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Chapter 5 Avian Embryos
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Avian Embryos: A Model for the Study of Primary Vascular Assembly in Warm-Blooded Animals Paul A. Rupp, Mike B. Filla, Cheng Cui, and Charles D. Little Contents 1. 2. 3. 4. 5. 6.
Introduction Culture Insert Preparation Culture Chamber Preparation Construction of Microscope Incubator Construction of Electroporation Chamber Embryonic Culture Insert/Dish Preparation 6.1. Solutions 6.2. Embryo staging and preparation 7. Cell Labeling 7.1. Using antibodies and microinjection for tagging endothelial cells or ECM fibers 7.2. Embryo whole-mount electroporation of DNA plasmids expressing fluorescent proteins 7.3. In ovo electroporation of fluorescent protein expressing constructs 8. Post-Incubation Fixation and Processing 9. Whole-Mount Immunolabeling 10. Plastic Embedding and Sectioning References
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Abstract The formation of a primary vascular bed is a dynamic process, aspects of which are readily amenable to time-lapse imaging in avian embryos. At early developmental stages, the body plan of avian embryos is very similar to mammals and has many properties that make it ideal for imaging. We devised labeling, culturing, and imaging techniques that capture high-resolution images of intact avian embryos in four dimensions over large length scales (1 to 5000 mm). Here, Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03005-X
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we describe multiple techniques for labeling and culturing avian embryos to study the cellular, tissue, and extracellular matrix dynamics of vascular morphogenesis.
1. Introduction We have devised novel methods for studying avian endothelial cell migration in vivo (Czirok et al., 2002; Rupp et al., 2003a,b), as well as the dynamics of extracellular matrix (ECM) assembly and tissue movements (Czirok et al., 2004, 2006; Filla et al., 2004; Zamir et al., 2005, 2006). Our studies have centered on the use of time-lapse microscopy to observe embryos from day 1 until early circulation stages. Avian embryos are excellent specimens for the study of an emergent vascular pattern in warm-blooded animals. Not only do avian embryos have body plans nearly identical to those of mammals at early stages, they are readily accessible, easy to stage, exhibit excellent optical properties, and are inexpensive. Avians possess high-performance cardiovascular systems with four-chambered hearts and complex vessel wall structure virtually identical to those in mammals. Tagging endothelial cells in quail embryos for fluorescence microscopy can be conveniently accomplished using two methods—one entails microinjection of fluorochrome-conjugated antibodies and the other takes advantage of introducing DNA plasmids encoding fluorescent proteins. Each method has advantages and disadvantages. When labeling with antibodies, only the antigen present at the time of microinjection is labeled. If the embryo is used for time-lapse imaging, it is therefore not possible to observe the entire vascular structure unless additional antibody is introduced at intervals throughout the image acquisition experiment. This can be an advantage, since not all cells within the vascular structure will be labeled; making it easier to track specific cells. An additional advantage of injecting fluorescently-conjugated primary antibodies that bears mentioning is that the surrounding ECM and/or extracellular growth factors (e.g., VEGF) can be tagged in the same specimen. This can be accomplished in conjunction with endothelial cell surface labeling for double or even triple labeling protocols. If motion analysis (time-lapse microscopy) is planned, labeling a relevant ECM component will permit analysis of cellular motion with respect to the ECM scaffold—that is, autonomous versus passive (tissue) motility (see Zamir et al. [2006] for similar analysis during gastrulation). Transfection of cells, via microinjection and electroporation, with fluorescent protein-expressing plasmids has its own benefits and pitfalls. Cells can be labeled at very early stages (as early as HH stage 1); however,
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this labeling is nonspecific. To label a distinct population of cells, such as the endothelial cells, the DNA plasmids must contain cell type–specific promoters. As with the microinjection of tagged antibodies, not all endothelial cells will be labeled with this technique. However, all progeny of cells that are transfected will likewise be labeled. In order to study cellular and tissue dynamics in vivo, several technical issues must be addressed, including a clear way to label the cells or molecules of interest, sustaining the specimen’s health, maintaining a clear optical path, and acquiring/processing images over wide length scales. The first three of these issues are discussed below, the latter is not. The imaging software has evolved (Czirok et al., 2001), with the latest software available as opensource code. Here we describe experimental methods to label quail embryos using the endothelial cell–specific QH1 antibody in conjunction with another extracellular epitope (singly or simultaneously). We also describe labeling chicken or quail endothelial cells, and other mesodermal derivatives, using electroporation. Perhaps most useful, we describe approaches proven to maintain the health of the embryos on a microscope stage.
2. Culture Insert Preparation A bed of parallel filaments is used to support an embryo during timelapse imaging and provide optimal differential interference contrast (DIC) microscopy. To create a culture insert, the permeable membrane of a Millicell cell culture insert (PICMORG-50, Millipore, Bedford, MA) is first removed. Parallel grooves approximately 3 mm apart were cut into the cell culture insert using a Dremel tool. Trilene XL Smooth Casting fishing line (6-lb test, 0.23-mm diameter, Berkley, Spirit Lake, IA) was woven through the notches (Fig. 5.1A). This structure provides support for the embryo on its vitelline membrane and allows for unobstructed imaging of early avian embryos (Fig. 5.1B). Two newer systems exists that use stainless steel suture bed inserts, Bioptechs DTculture dishes (Bioptechs, Butler, PA), and culture chambers of our design. (In Fig. 5.2A to D, version 1 is shown, from Rupp et al. [2003b].)
3. Culture Chamber Preparation Depending on the type of time-lapse to be performed, three different culture chambers are used. The first is created from a six-well culture dish, employed when the plastic culture insert (described above) is used for highresolution DIC optics. The chamber is abbreviated as 6-WDCC (6-well DIC
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Figure 5.1 The components of a six-well DIC culture chamber and system (6-WDCC). (A) An image of a plastic culture insert created from a tissue culture insert (Millicel) and fishing line.The filaments are 2 mm apart. (B) An avian embryo, ventral side up, resting on a culture insert within one well of the 6-WDCC. Note that the embryo can be situated such that the filaments do not interfere with the optical path. (C) The culture chamber with four wells available for imaging, and two wells for holding water (humidity control).The entire chamber is 21 mm thick in the vertical direction with the filament bed raising the embryo 3 mm from the bottom. (D) An image of a microscope incubator that encompasses the stage and associated optics of a Leica DMRXA2 TM upright microscope (Leica Microsystems, Wetzlar, Germany). Heated air is piped into the incubator through standard metal ducts and distributed via two manifolds within the incubator (see Czirok et al., 2002) to maintain the embryos at 37 C. (Modified from Rupp, P. A., Rongish, B. J., Czirok, A., and Little, C. D. (2003a). Culturing of avian embryos for time-lapse imaging. Biotechniques 34, 274^278.)
culture chamber), and was originally described by Rupp and colleagues (2003a). Using this arrangement, four embryos are typically imaged simultaneously. In order to image in DIC, the plastic within the optical path is replaced with glass (Fig. 5.1C). A 20-mm cork borer is heated over an open flame and a hole is bored through each of the wells within the base. The rough edges are sanded smooth. A no. 2 coverslip is then glued to the underside of the plate using MarineGoop (Eclectic Products, Pineville, LA) thinned with xylene. A watertight seal must be achieved. The plastic in the culture chamber lid is also replaced using 1-mm thick glass (see Fig. 5.1C). The 6-WDCC is approximately 21 mm thick (vertical height).
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Figure 5.2 The components of a four-well DIC electronic culture chamber and system (4-WDECC). (A) An image of the bottom half of a heated self-contained four-well chamber used for embryo culture and image acquisition.The system consists of a chamber of our design (Rupp et al., 2003b) used in conjunction with Bioptechs culture dishes (left corner) (Bioptechs Inc., Butler, PA), and stainless steel filament bed inserts (upper right corner). (B) An HH stage-10 quail embryo is shown within one chamber on a filament bed. The distance between the filaments is 3 mm. (C) A fully assembled electronic culture chamber for imaging up to four embryos in high-resolution DIC and epifluorescence microscopy. (D) The sealed culture chamber mounted on a Leica DMRXA2 upright microscope (Leica Microsystems,Wetzlar, Germany) and connected to the controller box. (Modified from Rupp, P. A., Czirok, A., and Little, C. D. (2003b). Novel approaches for the study of vascular assembly and morphogenesis in avian embryos.Trends Cardiol. Med. 13, 283^288.)
The second culture chamber, used with embryonic culture inserts (described below), is created from two six-well culture dishes (Fig. 5.3A-B). To begin with, the bottom of one of the culture dishes is sanded down from the top until level with the ridged base. A belt sander used in an area with adequate ventilation works well. The sanded dish is then flipped over and a Dremel tool is used to detach the inner wells from the ridged base. A 20-mm hole is created in the bottom of each detached well set using a heated cork borer, and the rough edges are sanded smooth. Holes are likewise bored through one of the lids and sanded smooth. The detached wells with holes should be used as a template to mark where the holes should be bored in the lid. Cover glasses (no. 2, 25 mm, Fisher Scientific) are glued to the topside of the modified lid using MarineGoop thinned with xylene. The glass-containing lid is then flipped over and the
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Figure 5.3 The components of 6-well narrow and 12-well narrow culture chambers (6-WNCC and 12-WNCC, respectively). (A) The lower half of a 6-WNCC used for imaging embryos on embryonic culture inserts after transfection by electroporation. Displayed to the left of the chamber is a Millicell tissue culture insert used to make embryonic culture inserts (albumen/agar plated) as well as the plastic culture inserts used for DIC optics (shown in Figure 5.1). An insert after the addition of albumen/agar is shown in the upper left well of the chamber. (B) An assembled chamber. Note how the lid is inverted from its usual orientation.This allows for increased objective lens working distance. (C) A 12-WNCC assembled from two 12-well dishes is shown without albumen/agar. (D) A single embryo laid dorsal side down on the solidified culture medium within a 12-WNCC. The entire assembly is 12 mm thick (vertically on the microscope stage).
set of detached wells are aligned within the lid and glued into place. The plastic within the second lid is removed and a sheet of glass glued in its place. The second lid is then flipped over and aligned with the new ‘‘base’’ creating a vertically thin 6-well narrow culture chamber (6-WNCC) with glass in all optical paths (Fig. 5.3B). As with the first chamber, all plastic in the optical path has been replaced. This system was designed to limit the amount of embryonic manipulation required after electroporation, and thus increase the success rate for time-lapse imaging. After electroporation, the embryo need only be flipped over (dorsal side down) upon the embryonic culture insert and then placed in the chamber for imaging. In addition, higher magnification objectives may be used due to the additional working distance created by this thinner chamber (12 mm). A third chamber contains 12 wells, and is created in an identical manner to the 6-WNCC except that 12-well culture dishes are used. The
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12-WNCC does not use the embryonic culture inserts; rather a thin layer of albumen/agar is coated directly onto the glass of each well. Embryos that have been labeled either by injection of antibody or electroporation can then be placed dorsal side down directly into the well (Fig. 5.3D). An advantage with this arrangement is that many more embryos can be imaged; provided there is sufficient image acquisition time for each specimen. Like the 6-WNCC, the 12-well version is vertically thinner (12 mm), providing a greater working distance.
4. Construction of Microscope Incubator To culture and dynamically image embryos, an incubator for use with the 6-WDCC, 6-WNCC, and 12-WNCC was fabricated from cardboard (4 mm thickness) to enclose the optics and stage of a Leica DMRXA2 upright microscope (Fig. 5.1D) (Leica Microsystems, Wetzlar, Germany). A portable heater and standard ductwork are used to direct heated air into the incubator and across the culture chamber. The temperature within the incubator is maintained at 38.5 C using a Love 1600 controller unit (Dwyer Instruments, Michigan City, IN) with thermocouple sensors (PT6, Physitemp Instruments, Clifton, NJ). The culture media within the 6-WDCC is kept at 37.5 C, as calibrated with liquid crystal thermometer foils (Edmund Industrial Optics, Barrington, NJ). The design is similar to that of Kulesa and colleagues (1999), and modified from Czirok and colleagues (2001) and Rupp and colleagues (2003a).
5. Construction of Electroporation Chamber Cui and colleagues (2006, 2007) provide a thorough description of the construction of an electroporation chamber (Fig. 5.4A) for whole-mount embryo electroporation. Briefly, a 100-mm Petri dish with a 20-mm hole is aligned underneath a 60-mm dish with a similar hole and glued together with xylene-thinned MarineGoop. A no. 2 cover glass is glued to the bottom of the 100-mm Petri dish. The glass bottom allows for precise positioning of the embryo above the anode. The anode (þ) is prepared by running a 76- 0.25-mm platinum wire (A-M Systems, Carlsborg, WA) along the bottom of the 60-mm Petri dish. The anode is bent such that a 3-mm section over the middle of the bored hole is raised 1 mm above the floor of the Petri dish (Fig. 5.4A to D and G). One end of the anode extends through holes at the base of both dishes and then extends up the outer wall of the Petri dish to be connected to the power supply. Clear nail polish is used to seal the holes, to fix the anode in place, and to insulate the anode
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Figure 5.4 Images and diagram of the electroporation chamber. (A) The electroporation chamber (60-mm dish) is contained within a100-mm dish.This allows for the placement of a weighted ring in the outer chamber to provide stability to the overall device. The platinum wire electrodes are insulated by clear nail polish (tinted cyan in these images). (B) The chamber during electroporation. (C) The cathode (^) positioned by a micromanipulator in the flat orientation for electroporation of a ‘‘large’’area. (D) The cathode (^) positioned by a micromanipulator in the point orientation to focally label a select group of cells. (E) An embryo 3.5 h post-electroporation of a nuclear-GFP expressing (H2B-GFP, Rusty Lansford, California Institute of Technology) DNA plasmid. The entire interstitial space between the vitelline membrane and the epiblast was flooded with plasmid.The cathode was used in the flat orientation with each side of the embryo being electroporated at different voltages (3 Von the right and 4 Von the left). Note how there are fewer GFP-labeled cells on the right side (3 V) than on the left side (4 V) demonstrating that differences in voltage will affect transfection efficiency. Scale bar, 250 mm. (F) A small group of clustered cells labeled when a small amount of plasmid DNA was introduced into a localized region. The cathode was in the flat orientation.
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except for the raised 3-mm ‘‘working’’ section above the windowed hole. A cathode (–) is prepared by bending a 3- 0.25-mm platinum wire (A-M Systems, Carlsborg, WA) at a 45-degree angle approximately 2 mm from one end. The shorter of the two ends is coated with clear nail polish to act as an insulator when mounted on a micromanipulator. The cathode is coupled to a thin stainless steel rod and mounted on a micromanipulator. Alternatively, the platinum wire can be fed through a glass microcapillary, which is glued within the base of an 18-gauge needle (steel needle has been removed; see Fig. 5.4B).
6. Embryonic Culture Insert/Dish Preparation 6.1. Solutions Agar-salt solution: Make and sterilize 0.6% agar (Becton Dickson, 214010) in 123 mM NaCl (Fisher Scientific, S271-10). Aliquots are stored at room temperature. Egg albumen preparation: Egg albumen isolated from fresh chicken eggs that have been incubated 12 to 24 h at 37.5 to 38.5 C is homogenized using a Wheaton 40-ml glass homogenizer. The homogenized egg albumen is heated to 56 C for 40 min prior to aliquoting and storing at – 20 C (good for 1 month). Egg albumen/glucose solution: Just prior to pouring plates, 1.5 ml 10% glucose is added to 48.5 ml of homogenized and heat-treated chicken albumen. The preparation of culture dishes for ex ovo incubation of embryos has been modified from the ‘‘EC culture’’ described by Chapman and colleagues (2001). Briefly, both the agar-salt and egg albumen/glucose solutions are heated to 56 C for at least 10 min. The equilibrated solutions are homogeneously mixed in a 1:1 ratio and immediately distributed as follows: 2 ml per 35-mm plate; 1 ml per Millicell tissue culture insert (PICMORG-50, Millipore, Bedford, MA); or 1 ml per well of the 12-WNCC. The thin (nonviscous) albumen is recommended for use when the embryo is to be imaged using time-lapse microscopy because it is more transparent. Store the solidified plates in a humidified chamber at 4 C for up to 1 week. Standard
Alternatively, the cathode could be positioned in the point orientation to achieve similar results. Scale bar, 250 mm. (G) A cross-section of the electroporation chamber used to electroporate pre-gastrulation stage avian embryos. (Modified from Cui, C., Lansford, R., Filla, M. B., Little, C. D., Cheuvront,T. J., and Rongish, B. J. (2006). Electroporation and EGFP labeling of gastrulating quail embryos. Dev. Dyn. 235, 2802^2810; and Cui, C., Rongish, B. J., Little, C.D., and Lansford, R. (2007). Ex ovo electroporation of DNA vectors into pre-gastrulation avian embryos. CSH Protocols doi:10.1101/pdb.prot4894.)
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35-mm embryonic culture dishes are used for microinjection of antibody followed by non-imaging incubations. The embryonic culture inserts (albumen/agar on tissue culture inserts) are used for electroporation of plasmid DNA and subsequent nonimaging incubations or for imaging within the 6-WNCC. The 12-WNCC with embryonic culture medium is used for time-lapse imaging when higher magnification is needed and to increase the number of samples.
6.2. Embryo staging and preparation Incubate fertile quail eggs (Coturnix coturnix japonica) or chicken eggs in a humidified incubator at 38 C until the appropriate stages as determined by Hamburger and Hamilton (HH) (1951, reprinted in 1992). The embryos are manipulated ex ovo by mounting them to paper rings as described by Chapman et al. (2001) and Rupp et al. (2003a). Briefly, the incubated eggs are opened carefully to ensure that the vitelline membranes on the dorsal aspect of the embryos remain intact. The albumen and yolk (with embryo) are gently poured into a sterile Petri dish. A transfer pipette is used to remove the viscous albumen exposing the vitelline membrane. Kimwipe Tissues (Kimtech Science, Kimberly-Clark Global Sales, Roswell, GA) may be used to remove any additional albumen from the surface of the yolk. An embryo is centered within a Whatman 52 filter paper ring (Whatman International, Maidstone, England) having an inner diameter slightly larger than the size of the embryo. The ring is allowed to adhere to the membrane for 1 min. The ring is cut along the perimeter of the ring beginning at the caudal end of the embryo using an angled iris scissors (Fine Science Tools, Foster City, CA). The embryo is gently pulled free by grasping the paper ring at the caudal end and pulling at a low degree of angle. The embryo is carefully submerged ventral side up into ePBS to remove yolk. A transfer pipette is used to help rinse any adhering yolk by gently pulsing ePBS across the surface of the embryo. Once free of yolk, the embryos are placed onto the embryonic culture dishes (Chapman et al., 2001) for labeling by antibody injection or onto embryonic culture inserts for microinjection and electroporation. Embryos are placed ventral side up for labeling with antibodies and ventral side down during electroporation.
7. Cell Labeling 7.1. Using antibodies and microinjection for tagging endothelial cells or ECM fibers The conjugation of primary antibodies with fluorochromes can be used to label your cell or molecule of interest. The technique is used for labeling embryos using the in vivo whole-mount cultures. We employ Alexa-488,
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Alexa-555, or Alexa-647 (Invitrogen, CA) fluorochrome-conjugated primary QH1 antibodies (Developmental Studies Hybridoma Bank, University of Iowa, Ames, IA) to study the behavior of vascular endothelial cells in the developing quail. QH1 is not effective in labeling chicken endothelial cells. For labeling ECM fibers in either quail or chicken, we use an Alexa488, Alexa-555, or Alexa-647 conjugated nonperturbing primary fibrillin2 antibody ( JB3) or a similarly conjugated fibronectin antibody (B3D6) (both from the Developmental Studies Hybridoma Bank). Depending on the stage of the embryo to be injected, the antibodies are delivered by microinjection of 5- to 25-nl volumes 1 to 16 times per embryo, at 1 ng/nl concentrations. Delivery of the antibodies into the interstitial space is accomplished using a micropipette (18-mm bore) and a pneumatically driven Pico-Injector (Harvard Apparatus, Holliston, MA) mounted to a hydraulic micromanipulator assembly (Narishige Scientific Instrument Laboratory, Tokyo). The needle enters the interstitial space at an acute angle (less than 45 degrees) and the antibody is introduced in 5- to 10-ms pulses with a pressure of 5.0 psi or less.
7.2. Embryo whole-mount electroporation of DNA plasmids expressing fluorescent proteins 7.2.1. Solutions Hanks balanced salt solution (HBSS): Fisher Scientific-CellGro, 21020-CV Embryonic phosphate buffered saline (ePBS, pH 7.4): 137 mM NaCl, 2.69 mM KCl, 8.1 mM Na2HPO4, 1.47 mM KH2PO4, 0.68 mM CaCl2, and 0.49 mM MgCl2 Buffered phenol red solution: 1.0 ml 10X PBS (1.37 M NaCl, 26.8 mM KCl, 81.0 mM Na2HPO4, 15.0 mM KH2PO4, pH 7.2), 10 ml of 1.0 M MgCl2, 5.0 ml of 0.4% phenol red (Fisher Cat no. P-391), and 3.890 ml endotoxin-free water DNA plasmid preparation: 1:1 ratio of buffered phenol red solution and DNA plasmid (endotoxin-free water) for a final concentration of 2.5 mg/ml. Microinjection and electroporation are used to transfect cells with DNA plasmids encoding a fluorescent protein as early as HH stage 1. A large area of the ectoderm can be transfected using a flat cathode in conjunction with the anode (Fig. 5.4C and E). Alternatively, a very focal labeling occurs if the cathode is rotated so that the point of the cathode is directed downward (Fig. 5.4D). The electroporation chamber is tested for continuity by placing an embryonic culture insert (minus any embryo) into it and performing a mock electroporation. The chamber is filled with HBSS to a level so that the anode is covered, but such that embryos will not be submerged when
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present. The cathode is placed parallel to the anode and approximately 3 mm above it (Fig. 5.4B and C). The leads of the electrodes are attached to a CUY-21 Square Wave power supply (BEX Co., Tokyo), which is programmed with the following pulse sequence: 5V, 40 ms power-on and 900 ms power-off for a series of four pulses. Bubbles should be observed on the cathode and recorded amperage of at least 0.01 A displayed. A glass micropipette is used to introduce the plasmid DNA/phenol red solution into the space between the epiblast/embryo and the vitelline membrane and should be prepared just prior to starting the injections (Fig. 5.4B). A concentration of 2.5 mg/ml is general guideline for microinjection of fluorescent protein expressing plasmids used for cell labeling. However, if constructs for overexpression or knockdown are used, the toxicity of the plasmids must be determined empirically. An embryo (quail or chicken) is positioned ventral side down on an embryonic culture insert and then placed into the electroporation chamber. The insert is positioned so that the anode is below the site of interest. As with the antibody injection, the needle penetrates the vitelline membrane at a low degree of angle (<45 degrees). The DNA plasmid (20 to 100 nl) is then introduced by adjusting the Pico-Injector’s P-balance setting to expel the DNA/phenol red into the area of interest. The P-balance allows the solution to be injected slowly (5 s) and with very low pressure (<1.0 psi) so as to not damage the delicate embryonic tissue. The needle is slowly withdrawn from the embryo and the cathode placed parallel to the anode. The cathode should be positioned as close to the embryo as possible without making contact. After placing two drops of HBSS on the vitelline membrane, the cathode is lowered until a slight tissue deformation is observed. The cathode is slowly raised until the deformation has disappeared. With the cathode in the flat orientation, the CUY-21 power supply delivers four square electrical pulses of 4 V, 40 ms each at 900-ms intervals. If the cathode is positioned in the point orientation for a more focal labeling, the pulse sequence is as follows: 3V, 40 ms power-on and 900 ms power-off for four to six pulses. Check for the formation of bubbles at the cathode as well as a change in the color of the phenol red from pink to red. The embryo culture insert may now be removed from the electroporation chamber. At this point, the embryo is flipped over (dorsal side down) onto the embryonic culture insert, placed in a six-well dish, and incubated at 37 C for the appropriate time (up to 40 h). If the embryos are to be used for time-lapse imaging, they are screened at 1 to 2 h of incubation to check for the presence of fluorescence. The selected embryos, on the embryonic culture inserts, may then be placed directly into the 6-WNCC for imaging. Alternatively, if more than six embryos are to be imaged, the 12-WNCC may be used. Well-labeled embryos are floated free of the embryonic culture inserts by placing them in ePBS and then positioned dorsal side down into the 12-WNCC. After filling any unused wells with sterile water,
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the edge of the culture chamber is sealed shut with electrical tape and placed onto the stage within the microscope incubator. It may take a few hours before sufficient levels of fluorescent protein are produced for imaging depending on the strength of the promoters used. This will also depend on the sensitivity of the image detector being used. In addition to the orientation of the cathode (flat or point), the amount and placement of the DNA plasmid determines the area/number of cells transfected. In Fig. 5.4E, the interstitial space between the vitelline membrane and epiblast was flooded with a nuclear-localizing GFP expressing plasmid and then each side electroporated with the cathode in the flat position. In contrast, a small population of cells was labeled in Fig. 5.4F when a small bolus of plasmid was present. Another caveat that alters the efficiency of transfection is the voltage setting: The right side of the embryo in Fig. 5.4E was electroporated with 3 V and the left side with 4 V. The electroporation as described will label epiblastic/ectodermal cells. In order to label a population of endothelial cells, embryos are electroporated at very early stages (HH stage 3 and younger) prior to gastrulation; that is, prior to formation of the mesoderm, this will eventually result in labeled endothelial cells as well as other mesodermal derivatives (unpublished observations). Labeling cells in the center to lower portion of the primitive streak prior to HH stage 3 is optimal for ensuring that endothelial cells will be labeled. DNA plasmids with endothelial cell–specific promoters would obviously label only the early vascular progenitor cells and their daughters.
7.3. In ovo electroporation of fluorescent protein expressing constructs In ovo electroporation of quail embryos is difficult due to the size of the egg. Therefore, use of chicken eggs is recommended, at least while learning the technique. Eggs incubated to the appropriate stage are rinsed with 70% ethanol and allowed to dry. Using a 5-ml syringe with an 18-gauge needle, 2 to 3 ml of albumen are removed from the top of one end of the egg. A window is cut into the top of the shell using an angled iris scissors (Fine Science Tools). Plasmid DNA is introduced between the vitelline membrane and the embryo as described in the embryo whole-mount cultures. India ink, diluted 1:10 in ePBS, can be injected below the embryo to aid in visualizing the structures. For in ovo electroporation, it is recommended that tungsten needles be used instead of platinum as the former appear to yield better labeling results (unpublished observations). Two electrodes are prepared by bending the last 2 mm of a 0.25-mm diameter tungsten wire (A-M Systems, Calsborg, WA) at a 45-degree angle. The wire can be insulated using heat-shrink tubing found at an electrical supply store. For more stability, the straight end of the electrode can be soldered to a male pin connector. Alternatively, the
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tungsten wire may be fed through a glass microcapillary tube with one end bent and the other folded down the outside of the capillary and taped in place with electrical tape. A third alternative is to purchase premade 0.25mm–diameter Tungsten Epoxy-Insulated Microelectrodes with tapered tips (A-M Systems, Carlsborg, WA). The surface of the vitelline membrane is moistened with a small amount of ePBS. The anode is then gently placed below the area of interest by inserting the wire through the vitelline membrane, inserting through the extraembryonic region, and rotating it into position. To get the correct depth, pull the anode up until it is visible under the embryo and then lower it until just visible. As with the embryo whole-mount electroporation the cathode can be placed in two orientations (flat or point). To label a broader area, lower the cathode in the flat orientation parallel to the just visible anode. The anode must be close to, but not touching, the embryo. The CUY-21 Square Wave power supply is used to deliver four pulses of 7 V, 40 ms each at 900- ms intervals. To label a very discrete narrow population of cells, the point orientation is used with the following pulse sequence: 4 V, 40 ms power-on and 900 ms power-off for six pulses. After the embryos have been injected and electroporated, the egg is sealed shut using a rectangular stretched piece of Parafilm (American National Can, Greenwich, CT). The eggs are returned to a 37 C incubator for the appropriate time.
8. Post-Incubation Fixation and Processing In order to preserve embryos post-incubation and allow further analysis, embryos are fixed in a solution of paraformaldehyde (PFA) in PBS. To begin, each specimen is transferred to an individual well of a 24-well plate containing 300 ml of 3% PFA at 4 C. Gently pipette additional cold PFA into each well and carefully submerge the embryos. Allow the embryos to fix for 30 min with gentle agitation at 4 C. If embryos have been incubated on the embryonic culture inserts, the embryos can be floated off of the albumen/agar by the addition of PBS. After fixation and removal from the vitelline membrane, successive dehydration and rehydration is essential for removing lipids and making the specimens permeable to antibodies and other labeling agents. After gently removing the PFA, the specimens are washed twice with PBS þ 0.05% azide (PBSa) at 4 C for 10 min each. The wash is replaced with 50% methanol in PBSa and then returned to 4 C for 15 min. The solution is aspirated off and replaced with 100% methanol at 4 C for a minimum of 30 min. The embryos are rehydrated through a series of decreasing ethanol concentrations beginning with 90%, followed by 70, 50, and 30%. Each step occurs at 4 C for a minimum of 10 min. When the 30% incubation is complete, an equal amount of PBSa is added to
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the wells and incubated for an additional 10 min at 4 C. The embryos are then washed two times in PBSa. For storage, the embryos can be kept in PBSa at 4 C, in the dark, for up to 1 year. Once the embryos are fixed and rehydrated, they may undergo whole-mount immunolabeling and then be subsequently embedded in plastic and sectioned (see following protocols).
9. Whole-Mount Immunolabeling Using the microinjection techniques described above, when a ‘‘pulse’’ of antibody or plasmid is introduced, only cells present at that time (or their daughters in the case of plasmids) will be marked. The entire embryo is often immunolabeled at the end of an experimental protocol in order to visualize the entire vasculature structure, matrix scaffold, or any marker of interest. The first step is to fix and process the embryo as described above, leaving them in PBSa. The embryos are then incubated in bovine serum albumen (BSA) ‘‘blocking’’ solution (3% BSA in PBSa) at 4 C for a minimum of 1 h while gently shaking. A primary antibody is diluted in the blocking solution at the appropriate ratio determined for that antibody. The blocking solution is replaced with the diluted primary antibody and incubated for 2 h at 4 C. The embryos are then washed three times with PBSa for at least 30 min each. If the primary antibody used was directly conjugated with a fluorochrome, the embryos can now be mounted on a slide and imaged. However if a secondary antibody is necessary for visualization, select and dilute the appropriate one. The embryos should be exposed to the secondary at 4 C for 2 h in the appropriate amount of BSA blocking buffer. The washes are then repeated as before. The embryos are now ready for imaging and can be mounted on a slide in a glycerol-based, anti-bleaching mounting solution or in PBSa. If high levels of nonspecific binding occur with the secondary antibody, alternative methods of blocking may be required. An important item to point out is that dehydration/delipidation with methanol will result in the loss of any green fluorescent protein (GFP) or red fluorescent proteins (RFP) signal. A solution of Triton X-100 (SigmaAldrich, St. Louis, MO) can be used as an alternative to methanol. A method to permeabilize fixed cells is found in the book ‘‘Antibodies: A Laboratory Manual’’ by Harlow and Lane (1988), and involves incubating the cells for 2 to 15 min at room temperature in 0.2% Triton X-100. We have found that to permeabilize fixed cells within an embryo, a higher concentration (up to 3.0%) of Triton X-100 is needed. The embryos are not as durable when treated with Triton X-100 as they are with the methanol treatment. A second option is to use the methanol protocol of dehydration/ delipidation, and then reacquire the GFP or RFP signal by using an anti-GFP or anti-RFP antibody to localize the protein.
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10. Plastic Embedding and Sectioning To view vascular labeling in cross-section, embryos may be embedded in plastic and sectioned. The embryos must be dehydrated for the embedding process. The specimens are exposed to a series of increasing ethanol concentration starting at 30%, followed by 70, 80, 95, and 100% for a minimum of 10 min. During the dehydration, the infiltration solution is prepared by adding 0.25 g dry catalyst (benzoyl peroxide, plasticized) to 20 ml of JB-4 Solution A (Electron Microscopy Sciences, Hatfield, PA) and mixed on a stir plate for 15 min. The infiltration solution may be stored for up to 2 weeks in the dark at 4 C. The embryos are individually soaked overnight in 1 ml infiltration solution in the dark at either room temperature or at 4 C. The embedding solution is prepared by adding 60 ml of JB-4 Solution B to 1.5 ml of cold infiltration solution per embryo and vigorously mixing for 30 s. Mark the cranial portion of each paper ring with a tissuemarking dye. The specimens are than submerged (paper rings and all) into the embedding solution within the mold plate wells. A fiduciary marker, in the form of a stiff dark-colored paintbrush hair, is placed 2 mm to the right of the embryo axis on the ventral side of each specimen. The embryos are incubated at room temperature in a vacuum oven in the dark for 1 h to allow the plastic to harden. Once hardened, a fresh razor blade is used to cut out 3 mm 3 mm 6 mm blocks of plastic containing the specimens. The blocks are placed into fresh wells of a mold plate, oriented appropriately, and covered with 2 mm of fresh embedding solution. The specimens are mounted on plastic microtome chucks (EBH2 block holder, Electron Microscopy Sciences, Hatfield, PA). Make sure that the central hole of the chuck is filled with embedding solution and that the orientation of the specimen does not change. Placing the samples at room temperature in a vacuum oven in the dark for 1 h once again hardens the plastic. The specimens are then sectioned using a microtome to produce 1.0- to 15.0-micron thick samples that are mounted on slides for viewing and imaging.
REFERENCES Chapman, S. C., Collignon, J., Schoenwolf, G. C., and Lumsden, A. (2001). Improved method for chick whole-embryo culture using a filter paper carrier. Dev. Dyn. 220, 284–289. Cui, C., Lansford, R., Filla, M. B., Little, C. D., Cheuvront, T. J., and Rongish, B. J. (2006). Electroporation and EGFP labeling of gastrulating quail embryos. Dev. Dyn. 235, 2802–2810. Cui, C., Rongish, B. J., Little, C. D., and Lansford, R. (2007). Ex ovo electroporation of DNA vectors into pre-gastrulation avian embryos. CSH Protocols doi:10.1101/pdb. prot4894.
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Czirok, A., Rupp, P. A., Rongish, B. J., and Little, C. D. (2001). Multi-field 3D scanning light microscopy of early embryogenesis. J. Microsc. 206, 209–217. Czirok, A., Rongish, B. J., and Little, C. D. (2004). Extracellular matrix dynamics during vertebrate axis formation. Dev. Biol. 268, 111–122. Czirok, A., Zamir, E. A., Filla, M. B., Little, C. D., and Rongish, B. J. (2006). Extracellular matrix macroassembly dynamics in early vertebrate embryos. Dev. Biol.Curr. Top. Dev. Biol. 73, 237–258. Filla, M. B., Czirok, A., Zamir, E. A., Little, C. D., Cheuvront, T. J., and Rongish, B. J. (2004). Dynamic imaging of cell, extracellular matrix, and tissue movements during avian vertebral axis patterning. Birth Defects Res. C Embryo Today 72, 267–276. Hamburger, V., and Hamilton, H. L. (1951). A series of normal stages in the development of the chick embryo. J. Morphol. 88, 49–92. Hamburger, V., and Hamilton, H. L. (1992). A series of normal stages in the development of the chick embryo: 1951. Dev. Dyn. 195, 231–272. Harlow, E., and Lane, D. (1988). ‘‘Antibodies: A Laboratory Manual.’’ Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Kulesa, P. M., and Fraser, S. E. (1999). Confocal imaging of living cells in intact embryos. Methods Mol. Biol. 122, 205–222. Rupp, P. A., Rongish, B. J., Czirok, A., and Little, C. D. (2003a). Culturing of avian embryos for time-lapse imaging. Biotechniques 34, 274–278. Rupp, P. A., Czirok, A., and Little, C. D. (2003b). Novel approaches for the study of vascular assembly and morphogenesis in avian embryos. Trends Cardiol. Med. 13, 283–288. Zamir, E. A., Czirok, A., Rongish, B. J., and Little, C. D. (2005). A digital image–based method for computational tissue fate mapping during early avian morphogenesis. Ann. Biomed. Eng. 33, 854–865. Zamir, E. A., Czirok, A., Cui, C., Little, C. D., and Rongish, B. J. (2006). Mesodermal cell displacements during avian gastrulation are due to both individual cell-autonomous and convective tissue movements. Proc. Natl. Acad. Sci. USA 103, 19806–19811.
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Avian Embryos: A Model for the Study of Primary Vascular Assembly in Warm-Blooded Animals Paul A. Rupp, Mike B. Filla, Cheng Cui, and Charles D. Little Contents 1. 2. 3. 4. 5. 6.
Introduction Culture Insert Preparation Culture Chamber Preparation Construction of Microscope Incubator Construction of Electroporation Chamber Embryonic Culture Insert/Dish Preparation 6.1. Solutions 6.2. Embryo staging and preparation 7. Cell Labeling 7.1. Using antibodies and microinjection for tagging endothelial cells or ECM fibers 7.2. Embryo whole-mount electroporation of DNA plasmids expressing fluorescent proteins 7.3. In ovo electroporation of fluorescent protein expressing constructs 8. Post-Incubation Fixation and Processing 9. Whole-Mount Immunolabeling 10. Plastic Embedding and Sectioning References
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Abstract The formation of a primary vascular bed is a dynamic process, aspects of which are readily amenable to time-lapse imaging in avian embryos. At early developmental stages, the body plan of avian embryos is very similar to mammals and has many properties that make it ideal for imaging. We devised labeling, culturing, and imaging techniques that capture high-resolution images of intact avian embryos in four dimensions over large length scales (1 to 5000 mm). Here, Anatomy and Cell Biology, University of Kansas Medical Center, Kansas City, Kansas Methods in Enzymology, Volume 445 ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)03005-X
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we describe multiple techniques for labeling and culturing avian embryos to study the cellular, tissue, and extracellular matrix dynamics of vascular morphogenesis.
1. Introduction We have devised novel methods for studying avian endothelial cell migration in vivo (Czirok et al., 2002; Rupp et al., 2003a,b), as well as the dynamics of extracellular matrix (ECM) assembly and tissue movements (Czirok et al., 2004, 2006; Filla et al., 2004; Zamir et al., 2005, 2006). Our studies have centered on the use of time-lapse microscopy to observe embryos from day 1 until early circulation stages. Avian embryos are excellent specimens for the study of an emergent vascular pattern in warm-blooded animals. Not only do avian embryos have body plans nearly identical to those of mammals at early stages, they are readily accessible, easy to stage, exhibit excellent optical properties, and are inexpensive. Avians possess high-performance cardiovascular systems with four-chambered hearts and complex vessel wall structure virtually identical to those in mammals. Tagging endothelial cells in quail embryos for fluorescence microscopy can be conveniently accomplished using two methods—one entails microinjection of fluorochrome-conjugated antibodies and the other takes advantage of introducing DNA plasmids encoding fluorescent proteins. Each method has advantages and disadvantages. When labeling with antibodies, only the antigen present at the time of microinjection is labeled. If the embryo is used for time-lapse imaging, it is therefore not possible to observe the entire vascular structure unless additional antibody is introduced at intervals throughout the image acquisition experiment. This can be an advantage, since not all cells within the vascular structure will be labeled; making it easier to track specific cells. An additional advantage of injecting fluorescently-conjugated primary antibodies that bears mentioning is that the surrounding ECM and/or extracellular growth factors (e.g., VEGF) can be tagged in the same specimen. This can be accomplished in conjunction with endothelial cell surface labeling for double or even triple labeling protocols. If motion analysis (time-lapse microscopy) is planned, labeling a relevant ECM component will permit analysis of cellular motion with respect to the ECM scaffold—that is, autonomous versus passive (tissue) motility (see Zamir et al. [2006] for similar analysis during gastrulation). Transfection of cells, via microinjection and electroporation, with fluorescent protein-expressing plasmids has its own benefits and pitfalls. Cells can be labeled at very early stages (as early as HH stage 1); however,
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this labeling is nonspecific. To label a distinct population of cells, such as the endothelial cells, the DNA plasmids must contain cell type–specific promoters. As with the microinjection of tagged antibodies, not all endothelial cells will be labeled with this technique. However, all progeny of cells that are transfected will likewise be labeled. In order to study cellular and tissue dynamics in vivo, several technical issues must be addressed, including a clear way to label the cells or molecules of interest, sustaining the specimen’s health, maintaining a clear optical path, and acquiring/processing images over wide length scales. The first three of these issues are discussed below, the latter is not. The imaging software has evolved (Czirok et al., 2001), with the latest software available as opensource code. Here we describe experimental methods to label quail embryos using the endothelial cell–specific QH1 antibody in conjunction with another extracellular epitope (singly or simultaneously). We also describe labeling chicken or quail endothelial cells, and other mesodermal derivatives, using electroporation. Perhaps most useful, we describe approaches proven to maintain the health of the embryos on a microscope stage.
2. Culture Insert Preparation A bed of parallel filaments is used to support an embryo during timelapse imaging and provide optimal differential interference contrast (DIC) microscopy. To create a culture insert, the permeable membrane of a Millicell cell culture insert (PICMORG-50, Millipore, Bedford, MA) is first removed. Parallel grooves approximately 3 mm apart were cut into the cell culture insert using a Dremel tool. Trilene XL Smooth Casting fishing line (6-lb test, 0.23-mm diameter, Berkley, Spirit Lake, IA) was woven through the notches (Fig. 5.1A). This structure provides support for the embryo on its vitelline membrane and allows for unobstructed imaging of early avian embryos (Fig. 5.1B). Two newer systems exists that use stainless steel suture bed inserts, Bioptechs DTculture dishes (Bioptechs, Butler, PA), and culture chambers of our design. (In Fig. 5.2A to D, version 1 is shown, from Rupp et al. [2003b].)
3. Culture Chamber Preparation Depending on the type of time-lapse to be performed, three different culture chambers are used. The first is created from a six-well culture dish, employed when the plastic culture insert (described above) is used for highresolution DIC optics. The chamber is abbreviated as 6-WDCC (6-well DIC
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Figure 5.1 The components of a six-well DIC culture chamber and system (6-WDCC). (A) An image of a plastic culture insert created from a tissue culture insert (Millicel) and fishing line.The filaments are 2 mm apart. (B) An avian embryo, ventral side up, resting on a culture insert within one well of the 6-WDCC. Note that the embryo can be situated such that the filaments do not interfere with the optical path. (C) The culture chamber with four wells available for imaging, and two wells for holding water (humidity control).The entire chamber is 21 mm thick in the vertical direction with the filament bed raising the embryo 3 mm from the bottom. (D) An image of a microscope incubator that encompasses the stage and associated optics of a Leica DMRXA2 TM upright microscope (Leica Microsystems, Wetzlar, Germany). Heated air is piped into the incubator through standard metal ducts and distributed via two manifolds within the incubator (see Czirok et al., 2002) to maintain the embryos at 37 C. (Modified from Rupp, P. A., Rongish, B. J., Czirok, A., and Little, C. D. (2003a). Culturing of avian embryos for time-lapse imaging. Biotechniques 34, 274^278.)
culture chamber), and was originally described by Rupp and colleagues (2003a). Using this arrangement, four embryos are typically imaged simultaneously. In order to image in DIC, the plastic within the optical path is replaced with glass (Fig. 5.1C). A 20-mm cork borer is heated over an open flame and a hole is bored through each of the wells within the base. The rough edges are sanded smooth. A no. 2 coverslip is then glued to the underside of the plate using MarineGoop (Eclectic Products, Pineville, LA) thinned with xylene. A watertight seal must be achieved. The plastic in the culture chamber lid is also replaced using 1-mm thick glass (see Fig. 5.1C). The 6-WDCC is approximately 21 mm thick (vertical height).
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Figure 5.2 The components of a four-well DIC electronic culture chamber and system (4-WDECC). (A) An image of the bottom half of a heated self-contained four-well chamber used for embryo culture and image acquisition.The system consists of a chamber of our design (Rupp et al., 2003b) used in conjunction with Bioptechs culture dishes (left corner) (Bioptechs Inc., Butler, PA), and stainless steel filament bed inserts (upper right corner). (B) An HH stage-10 quail embryo is shown within one chamber on a filament bed. The distance between the filaments is 3 mm. (C) A fully assembled electronic culture chamber for imaging up to four embryos in high-resolution DIC and epifluorescence microscopy. (D) The sealed culture chamber mounted on a Leica DMRXA2 upright microscope (Leica Microsystems,Wetzlar, Germany) and connected to the controller box. (Modified from Rupp, P. A., Czirok, A., and Little, C. D. (2003b). Novel approaches for the study of vascular assembly and morphogenesis in avian embryos.Trends Cardiol. Med. 13, 283^288.)
The second culture chamber, used with embryonic culture inserts (described below), is created from two six-well culture dishes (Fig. 5.3A-B). To begin with, the bottom of one of the culture dishes is sanded down from the top until level with the ridged base. A belt sander used in an area with adequate ventilation works well. The sanded dish is then flipped over and a Dremel tool is used to detach the inner wells from the ridged base. A 20-mm hole is created in the bottom of each detached well set using a heated cork borer, and the rough edges are sanded smooth. Holes are likewise bored through one of the lids and sanded smooth. The detached wells with holes should be used as a template to mark where the holes should be bored in the lid. Cover glasses (no. 2, 25 mm, Fisher Scientific) are glued to the topside of the modified lid using MarineGoop thinned with xylene. The glass-containing lid is then flipped over and the
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Figure 5.3 The components of 6-well narrow and 12-well narrow culture chambers (6-WNCC and 12-WNCC, respectively). (A) The lower half of a 6-WNCC used for imaging embryos on embryonic culture inserts after transfection by electroporation. Displayed to the left of the chamber is a Millicell tissue culture insert used to make embryonic culture inserts (albumen/agar plated) as well as the plastic culture inserts used for DIC optics (shown in Figure 5.1). An insert after the addition of albumen/agar is shown in the upper left well of the chamber. (B) An assembled chamber. Note how the lid is inverted from its usual orientation.This allows for increased objective lens working distance. (C) A 12-WNCC assembled from two 12-well dishes is shown without albumen/agar. (D) A single embryo laid dorsal side down on the solidified culture medium within a 12-WNCC. The entire assembly is 12 mm thick (vertically on the microscope stage).
set of detached wells are aligned within the lid and glued into place. The plastic within the second lid is removed and a sheet of glass glued in its place. The second lid is then flipped over and aligned with the new ‘‘base’’ creating a vertically thin 6-well narrow culture chamber (6-WNCC) with glass in all optical paths (Fig. 5.3B). As with the first chamber, all plastic in the optical path has been replaced. This system was designed to limit the amount of embryonic manipulation required after electroporation, and thus increase the success rate for time-lapse imaging. After electroporation, the embryo need only be flipped over (dorsal side down) upon the embryonic culture insert and then placed in the chamber for imaging. In addition, higher magnification objectives may be used due to the additional working distance created by this thinner chamber (12 mm). A third chamber contains 12 wells, and is created in an identical manner to the 6-WNCC except that 12-well culture dishes are used. The
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12-WNCC does not use the embryonic culture inserts; rather a thin layer of albumen/agar is coated directly onto the glass of each well. Embryos that have been labeled either by injection of antibody or electroporation can then be placed dorsal side down directly into the well (Fig. 5.3D). An advantage with this arrangement is that many more embryos can be imaged; provided there is sufficient image acquisition time for each specimen. Like the 6-WNCC, the 12-well version is vertically thinner (12 mm), providing a greater working distance.
4. Construction of Microscope Incubator To culture and dynamically image embryos, an incubator for use with the 6-WDCC, 6-WNCC, and 12-WNCC was fabricated from cardboard (4 mm thickness) to enclose the optics and stage of a Leica DMRXA2 upright microscope (Fig. 5.1D) (Leica Microsystems, Wetzlar, Germany). A portable heater and standard ductwork are used to direct heated air into the incubator and across the culture chamber. The temperature within the incubator is maintained at 38.5 C using a Love 1600 controller unit (Dwyer Instruments, Michigan City, IN) with thermocouple sensors (PT6, Physitemp Instruments, Clifton, NJ). The culture media within the 6-WDCC is kept at 37.5 C, as calibrated with liquid crystal thermometer foils (Edmund Industrial Optics, Barrington, NJ). The design is similar to that of Kulesa and colleagues (1999), and modified from Czirok and colleagues (2001) and Rupp and colleagues (2003a).
5. Construction of Electroporation Chamber Cui and colleagues (2006, 2007) provide a thorough description of the construction of an electroporation chamber (Fig. 5.4A) for whole-mount embryo electroporation. Briefly, a 100-mm Petri dish with a 20-mm hole is aligned underneath a 60-mm dish with a similar hole and glued together with xylene-thinned MarineGoop. A no. 2 cover glass is glued to the bottom of the 100-mm Petri dish. The glass bottom allows for precise positioning of the embryo above the anode. The anode (þ) is prepared by running a 76- 0.25-mm platinum wire (A-M Systems, Carlsborg, WA) along the bottom of the 60-mm Petri dish. The anode is bent such that a 3-mm section over the middle of the bored hole is raised 1 mm above the floor of the Petri dish (Fig. 5.4A to D and G). One end of the anode extends through holes at the base of both dishes and then extends up the outer wall of the Petri dish to be connected to the power supply. Clear nail polish is used to seal the holes, to fix the anode in place, and to insulate the anode
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Figure 5.4 Images and diagram of the electroporation chamber. (A) The electroporation chamber (60-mm dish) is contained within a100-mm dish.This allows for the placement of a weighted ring in the outer chamber to provide stability to the overall device. The platinum wire electrodes are insulated by clear nail polish (tinted cyan in these images). (B) The chamber during electroporation. (C) The cathode (^) positioned by a micromanipulator in the flat orientation for electroporation of a ‘‘large’’area. (D) The cathode (^) positioned by a micromanipulator in the point orientation to focally label a select group of cells. (E) An embryo 3.5 h post-electroporation of a nuclear-GFP expressing (H2B-GFP, Rusty Lansford, California Institute of Technology) DNA plasmid. The entire interstitial space between the vitelline membrane and the epiblast was flooded with plasmid.The cathode was used in the flat orientation with each side of the embryo being electroporated at different voltages (3 Von the right and 4 Von the left). Note how there are fewer GFP-labeled cells on the right side (3 V) than on the left side (4 V) demonstrating that differences in voltage will affect transfection efficiency. Scale bar, 250 mm. (F) A small group of clustered cells labeled when a small amount of plasmid DNA was introduced into a localized region. The cathode was in the flat orientation.
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except for the raised 3-mm ‘‘working’’ section above the windowed hole. A cathode (–) is prepared by bending a 3- 0.25-mm platinum wire (A-M Systems, Carlsborg, WA) at a 45-degree angle approximately 2 mm from one end. The shorter of the two ends is coated with clear nail polish to act as an insulator when mounted on a micromanipulator. The cathode is coupled to a thin stainless steel rod and mounted on a micromanipulator. Alternatively, the platinum wire can be fed through a glass microcapillary, which is glued within the base of an 18-gauge needle (steel needle has been removed; see Fig. 5.4B).
6. Embryonic Culture Insert/Dish Preparation 6.1. Solutions Agar-salt solution: Make and sterilize 0.6% agar (Becton Dickson, 214010) in 123 mM NaCl (Fisher Scientific, S271-10). Aliquots are stored at room temperature. Egg albumen preparation: Egg albumen isolated from fresh chicken eggs that have been incubated 12 to 24 h at 37.5 to 38.5 C is homogenized using a Wheaton 40-ml glass homogenizer. The homogenized egg albumen is heated to 56 C for 40 min prior to aliquoting and storing at – 20 C (good for 1 month). Egg albumen/glucose solution: Just prior to pouring plates, 1.5 ml 10% glucose is added to 48.5 ml of homogenized and heat-treated chicken albumen. The preparation of culture dishes for ex ovo incubation of embryos has been modified from the ‘‘EC culture’’ described by Chapman and colleagues (2001). Briefly, both the agar-salt and egg albumen/glucose solutions are heated to 56 C for at least 10 min. The equilibrated solutions are homogeneously mixed in a 1:1 ratio and immediately distributed as follows: 2 ml per 35-mm plate; 1 ml per Millicell tissue culture insert (PICMORG-50, Millipore, Bedford, MA); or 1 ml per well of the 12-WNCC. The thin (nonviscous) albumen is recommended for use when the embryo is to be imaged using time-lapse microscopy because it is more transparent. Store the solidified plates in a humidified chamber at 4 C for up to 1 week. Standard
Alternatively, the cathode could be positioned in the point orientation to achieve similar results. Scale bar, 250 mm. (G) A cross-section of the electroporation chamber used to electroporate pre-gastrulation stage avian embryos. (Modified from Cui, C., Lansford, R., Filla, M. B., Little, C. D., Cheuvront,T. J., and Rongish, B. J. (2006). Electroporation and EGFP labeling of gastrulating quail embryos. Dev. Dyn. 235, 2802^2810; and Cui, C., Rongish, B. J., Little, C.D., and Lansford, R. (2007). Ex ovo electroporation of DNA vectors into pre-gastrulation avian embryos. CSH Protocols doi:10.1101/pdb.prot4894.)
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35-mm embryonic culture dishes are used for microinjection of antibody followed by non-imaging incubations. The embryonic culture inserts (albumen/agar on tissue culture inserts) are used for electroporation of plasmid DNA and subsequent nonimaging incubations or for imaging within the 6-WNCC. The 12-WNCC with embryonic culture medium is used for time-lapse imaging when higher magnification is needed and to increase the number of samples.
6.2. Embryo staging and preparation Incubate fertile quail eggs (Coturnix coturnix japonica) or chicken eggs in a humidified incubator at 38 C until the appropriate stages as determined by Hamburger and Hamilton (HH) (1951, reprinted in 1992). The embryos are manipulated ex ovo by mounting them to paper rings as described by Chapman et al. (2001) and Rupp et al. (2003a). Briefly, the incubated eggs are opened carefully to ensure that the vitelline membranes on the dorsal aspect of the embryos remain intact. The albumen and yolk (with embryo) are gently poured into a sterile Petri dish. A transfer pipette is used to remove the viscous albumen exposing the vitelline membrane. Kimwipe Tissues (Kimtech Science, Kimberly-Clark Global Sales, Roswell, GA) may be used to remove any additional albumen from the surface of the yolk. An embryo is centered within a Whatman 52 filter paper ring (Whatman International, Maidstone, England) having an inner diameter slightly larger than the size of the embryo. The ring is allowed to adhere to the membrane for 1 min. The ring is cut along the perimeter of the ring beginning at the caudal end of the embryo using an angled iris scissors (Fine Science Tools, Foster City, CA). The embryo is gently pulled free by grasping the paper ring at the caudal end and pulling at a low degree of angle. The embryo is carefully submerged ventral side up into ePBS to remove yolk. A transfer pipette is used to help rinse any adhering yolk by gently pulsing ePBS across the surface of the embryo. Once free of yolk, the embryos are placed onto the embryonic culture dishes (Chapman et al., 2001) for labeling by antibody injection or onto embryonic culture inserts for microinjection and electroporation. Embryos are placed ventral side up for labeling with antibodies and ventral side down during electroporation.
7. Cell Labeling 7.1. Using antibodies and microinjection for tagging endothelial cells or ECM fibers The conjugation of primary antibodies with fluorochromes can be used to label your cell or molecule of interest. The technique is used for labeling embryos using the in vivo whole-mount cultures. We employ Alexa-488,
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Alexa-555, or Alexa-647 (Invitrogen, CA) fluorochrome-conjugated primary QH1 antibodies (Developmental Studies Hybridoma Bank, University of Iowa, Ames, IA) to study the behavior of vascular endothelial cells in the developing quail. QH1 is not effective in labeling chicken endothelial cells. For labeling ECM fibers in either quail or chicken, we use an Alexa488, Alexa-555, or Alexa-647 conjugated nonperturbing primary fibrillin2 antibody ( JB3) or a similarly conjugated fibronectin antibody (B3D6) (both from the Developmental Studies Hybridoma Bank). Depending on the stage of the embryo to be injected, the antibodies are delivered by microinjection of 5- to 25-nl volumes 1 to 16 times per embryo, at 1 ng/nl concentrations. Delivery of the antibodies into the interstitial space is accomplished using a micropipette (18-mm bore) and a pneumatically driven Pico-Injector (Harvard Apparatus, Holliston, MA) mounted to a hydraulic micromanipulator assembly (Narishige Scientific Instrument Laboratory, Tokyo). The needle enters the interstitial space at an acute angle (less than 45 degrees) and the antibody is introduced in 5- to 10-ms pulses with a pressure of 5.0 psi or less.
7.2. Embryo whole-mount electroporation of DNA plasmids expressing fluorescent proteins 7.2.1. Solutions Hanks balanced salt solution (HBSS): Fisher Scientific-CellGro, 21020-CV Embryonic phosphate buffered saline (ePBS, pH 7.4): 137 mM NaCl, 2.69 mM KCl, 8.1 mM Na2HPO4, 1.47 mM KH2PO4, 0.68 mM CaCl2, and 0.49 mM MgCl2 Buffered phenol red solution: 1.0 ml 10X PBS (1.37 M NaCl, 26.8 mM KCl, 81.0 mM Na2HPO4, 15.0 mM KH2PO4, pH 7.2), 10 ml of 1.0 M MgCl2, 5.0 ml of 0.4% phenol red (Fisher Cat no. P-391), and 3.890 ml endotoxin-free water DNA plasmid preparation: 1:1 ratio of buffered phenol red solution and DNA plasmid (endotoxin-free water) for a final concentration of 2.5 mg/ml. Microinjection and electroporation are used to transfect cells with DNA plasmids encoding a fluorescent protein as early as HH stage 1. A large area of the ectoderm can be transfected using a flat cathode in conjunction with the anode (Fig. 5.4C and E). Alternatively, a very focal labeling occurs if the cathode is rotated so that the point of the cathode is directed downward (Fig. 5.4D). The electroporation chamber is tested for continuity by placing an embryonic culture insert (minus any embryo) into it and performing a mock electroporation. The chamber is filled with HBSS to a level so that the anode is covered, but such that embryos will not be submerged when
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present. The cathode is placed parallel to the anode and approximately 3 mm above it (Fig. 5.4B and C). The leads of the electrodes are attached to a CUY-21 Square Wave power supply (BEX Co., Tokyo), which is programmed with the following pulse sequence: 5V, 40 ms power-on and 900 ms power-off for a series of four pulses. Bubbles should be observed on the cathode and recorded amperage of at least 0.01 A displayed. A glass micropipette is used to introduce the plasmid DNA/phenol red solution into the space between the epiblast/embryo and the vitelline membrane and should be prepared just prior to starting the injections (Fig. 5.4B). A concentration of 2.5 mg/ml is general guideline for microinjection of fluorescent protein expressing plasmids used for cell labeling. However, if constructs for overexpression or knockdown are used, the toxicity of the plasmids must be determined empirically. An embryo (quail or chicken) is positioned ventral side down on an embryonic culture insert and then placed into the electroporation chamber. The insert is positioned so that the anode is below the site of interest. As with the antibody injection, the needle penetrates the vitelline membrane at a low degree of angle (<45 degrees). The DNA plasmid (20 to 100 nl) is then introduced by adjusting the Pico-Injector’s P-balance setting to expel the DNA/phenol red into the area of interest. The P-balance allows the solution to be injected slowly (5 s) and with very low pressure (<1.0 psi) so as to not damage the delicate embryonic tissue. The needle is slowly withdrawn from the embryo and the cathode placed parallel to the anode. The cathode should be positioned as close to the embryo as possible without making contact. After placing two drops of HBSS on the vitelline membrane, the cathode is lowered until a slight tissue deformation is observed. The cathode is slowly raised until the deformation has disappeared. With the cathode in the flat orientation, the CUY-21 power supply delivers four square electrical pulses of 4 V, 40 ms each at 900-ms intervals. If the cathode is positioned in the point orientation for a more focal labeling, the pulse sequence is as follows: 3V, 40 ms power-on and 900 ms power-off for four to six pulses. Check for the formation of bubbles at the cathode as well as a change in the color of the phenol red from pink to red. The embryo culture insert may now be removed from the electroporation chamber. At this point, the embryo is flipped over (dorsal side down) onto the embryonic culture insert, placed in a six-well dish, and incubated at 37 C for the appropriate time (up to 40 h). If the embryos are to be used for time-lapse imaging, they are screened at 1 to 2 h of incubation to check for the presence of fluorescence. The selected embryos, on the embryonic culture inserts, may then be placed directly into the 6-WNCC for imaging. Alternatively, if more than six embryos are to be imaged, the 12-WNCC may be used. Well-labeled embryos are floated free of the embryonic culture inserts by placing them in ePBS and then positioned dorsal side down into the 12-WNCC. After filling any unused wells with sterile water,
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the edge of the culture chamber is sealed shut with electrical tape and placed onto the stage within the microscope incubator. It may take a few hours before sufficient levels of fluorescent protein are produced for imaging depending on the strength of the promoters used. This will also depend on the sensitivity of the image detector being used. In addition to the orientation of the cathode (flat or point), the amount and placement of the DNA plasmid determines the area/number of cells transfected. In Fig. 5.4E, the interstitial space between the vitelline membrane and epiblast was flooded with a nuclear-localizing GFP expressing plasmid and then each side electroporated with the cathode in the flat position. In contrast, a small population of cells was labeled in Fig. 5.4F when a small bolus of plasmid was present. Another caveat that alters the efficiency of transfection is the voltage setting: The right side of the embryo in Fig. 5.4E was electroporated with 3 V and the left side with 4 V. The electroporation as described will label epiblastic/ectodermal cells. In order to label a population of endothelial cells, embryos are electroporated at very early stages (HH stage 3 and younger) prior to gastrulation; that is, prior to formation of the mesoderm, this will eventually result in labeled endothelial cells as well as other mesodermal derivatives (unpublished observations). Labeling cells in the center to lower portion of the primitive streak prior to HH stage 3 is optimal for ensuring that endothelial cells will be labeled. DNA plasmids with endothelial cell–specific promoters would obviously label only the early vascular progenitor cells and their daughters.
7.3. In ovo electroporation of fluorescent protein expressing constructs In ovo electroporation of quail embryos is difficult due to the size of the egg. Therefore, use of chicken eggs is recommended, at least while learning the technique. Eggs incubated to the appropriate stage are rinsed with 70% ethanol and allowed to dry. Using a 5-ml syringe with an 18-gauge needle, 2 to 3 ml of albumen are removed from the top of one end of the egg. A window is cut into the top of the shell using an angled iris scissors (Fine Science Tools). Plasmid DNA is introduced between the vitelline membrane and the embryo as described in the embryo whole-mount cultures. India ink, diluted 1:10 in ePBS, can be injected below the embryo to aid in visualizing the structures. For in ovo electroporation, it is recommended that tungsten needles be used instead of platinum as the former appear to yield better labeling results (unpublished observations). Two electrodes are prepared by bending the last 2 mm of a 0.25-mm diameter tungsten wire (A-M Systems, Calsborg, WA) at a 45-degree angle. The wire can be insulated using heat-shrink tubing found at an electrical supply store. For more stability, the straight end of the electrode can be soldered to a male pin connector. Alternatively, the
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tungsten wire may be fed through a glass microcapillary tube with one end bent and the other folded down the outside of the capillary and taped in place with electrical tape. A third alternative is to purchase premade 0.25mm–diameter Tungsten Epoxy-Insulated Microelectrodes with tapered tips (A-M Systems, Carlsborg, WA). The surface of the vitelline membrane is moistened with a small amount of ePBS. The anode is then gently placed below the area of interest by inserting the wire through the vitelline membrane, inserting through the extraembryonic region, and rotating it into position. To get the correct depth, pull the anode up until it is visible under the embryo and then lower it until just visible. As with the embryo whole-mount electroporation the cathode can be placed in two orientations (flat or point). To label a broader area, lower the cathode in the flat orientation parallel to the just visible anode. The anode must be close to, but not touching, the embryo. The CUY-21 Square Wave power supply is used to deliver four pulses of 7 V, 40 ms each at 900- ms intervals. To label a very discrete narrow population of cells, the point orientation is used with the following pulse sequence: 4 V, 40 ms power-on and 900 ms power-off for six pulses. After the embryos have been injected and electroporated, the egg is sealed shut using a rectangular stretched piece of Parafilm (American National Can, Greenwich, CT). The eggs are returned to a 37 C incubator for the appropriate time.
8. Post-Incubation Fixation and Processing In order to preserve embryos post-incubation and allow further analysis, embryos are fixed in a solution of paraformaldehyde (PFA) in PBS. To begin, each specimen is transferred to an individual well of a 24-well plate containing 300 ml of 3% PFA at 4 C. Gently pipette additional cold PFA into each well and carefully submerge the embryos. Allow the embryos to fix for 30 min with gentle agitation at 4 C. If embryos have been incubated on the embryonic culture inserts, the embryos can be floated off of the albumen/agar by the addition of PBS. After fixation and removal from the vitelline membrane, successive dehydration and rehydration is essential for removing lipids and making the specimens permeable to antibodies and other labeling agents. After gently removing the PFA, the specimens are washed twice with PBS þ 0.05% azide (PBSa) at 4 C for 10 min each. The wash is replaced with 50% methanol in PBSa and then returned to 4 C for 15 min. The solution is aspirated off and replaced with 100% methanol at 4 C for a minimum of 30 min. The embryos are rehydrated through a series of decreasing ethanol concentrations beginning with 90%, followed by 70, 50, and 30%. Each step occurs at 4 C for a minimum of 10 min. When the 30% incubation is complete, an equal amount of PBSa is added to
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the wells and incubated for an additional 10 min at 4 C. The embryos are then washed two times in PBSa. For storage, the embryos can be kept in PBSa at 4 C, in the dark, for up to 1 year. Once the embryos are fixed and rehydrated, they may undergo whole-mount immunolabeling and then be subsequently embedded in plastic and sectioned (see following protocols).
9. Whole-Mount Immunolabeling Using the microinjection techniques described above, when a ‘‘pulse’’ of antibody or plasmid is introduced, only cells present at that time (or their daughters in the case of plasmids) will be marked. The entire embryo is often immunolabeled at the end of an experimental protocol in order to visualize the entire vasculature structure, matrix scaffold, or any marker of interest. The first step is to fix and process the embryo as described above, leaving them in PBSa. The embryos are then incubated in bovine serum albumen (BSA) ‘‘blocking’’ solution (3% BSA in PBSa) at 4 C for a minimum of 1 h while gently shaking. A primary antibody is diluted in the blocking solution at the appropriate ratio determined for that antibody. The blocking solution is replaced with the diluted primary antibody and incubated for 2 h at 4 C. The embryos are then washed three times with PBSa for at least 30 min each. If the primary antibody used was directly conjugated with a fluorochrome, the embryos can now be mounted on a slide and imaged. However if a secondary antibody is necessary for visualization, select and dilute the appropriate one. The embryos should be exposed to the secondary at 4 C for 2 h in the appropriate amount of BSA blocking buffer. The washes are then repeated as before. The embryos are now ready for imaging and can be mounted on a slide in a glycerol-based, anti-bleaching mounting solution or in PBSa. If high levels of nonspecific binding occur with the secondary antibody, alternative methods of blocking may be required. An important item to point out is that dehydration/delipidation with methanol will result in the loss of any green fluorescent protein (GFP) or red fluorescent proteins (RFP) signal. A solution of Triton X-100 (SigmaAldrich, St. Louis, MO) can be used as an alternative to methanol. A method to permeabilize fixed cells is found in the book ‘‘Antibodies: A Laboratory Manual’’ by Harlow and Lane (1988), and involves incubating the cells for 2 to 15 min at room temperature in 0.2% Triton X-100. We have found that to permeabilize fixed cells within an embryo, a higher concentration (up to 3.0%) of Triton X-100 is needed. The embryos are not as durable when treated with Triton X-100 as they are with the methanol treatment. A second option is to use the methanol protocol of dehydration/ delipidation, and then reacquire the GFP or RFP signal by using an anti-GFP or anti-RFP antibody to localize the protein.
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10. Plastic Embedding and Sectioning To view vascular labeling in cross-section, embryos may be embedded in plastic and sectioned. The embryos must be dehydrated for the embedding process. The specimens are exposed to a series of increasing ethanol concentration starting at 30%, followed by 70, 80, 95, and 100% for a minimum of 10 min. During the dehydration, the infiltration solution is prepared by adding 0.25 g dry catalyst (benzoyl peroxide, plasticized) to 20 ml of JB-4 Solution A (Electron Microscopy Sciences, Hatfield, PA) and mixed on a stir plate for 15 min. The infiltration solution may be stored for up to 2 weeks in the dark at 4 C. The embryos are individually soaked overnight in 1 ml infiltration solution in the dark at either room temperature or at 4 C. The embedding solution is prepared by adding 60 ml of JB-4 Solution B to 1.5 ml of cold infiltration solution per embryo and vigorously mixing for 30 s. Mark the cranial portion of each paper ring with a tissuemarking dye. The specimens are than submerged (paper rings and all) into the embedding solution within the mold plate wells. A fiduciary marker, in the form of a stiff dark-colored paintbrush hair, is placed 2 mm to the right of the embryo axis on the ventral side of each specimen. The embryos are incubated at room temperature in a vacuum oven in the dark for 1 h to allow the plastic to harden. Once hardened, a fresh razor blade is used to cut out 3 mm 3 mm 6 mm blocks of plastic containing the specimens. The blocks are placed into fresh wells of a mold plate, oriented appropriately, and covered with 2 mm of fresh embedding solution. The specimens are mounted on plastic microtome chucks (EBH2 block holder, Electron Microscopy Sciences, Hatfield, PA). Make sure that the central hole of the chuck is filled with embedding solution and that the orientation of the specimen does not change. Placing the samples at room temperature in a vacuum oven in the dark for 1 h once again hardens the plastic. The specimens are then sectioned using a microtome to produce 1.0- to 15.0-micron thick samples that are mounted on slides for viewing and imaging.
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