Using Xenopus Skin to Study Cilia Development and Function

2.1.1 mRNA injection into blastomeres

The use of mRNAs encoding cilia-related genes fused to fluorescent markers is an extremely powerful approach to visualize and characterize detailed aspects of cilia formation and function. Table 10.1 summarizes some common markers useful for labeling different cilia-related structures. Detailed protocols for harvesting, fertilizing and injecting Xenopus embryos have previously been published (Mimoto & Christian, 2011; Sive, Grainger, & Harland, 1998, 2007b, 2007c, 2007f, 2010). Additionally, the in vitro synthesis of mRNAs is well established (Sive et al., 1998) and is also feasible using established optimized kits (e.g., Ambion mMessage mMachine Cat # AM1340). However, to ensure effective protein localization in the embryonic skin and in MCCs, four separate injections are performed, one in each of the four quadrants of the animal pole of the embryo. The ciliated epithelium derives from the ectoderm, therefore injections should be targeted away from the midline and more laterally toward the center of the cell on the animal side of each blastomere, which enriches the ectodermal lineage in mRNA (Sive et al., 1998). Embryos are then allowed to develop until they reach the desired developmental stage for investigation. MCCs begin to express MCC specific genes at low levels as early as stage 11.5, with strong expression typically evident by stage 14. MCCs begin radial intercalation as early as stage 14 (ideally visualized between stages 17 and 18), begin to undergo ciliogenesis around stage 20 (ideally visualized between stages 20 and 22), and have mature oriented cilia by stage 28 (visualized between stages 23 and 28). Fluorescently tagged proteins can be visualized in live embryos (see Section 3.2.2) or fixed for further processing (e.g., antibody labeling, see Section 3.1).

2.1.2 Restricted expression to ciliated cells using the α-tubulin promoter

Targeted injection of mRNA allows for some spatial control of protein synthesis. However, for many applications, temporal control and precise localization of proteins specifically to differentiating MCCs is required to avoid off-target developmental defects. There are several approaches to achieve this that rely on the use of the α-tubulin promoter (Stubbs et al., 2006). This promoter drives gene expression specifically in MCCs starting at stage 13 when MCCs begin to form and continues to drive expression in MCCs during ciliated cell differentiation and throughout development. Simply, one can inject 10–50 pg of plasmid DNA containing the α-tubulin promoter driving expression of one’s gene of interest in a similar manner to the mRNA injections described above. For many cell biological analyses, the injection of plasmid DNA is sufficient; however, it has the drawback that it can result in mosaic and variable levels of expression that is problematic for tissue level analysis. For more consistent expression, it is preferable to make “transient transgenics,” a method that relies on injecting sperm nuclei along with the plasmid DNA into unfertilized eggs and achieving an F1 transgenic animal for immediate analysis. While this method requires slightly more effort, it will generate a more consistent level of expression across the tissue. There are currently numerous methods to achieve “transient transgenics,” and proper methods are beyond the scope of this chapter (Amaya & Kroll, 1999; Loeber, Pan, & Pieler, 2009).

2.2.1 Injection of antisense MOs to knockdown protein levels

MOs are usually designed to bind in close proximity to or overlapping with the mRNA start codon to prevent initiation of translation. Alternatively, MOs that are designed to overlap with splice sites interfere with proper pre-mRNA splicing (Draper, Morcos, & Kimmel, 2001). Splice MOs often yield a stronger reduction in the full-length protein and have the advantage that, in the absence of a good antibody, the extent of reduction of the full-length mRNA can be determined using qRT-PCR for mis-processed mRNA. However, splice MOs often generate partially truncated protein, as a result of alternative splicing. The mis-spliced protein may have residual biological functions or may act as a dominant negative or a dominant active form of the targeted protein. Generally, the use of more than one MO that cause the same phenotype as well as rescue experiments is advisable to minimize the risk that the observed phenotype is due to off-target effects of the MO (for a detailed review of the use of MOs in vertebrate embryos, see Eisen & Smith, 2008; Summerton, 2007). When working with an untested MO, it is necessary to empirically determine the optimal concentration. Most MOs seem to be effective at amounts ranging from 1 to 20 ng with doses of up to 60 ng used successfully in some reports. For coinjection with mRNA, it is critical that the mRNA be purified and resuspended in RNAse-free water that is not treated with DEPC. Qiagen’s RNeasy MinElute Cleanup kit (Cat # 74204) results in mRNA that is suitable for coinjecting with MOs.

2.2.2 Using mosaic embryos to circumvent potential lethality

Organism-wide knockdown of certain proteins using MOs may lead to broad developmental defects preceding formation of the ciliated epithelium. To circumvent such potential defects, MO injection can be restricted to a single cell at the 4-, 8-, or even 16-cell stage, thereby restricting the knockdown to only a subset of cells in the animal. This will often allow the development of a largely healthy embryo in which only a subset of cells are affected by the knockdown. In this case, it is necessary to label the cells that have received the MO either by using fluorescently labeled MOs or by coinjecting MOs with a marker mRNA such as membrane or nuclear targeted Red Fluorescent Protein. Injection of the MO with 0.1% phenol red can also be used to track the injection. This approach also allows the comparison of morphant and control tissue within the same embryo.

2.2.3 Rescue experiments

In order to validate the specificity of a phenotype associated with a MO-dependent protein knockdown, rescue experiments should be performed. Since MOs are often designed to target either parts of the 5′UTR when interfering with translation initiation or parts of an intron at intron–exon boundaries for splice MOs, in most cases, the MO phenotype can be rescued by simply coinjecting in vitro transcribed mRNA of the gene in question lacking the 5′UTR. In cases where the coding sequence is targeted, a rescue construct can be designed using alternate codon usage to maintain amino acid sequence but not DNA sequence.

3.2.1 Mounting of fixed embryos

Several different techniques have been described to mount whole embryos for fluorescent microscopy (Kieserman, Lee, Gray, Park, & Wallingford, 2010; Lee, Kieserman, Gray, Park, & Wallingford, 2008). For relatively flat fixed embryos, one can also sandwich the embryos between two large pieces of coverglass (Fig. 10.1A). Electrical tape is used as a spacer placing the embryos into holes cut into the tape using a scalpel or a razor blade. Generally, three layers of electrical tape are used for embryos stage 22 and younger, while two pieces are used after stage 22. This technique has the advantage that both sides of the embryos can be visualized, which is particularly important when performing mosaic or transplant experiments (see Sections 2.2.2 and 4.2):

1. Cut two or three 60-mm long pieces of electrical tape and place them one on top of the other on a 48 × 60 mm or 28 × 60 mm coverglass (Goldseal coverglass, VWR Inc., Cat # 48404 and 63770).

2. Using the edge of a sharp razor blade cut out an approximately 5 × 8 mm rectangle of electrical tape toward the middle of the coverglass to create a pocket that can accommodate up to three embryos. Multiple pockets can be cut into a single stack of labeling tape to accommodate additional samples per slide.

3. Transfer embryos to the pocket using a transfer pipette making an effort to minimize the amount of liquid that is transferred with the embryos.

4. Aspirate most of the liquid and quickly embed embryos in one drop of Fluorogel (EMS Cat # 17985-10) or Mowiol 4-88 (Polysciences, Inc., Cat # 9002-89-5).

5. Use forceps to position embryos within the pocket and carefully place a second 48 × 60 or 28 × 60 mm coverslip on top aligning the edges of both pieces coverglass.

3.2.2 Mounting of embryos for live imaging

Various mounting techniques for live imaging of Xenopus embryos have been described and can vary depending on the developmental stage of the embryo as well as the time period over which the embryo is to be imaged. Kieserman et al. have recently published a very comprehensive review covering several of these techniques, and we will refer to them for a detailed description (Kieserman et al., 2010). The key feature for mounting embryos for live imaging is keeping the embryos pressed against the coverglass for imaging while maintaining access to media and allowing for growth (for longer time course imaging). Additionally, a significant advantage for using Xenopus is the ability to culture explanted tissue (see Section 4.1) which can alleviate issues with three-dimensional growth and movement. One can use custom-made half-rose chambers (Wadsworth, 2007) attached to a 48 × 60 mm coverglass by applying a ring of silicone grease on the bottom of the chamber. Embryos can then be placed in the chamber and held in place by a smaller piece of coverglass with dabs of silicon on the sides (Fig. 10.1B). Customized microscope chambers as described here can be used for most applications and yield reliable results using extended imaging up to several hours. For imaging of shorter time intervals especially when high spatial resolution is required, such as imaging microtubule dynamics or cilia beating, it is preferable to use the mounting technique described in Section 3.2.1; however, this is not suitable for longer time periods especially if the embryo undergoes drastic shape changes during the imaging time frame. Additionally, for imaging of tailbud stage or older embryos, it is often advisable to immobilize embryos using anesthetics such as 0.02% tricaine (ethyl 3-aminobenzoate methanesulfonate salt, Sigma–Aldrich Inc., Cat # A5040) made up in 0.1 × Marc’s Modified Rinders (MMR, 1 × is 110 mM NaCl, 2 mM KCl, 1 mM MgCl₂, 2 mM CaCl₂, 2 mM NaHCO₃, 5 mM HEPES, pH 7.8) to prevent embryos from twitching during imaging.

3.3.1 Visualization of fluid flow using fluorescent or dyed microspheres

Visualization of fluid flow can be performed on most upright microscopes at fairly low magnification of 4–10×. A simple dissecting scope equipped with an eyepiece mounted digital camera (e.g., Scopetek DCM130E) is the most economical possibility. The velocity of fluid flow is generally measured indirectly by the displacement of microbeads along the surface of the embryo (Mitchell et al., 2007). If the microscope is equipped with a fluorescent light source, it is preferable to use fluorescently labeled microspheres since the recorded signal is brighter with minimal background making the recordings more amenable to particle tracking software:

1. Transfer 20 μl of resuspended fluorescent (Invitrogen Inc., Cat # F8836) or dyed (Polysciences Inc., Cat # 24294) microspheres to an eppendorf tube and dilute 1:20 with 0.1× MMR.

2. Spin down for 1 min at 5000 rpm and aspirate off most of the liquid. Repeat the wash step to remove any remaining toxins and detergent that is contained in the storage buffer of the beads.

3. Resuspend beads in 100 μl of 0.1 × MMR with 0.5% glycerol. Beads tend to float in the media; therefore, 0.5% glycerol is added to the beads to accelerate their sinking onto the embryos when added to the Petri dish.

4. Using vacuum grease, form several small open arrowheads in a Petri dish (see Fig. 10.1C) and fill dish with 0.1 × MMR. For analysis of older embryos, 0.02% tricaine can be added to the 0.1 × MMR to anesthetize the embryos and prevent them from swimming away.

5. Transfer embryos into the Petri dish and using forceps or a hair loop push individual embryos into the open arrowheads with the anterior side facing the inside tip of the arrowhead (Fig. 10.1C). This should prevent the embryo from gliding across the dish due to ciliary beating.

6. Start imaging and add beads dropwise on top or slightly anterior of the embryo using a pipette. Beads should be displaced along the embryo when sinking into close proximity of the skin.

7. Velocity of fluid flow can be determined by tracking the displacement of individual beads either manually or automatically using simple particle tracking software (e.g., ImageJ). Because many cameras are equipped with rather basic acquisition software, it may be necessary to calibrate the acquired recordings when performing particle tracking since this information may not be contained in the acquired movie or may not be compatible with the tracking software. The careful use of uniformly consistent magnifications in combination with a micrometer allows one to overcome such calibration issues. The amount of displacement is a factor of both cilia-driven fluid flow and the mass of the beads. Therefore, it is critical to treat the velocities as relative rather than absolute and identically sized beads should be compared from control and experimental animals in each experiment.

3.3.2 Imaging of coordinated cilia beating using high-speed microscopy

To visualize beat frequency and beat form using nonfluorescent light microscopy, one needs a fast camera that captures >1000 fps (e.g., Vision Research Phantom class cameras or Casio EX-F1). However, visualization of the surface of the embryos requires some considerations. First, to obtain good cilia waveforms, it is important to visualize cells on the side of the embryo that are projecting into the media. This allows ample light to pass from the microscope to the camera, which is the primary challenge with high-speed imaging. Conversely, one can image explanted tissue (see Section 4.1) from the top. However, for this application, it is important to use either albino or very lightly pigmented embryos to allow a significant amount of light to pass through the tissue.

3.3.3 Visualization of cilia beating for quantification of beat frequency using fluorescent confocal microscopy

An alternative to high-speed cameras is the use of high-speed confocal microscopy. In contrast to high-speed cameras, confocal microscopy requires that individual cilia are fluorescently labeled (e.g., Fig. 10.2A). We have previously described the use of a fluorescently tagged protein (Xl.16654) which strongly and specifically labels cilia. However, any other ciliary protein that associates with the ciliary axoneme or the tip of cilia and does not interfere with cilia function when overexpressed can be used as a fluorescent label. In general, this approach requires a relatively bright signal and light-sensitive objective with high numerical aperture. While it is difficult to obtain high-resolution waveforms of cilia beating, it is quite easy to generate kymographs that allow quantification of beat frequency (Fig. 10.2B):

1. Inject embryos with 0.1–1 ng of mRNA encoding a fluorescently tagged ciliary protein at the two- or four-cell stage. If testing the effects of MO knockdown or overexpression of a certain protein, MOs or experimental mRNA can be coinjected with the cilia marker.

2. Allow embryos to develop to the appropriate stage (preferably stage +28).

3. Prepare pockets of electrical tape on large coverslips as described above (see Section 3.2). The working distance of the objective dictates the number of layers of electrical tape to be used, and the ideal mounting condition must be determined empirically.

4. Transfer an embryo into the pocket and overlay with a second 48 × 60 mm coverslip.

5. Image individual ciliated cells at the side of the embryo away from the coverslip. Optimally, a subset of cilia toward the side of the cell give the best results for quantification, as it is easier to distinguish individual beat patterns. In our experience, acquisition of at least 200–250 fps for 5 s is necessary to obtain reliable and quantifiable recordings for scoring of beat frequency (Werner et al., 2011).

6. To quantify beat frequency, use kymographs aligned along the axis of beating of individual cilia. In polarized ciliated cells, this leads to a periodic pattern of lines in the kymograph representing each individual stroke of multiple cilia (Fig. 10.2B). Beat frequency can then be calculated from the amount of lines (strokes of cilia) over a given time period. When cilia polarity or cilia function is compromised, it is essential to image cilia at the side of a ciliated cell that are the least likely to come into contact with neighboring cilia which will interfere with the ciliary stroke. Statistically significant data can be obtained from analyzing five cilia per cell, on five cells from five embryos obtained from at least two independent experiments (Werner et al., 2011).

3.4.1 Visualizing and scoring polarity of individual cilia

The axonemes of motile cilia are anchored within the cell via basal bodies and the associated striated rootlet. The rootlet is well characterized by electron microscopy; however, little is known about its molecular composition and function. Additionally, an appendage of the basal body, the basal foot, is known to project away from the basal body in the direction of cilia beating and has been used via TEM to score cilia orientation (Mitchell et al., 2007). Importantly, the rootlet projects 180° opposite from the basal foot (Park et al., 2008). It has been proposed that the rootlet not only helps anchor individual cilia in the cytoplasm, but it may also help ensure proper temporal coordination of the ciliary stroke between neighboring cilia by connecting their basal bodies via the cortical actin cytoskeleton (Werner et al., 2011). It has been shown that the orientation of individual rootlets relative to the associated basal body is guided by polarity cues during establishment of cilia polarity (Mitchell et al., 2009). Fluorescently labeled rootlet markers (e.g., GFP-CLAMP and GFP-Mig12) and basal body markers (e.g., centrin2-RFP, RFP-HYLS1, and RFP-SAS6) can be used to visualize the orientation of cilia in both live and fixed embryos (Fig. 10.2C):

1. Coinject embryos with 200 pg of GFP-CLAMP and 35 pg centrin-RFP mRNA per blastomere at the two- or four-cell stage. Add any other mRNA or MO to the injection for experimental analysis.

2. Let embryos develop to the desired stage and fix with 3–4% PFA for > 2 h to preserve the fluorescent tag.

3. Mount embryos as described in Section 3.2. Use forceps or hair loops to align the embryos along the long axis of the coverslip. It is advisable to mount embryos in the same orientation in each experiment to allow for a direct comparison of each experimental condition. This is particularly important if the confocal scan head or camera is not perfectly aligned relative to the XY axis of the microscope stage.

4. Image individual ciliated cells at magnifications of 40× or higher, acquiring several Z-planes on the surface of a ciliated cell to capture both the basal body and the rootlet which extends ~ 0.5 μm into the cytoplasm.

5. To score cilia orientation use maximum intensity projections to create a single image from each Z-stack (Fig. 10.2C). Nikon’s Elements software or Metamorph can be used to score cilia orientation by measuring the relative angle composed of the vector pointing from the basal body along the associated rootlet and a reference vector aligned along the anterior–posterior (A–P) axis. Alternatively, it is possible to determine the relative XY coordinates of each basal body and associated distal tip of striated rootlet using ImageJ and then calculate the individual vector relative to the A–P axis using the atan2 function in Microsoft excel. Oriana 2.0 (Kovach Computing Services) is a useful program for the analysis of circular vector data sets and can be used to generate circular standard deviation, variance, and mean direction (Mitchell et al., 2007).

Day 1

• 1

  Prepare a 20 mg/ml dilution of fibronectin in 1 × PBS (stock solution aliquots of 1 g/ml can be stored at −20 °C).

• 2

  Place a sheet of wet Whatman paper in a 15-cm Petri dish for moisture and place a large glass plate on top.

• 3

  Flame 22 × 22 mm pieces of coverglass after immersing in 100% EtOH to sterilize and place on the glass plate.

• 4

  Place three to four small spacers cut from a sheet of plastic (1 mm thick) on the edges of each piece of coverglass (Fig. 10.3A).

• 5

  Add 200 μl of fibronectin to the center of the coverglass (Fig. 10.3A).

• 6

  Place a second coverslip on top of spacers. This evenly distributes the fibronectin and generates two usable coverslips with a smaller volume of fibronectin (Fig. 10.3A).

• 7

  Leave at RT for 30–45 min and then place at 4 °C ON.

Day 2

• 8

  Place each coverslip individually in a 35-mm Petri dish making sure that the surface exposed to fibronectin is up.

• 9

  Wash coverslips for 5 min in 1× PBS.

• 10

  Block coverslips for 1 h in 1× PBS+1% bovine serum albumin (BSA).

• 11

  Wash coverslips for 5 min in 1× PBS.

• 12

  Remove PBS and fill Petri dish with Danilchik’s for Amy (DFA) buffer+0.1% BSA+antibiotic (50 μg/ml gentamicin).

Preparation of explants

1. Transfer embryos into Petri dishes containing DFA buffer+0.1% BSA+antibiotic.

2. At the desired stage, carefully remove the vitelline membrane.

3. Using an eyebrow knife, remove a part of the ectoderm near the animal pole that will give rise to the ciliated epidermis (Fig. 10.3B) (Sive et al., 1998).

4. Make sure to trim away any nonectodermal tissue. With explants cut from stage 10 to 16 embryos, it is easy to separate the ectoderm from the underlying mesoderm and endoderm. In later explants, these tissues are more intimately associated and it is significantly more difficult to obtain pure ectodermal tissue.

5. Transfer the explant to the 35-mm Petri dish containing a fibronectin-coated coverslip using a wide bore transfer pipette. It is important to prevent the explant from coming into contact with the air–water interface as this will cause it to quickly disintegrate.

6. Carefully move the explant to the center of the coverslip with the outer ectodermal layer facing up.

7. Use a diamond knife to cut a small glass bridge from a 22×22 mm coverslip.

8. Add silicone grease to the ends of the glass bridge and carefully place the glass bridge over the explant. Carefully press down on the coverglass until the explant is held in place by the glass bridge (Fig. 10.3B).

9. Set aside control embryos that are the same developmental stage as the embryo from which the explant was obtained and keep them at the same temperature as the explant. These control embryos can later be used to determine the developmental stage of the explant.

10. Allow explants to attach to the coverslip for several hours at RT or 12 h at 16 °C.

11. Carefully remove the glass bridge using forceps without putting direct pressure on the explant.

12. When control embryos have reached the desired stage, explants can be fixed or used for live cell imaging.

Fixed explants can be imaged after IF staining by simply inverting the coverslip on which the explant is attached onto a second larger coverslip using dots of vacuum grease to hold the coverslip in place. Similarly, for live imaging of cilia or ciliated cell-associated proteins, coverslips can be inverted onto larger coverslips with a half-rose chamber (e.g., Fig. 10.1B) using dollops of vacuum grease to control the distance between the two coverslips. Fluid flow along explants can be visualized directly in Petri dishes using labeled microbeads (see Section 3.3.1). Alternatively, explants can be attached to glass bottom Petri dishes and displacement of fluorescent beads can be imaged through the relatively thin explant using an inverted microscope. Finally, the explants are amenable to studying the effects of external hydrodynamic forces using a customized flow chamber (Mitchell et al., 2007).