Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Summary

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Abstract

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Introduction

The vertebrate nervous system emerges from the neural plate as a homogeneous layer of neuroepithelial cells. Understanding how developmental programs are induced, encoded, and established during regionalization of the neural plate is, at present, a major goal in developmental biology. Compared to other systems, the experimentally amenable Xenopus embryo is a model of choice for analyzing early steps of neural development^1,2. It is easy to obtain large numbers of embryos, and external development gives access to the very first steps of neurulation³. Many tools are available to experimentally manipulate Xenopuslaevis (X. laevis) embryonic development. Micro-injection of mRNAs or morpholinos (MO), including inducible MOs, together with biochemical and pharmacological tools, allows controlled gain of function (GOF) and loss of function (LOF) and specific alteration of signaling pathways^4,5. The blastocoel roof ectoderm, located around the animal pole of a blastula, or a very early gastrula embryo, and referred to as the 'Animal Cap' (AC), is a source of pluripotent cells that can be programmed by manipulation of gene expression prior to explants preparation. In this manuscript are detailed protocols to use X. laevis AC explants to test in vitro and in vivo molecular mechanisms and cellular processes underlying neural development.

A technique is presented, allowing fine observation of gene expression patterns in a Xenopus tadpole neural tube, a preliminary step in the identification of fate determination cues. Whereas the observation of flat-mounted tissues is commonly used in the study of chick embryos⁶, it has not been properly described in Xenopus. Manipulation of gene expression by injecting synthetic mRNA or MO into the blastomeres of 2 or 4 cell stage embryos allows programming of AC explants⁴. For example inhibition of the Bone Morphogenetic Protein (BMP) pathway by expression of the anti-BMP factor Noggin, gives a neural identity to AC cells³. The protocol is detailed for performing local and time-controlled exposure of AC explants to extrinsic cues via direct contact with an anion exchange resin bead. Finally a technique is described for testing developmental features of neural progenitors in vivo by transplantation of mixed explants prepared from distinct programmed cells dissociated and re-associated.

The frog embryo is a powerful model to study early vertebrate neural development. Combining manipulation of gene expression to explant in vitro cultures provides important information in the study of neuroepithelium regionalization, proliferation, and morphogenesis^7-12. The programming of AC explants permitted development of a functional heart ex vivo^13,14. The use of explant grafting¹⁵ led to the identification of the minimal transcriptional switch inducing the neural crest differentiation program¹⁶. The zona limitans intrathalamica (ZLI) is a signaling center that secretes sonic hedgehog (Shh) to control the growth and regionalization of the caudal forebrain. When continuously exposed to Shh, neuroepithelial cells coexpressing the three transcription factor genes - barH-like homeobox-2(barhl2),orthodenticle-2 (otx2) and iroquois-3 (irx3) - acquire two characteristics of the ZLI compartment: the competence to express shh, and the ability to segregate from anterior neural plate cells. As a model system, the induction of a ZLI fate into neuroepithelial cells will be presented⁸.

These protocols aim at providing simple, cheap, and efficient tools for developmental biologists and other researchers to explore the fundamental mechanisms of key neural cell behaviors. These protocols are very versatile and allow the investigation of a large range of extrinsic and intrinsic neural determination cues. It permits long term in vivo analysis of neural lineage commitment, inductive interactions and cell behaviors.

Protocol

Experiments comply with National and European regulation on the protection of animals used for scientific purposes and with internationally established principles of replacement, reduction and refinement.

1. Flat-mounting of Xenopus laevis Tadpoles Anterior Neural Tube After Whole-mount In Situ Hybridization

1. Obtain X. laevis embryos according to standard procedures⁴ and age them until they reach neurula stage 26 and older (according to Nieuwkoop and Faber developmental table¹⁷.
2. Fix X. laevis tadpoles by incubating them in a solution of 4% paraformaldehyde (PFA). Rock and rotate the embryos in a 2 ml glass vial , 1 to 1.5 hr at RT for stage 26 to stage 38 embryos, 2 hr at RT for embryos at later stages.
   CAUTION: PFA is toxic by contact and a suspected carcinogen. It should be manipulated under a hood.
   NOTE: One X. laevis brood is usually fixed in a 2 ml glass vial but a larger glass vial can be used.
3. Wash the embryos in the same vial using 1x PBS with 0.1% Tween (PBT), 3 min at RT, twice.
   NOTE: Unless otherwise specify the PBS used is with or without calcium and magnesium.
4. Dehydrate the embryos in 100% methanol (MeOH) for at least 12 hr at RT in the same vial. Change the MeOH 100% at least twice under the hood using a plastic micropipette.
   NOTE: The MeOH turns slightly yellow as lipids dissolved in it. Caution the MeOH is toxic by inhalation and should be manipulated under a hood.
5. Rehydrate the embryos using the same vial through a graded series of MeOH baths: MeOH 75% in PBS, MeOH 50% in PBS, MeOH 25% in PBS, PBS twice, each bath 5-10 min at RT. The MeOH solutions are changed under the hood using a plastic micropipette.
6. Using a plastic pipette, transfer the embryo to be dissected in a 60 mm Petri dish filled to the top with PBT. Using two fine forceps carefully remove the eyes by inserting one fine forceps between the eyes and the neural tube, at the level of the optic stalk. Detach the eyes from the neural tube. Carefully introduce the forceps below the ectoderm overlying the neural tube. With care, peel off the ectoderm starting from behind the head and discard it.
7. Perform a double or single ISH using digoxigenin-labeled or fluorescein-labeled probes as previously described^18,19.
8. After the ISH transfer the embryos in a new 60 mm Petri dish filled to the top with PBT using a plastic pipette.
9. Using fine forceps carefully detach the neural tube from the rest of the embryo. At this stage, the anterior neural tube separates from the posterior neural tube. Carefully detach remaining parts of the ectoderm overlying the neural tube, the notochord that is loosely attached below the neural tube, the otic vesicles and remaining parts of the mesoderm. Ensure that the neural tube is devoid of any appendices at this stage.
   NOTE: To avoid damage to the neural tube perform all neural tube/embryo transfer with a plastic transfer pipette or a micropipette if necessary with a tip cut at its end. The diameter of the tip end is adjusted to the size of the neural tube.
10. Transfer the dissected neural tube (see note step 1.9) in a 1.5 ml tube filled with 50% glycerol diluted in PBS. Wait until the neural tubes have fallen to the bottom of the glycerol solution, usually O/N at 4 °C.
11. Remove the solution of 50% glycerol/PBS using a micropipette P1000 and add in the same 1.5 ml tube a solution of 90% glycerol/PBS using a micropipette P1000. Wait until the neural tubes fall to the bottom of the 90% glycerol/PBS solution, usually O/N at 4 °C.
    NOTE: for long term storage, use 90% glycerol/PBS plus antibiotics to prevent bacterial development.
12. Transfer the neural tubes on a glass plate, or a microscope slide with a plastic transfer pipette.
13. Dissect the neural tubes along the dorsal and ventral midlines using tungsten needles. During that step the neural tubes are kept in 90% glycerol/PBS.
14. Mount the two sides of the neural tube in 90% glycerol/PBS using reinforcement rings covered with a glass coverslip.
15. Fix the coverslips with varnish and keep at 4 °C.

2. Animal Cap Explants Induction Using Anion Exchange Resin Beads

1. Transfer 100 µl of anion exchange resin beads into a 2 ml tube using a micropipette P1000. Wash the beads at least five consecutive times with sterile distilled water. Allow the beads to sediment at the bottom of the tube and replace the sterile distilled water using a micropipette P1000. Do not touch the beads.
2. Let the beads soak O/N in a 2 ml tube filled with sterile distilled water with Bovine Serum Albumin (BSA) (10 mg/ ml) at 4 °C.
3. Two hr before collecting the ACs, place half of the beads into a new 1.5 ml tube using a micropipette P1000. Replace the sterile distilled water with 500 µl of the medium containing the molecule to be tested for its inductive potential, or known to induce a specific fate. Keep the other half of the beads, as they will be used as a negative control.
4. Incubate the beads at 4 °C for at least 2 hr.
5. Carry out all subsequent steps under the stereomicroscope and perform all AC transfer with a micropipette.
6. Transfer blastula or very early gastrula embryos into PBS with calcium and magnesium complemented with 0.2% BSA using a plastic transfer pipette.
   NOTE: Embryos developing at 12 °C reach blastula and gastrula stages in 20 to 24 hr.
7. Remove the embryo's vitelline membrane using forceps as previously described²⁰. Fix the embryo with one forceps. Pinch the vitelline membrane with the side of the other forceps. Holding the membrane, slowly peel it off the embryo. Perform this procedure on the ventral (vegetal) side of the embryo. Do not damage the animal side of the embryo.
8. Isolate small ACs as previously described²¹. The animal pole is the embryo's pigmented part. Using fine forceps cut out a small square of tissue out of the animal pole. The AC tissue contains only the ectodermal cells. Ensure that the tissue is of uniform thickness. If not, remove the AC from the analysis.
9. Place each AC in a Terasaki multiwell plates in 0.5x Modified Barth's Saline (MBS)⁴. Place the AC into the well with its pigmented animal side down, in contact with the round bottom of the well.
   NOTE: coating pipettes with a 0.1% BSA solution prevents the adhesion of small pieces of adhesive tissue to the plastic.
10. Place one bead on each cap using a micropipette P20. If needed, use forceps to carefully place the bead in the center of the cap.
    NOTE: When soaked in a conditioned medium, the beads are colored in pink. Select the most colored beads.
11. Incubate at RT (18 to 22 °C) for 6 hr without moving the Terasaki multiwell plate. The bead sticks to the AC in less than 2 hr.
12. Place the Terasaki multiwell plate on top of papers soaked in water. Cover the plate with a plastic container.
    NOTE: This 'humidity chamber' prevents premature evaporation of the wells.
13. Culture the ACs in 0.5x MBS or 3/4 NAM (see step 4.1) in the humidity chamber at 15-20 °C until the sibling embryos have reached the correct stage to test the effect of your factor.
14. Fix the ACs for 1 hr in freshly prepared 4% PFA. Dehydrate the ACs in 100% MeOH as previously described (Step 1.4).

3. Animal Cap Cell Dissociation and Reaggregation Before Grafting in a Xenopuslaevis Neurula

1. Coat petri dishes (60 mm) or 12 wells plates with 3% agarose in sterile water or in PBS without calcium and magnesium. Add enough agarose to cover the bottom of the petri dish or the well. Pre-warm the plates at RT (18-22 °C). Optionally, stored the plates for couple of days at 4 °C to prevent dehydration.
2. Prepare Calcium-free Holtfreter's saline (60 mM NaCl, 0.7 mM KCl, 4.6 mM HEPES, 0.1% BSA (A-7888 Sigma pH 7.6)) and Holtfreter's saline (60 mM NaCl, 0.7 mM KCl, 0.9 mM CaCl₂, 4.6 mM HEPES, 0.1% BSA pH 7.6)⁵. NOTE: the solution of CaCl₂ cannot be autoclaved.
3. Fill the agarose-coated well to the top with Calcium-free Holtfreter's saline.
4. Prepare blastula or very early gastrula embryos to isolate their ACs as previously described (steps 2.5 to 2.7).
5. Isolate at least 15 or up to 30, small ACs²¹. Using fine forceps cut out a small square of tissue out of the animal pole. The AC tissue only contains ectodermal cells and is therefore of uniform thickness. If not, remove the AC from the analysis.
   NOTE: The animal pole is the embryo's pigmented part.
6. Transfer the ACs into an agarose-coated well filled to the top with Calcium-free Holtfreter's saline. Place the ACs with their pigmented side facing upwards.
   NOTE: The dissociation process starts rapidly in calcium-free Holtfreter's saline.
7. Wait a few minutes for cells to start dissociating. Observe this through disaggregation of the tissues. Using fine forceps, separate the pigmented layer from the rest of the AC and discard them with a micropipette P20. Complete cell dissociation process indicates separation of cells from one another.
   NOTE: One or two pigmented layers can be left within the re-aggregated explant to help visualization during grafting.
8. Center the cells using circular movements of the plate. Using a micropipette P1000 carefully remove as much medium as possible. Be careful not to touch the cells.
9. Add 1 ml of Holtfreter's saline with calcium to the well. Transfer the dissociated ACs into a 1.5 ml tube.
   NOTE: 2 ml tubes cannot be used due to their round bottom.
10. Pellet the cells by centrifugation, 5 min at a maximum speed of 2,000 rpm for a bench centrifuge (500 x g). Remove carefully the supernatant with a micropipette P1000.
11. Add to the dissociated cells 20 µl of Holtfreter's saline with calcium (60 mM NaCl, 0.7 mM KCl, 0.9 mM CaCl₂, 4.6 mM HEPES, 0.1% BSA (A-7888 Sigma pH 7.6)⁵.
12. Keep the dissociated ACs, 3 to 6 hr at RT (18-22 °C).
    NOTE: This is necessary time for the cells to re-aggregate. The re-aggregation is visible as the cells form a small ball at the bottom of the tube.
13. Detach the explant carefully from the bottom of the tube by adding 1 ml of 0.5x MBS or of 1 ml of 3/4 NAM (see step 4.1). Transfer the explant in an un-coated plate using a plastic transfer pipette.
14. Stage the explants using their control siblings. For long-term culture use antibiotics: kanamycin (50 µg/ml), ampicillin (50 µg/ml) and gentamycine (50 µg/ml).

4. Grafting of Animal Caps Explants in the Neural Plate of X. laevis Embryo

1. Prepare 3/4 NAM (110 mM NaCl, 2 mM KCl, 1 mM Ca(NO₃)₂, 1 mM MgSO₄, 0.1 mM EDTA, 1 mM NaHCO₃, 0.2x PBS, 50 µg/ml gentamycin). Do not keep for more than 1 week for dissection and store at 4 °C. Older NAM can be used for preparing agarose-coated dissection dishes (below).
2. Coat petri dishes (60 mm) with 3% agarose in 3/4 NAM into which small holes have been made using either a silicone mold, or the cover of a table tennis racket.
   NOTE: The agarose covers the bottom of the petri dish. Leave some space so that the petri dish to fill with 3/4 NAM. Alternatively make dissection dishes with non-drying modeling clay. Within the clay, squeeze the embryos slightly during the dissection procedure. Dig small holes in the agarose using rounded forceps This 'dissection dish' helps to hold the embryos during the dissection procedure.
3. Collect dissection tools: plastic transfer pipettes, an 'eyebrow knife' (made with eyebrow hair embedded in paraffin at the tip of a glass Pasteur pipette)²², two fine dissection forceps, P1000 micropipette and tips, a stereosmicroscope with magnification 8-40X, a bright light source with optic fiber guides. Keep all the dissecting tools clean; after each experiment, rinse them twice with distilled water, once with 100% ethanol and let dry. Store away from dust and if necessary autoclave the metal dissecting tools.
4. Let the dissociated and re-aggregated ACs (protocol 3), and their siblings X. laevis embryos develop until they reach stage 13 (neurula) according to Nieuwkoop and Faber developmental table¹⁷. For transplantation within the neural plate stage 13 to stage 15 embryos are used.
5. Carry out all subsequent steps under the stereomicroscope.
6. Using a plastic transfer pipette, transfer the embryos into PBS with calcium and magnesium complemented with 0.2% BSA. Remove the embryo's vitelline membrane as previously described²⁰. Fix the embryo with one forceps. Pinch the vitelline membrane with the side of the other forceps. Holding the membrane firmly, slowly peel it off the embryo.
   NOTE: Do this procedure on the ventral (vegetal) side of the embryo. Do not damage the embryos neural plate. If embryos have been damaged during the vitelline membrane removal step, transfer the embryos into a 60 mm petri dish containing 3/4 NAM using a plastic transfer pipette and wait 15 min, a sufficient time for the healing process to occur.
7. Fill in the dissection dish to the top with fresh 3/4 NAM.
8. Using a plastic transfer pipette, transfer the embryos without their vitelline membrane into the dissection dish. Place the embryos dorsal side up into the wells. During the transfer process do not allow the embryo to enter in contact with the air-liquid interface since X. laevis are lysed by surface tension.
9. Transfer the AC explant to be grafted into the dissection plate using a plastic transfer pipette or a micropipette P1000. Once the pipette is inside the liquid, allow the explants to slowly sink down by gravity, or push very gently.
10. Maintain the embryos with rounded forceps. With the eyebrow knife make an incision into the neural plate where you intend to graft your explant's piece.
    NOTE: At stage 14 the anterior bending of the neural plate can be use as a landmark, it marks the diencephalon territory (Figure 4).
11. Cut out a small piece of neuroepithelium, using rounded forceps and an eyebrow knive. Cut a small piece of the explant with the eyebrow knife.
    NOTE: The piece of explant should be about the same size and shape as the neuroepithelium ablated area.
12. Place the piece of explant into the neuroepithelium incision using the eyebrow knife and fine forceps. Ensure that the explant rapidly attaches to the embryo.
13. Alternatively place a piece of glass coverslip onto the grafted embryo to maintain the graft in place. To do this, cut a fine glass coverslip into very small pieces using coarse forceps. Ensure that the size is approximately 1.5 mm². Immerse the pieces into a Petri dish containing 3/4 NAM or PBS using forceps. Choose a piece of glass bigger than the embryo to avoid damaging it. The embryo will be a little bit flattened.
14. Wait for at least 30 min without moving, or only move the dissecting plate gently. Let the embryo recover for 30 min to 2 hr and gently remove the coverslip if necessary. Carefully pipette the grafted embryos into a clean dish filled with 3/4 NAM, using the plastic transfer pipette.
    NOTE: Grafted embryos tend to develop bacteria or fungi contamination. For long-term culture use antibiotics: kanamycin (50 µg/ml), ampicillin (50 µg/ml) and gentamycine (50 µg/ml).

Results

Based on morphological considerations in different species, embryological manipulations, and the expression pattern of regulatory genes, a conceptual model holds that the neural plate is divided into transverse and longitudinal segments that define a developmental grid generating distinct histogenic fields. In the neural plate, the primordia of the forebrain, midbrain, hindbrain and spinal cord are all already established along the antero-posterior (AP) axis during gastrulation (reviewed ...

Discussion

Neural development is orchestrated by a complex interplay between cellular developmental programs and signals from the surrounding tissues (Reviewed in^3,31,32). Here we describe a set of protocols that can be used in X. laevis embryos to explore extrinsic and intrinsic factors involved in neural fate determination and neural morphogenesis in vitro and in vivo. These protocols can be used as such on X. tropicalis embryos, however X. tropicalis embryos are four times ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Paraformaldehyde	VWR	20909.290	Toxic
anion exchange resin beads	Biorad	140- 1231
Bovine Serum Albumin	SIGMA	A-7888	For culture of animal cappH 7.6
Gentamycine	GIBCO	15751-045	antibiotic
Bovine Serum Albumin	SIGMA	A7906	for bead preparation