesicles, epibranchial placodes, and heart. ES: epithelial somites; fb: fore-brain; fl: fore-limb; h: heart; hb: hind-brain; hl: hind limb; mb: mid-brain; nt: neural tube; psm: presegmented mesoderm.

POL Scientific
protocol TROUBLESHOOTING Potential problems that could arise during the electroporation protocol and suggestions to troubleshoot them are listed below in Table 1.