Confocal Microscopy of Drosophila Embryos
The genetically tractable organism Drosophila melanogaster is proving to be an excellent model system for cell biological analysis in the context of the whole organism. The relative ease with which embryos can be obtained in large numbers and processed for highresolution light microscopy has facilitated many recent advances at the interface between cell and developmental biology. Fine subcellular structures previously impossible to visualise by conventional fluorescence microscopy, on account of high noise resulting from out-of-focus signals, are revealed with clarity on a confocal microscope.
There are several reasons why scientists who have not used Drosophila before may wish to use Drosophila embryos for the analysis of protein localisation and expression. The embryo contains representatives of each cell type and is small enough (500 x 100 µm) to fit within the field of view of a 20X objective lens. The embryos are nearly transparent, permitting visualisation of all cells in whole mount preparations. These features allow one to assay the tissue distribution of a particular protein in a single specimen. The tissues have a relatively simple structure, with the epithelia being made up of a single layer of cells. In general, there are fewer copies of each protein encoded by the genome compared with vertebrates, e.g., one α-actinin rather than four, further simplifying the analysis of the distribution of a particular kind of protein. Injection of double-stranded RNA can be an effective way to knock down protein expression, provided the bulk of the protein in the embryo comes from new synthesis. Sophisticated manipulation of the proteins is possible using the powerful molecular genetic techniques available in this organism.
While Drosophila has many advantages for cell biological analysis, it also has some drawbacks. The cells are small: an embryonic epidermal cell, for example, has dimensions of only 2 x 5 µm compared to a vertebrate epidermal cell of 10 x 20µm, which can make it difficult to resolve different intracellular compartments. The embryo is the only stage where the whole animal can be stained in its entirety; at late stages of development, antibody penetration is blocked by the secreted exoskeleton [although the use of proteins tagged with green fluorescent protein (GFP) circumvents this problem]. Therefore, the most easily generated samples for analysis are the embryo or tissues, that are easily dissected from the larva or adult, such as the imaginal discs and ovaries. Only a small number of Drosophila cell lines are available for in vitro culture and experimentation. These represent just a few cell types and are also small. Finally, antibodies raised against vertebrate proteins rarely bind to Drosophila orthologues, even when they are highly conserved. Therefore, new antibodies need to be raised to see the distribution of a given protein in Drosophila. An important exception to this are antibodies raised to specific motifs of proteins, such as those recognising particular phosphorylated residues. A growing number of antibodies against Drosophila proteins are available, either from the Developmental Studies Hybridoma Bank or directly from the laboratory that has generated them; few are available commercially.
This article describes the techniques used to look at cell junctions and cytoskeletal structures in the developing embryo, using both antibodies against Drosophila proteins and GFP-tagged proteins expressed from transgenes.
The embryo is surrounded by two proteinaceous layers: the egg shell or chorion, which is removed easily; and the vitelline membrane, which is impermeable and more difficult to remove. The vitelline membrane can be permeabilised by treatment with heptane, permitting fixatives to reach the embryo. The vitelline membrane is generally removed by "popping" the embryos with a rapid osmotic shock produced by adding methanol to create a heptane-methanol interface and shaking vigorously. The fixation/devitellinisation steps are the part of the procedure most likely to go wrong, but with practice become routine. Once the embryos are fixed and the vitelline membrane removed, antibody staining is straightforward. Simple mounting of embryos is sufficient in most cases, which, due to the elongated shape of the embryo, allows many optical section views except a cross section of the embryo (i.e., cutting a slice from the middle of loaf of bread). As this can be very informative, we also describe a more labour-intensive method that allows such a view. We also make some recommendations for getting the best confocal images of Drosophila embryos. Additional references describing these methods are found in Davis (1999), Hazelrigg (2000), and Rothwell (2000).
II. INSTRUMENTATION AND MATERIALS
Fly bottles or vials (250-ml bottles and 30/40ml vials, Scientific Lab Supplies)
Egg collection cages (see Fig. 1, I)
Apple juice agar plates (see solutions)
Embryo collection baskets (see Fig. 1, I)
Scintillation vials (4ml, 908-054, Jencons; 20ml, 215/0079/00, BDH)
Rollers (Denley Spiramix 5 or equivalent)
Rotary shakers (Labinco BV or equivalent)
Microscope glass slides 76 x 26mm (Menzel Glaeser, Superfrost Color)
Coverslips 22 x 22mm, 22 x 40mm (Menzel Glaeser)
Forceps (Watkins and Doncaster)
A001 size dissecting tungsten needles (E6871, Watkins and Doncaster)
Needle holder (Watkins and Doncaster)
26G3/8, 0.45 x 10 syringe needle (Industrial and Scientific)
Artist's paintbrushes 00, 000
Heptane (27051-2, Sigma)
Methanol (M/4000/PC17, Fischer Chemicals)
Ethanol (E/0555DF/17, Fischer Chemicals)
Formaldehyde (37-40%, 101134-A, AnalaR/BDH, reagent grade)
Paraformaldehyde (40% EM grade, 15715, Electronmicroscopy Sciences)
Triton X-100 (T-9284, Sigma)
Bovine serum albumin (BSA) fraction IV (A-2153, Sigma)
NaCl (102415K, AnalaR/BDH)
KCl (101984L, AnalaR/BDH)
NaH2PO4 (301324Q, AnalaR/BDH)
Na2HPO4 (S-9390, Sigma)
KH2PO4 (102034B, AnalaR/BDH)
MgCl2 (63064 Fluka Biochemika)
EGTA (E-4378, Sigma)
PIPES (P-8203, Sigma)
SDS (Ultrapure, Melford Labs Ltd.)
Glycerol (101184K, AnalaR/BDH)
Sucrose (S-0389, Sigma)
Apple juice (commercial)
Bactoagar (0140-01, Difco laboratories)
Nipagin (Nipa Laboratories Inc.)
Baker's yeast blocks (Commercial)
Vectashield (H-1000,Vector Labs, Burlingame)
Voltalef H10S Halocarbon oil (Elf Atochem, France)
Fly food (Instant Drosophila medium, Phillip Harris Scientific)
A. Producing Embryos for Confocal Microscopy
B. Removal of Chorion (Dechorionation)
In this step, the embryos are collected from the plate, washed well to remove yeast and other detritus from the plate and the eggshell (chorion) removed. You should prepare vials ready for the fixation step (Cl) before starting this step.
C. Standard Fixation and Removal of Vitelline Membrane (Devitellinisation)
Embryos can be stored in methanol or ethanol at -20°C for several months and will continue to be successfully stained by most antibodies. However, detection of some antigens and GFP fluorescence are affected adversely by exposure to ethanol or methanol. In these cases, do steps 3-5 as rapidly as possible and proceed quickly to step D1. Some antibodies will not recognise their antigen in embryos fixed by this method, but will when one of the alternative methods described in Section III, G is used.
D. Antibody Staining of Embryos
E. Staining with Phalloidin to Visualise Actin
F. Standard Mounting on a Microscope Slide Steps
Almost every aspect of this procedure will vary from fly laboratory to fly laboratory and even within our laboratory there are a number of variations. None of the following are critical: which recipe is used for the egg collection plates (some laboratories prefer grape juice), the tubes used for fixation and staining, the solutions used for antibody staining, the timing of the incubations and the number of washes, and the medium for mounting the embryos. As mentioned at the beginning, the key steps are fixation and devitellinisation. Again there are many variants of this, and the important thing is to get a method that works well for the antibodies you are using. Once you have a method that works, try to do this step as identically as possible each time. This will give you reproducibly good staining. Generally, the first time a new person in the laboratory does this technique it does not work well. However, after a few times, it is hard to understand how you could have failed the first time. A simple check is to look at a few embryos at step C4 or D1 down a dissecting microscope. If the embryos are still surrounded by the transparent shiny vitelline membrane, throw them away and try again. Before moving onto microscopy, we will mention some useful variants of this technique.
G. Alternative Methods for Embryo Fixation and Devitellinisation
1. Alternative Fixation Solutions In these the method is identical to that described earlier, but instead of 1:1 4% formaldehyde in PBS: heptane, and 20-30min at room temperature, use the following alternatives.
1:1 4% formaldehyde in PEM:heptane, 20-30min at room temperature.
1:1 4% formaldehyde in PB :heptane, 20-30 minutes (Uemura et al., 1996).
1:1 37% formaldehyde:heptane, 2-5min at 25-37°C with gentle mixing
This can improve the preservation of cytoskeletal structures present in membrane compartments [e.g., scribble, β-heavy spectrin (Bilder et al., 2000)] and gives the best preservation of microtubules (Foe et al., 2000).
2. Heat Fixation (Muller et al., 1996)
Heat fixation improves the penetration of antibodies into late stage embryos and larvae (when the exoskeleton normally blocks antibodies).
3. Fixation Conditions for Visualising Nonmuscle Myosin (Foe et al., 2000)
4. Methanol Fixation
This is a good alternative when a new antibody fails to stain embryos fixed under standard conditions, as a number of antibodies have been found to only work on embryos prepared by this method. Other antibodies will work with both this method and standard fixation, while some will not stain with this method.
5. Fixation Conditions for Visualising GFP
If you are planning to visualise GFP fluorescence, keep exposure of the embryos to methanol or ethanol to a minimum and do not seal the slides with clear nail varnish. Organic solvents quench GFP fluorescence or severely affect its ability to fluoresce.
H. Embryo Thick Sections (Adapted from Grosshans and Wieschaus, 2000)
To get a good cross-sectional view of the embryo by confocal microscopy it is necessary to cut sections. They give far better resolution than optical X-Z scans (see later and Figs. 2E and 2F). We have used a relatively crude method that does not require a microtome and is compatible with our standard methods for antibody staining. These sections are particularly useful for the visualisation of morphogenetic events that occur in the midline and in the interior of the embryo, such as invagination of the mesoderm, formation of the midgut, and dorsal closure.
I. Confocal Microscopy
Drosophila embryos can be examined with a variety of confocal microscope setups. Both inverted and upright microscopes are appropriate. The key variable is the available laser lines; this will dictate the optimal choice of fluorochrome-conjugated secondary antibodies that one should use. A detailed description of confocal microscopy is clearly beyond the scope of this article (a good reference is Centonze and Pawley, 1995), but we hope to provide a few helpful hints.
Once one has obtained an initial quick view of the result of the staining, which is often possible by conventional fluorescence microscopy if the signal is strong, the usual goal is to produce a set of images for publication. The biggest challenge is to produce a set of comparable images of different embryos. For example, for a general description of the distribution of a new protein one would like comparable views of embryos at different stages; generally a lateral view. There is a strong convention in the fly field regarding images of embryos: they should always have anterior to the left and dorsal up. Similarly, for a comparison of staining in different mutants, it is essential to obtain a suitable micrograph from wild type and each mutant at the same stage and with the same view: this can often require looking through many embryos. Clearly, to be able to achieve this you must be able to tell the stage of the embryo and which way is up, which is not as easy as it sounds. The only way to achieve this is to spend time looking at embryos down the microscope. To learn your way around embryonic development, it helps to examine embryos that are stained with antibodies that highlight the development of a particular tissue. The following antibodies are recommended for this: Fasciclin III (available from the Developmental Studies Hybridoma Bank) and phosphotyrosine (available from Sigma). For an overall view of the embryo, phalloidin (rhodamine conjugated, Molecular Probes) is very useful (see Fig. 2).
In general, we start our confocal session by first scanning through the slide by conventional fluorescence, taking note of the approximate position on the slide of embryos of the right stage and orientation that have stained well. Next we find a well-stained embryo that is not one of the best ones and use it to work out the appropriate settings for each channel on the confocal. When it is essential that there is minimal mixing between the outputs of each channel, then it is advisable to do sequential scanning rather than simultaneous. Use the channel with the brightest signal to get started. In most cases we use a collection area of 1024 x 768 pixels. Using rapid scanning, find the best focal plane, adjusting the focus by hand (adjustments to the eyepieces often result in the view by eye being at a different focal plane than the resultant scan). Using intermediate scan rates, set the laser power, iris, gain, and background levels appropriately for the first channel and then proceed to the next channel. We keep the iris similar in the different channels so that they all capture the same depth of focus. Some confocal setups have this feature built into the software.
Once all channels are set up, adjust the focus to obtain the optimal image. Collect images using Kalman scanning to improve the signal-to-noise ratio. We use either 166 lines per second with three to five scans or 50 lines per second and two scans. Make a note of the orientation of the embryo, especially if you are just focused on a small region.
Because the embryo is curved, in many cases a single section will not capture all of the relevant tissue. In this case, collect a series of sections at different focal planes and combine them into a single image using the projection software. The first step is to find the top and bottom of the series you wish to collect. Turn on the focus motor, doing fairly rapid scans in a channel with the most robust signal, and vary the focal plane to find the top and bottom. Be aware that with an oil objective lens, the new focal plane can take a minute to "settle." Set up the software to collect the Z series. Generally we use a step of 1 µm, with a range from 0.5 to 2.0 µm. Collect the Z series and project the image. For some tissues, e.g., the somatic muscles, it helps if the embryos are mildly flattened by mounting in a minimal amount of Vectashield, as described in Section III,F,2; then fewer scans are required to visualise all the muscles (Figs.2A and 2B).
One other variable to keep an eye on is the zoom. We standardly keep this at 1 (no zoom), and it is helpful to check that it has not been varied by a previous worker, as your images from session to session will not be comparable. Some judicious use of the zoom function can improve resolution. In practice we have found that a zoom factor of greater than two provides no further improvement in resolution and may even lead to loss of resolution.
J. Examination of GFP in Living Embryos
IV. COMMON PROBLEMS
A. Nonstaining Embryos
The most common causes are (i) use of inappropriate fixation method, (ii) inefficient devitellinisation, (iii) inappropriate primary and/or secondary antibody, and (iv) faulty laser, either nonfunctioning or unsuitable for the fluorochrome coupled to the secondary antibody.
This is a common problem inherent to confocal microscopy, especially during the acquisition of a Z series at high laser power and high magnification. It can be reduced by ensuring that the laser power is set to the minimum value that is optimal for the image.
C. Low-Resolution Images
Low-Resolution Images can result from faulty image processing of the original image. Obtain the highest resolution image possible on the confocal. Information that does not exist in the acquired data cannot be added by image processing software. Too high a gain can result in too much noise. Such noisy images may be improved by increasing either the iris or the laser power. Out of focus signals resulting from inappropriate mounting and from inappropriate thickness of sections are the cause of low-resolution images from sections.
We thank Sylvia Erhardt, Anja Hagting, Jasmin Kirchner, Jordan Raft, Katja Roeper, Alex Sossick, and Christos Zervas for comments and suggestions.
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