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Table of Contents
Immunohistochemistry in C. elegans includes two major classes of techniques: immunocytochemical localization of proteins in situ and Western blot analysis of purified proteins. In this section, we will discuss a variety of techniques for immunocytochemistry, as well as simple Western blotting techniques for worm proteins.
Immunocytochemistry provides the most direct method for identifying both the cellular and subcellular distribution of your protein. Although GFP or other translational or transcriptional constructs can provide a relatively rapid indication of gene expression or protein distribution, there are a number of potential problems with these techniques (Mello and Fire, 1995). These include abnormal regulation of expression from multi-copy transgenic arrays and abnormal localization of the protein due to over-expression or the presence of the tag. If possible, the predicted distribution of your protein should be confirmed with the use of antibodies in situ. Specific antibodies also allow you to examine protein modifications and expression in vivo.
Although antibodies can be very useful tools, generating a specific antibody protein can be difficult, time-consuming, and expensive (see Harlow and Lane, 1988). Unfortunately, except for antibodies to a few highly conserved proteins (e.g., actin and tubulin), most antibodies generated against specific vertebrate or invertebrate proteins do not cross-react specifically with their C. elegans homologs. Furthermore, when you specifically generate an antibody against a C. elegans protein, the antibody may only work with a subset of immune techniques, e.g., Westerns but not in situ or vice versa. A brief Antigen preparation protocol highlights some considerations when designing antigens.
Whenever you use antibodies, it is important to test for specificity. In addition to the standard controls, such as pre-absorbance of your antibody, worm-specific methods may be available to test antibody specificity. If there are null mutants or RNAi worms for the gene and protein of interest, then compare antibody staining (of Westerns or in situ) in those strains with staining in wild-type worms. Nulls can also be used to affinity deplete your serum (see Depletion of primary antibody protocol). On the other hand, if there are transgenic over-expressing lines for the protein of interest, those can be used as positive controls. Generally, you will need to do some sort of affinity depletion before you can detect specific staining of Westerns or worms (see Affinity purification protocol).
C. elegans provides specific rewards and particular challenges for in situ analysis of protein distribution. If you have an antibody that works in situ, you can examine the distribution of the protein in the entire organism at all stages relatively rapidly. Other than the intestine, most tissues have relatively low levels of autofluorescence, so many different dyes and secondary antibodies may be used, allowing triple labeling of antibodies or more.
On the other hand, immunocytochemistry in worms is relatively difficult due to problems of access and size. Both the egg shell of embryos and the cuticles of larvae and adults are relatively impermeable and must be disrupted chemically or mechanically (or both) for the antibody to have access to the internal cells and proteins. The small size of the individual cells and their compact architecture can make visualization of the cellular and subcellular distribution of proteins in some cells quite difficult. A confocal fluorescent microscope or a wide-field fluorescent microscope with de-convolution software can provide increased spatial resolution. When detailed examination of subcellular distribution is necessary, then one can attempt the supreme challenge of immuno-electron microscopy. For a discussion of electron microscopy and immuno-EM in C. elegans (not covered in this section), see the Worm Atlas website.
There are several different techniques used to allow antibodies access to the tissue of interest. If you are specifically interested in the distribution of a protein within the cells of the gonad, the very early embryo, or the intestine, then the appropriate tissue can be dissected out of the adult (see Freeze-crack protocol). To examine protein in a larva or adult, the two general methods are to 1) collect worms in small tubes of liquid and to freeze and chemically treat them to disrupt their cuticles (see Tube fixation, Bouin's tube fixation, Peroxide tube fixation, and Picric acid + glutaraldehyde fixation protocols) or 2) compress and freeze worms between slides and mechanically pull off the cuticle layer (see Freeze-crack protocol). Different fixation conditions differentially preserve antigenicity and tissue morphology. The Fixation conditions protocol discusses how to determine the optimum fixation conditions for a particular antibody and protein combination. You can use a control antibody such as anti-DNA or anti-actin to monitor accessibility and morphology with the different fixation conditions.
Western blot analysis is useful for several reasons. It can be used to examine the relative levels of expression of different size forms of your protein, such as those due to alternative splicing. It can help identify post-translational modifications such as phosphorylation or glycosylation. Western blot analysis can be used to screen for changes in protein expression under different conditions or in different mutant backgrounds. Changes in protein expression may also be seen with immunocytochemistry (and examined in a cell-specific way). But, generally antibodies do not distinguish between protein variants in situ unless specifically designed to do so. And some antibody works well for Westerns but not in situ.
Preparation of denatured protein from C. elegans is relatively straight forward, since simple boiling of adults or embryos in a typical protein buffer frees many proteins from the body (see Worm protein preparation protocol). A major difficulty with Western blot analysis arises from the fact that the entire worm is usually used for protein preparation. Thus, even if your protein is of fair abundance in a particular tissue, e.g., the nervous system, its abundance relative to total worm protein may be quite low. Therefore, sensitive detection methods are generally required (see Western transfer protocol).
If you generated your antibody using a fusion protein or a coupled peptide, then first test your antibody for staining of a Western Blot using a very low concentration of this protein. If you can not detect the protein in this purified condition, you are unlikely to be able to detect it in a complex mixture of worm proteins. The antibody may still work on worms in situ, since the antigen will be in a different conformation in the fixed worm than in the protein gel.
When you test an antibody on a worm preparation, it is helpful if you can use a Western blot from a tissue or strain that is enriched for the protein. For example, if the protein is enriched in a particular stage, then preparation of synchronous populations is recommended. If a particular enriched tissue can be dissected out fairly rapidly (e.g., the gonad), that is also recommended as the starting preparation. If you have a transgenic strain or mutant that over-expresses your protein, then use it to test for specificity of antibody binding in Westerns.
C. elegans cell culture (Christensen et al., 2002) provides novel tools for immunohistochemistry. Cell culture provides an easy method to test antibodies for staining of cells without concerns about permeability and accessibility (see Fixation conditions protocol). It also may provide an enriched source of particular cell types (e.g., sorted GFP-positive cells) for use in Western analysis. Despite this, the intact organism continues as the major focus of immunohistochemical techniques, as described in the following protocols.
The figures illustrate antibody staining of C. elegans using the protocols covered in this section. The fixation conditions range from a very "light" fix (Figure 1, Protocols 6 and 13), to very "hard" fixes, followed by permeabilization by collagenase treatment (Figure 2, Protocols 7 and 14) or other chemical permeabilization (Figure 3, Protocols 8 and 14). Each antibody has its own optimum fixation conditions, see Protocol 5 for a discussion of how to determine those conditions. For staining in wild type versus transgenic strains, see Figure 4.
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Figure 1. Light fixation and staining of multiple antigens in the head of an adult OH99 mgIs18 [Pttx-3GFP] prepared using Protocol 6. Freeze-crack. (A) Staining of the vesicular monoamine transporter (VMAT = CAT-1), shown in red. This antigen is found in synaptic vesicles of monoaminergic neurons.(B) DNA, white, was labeled with DAPI. (C) The vesicular acetylcholine transporter (VAChT = UNC-17), found in cholinergic synaptic vesicles, is shown in blue. (D) Staining with an anti-GFP antibody is shown in green. The GFP fills the cytoplasm of the AIY neurons. (E) Merged image of anti-VMAT (red), anti-VAChT (blue), and anti-GFP (green). (F) Merged image of anti-VMAT (red), anti-GFP (green) and DAPI (blue). Fixation was for 2 minutes in methanol followed by 4 minutes in acetone. Staining followed Protocol 13: Staining slides. Primary antibodies were goat anti-VMAT (Goat 258 polyclonal antibody purified with Protocol 4: Affinity purification), rabbit anti-GFP (Molecular Probes) and mouse anti-VAChT (Monoclonal 1403). Secondary antibodies were Cy3-conjugated donkey anti-goat, Oregon Green 488-conjugated donkey anti-rabbit, and Cy5-conjugated donkey anti-mouse. See Protocol 13: Staining slides for a discussion of anti-GFP and secondary antibodies. mgIs18 [Pttx-3::GFP] was a gift of O. Hobert; unc-104(e1265) was provided by the CGC. Images are maximum projections of confocal series, contrast enhanced in Adobe Photoshop®. Anterior to left, ventral down, scale bar is 10 μm.
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Figure 2. Hard fixation in tubes to visualize neurotransmitters. Serotonin (red) and GFP (green) were labeled in the head of an adult RM2304 unc-104(e1265); mgIs18 [Pttx-3::GFP]. The bright red cells are the NSMs; the bright green cells are the AIYs. Worms were prepared using Protocol 7: Tube fixation with fixation for 24 hours in 4% formaldehyde, followed by incubation in collagenase for 6.5 hours. Staining was done using Protocol 14: Staining tube-fixed worms. Primary antibodies were rabbit anti-serotonin (H. Steinbusch, Free University) and mouse anti-GFP (Molecular Probes monoclonal 3E6); secondary antibodies were Cy3-conjugated donkey anti-rabbit and Oregon Green 488-conjugated donkey anti-mouse. mgIs18 [Pttx-3::GFP] was a gift of O. Hobert; unc-104(e1265) was provided by the CGC. Images are maximum projections of confocal series, contrast enhanced in Adobe Photoshop. Anterior to left, ventral down, scale bar is 10 μm.
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Figure 3. Bouin's tube fixation for muscle preservation. (A) Actin was labeled in a wild-type L2 larva fixed using Protocol 8: Bouin's tube fixation with 1 hour in Bouin's fixative with methanol and β-mercaptoethanol, followed by incubation in Borate Tris buffers. Staining was using Protocol 14: Staining tube-fixed worms with mouse anti-actin (Chemicon monoclonal C4), followed by Oregon Green 488-conjugated donkey anti-mouse. (B) Talin was labeled in a wild-type embryo with rabbit B547 anti-talin (gift of Bob Barstead and Gary Moulder) and Cy3-conjugated donkey anti-rabbit. (C) Double staining of talin, stained with rabbit B547, and vinculin, stained with monoclonal antibody MH24 (gift of Russ Francis) in a wild-type adult. Secondary antibodies were Oregon Green 488 donkey anti-rabbit and Cy3 donkey anti-mouse. Maximum projection of overlapping confocal series, contrast enhanced in Adobe Photoshop. All scale bars are 10 μm.
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Figure 4. Staining in wild-type versus transgenic strains. The transgenic strain RM1810 cha-1(md39); mdEx10 [F57G7] contains multiple copies of the cosmid that contains the cha-1 and unc-17 genes. This strain produces abnormally high levels of the ChAT and UNC-17/VAChT proteins. It is useful for testing the specificity of anti-ChAT and anti-VAChT antibody staining in situ, for increased ease of cell identification, and also for Westerns (see Protocol 17: Western transfer). (A) Staining of the VAChT, shown in red, in RM1810. (B) The same worm stained with DAPI to show nuclei. (C) A higher magnification view of the head of the same individual. (D) VAChT staining in a wild-type N2 shows the typical synaptic localization of the protein. Note that over-expression of VAChT leads to mislocalization of the protein in neuronal somas (compare panels C and D). The RM1810 and wild-type worms were prepared using Protocol 6: Freeze-crack. Fixation was for 2 minutes in methanol followed by 4 minutes in acetone. Staining followed Protocol 13: Staining slides, with mouse anti-VAChT (Monoclonal 1403) as the primary antibody, followed by Cy3-conjugated donkey anti-mouse. DNA was labeled by addition of DAPI in the mounting medium. Maximum projection of overlapping confocal series, contrast enhanced in Adobe Photoshop®. Anterior to left, ventral down, scale bar is 10 μm.
For a thorough discussion of antigens and antibody generation, see Harlow and Lane (1988).
There are a number of companies that will assist you in your antibody production, from the generation of fusion proteins or peptide antigens through serum purification. See the following for suggestions for peptide design and coupling (see Peptide coupling protocol) and affinity purification (see Antibody purification protocol).
The most frequently immunized animal is the rabbit. If you plan to do any double labeling, try generating your antibodies in one or two different species. Chickens are recommended: they are relatively inexpensive, the collection of antibody in eggs is humane, and they often generate robust immune responses. It is generally more economical to have the company deliver the eggs to you rather than the purified serum. Chicken antibodies are easy and inexpensive to partially purify from the egg yolks (see Chicken antibody purification protocol).
Generation of fusion proteins for immunization is much more laborious than buying peptides, but immunization with fusion proteins is more likely to lead to useful antibodies. If you have a company generate the fusion protein for you, be certain to request sufficient protein for testing, affinity purification, and affinity depletion of your serum.
Green Fluorescent Protein can be a wonderful protein to use in translational fusions (see Reporter gene fusions section of WormMethods). If expression of a translational GFP fusion leads to rescue of the mutant phenotype, then the location of GFP fluorescence in the rescued animals is likely to correspond at least partially to the normal localization of your protein. Of course, the localization should be checked with immunocytochemical localization of the endogenous protein in a wild-type animal. If your protein is not functional or stable in a GFP fusion protein, there are a number of very small epitope tags such as 6-His and Myc that can be fused to your protein and used for preliminary localization studies (Roehl et al., 1996).
Immunization with synthetic peptides is the quickest and easiest way to try and generate antibodies, although it is not always productive. Companies will predict immunogenic peptides or you can design the peptides yourself (http://immunax.dfci.harvard.edu/Tools/antigenic.html). If they are potentially immunogenic, then N-terminal or C-terminal peptides are recommended.
Be sure to order enough peptide for testing, affinity purification, and affinity depletion of your serum.
Use at least two different peptides from the same protein: peptides are inexpensive and many peptides are not immunogenic or are not exposed on the fixed protein. If cost is a major concern, two peptides from the same protein can be injected together during immunization; the serum can be specifically affinity purified as needed.
Peptides are generally conjugated to carrier proteins such as KLH (keyhole limpet hemocyanin), most easily using a kit and a peptide with an extra N-terminal cysteine. Peptides may also be coupled to serum albumin (see Peptide coupling protocol).
Peptides can be conjugated to two different carrier proteins (e.g., KLH and serum albumin). One conjugate can be used for immunization and the other conjugate can be used for affinity purification (see Antibody purification protocol).
This is a very simple and rapid procedure that uses glutaraldehyde to cross-link a peptide to serum albumin as the carrier protein (Harlow and Lane, 1988). The serum albumin should be from the species that will be immunized, e.g., if you plan to immunize a chicken you should use chicken serum albumin as the carrier protein. The basic premise is that the immunized animal will not generate many antibodies against its own serum albumin.
Dissolve 5 mg peptide in 1 ml sterile 100 mM NaPO4 buffer, pH 7 in a 1.5 ml tube.
Dissolve 20 mg serum albumin in 1 ml sterile 100 mM NaPO4 buffer in a separate 1.5 ml tube.
Mix peptide and albumin solutions in a tiny clear beaker and stir well with stir bar.
In hood: Stir constantly while adding drops of 20 mM glutaraldehyde (TOXIC). Add the drops at a rate of 100 ul per 5 minutes until up to 1 ml of glutaraldehyde has been added over a period of almost one hour.
Ideally, the solution should be slightly yellow when the protein is slightly cross-linked. If the solution turns yellow before the full 1 ml of glutaraldehyde has been added, stop adding the glutaraldehyde.
Dialyze at 4°C with PBS. Do at least 3 changes each with one liter of PBS for several hours.
Divide into 10 or 20 1.5 ml tubes and speedvac (no heat) until dry. Assume each tube has approximately 1 or 2 mg of protein-coupled peptide.
Store at -20°C in well-sealed aliquots.
This antibody enrichment procedure is based on the protocol of Gassmann et al. (1990). It uses serial PEG precipitations to enrich for the antibodies. The end product, although far from pure, is stable and is suitable for immunocytochemistry or Western blotting. Further purification is needed before immunoprecipitation.
Carefully break egg into palm of gloved hand or into an egg separator.
Using a squirt bottle of cold dd water, rinse the yolk.
Remove and discard as much of the white as possible.
Drop cleaned yolk into a beaker with a stir bar.
Break yolk with a pipette, if necessary.
Add cold buffer A, ~ 10 ml per yolk.
Stir on stir plate for >2 minutes.
Stir in 30 ml per yolk of cold 7% PEG.
When well mixed (>5 min), pour into centrifuge bottle (leave behind the scum on beaker).
Centrifuge at 14,000 g for 10 minutes.
Pour the top two layers (a thin layer of oil and a liquid layer) through a funnel with a Whatman #1 filter into a new centrifuge bottle. Discard the yellow pellet.
Add PEG 8,000 powder to a final concentration of 12g/100ml fluid.
Shake in centrifuge bottle on shaker at 4°C for 1-2 hours, until the PEG is in solution.
Centrifuge at 14,000 g for 10 minutes.
Pour off and discard fluid, leaving behind orange pellet.
Spin pellet at 5,000 rpm for 2 minutes.
Remove with any fluid with a pipette and discard.
Add 5 ml per yolk of buffer A.
Dissolve the pellet in the buffer by stirring it with a glass rod (laborious) or by letting it sit or shake overnight at 4°C.
If desired (not necessary for antibody for staining), remove residual PEG with dialysis or ammonium sulfate precipitation.
For maximum yield, have all solutions and do all work at 4°C.
Obtain and process pre-immune eggs. Pre-immune chickens may have anti-nematode antibodies.
Start by processing a single egg per beaker. Once a plateau of antibody production has been reached (after 3 immunizations), you may process together all the eggs produced in one week by one hen.
Intact eggs will last at least 3 weeks at 4°C.
| Buffer A: for 1 L of 10X |
| 13.6 g KH2PO4 (1X final 10mM) |
| 58.4 g NaCl (1X final 100mM) |
| 10 ml 10% Sodium Azide (1X final 0.01%) TOXIC |
Add to 900 ml dd water
Adjust pH to 7.2 with concentrated NaOH.
Top to 1 L with dd water.
| 7% PEG: for 100 ml |
| 10 ml 10X buffer A |
| 7 g PEG 8,000 |
Add dd water to 100 ml.
Time and stirring are required for the PEG to go into solution.
Pre-absorbing antibody with a bacterial acetone powder may improve the specificity of the binding of your antibody, especially if your antibody was generated against a bacterially expressed fusion protein (Harlow and Lane, 1988; Miller and Shakes, 1995). However, affinity purification has the potential for a much more significant improvement in specificity.
This protocol uses nitrocellulose membranes rather than columns for antibody purification (Smith and Fisher, 1984; Duerr et al., 1999). This technique minimizes the time that the antibodies spend in the elution buffer in order to maximize antibody recovery. It is also quick, simple, and inexpensive.
If you have used a KLH-coupled peptide as your antigen, you can affinity purify your antibody with a serum albumin-coupled peptide to increase specificity (or vice versa).
If you do not have a pure antigen protein preparation, you may still use this method to purify your antibody. Run your protein mixture (e.g., partially purified fusion protein) out on a gel and then Western blot it to a high-quality, high-binding nitrocellulose membrane. Cut out the area of the membrane containing the blotted protein of interest and use the membrane in the following procedure.
This protocol can also be used to affinity deplete your serum. For example, if you have used a partially purified fusion protein as your antigen, you can use a partially purified preparation for a different fusion protein to bind (and discard) antibodies that recognize the non-specific shared portion of the fusion protein or the bacterial contaminants.
Procedure 1. Coating of membrane with antigen
Add 10 ml NFDM to a 100 mm petri dish (to block any protein binding sites on the dish).
Shake at low speed (40–60 rpm) for 1 hour at room temperature (or longer at 4°C).
Rinse dish 3X with TBST (about 20 ml per wash) and drain well.
Add 15 ml Antibody Transfer Solution.
[Optional: save sample to check protein concentration in Antibody Transfer Solution.]
Place 1 nitrocellulose membrane into dish with forceps (not fingers).
Shake 30-60 minutes at room temperature (or longer at 4°C).
[Save unbound Antibody Transfer Solution and check for protein concentration.]
Rinse membrane in dish 3X with TBST.
Cover membrane with NFDM.
Put on shaker for 1 hour at room temperature (or longer at 4°C).
Procedure 2. Pre-elution of membrane
Rinse membrane in the coated dish 3X with TBST.
Rinse briefly with dd water.
Add 10 ml glycine buffer.
Swirl for exactly 60 seconds.
Pour off and discard glycine buffer.
Quickly rinse membrane 3X with TBST.
In hood, rinse briefly with dd water.
Add 10 ml TEA buffer (TOXIC).
Swirl for exactly 60 seconds.
Pour off and discard TEA buffer.
Quickly rinse 3X with TBST.
Cover membrane in dish with NFDM.
Shake 30 minutes at room temperature.
Procedure 3. Incubation with antibodies
Rinse membrane in dish 2X with TBST.
Add 10 ml TBST to dish.
Add 0.5 ml (or less) serum to dish.
Shake for 1 hour at room temperature (or longer at 4°C).
[Save antibody solution after binding (see comments below).]
Rinse membranes 3-6X for 5 minutes each with TBST.
Procedure 4. Elution of antibodies
Prepare a 15 ml tube with 150 ul 1M Tris pH 8.2 for glycine rinse.
Prepare a 15 ml tube with 800 ul 0.5M Tris pH 6.8 + 200 ul 1M HCl for TEA rinse.
Rinse membrane briefly with dd water.
Add 5 ml glycine buffer.
Swirl for exactly 20 seconds and pour off and save glycine buffer in tube with Tris pH 8.2.
Immediately add another 5 ml glycine buffer to dish with membrane.
Swirl membrane in second rinse of glycine buffer for 20 seconds.
Pour second glycine rinse in with the first rinse.
Quickly rinse membrane 3X with TBST.
In hood, rinse membrane briefly in dd water.
Add 5 ml TEA buffer (TOXIC).
Swirl for exactly 20 seconds and pour off and save TEA buffer in tube with Tris pH 6.8.
Immediately add another 5 ml TEA buffer to dish with membrane.
Swirl membrane in second rinse of TEA buffer for 20 seconds.
Pour second TEA rinse in with the first TEA rinse.
Quickly rinse membrane 3X with TBST.
Check pH of eluates and adjust to pH ~7.5 with appropriate pH Tris BEFORE STORING.
Since the volumes are so small, use pH strips to check the pH.
Cover used membrane with NFDM and store in dish at -20°C for later re-use.
Procedure 5. Optional concentration
Put glycine eluate in the top of a Centricon or other concentrator (mw cutoff <10-40,000D).
Add plain Phosphate Buffered Saline (PBS) to fill line.
Balance tubes and spin 20-30 minutes at 5,500 rpm at 4°C (or as directed) until concentrated.
Discard flow through.
Add PBS to 15 ml and repeat spin.
Discard flow through.
Add Antibody buffer to 15 ml and repeat spin.
Discard flow through.
Repeat spin if needed until volume <1.5 ml.
Store concentrate in a 1.5 ml capped tube at 4°C or -20°C.
Repeat for TEA eluate.
Nitrocellulose membranes must be high quality with high protein binding. We use Nitroplus transfer membranes 82 mm discs (MSI).
Never touch the membranes with your fingers; always transfer them with forceps.
Antigen-coated membranes may be used more than once. Store them in NFDM at -20°C. Membranes will show decreased purification effectiveness after 2-10 uses, depending upon the antigen and serum.
Be sure to have the Tris solutions ready to neutralize the buffers immediately after the elution steps. Elution and neutralization should take about 1 minute per buffer. Adjust pH to 7.5 before storing.
You may do several membranes simultaneously. However, be sure to do the elution step sequentially so that the antibody does not sit in the high or low pH buffer too long.
Glycine elutions have been more productive, but TEA elutions usually provide some specific antibodies in lower concentrations. Note that the glycine elution is a high salt and low pH elution.
The eluates may be stored at 4°C or -20°C; concentrate the antibody before storage.
If you are testing a new antibody, the eluates can be used after pH adjustment but before concentration for Westerns or immunocytochemistry. Try eluate diluted 1:1 - 1:50 in Antibody buffer with 1% BSA (for immunocytochemistry) or Antibody buffer with 1% NFDM (for Westerns).
The optimum amount of serum to use with each round of purification will vary. Start with 0.5 ml serum per membrane, but save the unbound serum and test it with another round of purification.
| Antibody transfer solution (per membrane) |
| 12 ml dd water |
| 3 ml Methanol (final 20%) |
| 375 ul 1.0 M Tris pH 8.0 (final 25 mM) |
Add ~1 mg antigen in its buffer to 15 ml transfer solution.
Take a sample to check the antigen concentration before and after binding to the membrane.
| Tris buffered saline with Tween-20 (TBST) for 1-2 membranes |
| 6.1 g Tris free base (final 50 mM) |
| 8.8 g NaCl (final 150 mM) |
| 500 ul Tween-20 (final 0.05%) |
| 5 ml 10% sodium azide in dd water (final 0.05%) |
| 900 ml dd water |
Add concentrated HCl to adjust pH to 8.0 (use pH paper).
Add dd water to final volume of 1 L.
Store at 4°C.
Make up and store 10X stock.
You can use PBST (PBS with 0.05% Tween-20, pH 8.0) instead of TBST.
| Tris buffered saline with nonfat dry milk (NFDM) for 1-2 membranes |
| 100 ml TBST (or PBST) |
| 3 g Nonfat dry milk (final 3%) |
Mix well with stirring and gentle heat, if needed.
Store at 4°C for up to 5 days.
| Glycine elution buffer for 10 membranes (20 ml per membrane) |
| 121 mg glycine (final 5 mM) |
| 25 mg BSA (final 0.01%) |
| 125 ul Tween-20 (final 0.05%) |
| 7.3 g NaCl (final 500 mM) |
| 200 ml dd water |
Use dilute HCl to adjust pH to 2.3.
Add dd water to 250 ml.
Store at 4°C (keeps many weeks).
| 1.0 M Tris pH 8.2 (~500 ul per membrane) |
| Store at 4°C. |
| TEA buffer (20 ml for 1 membrane): TOXIC |
| 175 ul Triethylamine (final 50 mM) |
| 25 ml dd water |
| 0.5 M Tris pH 6.8 (~1ml per membrane) |
Store at 4°C.
Phosphate buffered saline (PBS)
Antibody buffer
Purify your antibody before testing (see Antibody purification protocol), since non-specific staining of unpurified antibodies usually masks specific staining in C. elegans.
Although a few antibodies work with a wide range of tissue fixation conditions, your antibody is likely to work best (or only) with a particular fixation regime (see Harlow and Lane (1988) and Harlow (1999) for discussion). Major classes of fixatives are precipitating agents (e.g., methanol and acetone) or compounds that cross-link proteins (e.g., formaldehyde and glutaraldehyde).
After fixation, you must permeabilize the tissue to allow access of the antibodies. This is done by the inclusion of a non-ionic detergent in the incubation buffers. After or during particularly hard fixation conditions, treatment with an organic solvent such as methanol or acetone may be necessary for adequate permeabilization (Harlow and Lane, 1988).
In C. elegans, chemical treatments can be used to permeabilize the cuticle, including incubation in collagenase or reduction in β-mercaptoethanol (Finney and Ruvkun, 1990; Garriga et al., 1993; Miller and Shakes, 1995). However, these treatments may also destroy the ability of antibodies to recognize your protein. Therefore, initially test fixation conditions on freeze-cracked or otherwise mechanically disrupted worms or tissues. Or, if applicable, examine dissected tissue (see Freeze-crack protocol; Crittenden et al., 1994) or cultured cells (Christensen et al., 2002). Once the optimum fixation conditions for mechanically disrupted or dissected worms have been determined, then determine whether one of the Tube-fixation protocols that use chemicals to disrupt the cuticle might be more convenient.
You can monitor tissue preservation and accessibility at a gross level by examining the morphology of nuclei stained with the DNA-binding dye DAPI (see Staining slides protocol). DAPI permeates tissue more readily than antibodies; if you do not see DAPI labeling of your cells, then you are unlikely to see antibody staining.
For more specific (but more expensive) examination of tissue accessibility with any fixation method, use a control primary antibody. Good controls are an antibody to DNA (Chemicon mouse monoclonal antibody MAB030) or to a cytoskeletal protein (Amersham L9 anti-tubulin; Calbiochem Ab-1 anti-actin; ICN Biochemicals C4 anti-actin). All of these antibodies work on worms prepared with a variety of fixation conditions.
All fixatives are TOXIC.
Try the following with freeze-cracked slides or tissues or cells (See Freeze-crack protocol):
100% methanol: 2 minutes on ice.
100% acetone: 2 minutes on ice.
100% methanol 4 minutes on ice followed by 100% acetone 4 minutes on ice.
1% formaldehyde in 0.1 M Phosphate buffer + 50% acetone: 30 minutes on ice.
0.5% formaldehyde in 0.1M Phosphate buffer: 1 hour on ice.
4% formaldehyde in 0.1M Phosphate buffer: 1 hour on ice.
4% formaldehyde in 0.1M Phosphate buffer: overnight at 4°C.
4% formaldehyde + 0.1% glutaraldehyde in 0.1M Phosphate buffer: overnight at 4°C.
4% formaldehyde + 1% glutaraldehyde in 0.1M Phosphate buffer: 24 hours at 4°C.
Try the following with tube-fixed worms (See Tube fixation protocol):
1% formaldehyde: 1 hour on ice.
4% formaldehyde: overnight at 4°C.
See the Freeze-crack protocol for recipes for different fixatives. Formaldehyde and glutaraldehyde solutions should be made fresh.
Start with #3 and #6, and then try other conditions as needed. As you discover the optimum conditions, you may change the concentration of fixative, incubation duration, temperature, and the buffer (for example, you can try a buffer including picric acid and/or borate or methanol). Generally, fixation is done in cold solutions to prevent diffusion before fixation.
As you go down the list, the fixation conditions get harder, i.e., more precipitated and/or cross-linked. As this happens, the fixation conditions get more difficult to evaluate because the antibodies will not penetrate the tissue as well, autofluorescence will increase, and fewer worms will stick to the slides. If you need long formaldehyde and/or glutaraldehyde fixation conditions, you can switch from the Freeze-crack to the Tube fixation protocol. The tube fixation procedures are longer and have more variables than the freeze-crack procedure, but the morphology of hard fixed worms is much better with tube fixation.
If you are going to be testing multiple antibodies (or multiple batches of one antibody) you can keep stocks of unstained tube-fixed worms in the refrigerator or -20°C freezer (see Tube fixation protocol and Bouin's tube fixation protocols). Tube-fixed worms last months in the refrigerator or indefinitely at -20°C; although some antigenicity may be lost, fixed worms that are 5+ years old can show specific antibody binding.
If you do cell culture, you can fix and save your old preparations with a variety of fixatives. Store the fixed slides in PBS at 4°C or protected from dehydration at -20°C and use as needed to test antibodies. If you find a protocol that appears to give you specific staining, then use similar fixation conditions on freeze-cracked worms or dissected tissues.
Freeze-cracking provides a quick and simple way to remove portions of the worm's cuticle, allowing penetration of antibodies (Strome and Wood, 1982; Albertson, 1984; Miller and Shakes, 1995; Duerr et al., 1999). Dissection of worms (Crittenden et al., 1994) can increase antibody access to some tissues and early embryos.
The procedure generally begins with a large number of worms and ends with a few stained worms. The morphology of the worms is variable; preparation of relatively intact worms takes practice. However, the ease and speed of the technique and the minimal chemical disruption are very useful for evaluating antibodies and are optimum for localizing some antigens.
Procedure 6. Freezing plates of worms on slides
Use a plate of healthy worms.
Place large flat slabs of dry ice or place a flat metal plate firmly on top of crushed dry ice in a Styrofoam container.
Wash the worms from the plate with M9 buffer or dd water and collect them in a 1.5 ml tube.
Spin the worms at 2000 rpm (~400 g) for 2-3 minutes.
[Optional: To enrich for adults, let the worms settle for 5 minutes at room temperature.]
Remove the supernatant and repeat M9 wash until ~ all bacteria are gone and solution is clear.
Rinse worms with dd water, spin or let settle, and remove all but about 50 ul water.
Label lab-made poly-lysine slides with indelible ink and place on counter frosted side up.
Using a glass pipette, place a small drop (10-20 ul) of worms on the slide.
Let the worms settle briefly.
Use the pipette to spread out the worms and liquid so that individual worms are not overlapping.
Carefully press an unlabeled (commercial) poly-lysine slide down on the worms label side down so that only the unfrosted parts overlap.
Press straight down slightly, enough to immobilize but not rupture most of the adults.
Carefully (without sliding) put the slides on a flat piece of dry ice for 5-30 minutes.
Proceed to Methanol and/or acetone fixation or Formaldehyde and/or glutaraldehyde fixation.
Procedure 7. Freezing individuals or small numbers of worms
Transfer the worm(s) to a food-free plate and let them rid themselves of bacteria.
Place a small drop of dd water on the middle of a sticky lab-made poly-lysine slide.
Pick the worms up individually with a clean pick and shake them off in the water drop.
The worms should partially stick as they fall and touch the poly-lysine.
Multiple (up to ~50) worms of similar sizes can be done on the same slide.
The more worms you do on a slide, the larger the fraction that you will lose.
When you have transferred the worms, place a commercial poly-lysine slide over the worms.
For a few worms, the weight of the slide should provide sufficient compression.
With many worms, press down evenly and lightly to compress the worms.
The water droplet should not reach the edges of the slide after compression.
Immediately and smoothly transfer the slide to a flat piece of dry ice or metal on dry ice.
Proceed to Methanol and/or acetone fixation or Formaldehyde and/or glutaraldehyde fixation.
Procedure 8. Dissecting worms: for intestine, gonad, early embryos
Place a small drop of 0.2 mM levamisole in dd water onto a lab-made poly-lysine slide.
The levamisole paralyzes and contracts the worms and aids dissection.
Transfer several adults to the drop.
Slice the adults near the head, vulva, or tail with two fresh 25-26 gauge needles.
Options: Use dissection scissors, or a blade breaker/holder and fine razor blade.
With practice, the intestine and gonad will extrude and stick to the poly-lysine slide.
Early embryos can be pulled free from the rest of the worm.
Option: Bleach gravid adults, collect embryos and place in a drop of dd water on the slide.
When you have dissected several adults, place a commercial poly-lysine slide over the worms.
If necessary, press down lightly and evenly on the top slide until it touches the tissues.
Immediately and smoothly transfer the slide to a flat piece of dry ice or a metal plate on dry ice.
Proceed to Methanol and/or acetone fixation or Formaldehyde and/or glutaraldehyde fixation.
Procedure 9. Freezing older embryos (from a mixed population)
Pipette M9 over the surface of a plate containing worms in food.
Allow the adults and larvae to thrash free of the food and into the fluid.
Tilt plate and pipette off the M9 with thrashing worms and discard.
If adults and larvae are still present on the plate, repeat the M9 wash.
When the plate contains only embryos in food, use a stream of M9 to rinse off the embryos.
Collect embryos into a 1.5 ml tube.
Spin the worms at 2000 rpm (<800 g) for 2-3 minutes.
Replace supernatant with dd water.
Repeat spin and rinse as needed until embryos are clear of bacteria.
Transfer embryos in a small drop of water to a lab-made poly-lysine slide.
Press down lightly and evenly on the top slide until it contacts the embryos.
Immediately and smoothly transfer the slide to a flat piece of dry ice or metal plate on dry ice.
Proceed to Methanol and/or acetone fixation or Formaldehyde and/or glutaraldehyde fixation.
Procedure 10. Methanol and/or acetone fixation
See Fixation conditions protocol for variations with organic solvent fixatives.
Place a covered coplin jar of 100% acetone on ice (or in dry ice).
Place a covered coplin jar of 100% methanol on ice (or in dry ice).
Let chill until very cold (>15 minutes).
Crack the slide sandwich by swiftly twisting apart the two slides; discard the top (unlabeled) slide.
[Optional (for multiple slides): Immediately place the wormless sides of the slides (smooth sides) back-to-back]
Immediately immerse in ice cold methanol for 2+ minutes.
Immerse in ice cold acetone for 4+ minutes.
[Optional: Filter the acetone and methanol through a Whatman #1 filter and reuse 3-4 times.]
[Use cold methanol or acetone alone for <10 minutes (see Fixation conditions protocol).]
[Optional: Air-dry the slides for 1-5 minutes before proceeding. This may increase adhesion of the worms to the slide, but may decrease antigenicity.]
Place the slides in a new coplin jar.
Pour PBS down the side of the jar (not directly onto the worms).
Optional: Rehydrate the worms with a series of alcohol solutions (95%, 70%, 50%, and 30%).
Proceed with blocking and staining (see Staining slides protocol).
Procedure 11. Formaldehyde and/or glutaraldehyde fixation: TOXIC
See Fixation conditions protocol and solutions (below) for fixatives.
Prepare FRESH TOXIC formaldehyde and/or glutaraldehyde solution in hood.
Pre-chill fixative on ice.
Crack the slide sandwich by swiftly twisting apart the two slides; discard the top (unlabeled) slide.
Quickly place the slides (worms up) flat in a dish (with lid) in the cold room or refrigerator.
Cover the worms on each slide with 100-200 ul fixative.
Fix the worms for 30 minutes to 24 hours at 4°C.
[Optional: Incubate the slides back to back in a coplin jar containing cold fixative.]
[Incubation in the jar increases loss of worms from the slide, but is less messy.]
After fixation, rinse the slides gently several times with PBS (until there is no fixative odor).
Proceed with blocking and staining (see Staining slides protocol).
Use plates containing healthy well-fed worms since starved worms have increased autofluorescence in the gut. Also, dauers seldom crack open.
If you use a mixed population, then most of the worms that stick to the slides will be the fatter adults, since they adhere best to the top and bottom slides. You can use synchronized populations of worms if you wish to recover larvae of a particular age.
Only a fraction of embryos will be cracked open when you use a mixed population. To specifically stain embryos, use a population of embryos that have been transferred to a poly-lysine slide (Procedure 19 or 20).
Clean (bacteria-free) worms will stick the best to the poly-lysine slides. However, bacteria-free worms also stick well to clean plastic or glass pipettes. Use the same glass pipette for the initial and final rinses when cleaning the worms free of bacteria, so that the very thin coating of bacteria left on the inside of the glass prevents worms from sticking to the pipette.
Slides can be stored before ‘cracking’ at -80°C for days to weeks before staining with some (but not all) antibodies. Let slides from -80°C ‘warm up’ on dry ice for 10 minutes before cracking.
If you do not have slabs of dry ice available, then you can use a pre-chilled conductive (metal) plate on top of crushed dry ice. You may also use a metal plate on the shelf of a -30°C or -80°C freezer. However, this later technique may not work as well as a slab of dry ice (and is not good for your freezer).
Some people use a coverslip as the top of the slide sandwich, instead of a commercial poly-L-lysine slide. This allows more delicate compression of the worms, but removal of the cover slip can be more difficult than separating the two slides. Removal of the cover slip should only be done when wearing goggles to protect your eyes from flying shards of glass.
Problems with recovering intact worms on the poly-lysine slides can arise from:
| Insufficiently cleaned worms. |
| Insufficiently cleaned or poorly coated or poorly dried poly-lysine slides. |
| Too much or too little or sideways pressure during compression. |
| You can monitor worm compression by examining your slides with a dissection scope. |
| Harder fixation conditions lead to more loss of worms. |
| Air drying the slides may improve adhesion, but decreases staining with some antibodies. |
| Pouring solutions directly over the worms may wash them off of the slides. |
You can make extra sticky poly-lysine slides (for hard fixes or precious worms) as follows:
| Place a small drop of Lab-made poly-L-lysine Solution directly onto a clean slide. Either let the drop evaporate as is OR use a second slide to carefully spread the drop before drying. In either case, you should get a very sticky spot on your slide. Unfortunately, the poly-lysine spot will often bind non-specifically to your antibody and give high background staining. |
| M9 (for 1 L) |
| 3 g KH2PO4 |
| 6 g Na2HPO4 |
| 5 g NaCl |
| 1 ml 1M MgSO4 |
Add dd water to 1 L and stir to dissolve fully.
Store at 4°C.
| "Lab-made" poly-L-lysine solution |
| 400 mg Sigma P1524 poly-L-lysine (very high molecular weight) |
| 200 ml dd water |
| 0.2 g Sodium azide (0.1%) TOXIC |
Mix together and store at 4°C.
Reuse solution to coat slides until it becomes noticeably less sticky.
"Lab-made" poly-L-lysine Slides:
Wipe ‘pre-cleaned’ frosted edge slides very clean with kimwipes.
Place slides back-to-back (frosted sides exposed) in slide holder.
Shake 30 minutes in dilute ionic detergent (e.g., squirt of dishwashing soap in water).
Rinse in running water > 1 minute.
Rinse in running distilled water > 1 minute.
Shake in 70% ethanol + 1% HCl for > 5 minutes.
Rinse in running distilled water > 5 minutes.
Dry in 60°C oven > 5 minutes.
Shake in "Lab-made" poly-L-lysine Solution for 5 minutes.
Dry in 60°C oven > 1 hr to overnight.
Wear safety goggles and work over a pad to avoid glass shards.
Crack the slides apart by sliding a scalpel between the slides.
Coated slides will last months if kept in a clean slide box at 4°C.
Commercially prepared poly-L-lysine slides:
Brands include Sigma poly-prep and ESCO polysine.
They are not as sticky as the "lab-made" slides so are used for the top slide of the sandwich.
| 10X PBS (for 1 L) |
| 80 g NaCl |
| 2.0 g KCl |
| 27 g Na2HPO4:7H2O |
| 2.4 g KH2PO4 |
| 2 g Sodium Azide TOXIC |
Allow salts to dissolve (with gentle heat and stirring) in 900 ml dd water.
Adjust pH to 7.2 with 10M NaOH.
Top to 1 L with dd water.
Autoclave and store at room temperature.
Formaldehyde solutions (for 40 ml of 4% formaldehyde) from liquid: TOXIC
Open a fresh 10 ml ampoule of 16% formaldehyde.
Use reagent grade 16% formaldehyde (e.g., Ted Pella brand for electron microscopy).
Pour into covered coplin jar on ice in hood.
Add 30 ml 0.1 M phosphate (Na2PO4-H2O, pH 7.2).
Use as soon as chilled or later the same day.
Discard used fix appropriately.
OPTIONS:Vary formaldehyde from 0.5% to 4%.
Mix formaldehyde with 50% methanol.
Mix 4% formaldehyde with 0.5-1.0% glutaraldehyde.
| Glutaraldehyde solutions: TOXIC |
| Dilute 25% glutaraldehyde in 0.1 M phosphate (Na2PO4-H2O, pH 7.2). |
| Pour into covered coplin jar in hood and pre-chill on ice. |
| Use as soon as chilled or later the same day. |
This protocol can produce worms with fairly permeable cuticles and nice morphology (Li and Chalfie, 1990, Garriga et al., 1993). The related Peroxide tube fixation protocol uses additional reducing and oxidizing agents and omits the collagenase treatment (Finney and Ruvkun, 1990). In either case, the fixed and treated worms can be stored for months at -20°C.
Treatment with collagenase can be quite variable and the fixation and permeabilization conditions disrupt many antigens. However, the overall structure of the worm is often well preserved.
Procedure 12. Fixation
Use 1-10 plates of healthy, well-fed worms.
Wash the worms from the plates with M9 buffer and collect in a 1.5 ml or 15 ml tube.
Spin the worms at 2000 rpm (~400 g) for 2-3 minutes.
[To enrich for adults, let the worms settle for 5 minutes at room temperature.]
Remove the supernatant and repeat M9 wash until ~ all bacteria are gone and solution is clear.
Transfer worms to a new screw cap 1.5 ml tube, using dd water.
Top tube with dd water.
Spin 2 minutes at 2,000 rpm (~400 g).
Remove all but 50-100 ul of solution.
Put on ice for 2-3 minutes.
Add 1 ml TOXIC Fix solution.
Put in beaker containing dry ice + 95% ethanol for 2-3 minutes to freeze worms solid and crack their cuticles.
[Optional: Use liquid nitrogen.]
Defrost with warm water.
Repeat the freeze and defrost steps 1-2 times (to crack as many worms as possible).
Put on shaker at 4°C to 37°C for 1-24 hr.
Spin at high speed, remove fixative (discard safely), and top with PBST.
Repeat PBST rinse 3-5 times, until the fixative is no longer detectable.
[Optional: Worms can be stored in PBST for up t
