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Table of Contents
Although C. elegans is primarily touted for its facile genetics, there has been a burgeoning interest in studying cell biological processes in this organism. Strong genetics (Brenner, 1974; Jorgensen and Mango, 2002), the development of fluorescent protein tags (Chalfie et al., 1994; Yang et al., 1996; Zhang et al., 2004), the availability of RNA interference strategies to disrupt gene function (Fire et al., 1998; Timmons and Fire, 1998), and the ability to perform studies on primary cultures of embryonic cells (Christensen et al., 2002; Zhang et al., 2002), have all led to an increase in the number of cell biological problems addressed in the worm. Furthermore, the transparency of the organism affords a unique opportunity to study the roles of cell biological processes in a living multicellular animal.
A serious obstacle to studying cell biological phenomena in C. elegans is the small size of its cells. However, advances in imaging techniques have allowed faint signals to be significantly amplified, and small structures to be visualized, allowing examination of transport, export, and import processes, as well as examination of cytoskeletal and chromatin structure.
Here we have compiled a set of protocols that broadly fit under the category of Cell Biology. We begin with a brief discussion of various microscopical techniques employed by C. elegans researchers to study aspects of the cell. We then describe methods for studying protein-protein and protein-DNA interactions in C. elegans. We also describe methods used to study specific cell biological problems (e.g. endocytosis, chromatin, programmed cell death). Finally, we conclude this chapter with a discussion of primary embryonic cell culture and its uses.
Contributors of sections or protocols are acknowledged in parentheses following the section or protocol titles. In some cases, protocols are based on previously published work, which is then cited after the protocol title. The on-line format of this chapter should easily allow for revisions and additions to the protocols presented here. Researchers willing to share protocols not presented here or with comments on existing protocols are encouraged to submit these to shaham@rockefeller.edu.
To study objects we must interact with them. Generally, the technique of interaction is determined by the size of the object. Thus, an object of macroscopic size can be studied by direct contact. However, microscopic objects, such as the cell and its organelles, must be studied with agents of similar or smaller size. Cells in C. elegans are roughly 3–30 microns in diameter, thus, light with wavelength in the visible range (~500 nm) is an ideal interacting agent. The set up of light microscopes affords a resolution that is about half the wavelength of light employed. Thus, light microscopy is useful for examining cells and cellular substructures on the order of 200–300 nm or larger. However, vesicles (often 50 nm in diameter), and other objects of similar or smaller scale cannot be resolved using current setups. Although, in principle, light of very short wavelength (e.g. X-rays) could be used to resolve smaller cellular structures, such light is too energetic, damaging the cell upon contact. In addition, lenses to focus high energy photons do not exist. High resolution can be obtained using electron microscopy. Moving electrons in an electron microscope possess wavelengths on the order of 0.3 nm. Using an electron microscope, the electrons can be used to form resolved images of cellular structures of about 3 nm in size.
Visible light can be used to examine C. elegans, however, in general, bright field and phase-contrast microscopy offers little contrast- making cells and their major components difficult to see. DIC microscopy, however, allows high contrast images to be formed, and is ideal for examining nuclei, nucleoli, and granular structures within C. elegans cells (Sulston and Horvitz, 1977; Sulston et al., 1983). In DIC microscopy light passes through a plane polarizer. The resulting polarized light is split into two components using a prism. These components interact with the sample, and are combined above the objective using a second prism. Finally, the transmitted light passes through an analyzer (essentially a polarizer that is polarized at 90 degrees with respect to the original plane of polarization) and on to the observer. The interaction of polarized light with the sample changes its plane of polarization. Thus, only light whose polarization is changed by the sample will be detected by the observer. In this way, highly contrasted images are formed. It should be remembered that shadowing effects seen using DIC microscopy do not reflect three-dimensional features of the sample. Rather, they reflect the ability of the specimen to interact with polarized light. Current microscopes allow the use of a 100X objective with DIC, giving high magnification and resolution. Some microscopes are equipped with additional lenses, however, these do not afford better resolution.
A protocol for mounting C. elegans for observation using DIC microscopy is presented below. This protocol can be used to mount animals for all microscopic techniques utilizing a compound microscope.
Protocol 1: Mounting animals for observation with DIC optics (Monica Driscoll)
1. Preparation of Agar Pads:
a. Materials
5% agar solution in water, melted and kept molten by placing the tube in a heat block at 65°C
Pasteur pipette and bulb
2 glass slides with pieces of labeling tape (for example, Fisher #11-880-5-D) taped over both ends to serve as spacers
2 clean glass slides
b. Preparation of Pads (See Figure 1):
Place the two taped slides with a clean slide sandwiched between them on a flat surface. Using the Pasteur pipette, place a drop of agar onto the clean slide. Cover the agar with another clean slide placed on top of the three slides in a perpendicular fashion. Press gently so the agar drop is flattened to a circle about 0.4 mm thick (the thickness of the tape spacers). Avoid getting bubbles in the agar since worms will get stuck in them. After the agar solidifies, gently pull out the taped slides, then separate the remaining two slides by sliding one relative to the other. The agar pad should adhere to one of the slides (usually the bottom one). Rest the slide, agar side up, on the bench top.
Note: The agar pad should be prepared just before use so that it does not dry out. Alternatively, once you pull out the taped slides, you can wrap the cross shaped slide-agar-slide in a piece of Saran wrap. This way you can keep the pad moist for a couple of hours.
2. Mounting Live Animals:
Place a 1–2 ul drop of M9 containing 10–25 mM sodium azide (NaN3) onto the center of the agar pad. NaN3 anesthetizes the worms so that they will not move. The agar pad can also be prepared with anaesthetic included for a final concentration of 2–10 mM NaN3 in the agar, instead of in the drop. This generally results in faster anesthetic action. If live worms are needed, NaN3 can be omitted and bacteria can be added to the drop to allow the worms to feed with little motion.
Transfer animals to be observed into the drop. Animals can be transferred using a worm pick or an eyebrow hair fastened to a toothpick with wood glue or clear nail polish. When the hair or pick is moved into the drop, animals float off easily. Alternatively, animals can be spun down in M9 in an eppendorf tube at 1200–1500 rpm with slow acceleration/deceleration. After the supernatant is discarded, the animals are suspended in 10 ul M9-NaN3. 2–3 ul of the solution with the animals can then be transferred to the pad.
Gently lay a coverslip over the animals. Most animals will lie on their left or right sides. Embryos also generally assume stereotypic orientations on agar pads. Between the 4-cell stage and 100–150 minutes the embryos display either the left or right sides. At gastrulation (150 min) they turn from left to dorsal or from right to ventral. At 350–400 minutes they return to display left or right sides (Sulston et al., 1983).
Animals will stop moving within a minute or two. Animals in sodium azide can recover following incubation in sodium azide of one-hour or less.
Polarized light can be used to examine repeated or crystalline structures. In a manner similar to DIC, plane polarized light that passes through a specimen will interact with the specimen and change its plane of polarization, allowing light to be detected through an analyzer located above the specimen and oriented at 90 degrees to the initial plane of polarization (for a more complete description see http://www.microscopyu.com/articles/polarized/ polarizedintro.html).
The muscles used for locomotion in C. elegans reside in the body wall. In the adult, there are 95 spindle shaped cells divided among four quadrants just underlying a basement membrane, hypodermis and cuticle. In each quadrant, the cells are arranged in interlocking pairs. In these muscle cells, myofilaments form a lattice that is restricted to a narrow zone of ~1.5 microns, just underlying the basement membrane and hypodermis. By polarized light microscopy, obvious striations are seen; bright (“birefringent”) A-bands alternate with dark I-bands; each I-band contains a row of dense bodies, which are the analogs of Z-discs of vertebrate striated muscle (Figure 2). Because the striations lie at a slightly oblique angle with respect to the long axis of the worm, this muscle is called “obliquely striated”. Polarized light is also useful for evaluating the second largest set of muscles, those in the pharynx. Below we present a protocol for observing C. elegans muscle using polarized light.
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Figure 2. The top three panels show polarized light microscopy of adult body wall muscle from wild type and from two muscle-affecting mutants. Each image was captured with a 40X objective and shows portions of about three muscle cells. The inset shows an enlarged view. Wild type muscle shows a high degree of organization with birefringent A-bands alternating with dark I-bands, each containing a row of dense bodies. unc-89(su75) exhibits the phenotype of a typical unc-89 mutant: reduced organization of the myofilament lattice with an almost “basketweave” pattern. unc-98(sf19) displays disorganized myofibrils with characteristic birefringent “needles” at the ends of the muscle cells. The bottom panel is an illustration showing the position of the four muscle quadrants (right) and a portion of a single muscle cell emphasizing the myofilament lattice. The oval indicates the plane of view seen by polarized light (Denise Flaherty and Guy Benian).
Cellular structures and components can be labeled with a variety of fluorescent dyes that can be visualized using a fluorescence microscope. In response to incident light of specified wavelength (produced by a laser or using appropriate filters), fluorescent dye molecules are excited. Decay of the molecule from the excited state to an intermediate excited state produces a photon of a wavelength longer than that of the excitation source, which can easily be detected and separated, using appropriate filters, from the incident light (for a more detailed description see http://www.microscopyu.com/articles/fluorescence/fluorescenceintro.html). Below are a number of protocols for using dyes to visualize DNA, proteins and cellular structures. Dyes coupled to antibodies are particularly useful for visualizing the presence and localization of specific proteins within the cell, and several methods for antibody staining are discussed in the chapter on gene expression. Often it is of interest to study whether two cellular molecules physically interact. Fluorescence resonance energy transfer (FRET) has been used to indicate the proximity of two labeled fluorophores. However, this method has not been significantly used in C. elegans to date.
A number of fluorescent dyes can be used to visualize DNA in fixed animals. 4',6'-diamidino-2-phenylindole hydrochloride (DAPI), is by far the most commonly used DNA dye in C. elegans (Figure 3).
Amphid and phasmid neurons can take up lipophilic dyes from the environment (Hedgecock et al., 1985; Perkins et al., 1986). These dyes will label all parts of the neuron. The mechanism of dye uptake is unclear, but seems to correlate with some aspect of neuronal function. Thus, these dyes can be used to visualize neurons and perhaps to study their physiology (Figure 4). The dye fluorescein isothiocyanate (FITC) can be used to label the ADF, ASH, ASI, ASJ, ASK, and ADL cells of the amphid sensilla, and the PHA and PHB neurons of the phasmid sensillum. DiI, DiO, and DiD (Molecular Probes) can be used to visualize the ASI, ADL, ASK, AWB, ASH, and ASJ amphid neurons, as well as the PHA and PHB phasmid neurons (Collet et al., 1998).
Filamentous actin can be readily observed in C. elegans muscle by staining with fluorescin-conjugated phalloidin (Strome, 1986). Other actin structures can also be visualized using this method, however, Protocol 7, below, should be used.
The use of fluorescent proteins to tag cellular proteins has revolutionized cell biology (Zhang et al., 2002). In C. elegans, green fluorescent protein (GFP) and its derivatives, and more recently DsRed, have been employed to visualize protein localization, movement, and conformational changes in vivo (Chalfie et al., 1994; Miyawaki, 2002). Fusions to any protein can be generated using sets of vectors constructed by Andy Fire and others (Miller et al., 1999). In addition, labeling of specific organelles can be accomplished using pre-made vectors containing GFP fused to specific subcellular targeting sequences. Localization signals that are frequently used include those for the nucleus, mitochondria, and plasma membrane. Details regarding fluorescent proteins can be found in the gene expression chapter.
Examining protein localization using GFP or DsRed fusion proteins requires injection of corresponding transgenes into animals. Thus, a number of caveats exist in interpreting results from such experiments. For example, expression from transgene arrays is often higher than from the endogenous locus. Furthermore, fluorescent proteins are fairly bulky, thus, fusion proteins may not display appropriate localization. In addition, promoter elements used to drive expression of GFP transgenes are often expressed in cells where the endogenous protein is normally not made, or vice versa.
Fluorescently-labelled antibodies generated against a protein of choice allow direct examination of endogenous unmodified proteins, giving a clearer assessment of where endogenous expression is localized. Antibodies coupled to gold particles can also be used for electron microscopy to examine subcellular localization in great detail. Antibodies can only label fixed tissues, thus, they cannot be used for real time observation of protein localization dynamics. In addition, fixation often destroys cellular structure, limiting the resolution obtained by immunostaining. Detailed protocols for use of antibodies can be found in the gene expression chapter.
A number of microscopes are now commercially available for generating and examining fluorescent signals. All are able to resolve fluorescent entities both in space and in time. Differences among these microscopes center on spatial and temporal resolving power.
This binocular microscope allows visualization of highly expressed fluorescent signals at low magnification. The major use of the instrument is to visualize GFP and DsRed-labelled proteins in living worms on the surface of standard agar plates. Generally, use of this microscope allows selection of animals expressing a fusion protein of interest, as well as examination of very coarse cellular features (e.g. presence or absence of a neuronal process) for genetic screens (see for example Shaham and Bargmann, 2002). This method does not allow fine structure to be examined. The microscope consists of a standard binocular dissecting microscope equipped with epifluorescence.
This microscope has significantly higher resolution and magnification compared to the dissecting fluorescence microscope, and observation can be combined with DIC (Figure 5). Objectives of 100X are often used, and allow examination of gross subcellular localization of proteins. Images collected from such a microscope at high magnification suffer from an extensive background of fluorescent light emitted by cells not in the plane of focus and of scattered light. This often severely cuts down on the resolving power of this microscope. However, for first pass examination of samples, or for examining highly localized signals, this microscope is often the instrument of choice (for a more detailed description see http://www.microscopyu.com/articles/fluorescence/fluorescenceintro.html).
This instrument is essentially a compound fluorescence microscope equipped with a sensitive light detection system. Light collected from optical sections through the sample is processed using sophisticated computer software, which assigns out-of-focus light to its correct focal plane (Wallace et al., 2001). Images generated by this microscope consist of optical sections through a sample, and allow 3-dimensional reconstruction of the fluorescent signal with little background. Sections can also be projected onto a single image, giving highly resolved artificial two-dimensional images (Figure 6). All images are viewed on a computer screen. Time-resolution of the technique is, in general, inversely proportional to spatial resolution. Thus, the more optical sections imaged, the larger the time intervals between image collections at any given focal plane. This type of microscope is not efficient for examining processes that occur over intervals of seconds or faster.
This instrument consists of a compound microscope equipped with a laser for fluorophore excitation, and a special detection set up consisting of a small pinhole through which emitted fluorescence light produced only near the focal plane of observation can pass. Light passing through the pinhole is collected in a sensitive light detector, and images are produced on a computer screen. This technique eliminates out-of-focus fluorescence, giving sharp images with little background (for more details see http://www.microscopyu.com/articles/confocal/). As with the deconvolution microscope, images consist of optical sections through the sample. Time-resolution of this microscopy is affected by several factors, including the time required for the laser to scan the field of view, and the amount of light that needs to be collected to visualize fluorescence. Unlike the deconvolution microscope, which uses all the light emitted by the sample to calculate the point of emission, a confocal microscope only collects light traveling through the pinhole, necessitating longer exposures or more sensitive light-detection systems.
This instrument is a variation of the laser scanning confocal microscope. Here, many pinholes are arranged in a spiral pattern on a disc that can rotate at high speed. Thus, time resolution can be excellent (on the order of l00 msec or less), and photobleaching is generally not a significant issue as with other microscopes (Nakano, 2002). Image display is essentially as with the standard confocal microscope (Figure 7).
The output from this microscope is similar to that from deconvolution or confocal microscopes. Thus, optical slices of images are generated that can be reconstructed to form a three dimensional image. Multiphoton microscopy eliminates out-of-focus light by directing excitation to the focal plane. This is accomplished by shining photons of long wavelengths and at high density onto the sample. At the focal plane long wavelength photons can superimpose to become, effectively, a shorter wavelength photon. This photon can now excite the fluorophore and induce fluorescence. Superposition of photons occurs significantly only at the focal plane. Multiphoton microscopy is also useful for looking at samples that are much thicker than confocal or deconvolution samples, with nearly equivalent resolution (Helmchen and Denk, 2002; Michalet et al., 2003).
Electron microscopy (EM) is currently the method of choice for examining subcellular structures in C. elegans. The resolution offered by the technique is unparalleled, allowing objects as small as, or even smaller than ribosomes to be viewed. Although EM is technically demanding, it is well worth the effort if resolution of small structures is of importance. Transmission EM (TEM) generally involves fixation of animals with any combination of gluteraldehyde, paraformaldehyde, or osmium tetraoxide (OsO4), embedding of fixed animals in a special resin, sectioning animals (sections are usually 50–100 nm thick), staining, and imaging on an electron microscope (Hall, 1995; Figures 8 and 9). Fixation conditions are generally the key to a successful EM experiment. Standard treatments with fixatives are adequate for most applications, however, they have the disadvantage that cellular structures are often slightly distorted because of osmotic imbalance and other damage that occurs during the fixation procedure. High Pressure Freezing (HPF) has recently been employed to avoid these issues. Tissues and cells are generally rounder, and the ultrastructure seems to reflect the in vivo structures more accurately (Figure 10). Because HPF fixation has only recently come to prominence in C. elegans, a number of issues still remain to be addressed with the technique. Specifically, results are often more variable than using standard fixation. In addition, embryos and L1 larvae seem to fix better than older stages. Interestingly, embryo fixation using standard methods is fairly difficult, thus, HPF and standard techniques are somewhat complementary.
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Figure 8. TEM micrograph of a longitudonal section of the C. elegans nose (Erika Hartwieg and Bob Horvitz).
Because fixation is so critical for good ultrastructure, we present below several alternative protocols for TEM. At the end of the section we present a method for scanning EM (SEM) of worms. In this method, whole-mount animals or structures are viewed by EM, offering both broad-scale and highly resolved images (Figure 11).
The protocols below are intended to give a sense of the types of fixation procedures used for EM and should be used as guidlines for those with previous EM experience. These protocols are not intended for teaching EM from scratch.
