Name: xenotransplantation of human stem cells into the chicken embryo
Uploaded: 2010-07-11T19:00:00
Duration: 15 min 42 s
Description: Watch this Scientific Journal Video about Xenotransplantation of Human Stem Cells into the Chicken Embryo at JoVE.com

The work was supported by grants from Helse og Rehabilitering (JLB), The Norwegian Research Council (GH and JCG) and the University of Oslo (NK and JCG).

1. Isolation and culture of human stem cells and preparation for injection into embryos
<ol>
	<li>Human stem cells used as examples in this protocol (brain, adipose, bone marrow) are isolated in different ways from the different tissues. For example, adult neural stem cells are isolated from tissue pieces collected from the ventricular wall of the brain during endoscopic neurosurgery, which are then dissociated to individual cells that are cultured in vitro. The neural stem cells spontaneously form neurospheres that float in the culture medium while other cell types adhere to the bottom of the dish7. Adipose-derived mesenchymal stem cells are obtained from liposuction material that is enzymatically digested and strained to remove fibrous tissue and then centrifuged to remove mature adipocytes. The resulting cells are resuspended in culture medium supplemented with serum to promote the proliferation of stem cells over other cell types8 Hematopoietic stem and progenitor cells can be isolated from bone marrow aspirates by positive selection using specific antibodies conjugated to magnetic particles (Myltenyi Biotec, Germany or Dynal-Invitrogen, USA) and magnetic cell separation, and/or using antibodies conjugated to fluorophores and fluorescence activated cell sorting (FACS)9.</li>
	<li>To facilitate later identification, label the stem cells with a fluorescent marker prior to implantation into chicken embryos. Stem cells can be labeled with fluorescent lipophilic dyes such as DiI (Invitrogen, USA) or PKH26 (Sigma, USA) using supplier-recommended protocols up to shortly before implantation, or, for a permanent label with essentially no danger of contaminating host cells, they can be transduced with the gene for a fluorescent protein like Green Fluorescent Protein, using lentiviral vectors, while still in culture10.</li>
	<li>Protocols for the culture and propagation of human stem cells vary. Detailed protocols can be found in the literature. Briefly, sphere-forming stem cells are typically cultured in flasks, whereas adherent stem cells are typically cultured in petri dishes (which facilitate trituration when splitting, see below), in appropriate medium. For splitting or harvesting, pellet sphere-forming cells by 5 min centrifugation at 1200rpm and resuspend in pre-warmed trypsin/EDTA solution for no more than 5 min at 37 degrees Celsius. For adherent cells add the trypsin/EDTA solution to the petri dish, and triturate at the end of 5 minutes. Terminate the trypsinization by adding Ca2+-containing medium, sometimes complemented with albumin. To remove the trypsin/EDTA, centrifuge the cells (5 min at 1200rpm), remove the supernatant, resuspend in cold injection vehicle (typically medium or phosphate-buffered saline), and centrifuge again (5 min at 1200rpm). Remove most of this supernatant and gently resuspend the cells to create a highly concentrated cell suspension.</li>
	<li>Keep the suspension of human stem cells in a small, capped vial or microfuge tube on ice for up to several hours prior to injection. Survival in this state may be cell type-dependent. The injection vehicle must be optimized according to the properties of the cells, including their adhesivity (e.g. a low-Ca2+ vehicle may help prevent clumping) and resistance to hypoxia and pH changes.</li>
</ol>
2. Egg incubation and preparation
<ol>
	<li>Store fertilized eggs in a cooling chamber (for example: Termaks, Bergen, Norway) at 14-15&deg;C prior to incubation. At this temperature development is arrested until incubation begins. Fertilized eggs stored for more than about 10 days or at lower temperatures have a lower rate of subsequent development and survival.</li>
	<li>To reinitiate development, place the eggs in a humidified, forced draft incubator (for example: Binder, New York, USA) at 38-39&deg;C, and incubate until the desired stage of development is reached (for staging, see ref. 11).</li>
	<li>Before opening the egg, create an air pocket above the embryo by inserting an 18 gauge syringe needle through the egg shell (not too deeply to avoid piercing the yolk) and evacuating 4 ml of albumin (Figure A). This creates a pressure drop that within several minutes is equalized by the entry of air through the porous shell. As the air enters it collects at the highest point within the shell, thus separating the embryo from the inner surface of the shell.</li>
	<li>Seal the hole created by the syringe needle with conventional plastic tape. </li>
</ol>
3. Pulling glass micropipettes
<ol>
<li>Pull glass micropipettes from borosilicate glass (for example, 1.2mm OD x 0.94 mm ID, Harvard Apparatus, USA) on a conventional electrode puller (for example, Model P-2000, Sutter Instruments, USA). Adjust puller settings to obtain micropipettes that have high input resistance when used as microelectrodes. Tips should be approximately 1-1.5 cm long. The shafts can be marked at 1 mm intervals for calculating volumes ejected.</li>
<li>Break the micropipette tips to a workable size by bending them with a forceps or pushing them against a solid surface while viewing under a microscope. The break should be as even as possible around the perimeter of the glass and the resultant inner diameter at the tip should accommodate the size of the cells to be used without getting clogged. Typically a 20-30 micron inner diameter works well.</li>
</ol>
4. Preparation of Fast Green dye solution (contrasting reagent)
<ol>
<li>Prepare a 0.1 % (w/v) solution of Fast Green dye (Sigma-Aldrich, USA) by dissolving in chicken physiological saline (137 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 1 mM Na-phosphate, 5 mM HEPES, 11 mM glucose, pH 7.4).</li>
<li>Filter the Fast Green solution through a 0.22 &mu;m Millex GP filter (Millipore Co., Bedford, USA). </li>
</ol>
5. Opening the eggs and contrasting the embryo
<ol>
<li>Place two pieces of tape on the top of the egg, covering the dimensions of the air pocket. Keeping the egg oriented so that the taped area (and thus the air pocket) is uppermost, cut a small window within the borders of the taped area using a sturdy dissection scissors. The tape adds support and helps prevent eggshell fragments from falling onto the embryo. Air bubbles associated with the air pocket can be popped using the back end of a plastic pipette tip or other blunt probe.</li>
	<li>Draw some Fast Green solution into a 1 ml syringe with a 25 gauge needle. Remove any air bubbles in the syringe and the needle. Bend the needle to a 45-degree angle, penetrate at a point outside the embryonic plate (identified by the ring of blood vessels that circumscribe it; piercing within the embryonic plate increases mortality) and carefully guide the needlepoint under the embryo. Inject the dye until desired contrast is achieved. The embryo, which is normally transparent, should now stand out as a whitish ghost against the dark green color (Figure B). Many investigators use India Ink (diluted to 10% v/v) instead of Fast Green. In this case it is important to use India Ink that is non-toxic, such as Pelikan Fount India Ink.</li>
</ol>
6. Making hindbrain and spinal cord neural tube lesions
<ol>
<li>Produce sharpened tungsten needles by flame-etching short (2-3 cm) pieces of 100 &mu;m diameter tungsten rod (catalog number 719000, A&amp;R Systems, Seattle, Washington), with an acetylene/oxygen torch. This is done by inserting the end of the rod very briefly into the torch flame until a spark is seen, which represents the explosive erosion of the outer layers of tungsten, leaving behind a sharp tip.</li>
<li>Using a sharpened tungsten needle, slice through and deflect the vitelline membrane covering the area for microsurgery.</li>
	<li>Using a second sharpened tungsten needle (the tip of the first needle is often damaged after slicing through the vitelline membrane and not fit for subsequent microsurgery), cut carefully through the epithelium overlying the neural tube and deflect it, then cut around and under the piece of the neural tube to be removed, using short, quick upward strokes (Figure C). Making the transverse cuts first and then the longitudinal cuts is usually most successful. Do not cut too deep or you risk penetrating underlying blood vessels, which typically is fatal. </li>
	<li>Gently flip or drag the resected tissue piece away from the lesion site using the tungsten needle. If this proves difficult, it can also be removed by mouth suction using a glass micropipette attached to flexible tubing.</li>
</ol>
7. Injection of human stem cells
a. Injection into a lesion of the neural tube
Draw a small amount of cells into the tip of a glass micropipette by mouth suction using a plastic tube. The working concentration of the cell suspension must be optimized for the cell type to be injected. Mount the micropipette on a micromanipulator and connect it to a microinjector pump (for example: PV830 Pneumatic PicoPump, WPI, Washington, USA). Under visual control using a dissection microscope, guide the micropipette tip into the lesion and carefully expel the cells by increasing the air pressure (either by pulses of constant pressure or by gradually ramping up the pressure) (Figure C). When the desired amount of cells is deposited within the lesion site, carefully withdraw the micropipette.

b. Injection into the lumen of the neural tube
In some cases it may be desirable to inject cells into the developing brain ventricles, for example if the cells are invasive enough to gain access to the neural tissue in the absence of lesions. Injection of cells into the lumen of any brain region requires using a developmental stage at which the lumen provides a large enough receptacle and is easily accessed; HH stages 12 to 18 are favorable in this regard. A lesion is not necessary. Simply pierce the wall of the neural tube with the glass micropipette prior to injection of cells. The key elements for the success of this method are the sharpness of the micropipette and the abruptness of the penetration; too slow a movement and the micropipette tip may only dimple the neural tube wall without penetrating.
c. Systemic injection
Systemic injections can be made via the extra-embryonic blood vessels (starting from HH stage 15, when these vessels are well developed). This is not uncommonly accompanied by some bleeding and since there are many small arteries and fewer larger veins, intra-arterial injections are generally safer when it comes to the embryo's survival. On the other hand, intravenous injections provide quicker access of the cells to the heart and thus probably a more even distribution of the cells. A successful injection requires a sharp micropipette with a diameter smaller than the vessel targeted. Approach the vessel along its axis at a small angle from the horizontal (about 30 degrees). Push the micropipette against the vessel until the vessel begins to occlude. Then push further in a firm, abrupt movement to penetrate. Retract slowly until blood is seen to draw by capillary action into the tip of the micropipette. Injection of cells should then be performed using constant pressure, after which the micropipette is retracted with a quick movement.
8. Sealing the egg
<ol>
	<li>After the injection, let the egg stand still for a few minutes to allow the injected cells to settle and adhere. To avoid dehydration (which can occur quickly and can prove fatal), lay a piece of moistened tissue or filter paper over the window during this rest period.</li>
	<li>After a few minutes and the cells have settled, dry the shell around the window and seal the window with conventional plastic tape. Make sure this seal is tight and that no albumin can leak through the opening or between the shell and the tape (this often proves fatal during subsequent incubation).</li>
	<li>Replace the eggs to the forced-draft incubator for further development. Observation can be made at intervals by removing the tape over the window.</li>
</ol>
9. Embryo dissection
<ol>
	<li>Once the embryo has reached the desired stage, crack open the egg into a bowl of ice-cold physiological saline or phosphate-buffered saline (the tape covering the window should be cut first to allow the two halves of the eggshell to separate). The ice-cold saline serves to anesthetize the embryo by hypothermia. The heart should quickly slow and stop within a couple of minutes.</li>
	<li>Separate the embryo from the extraembryonic membranes and stage it using ref. 11.</li>
	<li>Decapitate the embryo, transfer it to a dissection dish that has a silicone rubber floor and contains oxygenated physiological saline supplemented with glucose (137 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 1 mM Na-phosphate, 5 mM HEPES, 11 mM glucose, pH 7.4) and pin it fast ventral side up. Remove the abdominal and thoracic viscera. Using an angled scissors, make longitudinal incisions along each ventrolateral aspect of the vertebral column. Particularly at later stages (after day 6 of development), the initial incision is best made in the lumbosacral region where the space between the vertebrae and the spinal cord is greatest. Remove the ventral aspect of the vertebral column to reveal the spinal cord. Make sure that the physiological saline remains oxygenated by bubbling at approximately 10-minute intervals with pure medical oxygen.</li>
	<li>Carefully cut the nascent dorsal and ventral roots on either side, transect the spinal cord at the desired levels, and gently lift it out of the vertebral canal. The spinal cord will survive for many hours in this state and can be subjected to anatomical12 and physiological13,14 experiments.</li>
</ol>
<img src="/files/ftp_upload/2071/2071fig1.jpg" alt="Figure 1" /> Figure 1. A) The appropriate approach for evacuation of albumin. Note the position of the embryo and the placement of the needle. B) Contrasting of the embryo. Note the penetration point of the needle outside the embryonic disc. C) Schematic representation of the procedure for unilateral spinal neural tube lesion followed by cell transplantation.

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No conflicts of interest declared.

The chicken embryo is a convenient and versatile animal model for xenotransplantation experiments. The lack of a functional immune system during embryonic and fetal development permits the survival and development of cells from any species that tolerates the incubation temperature. Chicken embryos have been used to test the regenerative potential of various types of stem cells (reviewed in ref. 15). Mammalian stem cells can integrate into chicken embryo tissues including the central nervous system, where they can give rise to neurons whose axonal projections, synaptic connectivity and electrophysiological properties can be assessed in the context of functional neural circuits1,2.
Because the surgical procedures are relatively easy, and injections are made under visual instead of stereotaxic control, the number of embryos can easily be increased to accommodate different experimental conditions within a single experiment. Typically cells can be injected into 20-30 embryos within a 4-hour experiment. The post-injection mortality of the embryos is the principal limiting factor. Precautions must be taken to avoid hemorrhaging and dehydration. Be aware that there are certain critical developmental stages at which the natural mortality rate increases. Supplementing the embryo with a few mL of warm, sterilized chicken ringer following injection and the day before critical developmental stages, as well as ensuring that the window in the egg is well-sealed and oriented to avoid leakage, improves survival. If leakage through the tape seal should occur, remove the tape, dry the surfaces and re-tape. Some investigators use wax and glass or plastic coverslips to seal the window. The mortality should not be substantially different in injected versus sham-operated embryos.
Although we have focused here on the injection of human stem cells into the anlage of the brain and spinal cord, stem cells can also be injected into the anlage of other organs (see for example ref. 4). Each organ poses its own challenges. For example, injection into the heart can be challenging because of the high mechanical resistance to pipette penetration and because of movements, which on the other hand can be slowed by cooling the embryos to room temperature prior to injection. Organs that lack a lumen may not accommodate the injection of sufficient volumes and may therefore require surgical lesion to create a receptacle. In our experience, post-lesion tissue regeneration is an advantage because it promotes the integration of the human stem cells into the tissue. Cells can also be injected into embryonic mesenchyme such as the dispersing somitic mesenchyme or the migrating neural crest as a way to achieve incorporation into mesenchymal derivatives, including neural crest-derived peripheral ganglia16,17.

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xenotransplantation of human stem cells into the chicken embryo

The chicken embryo is a classical animal model for studying normal embryonic and fetal development and for xenotransplantation experiments to study the behavior of cells in a standardized in vivo environment. The main advantages of the chicken embryo include low cost, high accessibility, ease of surgical manipulation and lack of a fully developed immune system. Xenotransplantation into chicken embryos can provide valuable information about cell proliferation, differentiation and behavior, the responses of cells to signals in defined embryonic tissue niches, and tumorigenic potential. Transplanting cells into chicken embryos can also be a step towards transplantation experiments in other animal models. Recently the chicken embryo has been used to evaluate the neurogenic potential of human stem and progenitor cells following implantation into neural anlage1-6. In this video we document the entire procedure for transplanting human stem cells into the developing central nervous system of the chicken embryo. The procedure starts with incubation of fertilized eggs until embryos of the desired age have developed. The eggshell is then opened, and the embryo contrasted by injecting dye between the embryo and the yolk. Small lesions are made in the neural tube using microsurgery, creating a regenerative site for cell deposition that promotes subsequent integration into the host tissue. We demonstrate injections of human stem cells into such lesions made in the part of the neural tube that forms the hindbrain and the spinal cord, and into the lumen of the part of the neural tube that forms the brain. Systemic injections into extraembryonic veins and arteries are also demonstrated as an alternative way to deliver cells to vascularized tissues including the central nervous system. Finally we show how to remove the embryo from the egg after several days of further development and how to dissect the spinal cord free for subsequent physiological, histological or biochemical analyses.

Watch this Scientific Journal Video about Xenotransplantation of Human Stem Cells into the Chicken Embryo at JoVE.com

Xenotransplantation of Human Stem Cells into the Chicken Embryo

In this paper we present a method for transplanting human stem cells into various regions of the central nervous system of the chicken embryo. This provides an in vivo model for assessing the proliferation and differentiation of various types of human stem cells in embryonic tissue environments.

The chicken embryo is a classical animal model for studying normal embryonic and fetal development and for xenotransplantation experiments to study the ...

xenotransplantation-of-human-stem-cells-into-the-chicken-embryo

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Biology

Transfecting Human Neural Stem Cells with the Amaxa Nucleofector

Transfection of primary mammalian neural cells, such as human neural stem/precursor cells (hNSPCs), with commonly used cationic lipid transfection reagents has often resulted in poor cell viability and low transfection efficiency.  Other mechanical methods of introducing a gene of interest, such as a &#8220;gene gun&#8221; or microinjection, are also limited by poor cell viability and low numbers of transfected cells.  The strategy of using viral constructs to introduce an exogenous gene into primary cells has been constrained by both the amount of time and labor required to create viral vectors and potential safety concerns.  We describe here a step-by-step protocol for transfecting hNSPCs using Amaxa's Nucleofector device and technology with electrical current parameters and buffer solutions specifically optimized for transfecting neural stem cells.  Using this protocol, we have achieved initial transfection efficiencies of ~35% and ~70% after stable transfection.  The protocol entails combining a high number of hNSPCs with the DNA to be transfected in the appropriate buffer followed by electroporation with the Nucleofector device.

Introducing a gene of interest into a cell is a powerful method for elucidating its function in vivo. This protocol describes an efficient method of transfecting a culture of human neural stem/precursor cells (hNSPCs) using the Nucleofector electroporation apparatus made by Amaxa.

Transfection of primary mammalian neural cells, such as human neural stem/precursor cells (hNSPCs), with commonly used cationic lipid transfection ...

Passaging Human Neural Stem Cells

The ability to manipulate human neural stem/precursor cells (hNSPCs) in vitro provides a means to investigate their utility as cell transplants for therapeutic purposes as well as to explore many fundamental processes of human neural development and pathology.  This protocol presents a simple method of culturing and passaging hNSPCs in hopes of standardizing this technique and increasing reproducibility of human stem cell research.  The hNSPCs we use were isolated from cadaveric postnatal brain cortices by the National Human Neural Stem Cell Resource and grown as adherent cultures on flasks coated with fibronectin (Palmer et al., 2001; Schwartz et al., 2003).  We culture our hNSPCs in a DMEM:F12 serum-free media supplemented with EGF, FGF, and PDGF and passage them 1:2 approximately every seven days.  Using these conditions, the majority of the cells in the culture maintain a bipolar morphology and express markers of undifferentiated neural stem cells (such as nestin and sox2).

The ability to manipulate human neural stem/precursor cells (hNSPCs) in vitro allows to investigate their utility as cell transplants for therapeutic purposes and to explore human neural development. This protocol presents a method of culturing and passaging hNSPCs in hopes of increasing reproducibility of human stem cell research.

The ability to manipulate human neural stem/precursor cells (hNSPCs) in vitro provides a means to investigate their utility as cell transplants for ...

Immunocytochemistry: Human Neural Stem Cells

Immunocytochemistry is a very powerful and fairly straightforward method for determining the presence, subcellular localization, and relative abundance of an antigen of interest, most commonly a protein, in cultured cells. This protocol presents an easy-to-follow series of steps that will enable researchers to conserve primary and secondary antibodies while getting high quality, reproducible qualitative and quantitative data out of their staining. There are two aspects of this protocol that help to conserve the volume of antibody necessary for staining. For one, the cells are grown on small, circular coverslips that are placed in wells of a tissue culture plate. After fixation, the cells on coverslips can be removed from the wells of the plate. For antibody staining, the coverslip with cells is inverted onto a small drop of antibody solution on parafilm and is covered with a second piece of parafilm to prevent drying. Using this method, only ~25 &mu;l of antibody solution is needed for each coverslip (or sample) to be stained. This protocol describes immunostaining of human neural stem/precursor cells (hNSPCs), but can be used for many other cell types.

Immunocytochemistry is a powerful method to determine the presence, subcellular localization, and relative abundance of an antigen of interest in cultured cells. This protocol presents an easy-to-follow series of steps that will enable one to conserve antibodies and get the most out of one's staining.

Immunocytochemistry is a very powerful and fairly straightforward method for determining the presence, subcellular localization, and relative abundance of ...

In Ovo Electroporations of HH Stage 10 Chicken Embryos

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of molecular biological techniques, the chick has become a useful system in which to study gene regulation and function.  By electroporating DNA or RNA constructs into the developing chicken embryo, genes can be expressed or knocked down in order to analyze in vivo gene function.  Similarly, reporter constructs can be used for fate mapping or to examine putative gene regulatory elements.  Compared to similar experiments in mouse, chick electroporation has the advantages of being quick, easy and inexpensive.  This video demonstrates first how to make a window in the eggshell to manipulate the embryo.  Next, the embryo is visualized with a dilute solution of India ink injected below the embryo.  A glass needle and pipette are used to inject DNA and Fast Green dye into the developing neural tube, then platinum electrodes are placed parallel to the embryo and short electrical pulses are administered with a pulse generator.  Finally, the egg is sealed with tape and placed back into an incubator for further development.  Additionally, the video shows proper egg storage and handling and discusses possible causes of embryo loss following electroporation.

Chick in ovo electroporation is a technique which allows genetic manipulation of the avian embryo. Common applications of this technique include functional analysis of genes and putative enhancer elements. This video demonstrates neural tube electroporation in HH 10 chick embryos. Injection technique and proper egg handling are discussed.

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of ...

Differentiation of Embryonic Stem Cells into Oligodendrocyte Precursors

Oligodendrocytes are the myelinating cells of the central nervous system. For regenerative cell therapy in demyelinating diseases, there is significant interest in deriving a pure population of lineage-committed oligodendrocyte precursor cells (OPCs) for transplantation. OPCs are characterized by the activity of the transcription factor Olig2 and surface expression of a proteoglycan NG2. Using the GFP-Olig2 (G-Olig2) mouse embryonic stem cell (mESC) reporter line, we optimized conditions for the differentiation of mESCs into GFP+Olig2+NG2+ OPCs. In our protocol, we first describe the generation of embryoid bodies (EBs) from mESCs. Second, we describe treatment of mESC-derived EBs with small molecules: (1) retinoic acid (RA) and (2) a sonic hedgehog (Shh) agonist purmorphamine (Pur) under defined culture conditions to direct EB differentiation into the oligodendroglial lineage. By this approach, OPCs can be obtained with high efficiency (&gt;80%) in a time period of 30 days. Cells derived from mESCs in this protocol are phenotypically similar to OPCs derived from primary tissue culture. The mESC-derived OPCs do not show the spiking property described for a subpopulation of brain OPCs in situ. To study this electrophysiological property, we describe the generation of spiking mESC-derived OPCs by ectopically expressing NaV1.2 subunit. The spiking and nonspiking cells obtained from this protocol will help advance functional studies on the two subpopulations of OPCs.

We describe a small molecule-based protocol for differentiation of mouse embryonic stem cells into oligodendrocyte precursor cells (OPCs). This protocol generates Olig2+NG2+ OPCs with high efficiency by 30 days of differentiation. We also describe a method to generate "spiking" OPCs that can fire action potentials.

Oligodendrocytes are the myelinating cells of the central nervous system. For regenerative cell therapy in demyelinating diseases, there is significant ...

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells (HSCs) differentiate into oligopotent progenitors, committed precursors and mature red blood cells 1. This process is regulated for a large part at the level of gene expression, whereby specific transcription factors activate lineage-specific genes while concomitantly repressing genes that are specific to other cell types 2. Studies on transcription factors regulating erythropoiesis are often performed using human and murine cell lines that represent, to some extent, erythroid cells at given stages of differentiation 3-5. However transformed cell lines can only partially mimic erythroid cells and most importantly they do not allow one to comprehensibly study the dynamic changes that occur as cells progress through many stages towards their final erythroid fate. Therefore, a current challenge remains the development of a protocol to obtain relatively homogenous populations of primary HSCs and erythroid cells at various stages of differentiation in quantities that are sufficient to perform genomics and proteomics experiments.
Here we describe an ex vivo cell culture protocol to induce erythroid differentiation from human hematopoietic stem/progenitor cells that have been isolated from either cord blood, bone marrow, or adult peripheral blood mobilized with G-CSF (leukapheresis). This culture system, initially developed by the Douay laboratory 6, uses cytokines and co-culture on mesenchymal cells to mimic the bone marrow microenvironment. Using this ex vivo differentiation protocol, we observe a strong amplification of erythroid progenitors, an induction of differentiation exclusively towards the erythroid lineage and a complete maturation to the stage of enucleated red blood cells. Thus, this system provides an opportunity to study the molecular mechanism of transcriptional regulation as hematopoietic stem cells progress along the erythroid lineage. 
Studying erythropoiesis at the transcriptional level also requires the ability to over-express or knockdown specific factors in primary erythroid cells. For this purpose, we use a lentivirus-mediated gene delivery system that allows for the efficient infection of both dividing and non-dividing cells 7. Here we show that we are able to efficiently knockdown the transcription factor TAL1 in primary human erythroid cells. In addition, GFP expression demonstrates an efficiency of lentiviral infection close to 90%. Thus, our protocol provides a highly useful system for characterization of the regulatory network of transcription factors that control erythropoiesis.

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

An ex vivo protocol to generate mature human red blood cells from hematopoietic stem/progenitors is described. Additionally we describe an efficient lentiviral-delivery method to knockdown the transcription factor TAL1 in primary erythroid cells. The efficiency of lentivirus mediated gene delivery is demonstrated using GFP expressing viruses.

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells ...

Reprogramming Human Somatic Cells into Induced Pluripotent Stem Cells (iPSCs) Using Retroviral Vector with GFP

Human embryonic stem cells (hESCs) are pluripotent and an invaluable cellular sources for in vitro disease modeling and regenerative medicine1. It has been previously shown that human somatic cells can be reprogrammed to pluripotency by ectopic expression of four transcription factors (Oct4, Sox2, Klf4 and Myc) and become induced pluripotent stem cells (iPSCs)2-4 . Like hESCs, human iPSCs are pluripotent and a potential source for autologous cells. Here we describe the protocol to reprogram human fibroblast cells with the four reprogramming factors cloned into GFP-containing retroviral backbone4. Using the following protocol, we generate human iPSCs in 3-4 weeks under human ESC culture condition. Human iPSC colonies closely resemble hESCs in morphology and display the loss of GFP fluorescence as a result of retroviral transgene silencing. iPSC colonies isolated mechanically under a fluorescence microscope behave in a similar fashion as hESCs. In these cells, we detect the expression of multiple pluripotency genes and surface markers.

Reprogramming Human Somatic Cells into Induced Pluripotent Stem Cells iPSCs Using Retroviral Vector with GFP

A method to generate human induced pluripotent stem cells (iPSCs) via retrovirus-mediated ectopic expression of OCT4, SOX2, KLF4 and MYC is described. A practical way to identify human iPSC colonies based on GFP expression is also discussed.

Human embryonic stem cells (hESCs) are pluripotent and an invaluable cellular sources for in vitro disease modeling and regenerative medicine1. It has ...

Differentiation of Newborn Mouse Skin Derived Stem Cells into Germ-like Cells In vitro

Studying germ cell formation and differentiation has traditionally been very difficult due to low cell numbers and their location deep within developing embryos. The availability of a "closed" in vitro based system could prove invaluable for our understanding of gametogenesis. The formation of oocyte-like cells (OLCs) from somatic stem cells, isolated from newborn mouse skin, has been demonstrated and can be visualized in this video protocol. The resulting OLCs express various markers consistent with oocytes such as Oct4 , Vasa , Bmp15, and Scp3. However, they remain unable to undergo maturation or fertilization due to a failure to complete meiosis. This protocol will provide a system that is useful for studying the early stage formation and differentiation of germ cells into more mature gametes. During early differentiation the number of cells expressing Oct4 (potential germ-like cells) reaches ~5%, however currently the formation of OLCs remains relatively inefficient. The protocol is relatively straight forward though special care should be taken to ensure the starting cell population is healthy and at an early passage.

Differentiation of Newborn Mouse Skin Derived Stem Cells into Germ-like Cells In vitro

In recent years the formation of germ cells using in vitro culture methods has been demonstrated using several different types of somatic stem cells. This article will visually present the differentiation of newborn mouse derived stem cells to early oocyte-like cells.

Studying  germ cell formation and differentiation has traditionally been very difficult  due to low cell numbers and their location deep within developing ...

Generation of Transgenic Hydra by Embryo Microinjection

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among multicellular animals. Hydra is used as a model for the study of regeneration, pattern formation, and stem cells. However, research efforts have been hampered by lack of a reliable method for gene perturbations to study molecular function. The development of transgenic methods has revitalized the study of Hydra biology1. Transgenic Hydra allow for the tracking of live cells, sorting to yield pure cell populations for biochemical analysis, manipulation of gene function by knockdown and over-expression, and analysis of promoter function. Plasmid DNA injected into early stage embryos randomly integrates into the genome early in development. This results in hatchlings that express transgenes in patches of tissue in one or more of the three lineages (ectodermal epithelial, endodermal epithelial, or interstitial). The success rate of obtaining a hatchling with transgenic tissue is between 10% and 20%. Asexual propagation of the transgenic hatchling is used to establish a uniformly transgenic line in a particular lineage. Generating transgenic Hydra is surprisingly simple and robust, and here we describe a protocol that can be easily implemented at low cost.

Generation of Transgenic Hydra by Embryo Microinjection

Stably transgenic Hydra are made by microinjection of plasmid DNA into embryos followed by random genomic integration and asexual propagation to establish a uniform line. Transgenic Hydra are used to track cell movements, overexpress genes, study promoter function, or knock down gene expression using RNAi.

As a member of the phylum Cnidaria, the sister group to all bilaterians, Hydra can shed light on fundamental biological processes shared among ...

Generation and Maintenance of Primate Induced Pluripotent Stem Cells Derived from Urine

Cross-species approaches studying primate pluripotent stem cells and their derivatives are crucial to better understand the molecular and cellular mechanisms of disease, development, and evolution. To make primate induced pluripotent stem cells (iPSCs) more accessible, this paper presents a non-invasive method to generate human and non-human primate iPSCs from urine-derived cells, and their maintenance using a feeder-free culturing method.
The urine can be sampled from a non-sterile environment (e.g., the cage of the animal) and treated with a broad-spectrum antibiotic cocktail during primary cell culture to reduce contamination efficiently. After propagation of the urine-derived cells, iPSCs are generated by a modified transduction method of a commercially available Sendai virus vector system. First&#160;iPSC colonies may already be visible after 5 days, and can be picked after 10 days at the earliest. Routine clump passaging with enzyme-free dissociation buffer supports pluripotency of the generated iPSCs for more than 50 passages.

The present protocol describes a method to isolate, expand, and reprogram human and non-human primate urine-derived cells to induced pluripotent stem cells (iPSCs), as well as instructions for feeder-free maintenance of the newly generated iPSCs.