“Freeze-cracking,” a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.
To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. “Freeze-cracking” is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.
To determine the cellular and subcellular localization of proteins, scientists have traditionally labeled tissues with antibodies selected to specifically recognize particular proteins1. In some model organisms such as C. elegans, antibody staining has often been replaced by molecular genetic techniques, which yield results more quickly. These include transforming organisms with constructs consisting of gene fusions between the promoter and coding regions of the gene of interest and green fluorescent protein. However, molecular techniques are subject to a number of artifacts, including problems with knowing the true promoter and changes in expression of constructs at high copy number (the usual techniques in C. elegans)2,3. Therefore, staining with antibodies remains a goal for many scientists studying protein function in vivo.
Staining tissues with antibodies can be difficult, since antibodies may only recognize their antigen in a particular conformation1. For example, antibodies may recognize only denatured or intact antigen fixed in a particular way and may not recognize the antigen in situ. The problem of antibody staining in nematodes is exacerbated by the fact that the cuticle of the nematode forms a relatively impermeable barrier, blocking access of antibodies to tissues.
There are several different methods used for antibody staining in C. elegans (reviewed in our previous work4). To stain intact 'worms,' methods were developed to freeze and thaw in a relatively hard fixative (formaldehyde or glutaraldehyde); the freeze-thaw cycles help to crack the cuticle to allow rapid penetration of the fixative5. After fixation, the cuticle was permeabilized to allow penetration of the antibody; methods included treatment with reducing agents, collagenase, or both5-7. These treatments preserved morphology, but often reduced or destroyed antibody recognition. Alternative methods include dissection8 to allow antibody penetration.
"Freeze-cracking" is one way to gain access to the interior of the semi-intact worm to allow antibody staining9-10. Freeze-cracking can be performed with a variety of fixation conditions while avoiding collagenase and reduction treatments. The nematodes are placed between two adhesives slides, frozen, and then the slides are separated, leaving most of the nematode on the bottom slide and much of the cuticle on the top slide. The bottom slide can be placed in fixative and the entire slide with adhered nematodes is transferred through the antibody staining procedure. The method does present two major difficulties. First, it is difficult to apply the correct amount of pressure necessary to split the nematodes without seriously deforming them. Second, many of the worms will not stick to either slide, and will be lost in the fixative or rinses. However, with the proper slides and practice, the method will rapidly yield nematodes with reasonable morphology which can be used with a wide variety of fixatives and antibodies.
The method may be varied slightly, depending upon the goal of the experimenter. If staining of single nematodes is desired (e.g. to determine whether a single transformant has altered antibody staining), then slides with maximal adhesion (but higher background staining) can be used, such as laboratory-prepared slides with extra polylysine (see below). For formaldehyde or glutaraldehyde fixation, higher adhesion slides (laboratory-prepared polylysine slides) should be used, since nematodes adhere more poorly to polylysine after these fixations. If wild-type nematodes are being fixed with methanol and/or acetone, then slides with lower adhesion and lower background staining should be used (commercial or laboratory-prepared slides). If a variety of antibodies and fixatives are being used in the lab, a selection of slides may be prepared ahead of time and stored until needed.
After access to the nematode tissue has been gained, antibody staining procedures follow standard methods (with longer incubations characteristic of tissues rather than cells1,11).
NOTE: Multiple protocol steps are presented with options for set-up and preparation depending on the specific conditions of the experiment. For these cases, alternative steps are presented as A, B, C, etc. The protocol with alternative steps is outlined in Figure 1.
1. Preparation of Polylysine Coated Slides
Three different types of slides may be used, depending upon the desired trade-off between ease of preparation, relative adhesion, and non-specific binding of the antibody. Details of slide preparation alternatives are given below in steps 1A-1C in order of increasing adhesion and complexity. Slides prepared by these different methods can be used together for a single experiment.
1A. No slide preparation
Commercially available polylysine coated slides provide low adhesion and low background.
1B. Slide preparation optimum for methanol-acetone fixation
Medium adhesion and low background.
1C. Slide preparation optimum for formaldehyde fixation
Highest adhesion but high background.
2. Preparation of Frozen Nematode Slides
Prepare nematodes for staining by completely rinsing free of bacteria using method 2A (for plates of nematodes) or 2B (for individual nematodes).
2A. Preparation of slides from plates of nematodes
2B. Preparation of slides of individual nematodes
3. Fixation
Fixatives are necessary to 'fix' the antigen in place in the cell, by either precipitating ("light fixation") or cross-linking ("hard fixation") the antigen. Fixation may disrupt antigenicity; most known antibodies work best with a particular fixation condition. For new antibodies, a range of fixation conditions should be tested.
For any fixation, the first step is to make phosphate buffered saline solution. Next fix slides with nematodes for antibody staining using method A (light fix using methanol and acetone) or B (harder fix using formaldehyde or glutaraldehyde).
3A. Light fix using methanol-acetone
3B. Hard fix using formaldehyde or glutaraldehyde
4. Antibody Staining
The antibody staining protocol is similar to standard protocols for any tissue on slides. This protocol is the same for all slides regardless of the preparation in steps 1-3.
When worms are properly compressed, cracked, and lightly fixed, virtually the entire worm can be accessible to antibody staining (see Figure 2). The location of the nuclei as indicated by DAPI staining indicates which parts of the worm are intact When the worms are subjected to a harder fixative, such as formaldehyde, the morphology of the worm may become distorted (as seen by the unnaturally wavy appearance of the muscles in Figures 3A-D). Another common problem is uneven fixation or penetration of particular tissues. An example of this is shown in Figures 3E-H, where the muscle is stained only in a portion of the worm, despite the fact that the presence of other staining, including DNA staining, indicates that at least part of the body was present (i.e., not removed during freeze-cracking). The resulting pattern of partial staining can be readily identified with practice, so that the entire distribution of staining can be determined even if any single worm shows uneven staining.
Common problems encountered with freeze-cracking using any fixation condition are illustrated in the final figure (Figure 4). The most common problem is twisting of the sample due to relative motion of the two slides during compression. The fact that the body of the worm is twisted just posterior of the pharynx can be seen by the morphology of the stained tissues. This image also shows the problem with background staining, due to antibody sticking to the poly-lysine coating the slide. Note, however, that all of these images were collected using slides made during the first antibody staining attempts of undergraduates in a cell biology laboratory course. Even with practice, many individual worms will show the problems seen in Figure 4. However, on any one slide, worms like those seen in Figures 2 and 3 can be found and examined.
Figure 1. Outline of procedures. Flow chart indicating steps to perform the complete freeze-cracking procedure. Alternatives for varied fixation conditions and numbers of worms are indicated.
Figure 2. Lightly fixed, well-stained worms. Heads of two adult hermaphrodites fixed with methanol-acetone and labeled with multiple antibodies and dyes: A) Primary antibody to ChAT (choline acetyltransferase) and Cy3-conjugated secondary antibody, B) DAPI (blue DNA), C) Oregon Green 488-anti-GFP (green fluorescent protein), D) shows the overlay (with two cholinergic markers, Cy3-anti-ChAT and Cy5-anti-VAChT (vesicular acetylcholine transporter) shown as red). This transgenic strain (PS4657) expresses a translation fusion between GFP and AJM-1, found at pharyngeal apical junctions. The images are maximal projections of series of confocal images; scale bar is 50 μm.
Figure 3. Formaldehyde-associated distortions. Formaldehyde fixed adults stained with Cy3-phalloidin (binds filamentous actin), DAPI (blue DNA), and Oregon Green 488-anti-GFP. A-D) Fours images of the body just posterior of the vulva (far left) in the same nematode, showing an unnaturally wavy morphology due to distortion induced by fixation. A) Red-phalloidin shows slightly irregular muscle morphology. B) Nuclei (blue) are evenly spread throughout the worm. C) Green shows the distribution of a CED-1::GFP fusion protein in the plasma membrane of the gonadal sheath cell, driven by the lim-7 promoter (strain MD701). D) Shows the overlay of these three dyes. E-H) Labeling can vary with different dyes on the same nematode. E) Phalloidin (red) labels only a portion of the muscle on one side of the worm. F) Nuclei (blue) are present throughout the worm (although stained with uneven intensity). G) The OG488-anti-GFP labels collagen-19::GFP in the cuticle on a side of the worm that lacks staining of the more central muscle tissue, indicating that muscle is present but not staining on one surface. H) Shows the overlay of the three dyes. The images are maximal projections of series of confocal images; scale bars are 50 μm.
Figure 4. Compression-associated distortions. Formaldehyde fixed larva stained with A) Cy3-phalloidin (binds filamentous actin), B) DAPI (blue DNA), C) Oregon-Green 488-anti-GFP, D) overlay. The twist in the body just posterior of the pharynx is apparent, as the green excretory cell and ventral nerve cord cross over the rest of the body. High background staining is also apparent with Cy3-phallodin. This strain expresses a VHA-8::GFP fusion protein, located predominantly in the excretory cell. The images are maximal projections of series of confocal images that extended to the surface of the slide (leading to high background staining); scale bar is 50 μm.
The freeze-cracking protocol is one of several methods for antibody staining in C. elegans. It provides a relatively simple way to stain worms, but does require specific reagents and practice for optimum results. Critical steps (outlined in Figure 1) include: 1) slide preparation (see steps 1 and 2) manual compression of the nematodes (see steps 2 and 3) rapid fixation (see step 3). First, to maximize adhesion of the nematodes to the slides, it is best to prepare slides with high molecular weight (long polymer) polylysine, instead of relying on commercially prepared slides. Commercial slides are coated with polylysine to allow cells to adhere, while freeze-cracking slides are designed to stick to the cuticle of entire nematodes. Second, the procedure for the correct compression of the worms between the slides before freezing is critical. It requires steady hands and practice to press the slides together using the correct amount of pressure so that individual worms contact both slides without destroying their morphology. Either too much or too little compression leads to increased loss of worms during fixation. Further care is needed to move the slides onto the dry ice chilled surface for freezing without moving the slides relative to one another. Any such motion will cause the worms to tear or twist. Third, as with any fixation technique, the frozen worms should be placed directly into fixative before defrosting. Otherwise, the antigen may diffuse, providing misleading information about subcellular localization. If these critical steps are all done properly, for best results it is still necessary to scan several stained slides to find worms with the best morphology.
One unavoidable aspect of this technique is that the worms will be compressed in one dimension. The compression is most noticeable in adults, where the apparent diameter of the round body in the z-axis may be only one-fourth that seen in the x- and y-axes. This compression tends to decrease in smaller larvae, and can be negligible in embryos. Due to the way the living nematodes move during slide preparation, the stained worms are often on their sides. This mimics the orientation of worms on agar dishes, with the lateral midline lying extending down the middle of the stained worm (see Figures 2-4). Thus, the compression is generally in the lateral dimension. While this actually makes standard fluorescent microscopy easier (more of the specimen will be in focus simultaneously), it means that 3-D reconstructions will not be accurate.
As with any antibody staining, it is important to check specificity of binding. In most species, this is done by controls such as affinity depleting your antibody or using no primary. In C. elegans, more specific means of checking for specificity are often available. Null mutants for the gene and protein of interest provide an excellent control for staining specificity. Both strains should be fixed under identical conditions before comparison, since staining is usually fixation dependent. If no null mutant is available, then it is relatively easy in C. elegans to decrease protein levels with RNAi (RNA inhibition12) and look for decreased staining in RNAi treated worms.
Polyclonal antibodies often show non-specific staining, even if affinity-purified with the antigen. Often, light fixation using methanol and acetone (which fix proteins by precipitating them rather than crosslinking them) gives lower non-specific staining than formaldehyde or glutaraldehyde (which generate more variant epitopes due to varied cross-linking)1,11. However, in C. elegans non-specific staining is very common under any fixation condition, especially in the gut. If your antigen is expressed in the gut, then this non-specific staining may mask your specific staining. If null mutants are available for your protein, then worms fixed with this method can be used to affinity deplete your serum. Since antigenicity depends upon fixation conditions, the worms that you use to deplete your antibody should be prepared in the same way as the wild-type worms you are staining. After one round of depletion of non-specific staining, mutant and wild-type worms can be stained in parallel with the depleted serum for an excellent control. Additional round of depletion with mutants can be done as necessary.
This technique is very useful for testing new antibodies for specific staining under a variety of conditions. These may be antibodies generated against C. elegans proteins or antibodies generated against conserved proteins from other species. While this method gives the best morphology with lightly fixed nematodes, it is relatively simple to try a range of fixatives to determine which conditions, if any, give specific results. Unlike other methods for hard fixation using formaldehyde or glutaraldehyde, enzymatic or reducing treatments are not needed for sample preparation. Since these treatments may decrease antigenicity, this increases the ability to identify specific antibodies and their optimum fixation conditions. If hard fixation conditions work best, the experimenter can then turn to methods performed on intact worms to obtain better morphology (summarized in our previous work 4). If light fixation preserves antigenicity, then this technique may be used for all further staining for light microscopy.
The authors have nothing to disclose.
Funding was provided by NSF CCLI#0633618 and Ohio University. Some strains were provided by the Caenorhabditis Genetics Center. Graduate student Reetobrata Basu appears in the video.
Name of Reagent/Material | Company | Catalog Number | Comments |
poly-L-lysine slide (ESCO brand) | Fisher | 12-545-78 | Sigma brand slides also suitable |
poly-L-lysine hydrobromide | Sigma | P1524 | IMPORTANT: Brand and high molecular weight of polylysine critical |
single frosted edge slides | Fisher | 48312-002 | Other brands are suitable |
sodium azide | Sigma | S2002-25G | TOXIC, wear gloves and mask while weighing out powder to make 10% stock. Wear gloves when pipeting stock solution |
coplin staining jar | VWR | 47751-792 | Other brands are suitable |
5-slide mailer | Electron Microscopy Science | 71549-01 | One of many different brands of small slide holder, suitable for staining |
Paraformaldehyde | VWR | MK262159 | IMPORTANT: Regular formalin solutions (37% formaldehyde) are NOT suitable. Either 1) Dissolve crysatlaline paraformaldehyde in phosphate buffer and store in small aliquots frozen. Or 2) buy electron microscopy grade paraformaldehyde in aliquots. TOXIC -wear gloves; dispose of excess down sink with water. |
Glutaraldehyde solution, 25% in water | Sigma | G 5882 | IMPORTANT: purchase electron microscopy grade glutaraldehyde in ampules; TOXIC -wear gloves; dispose of excess down sink with water. |
Triton X-100 | VWR | EM-9410 | Other brands are suitable |
Bovine Serum Albumin | Fisher | ICN820451 | Other brands are suitable |
Donkey Serum, lyophilized | Jackson Immunoresearch | 017-000-121 | Other brands are suitable |
Cy3 Donkey anti-Mouse IgG multi-labeling | Jackson Immunoresearch | 715-165-150 | This company is recommended for high quality multi-label (affinity depleted) antibodies. Select appropriate dye and antigen (animal used to raise primary antibody). |
n-propyl gallate | Fisher | ICN10274750 | Other brands are suitable |
DAPI, 4′,6′-diamidino-2-phenylindole dihydrochloride | Sigma | D 9542 | TOXIC. Wear gloves and dispense into small aliquots and feeze to minimize exposure |
Whatman #1 15 cm diameter filters | Fisher | 09805G | |
Coverslip #1 1/2 rectangle 24 x 60 mm | VWR | 48393-252 | #1 1/2 thickness is optically best |
Microscope slide folder | VWR | 48429-092 | Other brands are suitable |