Name: chicken embryo spinal cord slice culture protocol
Uploaded: 2013-03-25T13:00:00
Duration: 10 min 57 s
Description: Watch this Scientific Journal Video about Chicken Embryo Spinal Cord Slice Culture Protocol at JoVE.com

We would like to thank Sharon Boast for many insightful discussions and B. Novitch for the kind gift of the antibodies to Foxp1. This work as funded by a studentship from the Wellcome Trust to KCT and a project grant from the Wellcome Trust to SRP (Grant reference number 094399/B/10/Z).

1. Make Required Solutions
 <ol>
 <li>1 M Phosphate buffer: Weigh out 109.4 g Na2HPO4 and 32 g NaH2PO4.H20, mix and dissolve in water to total volume of 1 liter.</li>
 <li>Phosphate Buffered Saline (PBS): Prepare 0.1 M Phosphate buffer, 0.15 M NaCl. 100 ml of PBS should suffice for the processing of slices from 10 embryos.</li>
 <li>Agarose: Heat PBS solution to around 70 &deg;C and add solid low melting temperature agarose slowly whilst stirring continuously. Make the solution to a final concentration of 4% (w/v) agarose. Make a stock of 50 ml of this solution. Leave this solution in a waterbath kept at 48 &deg;C until required. Any agarose solution unused after the experiment can be stored at 4 &deg;C for several weeks.</li>
 <li>Antibody block solution: Add foetal calf serum and triton X-100 to a final concentration of 1% (v/v) foetal calf serum, 0.1% (v/v) Triton X-100 in PBS. 10 ml of block solution will suffice for the immunostaining of 5 slides of cryostat sections.</li>
 <li>Fixative: Prepare 0.75 mm NaOH, 4% (w/v) paraformaldehyde (CAUTION). To prepare fixative heat required amount of water to 70 &deg;C, add concentrated NaOH to required final concentration, add solid paraformaldehyde and mix until dissolved. Cool on ice to 4 &deg;C. 10 ml of this solution will suffice for 20 wells of embryo slices.</li>
 <li>Sucrose Solution: Prepare 10 ml of a 30% (w/v) sucrose solution containing 0.1 M Phosphate buffer. 10 ml of this solution will suffice for 10 wells of embryo slices.</li>
 <li>70% (v/v) Ethanol. Dilute 70 ml of absolute ethanol with 30 ml of water.</li>
 <li>Culture Medium: Prepare a solution of 1% chicken embryo extract, 1% Penicillin/ Streptomycin, 0.35% L-Glutamine, 0.1% 2-mercaptoethanol, 4% chicken serum, 2% B27 supplement and 100 ng/ml ciliaryneurotrophic growth factor (CNTF) all diluted in neurobasal medium; prepared on ice and used the same day. 10 ml of culture medium is required for 20 wells of embryo slices.</li>
 </ol>
 2. Chick Embryo Preparation
 <ol>
 <li>Place fertilized hens eggs with the longest side horizontally in a forced draft incubator at 38 &deg;C until they develop to stage15 24. This usually requires approximately 4 days of incubation.</li>
 <li>On the second day of incubation, sterilize the egg shell by wiping with a tissue paper soaked in 70% ethanol. Carefully pierce the flat end of the egg shell using a 21 gauge needle attached to a 5 ml syringe and remove 5 ml of albumin. This lowers the embryo away from the shell so the embryo is not damaged when the shell is opened. Return the eggs to the incubator and incubate them for a further 2 days.</li>
 <li>Using blunted dumont #5 forceps carefully pierce the top middle of the shell and remove around 9 cm2 of shell. Cut around the embryo and carefully lift out the embryo and place in HBSS during dissection. The embryos should be staged according to Hamburger and Hamilton (1992)15. All embryos used should be at close to stage 24. Any embryos that show any signs of abnormality should be discarded.</li>
 <li>Remove the amnion membrane, chorio-allantoic membrane, the head, and allantois using two dumont #5 forceps. Eviscerate the embryo to remove all internal organs. To do this, cut open the ventral midline of the embryo with micro dissecting scissors, hold the embryo around the rostral trunk region with one pair of forceps, grab the rostral part of the gut and the heart and pull caudally. It is important that this is done cleanly. You should be able to see the embryo somites clearly. Cut the embryo in half, transversely between the upper and lower limb buds.Now trim the thoracic body wall using microdissection scissors.</li>
 </ol>
 3. Embryo Slice Preparation
 <ol>
 <li>Remove the agarose solution from the water bath. Cut 5 cm from the bottom of a disposable 3 ml plastic Pasteur pipette.</li>
 <li>Gently remove the lumbar half of the dissected embryo using two forceps to cradle the embryo out of the dissection solution and place in a dry Petri dish. Using the Pasteur pipette, remove around 2 ml of agarose, pipette approximately 0.5 ml on top of the embryo and then suck up the embryo slice into the pipette. Transfer the embryo and agarose to a peel away plastic mould take care not to introduce any bubbles in the agarose. Gently lower the embryo to the bottom of the agarose in the plastic mould with the thoracic section facing downwards. The embryo should be positioned to be straight using a 21 gauge needle. The slices will also contain part of the developing limb and it is important that the orientation of the embryo in the agarose will allow this. Place the boat on ice to set the agarose. Take care that the embryo does not change orientation during this period (around 2 min).</li>
 <li>The Leica VT1000S vibratome (Speed 3, Frequency 4, 400 &mu;m thick slices) should be set up with the slicing chamber filled with HBSS and surrounded by ice-cold water. The agarose block is then stuck to the vibratome platform with super glue. The agarose block is trimmed with a razor blade to leave the embryo piece with 1-3 mm of agarose surrounding it. The slices that include limb bud sections with a clear apical ectodermal rigde are selected and these slices are then placed in HBSS. Generally, there are only one to two suitable slices per embryo. Gently remove the agarose around the tissue slices using two needles. It is essential that this is done thoroughly and carefully.</li>
 </ol>
 4. Culturing Embryo Slices
 <ol>
 <li>Add 500 ml of culture solution (outlined above) to the required number of wells of a 24-well ultra low attachment treated tissue culture plate. Transfer the slices carefully from the HBSS solution into the well. In general, we try to have two to three slices in each well. Place the 24-well plate in the tissue culture incubator at 37 &deg;C with 5% CO2 for 24 hr.</li>
 </ol>
 5. Sectioning the Slices for Immunostaining
 <ol>
 <li>Following the culture period, add 500 &mu;l of unbuffered fix to each well of embryo slices and leave on ice for 20 min. The total solution is then removed carefully and three PBS washes are carried out, with 5-min intervals. The slices are then left in sucrose solution at 4 &deg;C, until the slices sink, around 6 hr but can be left for longer (e.g. overnight).</li>
 
 <li>Remove the slices and dab off extra sucrose solution and equilibrate in around 1 ml O.C.T for around 5 min at 20 &deg;C. Mount each slice flat in an O.C.T.-filled peel away plastic mold. After mounting, place the molds vertically on dry ice to solidify.</li>
 
 <li>Mount the O.C.T. blocks into the cryostat and equilibrate the blocks at -24 &deg;C for around 20 min. Cut each slice with 15 mm sections mounted on Superfrost-Plus slides. Slides can be stored at -80 &deg;C until they are needed.</li>
 </ol>
 6. Immunofluorescence
 <ol>
 <li>Pipette 1-2 ml of PBS on the horizontal slides held in a humidified chamber with the slides raised from the bottom of the chamber (we use 1 or 2 ml plastic pipettes fixed with autoclave tape). Incubate the sections in PBS for 5 min at 20 &deg;C to remove excess O.C.T. Remove the PBS solution from the slide and add 500 ml of block solution over the sections, incubate for 30 min at 20 &deg;C. Remove the block solution and immediately add 500 ml of diluted primary antibody added to the slides, incubate at 4 &deg;C for 18-22 hr.</li>
 <li>Remove primary antibody solution and immediately wash the slides with 1 ml of PBS for 5 min, three times at 20 &deg;C to remove excess primary antibody solution. Add 500 ml of diluted secondary antibody over the sections. The slides are then left at 20 &deg;C for 30 min. The secondary antibody solution is then removed and the slide washed 3 times 5 min in PBS at 20 &deg;C.</li>
 <li>Following these washes, remove the final PBS wash, dab the edge of the slide to remove excess PBS, add two drops of vectashield on the slides and gently lay a glass coverslip over the sections, avoiding bubble formation. Remove excess Vectashield by blotting from the edge of the slides before imaging.</li>
 </ol>
 7. Imaging
 <ol>
 <li>Image the immunostained sections. We use a Nikon Eclipse E80i fluorescence microscope equipped with a Hamammatsu ORCA ER digital camera and 10 X, 20X and 40 X objective lenses.</li>
 </ol>

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryo+slices.">Embryo slices.</a> Methods in Cell Biology. 51 (51), 109-124 (1996).</li><li>Hotary, K. B., Tosney, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+interactions+that+guide+sensory+and+motor+neurites+identified+in+an+embryo+slice+preparation.">Cellular interactions that guide sensory and motor neurites identified in an embryo slice preparation.</a> Developmental Biology. 176 (1), 22-35 (1996).</li><li>Krull, C. 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None of the authors have competing interests or conflicts of interest.

The protocol described here allows a chicken embryo slice to be incubated for more than one day whilst keeping motor neurons alive and continuing to migrate during the culture period. In elucidating the conditions to keep motor neurons alive, we have identified several key features required. It seems that up to 4% chick serum is critical to the survival of motor neurons and its replacement with serum from a different species is not tolerated. Interestingly, we found that the presence of higher concentrations of serum were detrimental to the slices as they promoted overgrowth of the tissue leading to gross distortions in the slice shape. More importantly, the presence of cilliary neurotrophic factor also proved critical. It remains a distinct possibility that the slices may be able to be cultured for considerably longer periods than just 24 hr, perhaps even allowing the visualization of sensory afferent input to the spinal cord. Additionally, the culture conditions here may also prove useful to slice cultures of the developing brainstem and midbrain/ cerebellum.
	Perhaps the biggest potential benefit in the use of these slice culture conditions is that they facilitate the possibility for pharmacological manipulation of protein function whilst minimizing the potential for adverse effects on the rest of the embryo, such as the heart. A large number of inhibitors and agonists of different signalling pathways can be used to assess the migration of different spinal neuron subtypes and, potentially, their effect of the formation of local spinal circuits during development. Additionally, some genetic perturbations of protein expression, such as those that make use of siRNAs and morpholino oligonucleotides, suffer from the fact that they can only be introduced early in development (owing to limitations in in ovo electroporation procedures) and the effects only last for a short period of time (typically one day). Our slice culture is started at later stages and affords the possibility to use this technology later in development at the slicing stage to view their effects during the period of culture of the slice.
	The slice culture protocol could also allow the visualization of both spinal motor neuron as well as dorsal interneuron migration in real time via time-lapse video microscopy. This could be achieved using phase contrast microscopy under white light or through the use of fluorescence imaging. If the latter technique were to be used then the introduction of fluorescent labels is needed. This could be achieved either by introduction of fluorescent dyes such as DiI or DiO via either retrograde labelling from the limb or anterograde labelling from the ventricle or in ovo electroporation of constructs to drive fluorescent proteins in a subset of neurons.

During normal embryo development, many different cell types are generated which must migrate from their point of origin to where they will eventually function in the mature organism. Within the developing spinal cord, progenitor cells are located in the ventricular zone at the midline and generate many subtypes of postmitotic neurons and glial cells which must migrate to integrate into functional neuronal circuits required for sensory processing and motor outputs1. Spinal motor neurons that control the contraction of limb muscles have been extensively studied and much is known of the molecules and mechanisms that drive the formation of their different subclasses. However, much less is known of the mechanisms by which motor neurons migrate, segregate and receive synaptic inputs.
	Motor neuron subtype identity is acquired during development in a hierarchical fashion. Motor neuron subtypes can be broadly classed into distinct columns, divisions and pools which are defined by their axon projections. Motor columns project axons to either axial, visceral or limb muscles. Motor divisions subdivide the limb projecting motor neurons of the lateral motor column (LMC) into those projecting to ventral or dorsal muscles2. Within the divisions, clusters of motor neurons termed motor pools project to individual muscles in the limb2, 3. Limb-projecting motor neurons are defined by their expression of the transcription factor Foxp14, 5, whereas divisional identity is characterized by the expression of the homeodomain transcription factors Islet-1 (ventrally projecting) or Lhx-1 (dorsally projecting)6. Indeed, Lhx-1 expression is required for the appropriate dorsal projections of the motor neurons7. Each of these subtypes occupy distinct positions within the spinal cord; ventral projecting motor neurons come to reside medial to dorsally projecting motor neurons. This results in the LMC being segregated into so called LMCmedial (LMCm) and LMClateral (LMCl) divisions. Divisional and motor pool organization is acquired in two phases8. In the first phase, divisional segregation is achieved via a migration of motor neurons in a characteristic medial to lateral fashion. The LMCl cells are generated after the LMCm cells and so LMCl migrates through the LMCm to achieve its final settling position. The second phase takes motor neurons within a division and clusters them into motor neuron pools, presumably via a dorsal-ventral sorting of the motor neurons. Members of the catenin-dependent classical cadherin family have been shown to be crucial to both phases of motor neuron organization8-10. However, how cadherin function controls cell movement is poorly understood.
	The acquisition of columnar, divisional and pool identities requires extrinsic signals that pattern intrinsic expression of transcription factors11. Additionally, around 50% of all motor neurons die via apoptosis during normal development in a process that requires extrinsic signals from the limb, largely through neurotrophic support of motor neuron survival. Thus, motor neuron development requires the appropriate environment to be maintained over a relatively long timecourse. For this reason, the practicalities of generating spinal cord slice cultures to maintain motor neuron survival require the elucidation of appropriate culture conditions. Previous embryo slice cultures have been used to follow the behavior of extrinsically added purified neuronal populations12-14. However, to our knowledge culture conditions that facilitate in situ motor neuron survival and migration within the slice itself have not been published. We modified a standard culture condition that facilitates the survival of purified, dissociated cranial motor neurons to facilitate motor neuron development within an embryo slice culture system. We also monitored the positioning of the surviving motor neurons and demonstrate that largely normal positions of the motor neuron divisions is acquired during the culture period.

<table border="1">
 <tbody>
 <tr>
 <td>Eggs</td>
 <td>Henry Stewart Farms, UK</td>
 <td></td>
 <td>Fertilised Brown Bovan Gold Hen's eggs (Henry Stewart Farms, UK), stored at 14 &deg;C until incubation</td>
 </tr>
 <tr>
 <td>21 Gauge needle</td>
 <td>BD-Microlance-3</td>
 <td> 304432</td>
 <td></td>
 </tr>
 <tr>
 <td>5 ml syringe</td>
 <td>BD Plastipak</td>
 <td> 302187</td>
 <td></td>
 </tr>
 <tr>
 <td>#5 Dumont forceps</td>
 <td>Roboz</td>
 <td> RS 5045</td>
 <td></td>
 </tr>
 <tr>
 <td>Micro Dissecting scissors</td>
 <td>Roboz</td>
 <td> RS 5910SC</td>
 <td>(3.5 in straight)</td>
 </tr>
 <tr>
 <td>Embryo embedding moulds: Peel away moulds</td>
 <td>Agar Scientific</td>
 <td> G3764</td>
 <td>Pyramidal shaped 22 mm to 12 mm</td>
 </tr>
 <tr>
 <td>Super glue</td>
 <td></td>
 <td></td>
 <td>Power Flex, Loctite</td>
 </tr>
 <tr>
 <td>Ethanol</td>
 <td>SIGMA</td>
 <td> 32221</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium di-hydrogen Phosphate, mono-hydrate</td>
 <td>BDH</td>
 <td> 102454R</td>
 <td></td>
 </tr>
 <tr>
 <td>Di Sodium mono-hydrogen Phosphate</td>
 <td>VWR/ BDH</td>
 <td> 102494C</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium Chloride</td>
 <td>VWR/BDH</td>
 <td> 27810.295</td>
 <td></td>
 </tr>
 <tr>
 <td>Low melting temperature agarose</td>
 <td>Invitrogen</td>
 <td> 16520</td>
 <td></td>
 </tr>
 <tr>
 <td>Fetal Bovine serum</td>
 <td>Gibco</td>
 <td> 10500-064</td>
 <td></td>
 </tr>
 <tr>
 <td>Triton X-100</td>
 <td>SIGMA</td>
 <td> T8787</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium Hydroxide</td>
 <td>FLUKA</td>
 <td> 71689</td>
 <td></td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>VWR/BDH</td>
 <td> 28794.295</td>
 <td></td>
 </tr>
 <tr>
 <td>Plastic disposable pasteur pipettes</td>
 <td>ElKay Precision laboratory consumables</td>
 <td> 127-P503-000</td>
 <td>Liquipette 3 ml volume</td>
 </tr>
 <tr>
 <td>Hanks Balanced Salt Solution (HBSS)</td>
 <td>Gibco</td>
 <td> 14170</td>
 <td></td>
 </tr>
 <tr>
 <td>Sucrose</td>
 <td>SIGMA</td>
 <td> S0389</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium Hydrogen Carbonate</td>
 <td>SIGMA</td>
 <td> S5761</td>
 <td></td>
 </tr>
 <tr>
 <td>Chicken Embryo Extract</td>
 <td>Seralabs</td>
 <td> CE650-J
 </td>
 <td></td>
 </tr>
 <tr>
 <td>Penicillin/ Streptomycin solution</td>
 <td>Gibco</td>
 <td> 15070</td>
 <td></td>
 </tr>
 <tr>
 <td>L-Glutamine solution</td>
 <td>Gibco</td>
 <td> 25030</td>
 <td></td>
 </tr>
 <tr>
 <td>2-mercaptoethanol</td>
 <td>Gibco</td>
 <td> 21985</td>
 <td></td>
 </tr>
 <tr>
 <td>Horse serum</td>
 <td>Southern Group Labs</td>
 <td> 9028B</td>
 <td></td>
 </tr>
 <tr>
 <td>B27 supplement</td>
 <td>Invitrogen</td>
 <td> 17504-044</td>
 <td></td>
 </tr>
 <tr>
 <td>Neurobasal medium</td>
 <td>Gibco</td>
 <td>Cat ~ 2103</td>
 <td></td>
 </tr>
 <tr>
 <td>CilliaryNeurotrophic factor (CNTF)</td>
 <td>R and D Systems</td>
 <td> 557-NT-010</td>
 <td></td>
 </tr>
 <tr>
 <td>Chick Serum</td>
 <td>SIGMA</td>
 <td> C5405</td>
 <td></td>
 </tr>
 <tr>
 <td>Primary antibodies</td>
 <td></td>
 <td></td>
 <td>were purchased from Developmental Studies Hybridoma Bank, with the exception of anti-foxP1 antibodies, which were a kind gift from Bennett Novitch of UCLA. Primary antibodies dilutions were Rabbit anti-FoxP1 (1/32000), Mouse anti-Lhx1/2 (4F2) (1/50), Guinea Pig anti-FoxP1 (1/1000). 
 All antibodies were stored at 4 &deg;C.</td>
 </tr>
 <tr>
 <td>Secondary antibodies</td>
 <td></td>
 <td></td>
 <td>Donkey anti-Guinea Pig (cy3) (1:1000) purchased from Jackson Immunoreseach Laboratories Inc. Donkey anti-Mouse (Alexa Fluor 488) (1:1000), Donkey anti-Rabbit (Alexa Fluor 594) (1:1000) purchased from Invitrogen 
 All antibodies were stored at 4 &deg;C.</td>
 </tr>
 <tr>
 <td>24-well plate</td>
 <td>Corning COSTAR</td>
 <td> 3473</td>
 <td>ultralow attachment 24-well plate</td>
 </tr>
 <tr>
 <td>O.C.T.</td>
 <td>Tissue Tek, Sakura</td>
 <td> 4583</td>
 <td>Optimum Cutting Temperature</td>
 </tr>
 <tr>
 <td>Vectashield</td>
 <td>Vector Labs</td>
 <td> H1000</td>
 <td>Vectashield fluorescent mounting medium</td>
 </tr>
 <tr>
 <td>Slides</td>
 <td>Thermo Scientific</td>
 <td></td>
 <td>Superfrost-Plus positively-charged slides, 25 x 75 x 1.0 mm</td>
 </tr>
 <tr>
 <td>Cover Glass</td>
 <td>VWR International</td>
 <td> 631-0167</td>
 <td>25x60 mm</td>
 </tr>
 <tr>
 <td>Forced draft incubator: </td>
 <td>Lyon Electric Company</td>
 <td></td>
 <td>kept humid by ensuring water levels always sufficient, maintained at 38 &deg;C</td>
 </tr>
 <tr>
 <td>Cryostat</td>
 <td>Brite, Huntingdon, UK</td>
 <td></td>
 <td>-36 &deg;C</td>
 </tr>
 <tr>
 <td>Tissue Culture Incubator</td>
 <td>DH Autoflow</td>
 <td></td>
 <td>Nuaire 38 &deg;C, 5% CO2</td>
 </tr>
 <tr>
 <td>Vibratome</td>
 <td>Leica</td>
 <td></td>
 <td>Leica VT1000-S</td>
 </tr>
 <tr>
 <td>Microscope</td>
 <td>Nikon</td>
 <td></td>
 <td>Nikon Eclipse E80i fluorescence microscope</td>
 </tr>
 <tr>
 <td>Digital Camera</td>
 <td>Nikon</td>
 <td></td>
 <td>Nikon DS5M and Hamammatsu ORCA ER</td>
 </tr>
 </tbody>
</table>

chicken embryo spinal cord slice culture protocol

Slice cultures can facilitate the manipulation of embryo development both pharmacologically and through gene manipulations. In this reduced system, potential lethal side effects due to systemic drug applications can be overcome. However, culture conditions must ensure that normal development proceeds within the reduced environment of the slice. We have focused on the development of the spinal cord, particularly that of spinal motor neurons. We systematically varied culture conditions of chicken embryo slices from the point at which most spinal motor neurons had been born. We assayed the number and type of motor neurons that survived during the culture period and the position of those motor neurons compared to that in vivo. We found that serum type and neurotrophic factors were required during the culture period and were able to keep motor neurons alive for at least 24 hr and allow those motor neurons to migrate to appropriate positions in the spinal cord. We present these culture conditions and the methodology of preparing the embryo slice cultures using eviscerated chicken embryos embedded in agarose and sliced using a vibratome.

The use of cultured slices to follow the development of interneuron and projection neurons has been influential in progressing our understanding of the generation of organization within the cortex16. In the spinal cord, motor neuron development has been followed using cultures of whole zebrafish embryos17. However, spinal motor neuron organization in zebrafish is relatively simple (owing to the lack of large limb muscles in fish). The molecular mechanisms that drive a hierarchy of spinal motor neuron organization during the development of higher vertebrates are currently poorly understood. Culture conditions that facilitate neuron survival and cell body migration in embryo slice cultures of higher vertebrates are thus required. Currently, spinal cord slice cultures of higher vertebrates have been used to seed dissociated neurons on top of the slices12-14, investigate fluorescently labelled axons in the periphery of the slice18, or of progenitor cell behavior in early spinal cord development19. Slice culture conditions for later spinal cords, particularly those that maintain motor neuron development are currently lacking.
To attempt to follow spinal neuronal development, particularly that of spinal motor neurons, we investigated various culture conditions that might promote motor neuron survival and the acquisition of initial order in motor organization through lateral migration of the neurons. Initial trials using stage 18 to stage 20 chick embryo spinal cord slices proved fruitless; we were not able to keep motor neurons alive for longer than a few hours and were unable to demonstrate the generation of substantial numbers of LMCl cells over the culture period (data not shown). We therefore investigated slice cultures of stage 24 embryos, a timepoint when the majority of LMCl and LMCm neurons have been generated, but when LMCl neuron migration is still in its infancy (Figure 1a-c). This lateral migration of LMCl neurons is largely complete by stage 27, around 24 to 36 hr later (Figure 1 d-f). Our assay could thus be simplified to being able to keep the neurons alive and to facilitate their migration laterally into the ventral horn. We started our experiments with relatively simple culture conditions containing just Hank's balanced salt solution with or without chicken serum or chicken embryo extract. In all cases, although we were able to maintain LMC neurons in the slice, we found little evidence of LMCl neurons being maintained (at least by expression of Lhx-1). Further, we found inappropriate expression of genes such as HB9 in the dorsal spinal cord, a situation that never occurs in vivo (data not shown). Thus, these simple culture conditions are not suitable for the investigation of the acquisition of motor neuron organization.
We thus sought out more complex conditions and as a base used a formulation that has been published to facilitate the survival of dissociated embryonic chicken cranial motor neurons. This condition20 (1% chick embryo extract, 1% Pen Strep, 0.35% L-Glutamine, 0.1% 2-mercaptoethanol, 2% horse serum, 2% B27 supplement and 50 ng/ml ciliaryneurotrophic growth factor (CNTF) in neurobasal medium (here called Guthrie medium) did keep more motor neurons alive than the more simple media. Additionally, it did not result in spurious expression of transcription factors in the dorsal spinal cord. However, the survival of LMCl neurons was poor (Figure 1 g-i, p). We therefore varied what we considered to be key factors in the Guthrie medium, namely the animal of origin of the serum, the concentration of the serum and the concentration of CNTF in the medium. We found that changing the serum from Horse serum to Chick serum and an increase in concentration of CNTF from 50 ng/ml to 100 ng/ml substantially increased the survival of the LMCl and LMCm cells and also allowed their migration into the ventral horn over the 24 hr of the culture conditions (Figure 1 j-l, p). The total number of motor neurons, the ratio of LMCm to LMCl and, indeed, the migration of the LMCl cells into the ventral horn appeared very similar to sections of embryos that had been allowed to develop in ovo rather than in the slice culture (Figure 1 k-o, p). Thus, we believe that based on transcription factor expression, cell number and position of motor neurons, our slice culture conditions recapitulate normal spinal motor neuron development at least over a 24 hr period.
<img alt="Figure 1" fo:content-width="2in" fo:src="/files/ftp_upload/50295/50295fig1highres.jpg" src="/files/ftp_upload/50295/50295fig1.jpg" /> 
Figure 1. a-c. Status of generation of LMC (Foxp1 staining in b) and LMCl (Lhx-1 staining in a) at stage 24. c is a merge of the two channels. The midline is shown as a dotted line in a and D, V shows the orientation of the dorsoventral (D, V) axis of the spinal cord. d-f. Status of LMC (Foxp1 staining in d) and LMCl (Lhx-1 staining in e) organization at stage 27. f is a merge of the two channels. g-i. Lhx-1 (g) and Foxp1 (h) expression in sections of a slice cultured in a medium that supports cranial motor neuron cell survival (Guthrie medium, our "initial medium" see reference 20). Lhx-1 is no longer expressed in motor neurons, although there is some survival of LMC cells evidenced by Foxp1 expression. D, V in (g) shows the orientation of the dorsoventral (D, V) axis of the spinal cord. M-L shows the orientation of the mediolateral (M, L) axis of the spinal cord j-l. LMCl cells (Lhx-1 in the ventral horn in j) and the LMC in general (Foxp1 in k) can be observed in slices cultured in medium containing 4% chick serum and 100 ng/ml CNTF. Note that some LMCl cells are found in a lateral position in the ventral horn. The arrow in (j) shows the lateral position of some of the the LMCl cells. (l) is a merge of the two channels. m-o. Status of expression of Lhx-1 (m) and Foxp1 (n) in sections of an embryo kept in ovo during the slice culture of the embryo shown in j-l. (o) is a merge of the two channels. p. Quantitation of the percentage of LMCl (Foxp1+ve/ Lhx1+ve) cells versus LMC (Foxp1+ve/Lhx1-ve) cells found in slice cultures in different conditions compared to control embryos kept in ovo (which represent 100%). Student's t-test *** represents p&lt;0.01. <a href="https://www.jove.com/files/ftp_upload/50295/50295fig1large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about Chicken Embryo Spinal Cord Slice Culture Protocol at JoVE.com

Chicken Embryo Spinal Cord Slice Culture Protocol

Slice cultures facilitate the manipulation of embryo development by gene and pharmacological perturbations. However, culture conditions must ensure that normal development can proceed within the reduced environment of the slice. We illustrate a protocol that facilitates normal spinal cord development to proceed for at least 24 hr.

Slice cultures can facilitate the manipulation of  embryo development both pharmacologically and through gene manipulations. In  this reduced system, ...

chicken-embryo-spinal-cord-slice-culture-protocol

Research

JoVE Journal

Neuroscience

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

Spinal Cord Neurons Isolation and Culture from Neonatal Mice

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on postnatal day 1-3. A mouse litter, usually 4-10 pups born from one breeding pair, is gathered for one experiment, and spinal cords are collected individually from each mouse after euthanasia with isoflurane. The spinal column is dissected out and then the spinal cord is released from the column. The spinal cords are then minced to increase the surface area of delivery for an enzymatic protease that allows for the neurons and other cells to be released from the tissue. Trituration is then used to release the cells into solution. This solution is subsequently fractionated in a density gradient to separate the various cells in solution, allowing for neurons to be isolated. Approximately 1-2.5 x 106 neurons can be isolated from one litter group. The neurons are then seeded onto wells coated with adhesive factors that allow for proper growth and maturation. The neurons take approximately 7 days to reach maturity in the growth and culture medium and can be used thereafter for treatment and analysis.

This study presents a technique for the isolation of neurons from WT neonatal mice. It requires the careful dissection of the spinal cord from the neonatal mouse, followed by the separation of neurons from the spinal cord tissue through mechanical and enzymatic cleavage.

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

A Minimally Invasive, Fast Spinal Cord Lateral Hemisection Technique for Modeling Open Spinal Cord Injuries in Rats

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique eliminates laminectomy by removing two spinous processes and lifting, then tilting the caudal vertebral arch. The surgical area opens up without the need for laminectomy. Lateral hemisection is then performed with direct visible control under a microscope. The trauma is minimized, requiring only a small bone wound. 
This technique has several advantages: it is faster and, therefore, less of a burden for the animal, and the bone wound is smaller. Because the laminectomy is eliminated, there is less chance for unwanted injury to the spinal cord, and there are no bone splinters that can cause problems (bone splinters embedded in the spinal cord can cause swelling and secondary damage). The vertebral canal remains intact. The main limitation is that the hemisection can only be performed in the intervertebral spaces. 
The results show that this technique can be performed much faster than the traditional surgical approach, using laminectomy (11 min vs. 35 min). This technique can be useful for researchers working with animal models of open spinal cord injury as it is widely adaptable and does not require any additional specialized instrumentation.

Here, we describe a new, fast technique modeling open spinal cord injury in rats that eliminates laminectomy. Lateral hemisection is performed while viewing through a microscope. The technique is versatile and can also be used in the cervical, thoracic, and lumbar regions of the spinal cord of other animals.

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique ...

Electromagnetic Controlled Closed-Head Model of Mild Traumatic Brain Injury in Mice

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and understanding the mechanisms of how a TBI alters brain function. The availability of multiple animal models of TBI is necessary to model the different aspects and severities of TBI seen in people. This manuscript describes the use of a midline closed head injury (CHI) to develop a mouse model of mild TBI. The model is considered mild because it does not produce structural brain lesions based on neuroimaging or gross neuronal loss. However, a single impact creates enough pathology that cognitive impairment is measurable at least 1 month after injury. A step-by-step protocol to induce a CHI in mice using a stereotaxically guided electromagnetic impactor is defined in the paper. The benefits of the mild midline CHI model include the reproducibility of the injury-induced changes with low mortality. The model has been temporally characterized up to 1 year after the injury for neuroimaging, neurochemical, neuropathological, and behavioral changes. The model is complementary to open skull models of controlled cortical impact using the same impactor device. Thus, labs can model both mild diffuse TBI and focal moderate-to-severe TBI with the same impactor.

The protocol describes mild traumatic brain injury in a mouse model. In particular, a step-by-step protocol to induce a mild midline closed head injury and the characterization of the animal model is fully explained.

Highly reproducible animal models of traumatic brain injury (TBI), with well-defined pathologies, are needed for testing therapeutic interventions and ...