Biotech Glossary | Bioinformatics | Lab Protocol | Notes | Malaysia University |
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In Situ Hybridization to Drosophila Embryo Sections Using α-[35S]-Riboprobes |
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| Overview |
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| This protocol describes the detection of mRNA sequences in sectioned Drosophila embryos. | |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| Procedure |
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A. Fixing the Embryos 1. Use as many embryos as is practical to collect. Working with less than 500 to 1,000 embryos will probably be difficult (a gram or two should suffice). 2. Wash the embryos from the collection plate into a sieve with a fine enough mesh to retain the embryos. Rinse them with ddH2O to remove excess yeast. 3. Dechorionate (remove the outer layer) the embryos in 50% Bleach Solution for approximately 2 min. 4. Rinse the dechorionated embryos thoroughly with ddH2O containing 0.1% Triton X-100. 5. Transfer the embryo to a tube, with a tight-fitting lid, that contains a 1:1 mixture of freshly made Formaldehyde Solution and Heptane. For a gram of embryos, use a 50 ml tube with 20 ml each of Formaldehyde Solution and Heptane. For fewer embryos, scale down the volumes appropriately. 6. Shake the tube vigorously for 15 min. 7. Remove as much of the aqueous (bottom) layer as possible with a pipette. Be careful not to remove the bulk of the embryos that will be at the interface. 8. Add a volume of Methanol equal to the volume of the Heptane layer in the tube. 9. Shake the tube very hard for approximately 30 sec and allow the two phases to separate. The devitellinized embryos will sink to the bottom of the tube. The rest will remain at the interface with the empty vitelline membranes. The yield at this stage is variable (greater than 50% to 60% is good). If there are fewer than 50%, shake the tube harder and/or longer. 10. Remove the entire interface and as much liquid as possible, leaving the embryos at the bottom of the tube. 11. The rest of the treatments are carried out in the same tube the embryos are now in. For steps that call for washing, each liquid is added to the tube, the embryos are allowed to settle, and the liquid is removed with an aspirator. 12. Rinse the embryos in Methanol for 2 min. 13. Rehydrate the embryos with the following schedule: Wash for 2 min in (9:1) Methanol:Formaldehyde Solution Wash for 2 min in (7:3) Methanol:Formaldehyde Solution Wash for 2 min in (1:1) Methanol:Formaldehyde Solution Wash for 2 min in (3:7) Methanol:Formaldehyde Solution 14. Post-fix the embryos in Formaldehyde Solution for 20 min. 15. Wash the embryos once in PBS. 16. Dehydrate the embryos in a series of ethanol solution washes as follows: Wash for 5 min in 30% Ethanol Wash for 5 min in 50% Ethanol Wash for 5 min in 70% Ethanol 17. The embryos can be stored in 70% ethanol at -20°C for several weeks. B. Embedding the Embryos 1. Prepare the Paraplast Plus Embedding Medium by melting some in a conical flask in a water bath and add an equal volume of pre-warmed xylenes (see Hint #3). 2. Fully dehydrate the embryos as follows: Wash for 5 min in 90% Ethanol twice Wash for 5 min in 95% Ethanol twice Wash for 5 min in 100% Ethanol four times 3. Transfer the embryos to a glass test tube and incubate them in Xylenes for 10 min. 4. Remove the Xylenes, add fresh Xylenes, and incubate for 10 min. 5. Incubate the embryos in a 1:1 mixture of Paraplast Plus Embedding Medium:Xylenes at 61°C for 10 min. 6. Repeat the 10 min incubation with fresh Paraplast Plus Wax:Xylene mixture. 7. Keep the tubes in a 60 to 61°C hot block for the rest of the procedure. 8. Incubate the embryos in the Paraplast Plus wax at 61°C for 10 min twice. 9. Set up the Electron Microscopy Science's Peel-A-Way molds on a hot plate set at a temperature that will ensure that the Paraplast Plus wax does not set when a little is put in the bottom of the mold. 10. Quickly dispense some of the embryos in wax into the bottom of the mold. Use a Pasteur pipette with the end broken off for this operation. Warm the broken end briefly in the flame of a Bunsen burner to bring it to the temperature of the wax in the water bath and then pipette the embryos. Put enough wax/embryos solution in the mold to cover the base of the mold in a thin layer. The number of embryos should be enough to form a monolayer in the mold; fewer embryos will result in fewer embryos in each section, and more embryos will prevent the embryos from settling horizontally in the wax. 11. Allow the embryos to settle for about 1 min and then try to make them move into the center of the mold. Do this by blowing the surface of the wax with a Pasteur pipette, pushing the embryos away from the sides of the mold. (This takes a little practice to perfect the skill; however, it really is not critical to the experiment if the embryos are spread around the base of the mold.) 12. Fill the mold with Paraplast Plus wax by slowly and gently running more wax down the side of the mold. Try not to disturb the embryos. The wax should fill the mold to approximatley two-thirds of its volume. Do not let the wax solidify before adding more to the mold. This will form a weak region, and the block may break. 13. Carefully remove the mold from the hotplate and allow it to stand undisturbed for about 15 min at room temperature to set. 14. The blocks can now be stored at 4°C for several months before cutting sections. C. Subbing the Slides (see Hint #4) 1. Wash the slides by soaking them in a large container of hot water with some detergent and gentle agitation. Rinse the slides thoroughly (i.e., in running water for 30 min) and perform a final rinse with ddH2O. 2. Load the slides into racks and dry them in an oven. 3. Soak each rack of slides in a solution of 50 μg/ml poly-D-lysine in 10 mM Tris, pH 8.0 for 10 min (see Hint #5). 4. Air-dry the slides in a dust-free environment for 24 hr. Store the slides in boxes at room temperature. D. Cutting Sections 1. The exact procedure for cutting sections will depend on the microtome used. This portion of the protocol should serve more as a guide than a strict step-by-step procedure. 2. Push the wax block out of the mold and mount it into a wooden microtome block. Use molten Paraplast Plus to attach it. 3. Trim the block with a razor blade so that the face to be cut is a trapezoid shape. 4. Mount the block onto the microtome and cut 6 micron-thick sections. 5. Place a subbed slide onto a hot plate set at 45°C. 6. Put a drop of clean water onto the slide and carefully place a section onto the water drop (a small paintbrush is an easy way to handle the sections). It is important to place the newly cut (shiny) side of the section downwards. The section will spread flat over the water and will be slowly lowered to the slide surface as the water evaporates. After you have put sufficient sections on the slide, allow the slide to sit on the hotplate undisturbed. The number of sections that can fit onto each slide depends on the size of the sections and the size of the coverslip used for the hybridizations. About four sections will fit under 22 mm2 coverslips. 7. Allow the slides to sit on the 45°C hot plate overnight (or in a 45°C oven). 8. The sections can be stored at room temperature in a dry box for several weeks prior to hybridization. E. Transcribing the α-[35S]-Riboprobe 1. Linearize the gene template DNA downstream of the insert with a restriction digest. 2. In order to remove any contaminating RNases, treat the digest with 200 μg/ml Proteinase K for 30 min at 37°C. 3. Add an equal volume of Phenol:Chloroform and vortex. 4. Centrifuge for 3 min at full speed in a microcentrifuge to separate the phases. 5. Recover the aqueous phase, add an equal volume of chloroform, and vortex. 6. Centrifuge for 3 min at full speed in a microcentrifuge to separate the phases and recover the aqueous phase. 7. Add one-tenth volume of 3 M Sodium Acetate and 2 volumes of Ethanol to precipitate the DNA. 8. Centrifuge for 5 min at full speed in a microcentrifuge to pellet the DNA. 9. Resuspend the DNA to a final concentration of 1 μg/ml DNA. 10. Set up a transcription reaction with the following components (see Hint #6, Hint #8, and Hint #1): 1 μg of DNA templated 1 μl 10 mM rATP 1 μl 10 mM rCTP 1 μl 10 mM rGTP 1 μl RNase block (RNasin) 1 μl 0.75 M DTT 4 μl 5X transcription buffer 5 μl 40 μCi/ml α-[35S]-UTP 1 μl of a 1:5 dilution of T3 or T7 RNA polymerase in 1X transcription buffer final volume should be 20 μl 11. Incubate at 30°C for 1 hr. 12. Add 2.2 μl 10X MS and 1 μl of 1 mg/ml RNAse-free DNase. 13. Incubate at 37°C for 30 min. 14. Trichloroacetic Acid precipitate a tiny aliquot and determine the percentage of label incorporated (see Protocol ID#1945). This will allow you to calculate the amount of probe transcribed. Since new RNA is being transcribed in the reaction, the total amount of radiolabeled RNA probe will depend on the amount of the radioactive (limiting) nucleotide and the efficiency of incorporation. 15. Add tRNA as carrier. To determine the amount of tRNA to add to arrive at the correct specific activity of RNA in the hybridization, use the following example as a guide. For 75% incorporation of radiolabel into the RNA probe using 200 μCi of UTP in the transcription reaction, add 500 μg tRNA. For different efficiencies of incorporation, add an amount of tRNA that will give the same ratio of label to tRNA. (e.g., for 50% incorporation of the same amount of label, add 333 μg tRNA). 16. Add an equal volume of Phenol and vortex. 17. Centrifuge for 3 min at full speed in a microcentrifuge to separate the phases. Recover both the aqueous phase and the organic phase in separate tubes. 18. Add an equal volume of 10 mM DTT to the organic phase and vortex. 19. Centrifuge for 3 min at full speed in a microcentrifuge to separate the phases and recover the aqueous phase. Add this to the first aqueous phase. 20. Add an equal volume of chloroform and vortex. 21. Centrifuge for 3 min at full speed in a microcentrifuge to separate the phases and recover the aqueous phase. 22. Add an equal volume of 4 M Ammonium Acetate and 2.5 volumes of Ethanol to precipitate the RNA. 23. Chill the sample to -20°C. 24. Centrifuge for 10 minutes at full speed in a microcentrifuge to pellet the RNA. 25. Resuspend the RNA pellet in 70% Ethanol. 26. Centrifuge again for 10 minutes at full speed in a microcentrifuge to pellet the RNA. Resuspend the RNA in 50 μl of 10 mM DTT. 27. Add 50 μl of 2X Carbonate Buffer. 28. Incubate at 60°C for 140 min. This will give transcripts with a mass average of 100 bases (See Hint #9). 29. Add an equal volume of 0.2 M Sodium Acetate/1% Acetic Acid and 2.5 volumes of Ethanol. Chill at -20°C. 30. Centrifuge for 10 minutes at full speed in a microcentrifuge to pellet the RNA and remove the supernatant. 31. Resuspend the RNA in 70% Ethanol and centrifuge again to pellet the RNA. 32. Resuspend the RNA pellet in 50% high-quality Formamide. Aim for 500,000 cpm/ul in the formamide (see Hint #7). F. Prehybridization Treatment of the Slides 1. Load the slides into a rack. The following treatments should be carried out in 400 ml glass dishes. 2. Dewax the slides by incubating twice in Xylenes for 10 min each. 3. Rehydrate the slides in a series of Ethanol washes Wash for 2 min in 100% Ethanol Wash for 2 min in 95% Ethanol Wash for 2 min in 80% Ethanol Wash for 2 min in 60% Ethanol Wash for 2 min in 30% Ethanol 4. Incubate the slides in 0.2 M HCl for 20 min at room temperature. 5. Rinse in ddH2O for 5 min. 6. Incubate the slides in 2X SCC for 30 min at 70°C. 7. Rinse in ddH2O for 5 min. 8. Digest with 0.125 mg/ml Protease in P Buffer at room temperature for 10 min. 9. Incubate the slides in 0.2% Glycine in PBS for 1 min. 10. Rinse in PBS twice for 1 min each. 11. Postfix in Formaldehyde Solution for 20 min at room temperature. 12. Rinse once in PBS for 1 min. 13. Acetylate the sections by incubating the slides in 0.5% Acetic Anhydride in 0.1 M Triethanolamine, pH 8.0 for 10 min at room temperature with vigorous stirring in a fume hood. Acetic Anhydride is very unstable in water. Add it to the Triethanolamine Solution at the same time as the slides. 14. Rinse slide in PBS for 2 min. 15. Dehydrate the slides in a series of ethanol washes: Wash for 2 min in 30% Ethanol Wash for 2 min in 60% Ethanol Wash for 2 min in 80% Ethanol Wash for 2 min in 95% Ethanol Wash for 2 min in 100% Ethanol 16. Air-dry the slides. The hybridizations should be set up as soon as possible after this pretreatment, as the sections are not very stable at this stage. G. Hybridization of the Radiolabeled Riboprobe to the Sections 1. The RNA probe concentration during the hybridization should be around 100,000 cpm/ul. (i.e., diluted five-fold from the stock made in step #D32) 2. Boil the radiolabeled probe for 2 min. (Use 3 μl for each slide to be hybridized.) 3. Immediately chill the probe on ice and add four volumes of Hybridization Buffer. 4. Mix well and centrifuge for 30 sec in a microcentrifuge. 5. Place 15 μl radiolabeled probe onto each slide near the edge of the tissue sections. 6. Carefully lower a clean (but not necessarily silanized) 22 mm2 coverslip over the so that the probe covers the sections. Try to minimize the introduction of air bubbles. 7. Seal the edge of the coverslip with ample rubber cement dispensed from a 5 ml syringe. 8. Hybridize overnight in a humidified chamber at 50°C. H. Washing the Sections 1. Carefully peel off the rubber cement with a sharp pair of forceps. Do not dislodge the coverslips during this step. 2. Place the slide in a rack in a tank of Wash Buffer at 50°C. Perform this and all subsequent washes in a fume hood (the Wash Buffer has a foul odor). Suspend the rack so that there is space for the coverslips to slide off, which should happen in a few minutes. If needed, stir the Wash Buffer gently. 3. When the coverslips come off, move the rack into a fresh tank of Wash Buffer. Wash for approximately 4 hr at 50°C, changing the Wash Buffer every hour. 4. Wash the slides in 1X NTE at 37°C for 5 min. 5. Incubate the slides in 20 μg/ml RNaseA (dilute RNAse from a 10 mg/ml stock) in NTE at 37°C for 30 min to remove any single-stranded RNA and reduce the background. 6. Wash in 1X NTE at 37°C for 1 hr with four changes of buffer. 7. Dehydrate the slides in a series of ethanol washes: Wash for 2 min in 30% Ethanol in 0.3 M Ammonium Acetate Wash for 2 min in 60% Ethanol in 0.3 M Ammonium Acetate Wash for 2 min in 80% Ethanol in 0.3 M Ammonium Acetate Wash for 2 min in 95% Ethanol in 0.3 M Ammonium Acetate Wash for 2 min in 100% Ethanol in 0.3 M Ammonium Acetate 8. Air-dry the slides. I. Autoradiography 1. Use Kodak NTB2 Emulsion previously diluted to 1:1 with water (see Hint #10), placed in aliquots in 5 ml scintillation vials, and stored in absolutely light-proof and radiation-free conditions at 4°C (see Hint #10). Select a container to use as a dipping chamber that is large enough to allow the sections to be covered by emulsion when the slide is dipped into the dipping chamber. 2. Set up a water bath at 45°C in a dark room and melt an aliquot of the emulsion. Put the vial inside a lightproof film-developing tank filled with an adequate amount water and put the whole thing into the water bath. Alternatively, a dark room with a rotating door can also be used. The emulsion takes about 15 min to melt. 3. Pour the melted emulsion into the dipping chamber, which should be sitting in the water bath to keep it warm and to prevent the emulsion from solidifying. Dip each slide briefly into the dipping chamber and remove it without rubbing off the layer of liquid Emulsion. Place the slide into a rack so that it stands vertically. This allows excess Emulsion to drain away from the sections and causes a thin, uniform layer to form over them (see Hint #11). Also, dip a clean slide and develop it as a negative control. Very few silver grains should appear. 4. Allow the slides to dry for at least 2 hr before putting them into light-proof boxes containing Drying Gel (e.g., Drierite) (see Hint #12). Tape the boxes shut and wrap them extensively in aluminum foil. Expose the slides at 4°C away from other sources of radioactivity. 5. Develop the first set of slides after 4 or 5 days. Moderate to strong signals should be adequately exposed by then. Determine the amount of time appropriate for the other slides from the amount of signal determined at this time. J. Developing the Slides 1. It is critical that all the solutions used in the developing and staining of the slides are at the same temperature, which should be below 20°C. If they are not, the emulsion, which is very delicate when wet, may come off during their treatment and ruin the entire experiment. To avoid this, make up the solutions the day before developing and keep them in an 18°C cold room. When ready for use, you can move them into the dark room and allow then to warm up slowly together. 2. Allow the box of slides to come to room temperature for 1 hr while still sealed. 3. In a dark room with appropriate safe light, load the slides into a rack. 4. Immerse them in Developer for 2 min (CAUTION! See Hint #1). 5. Transfer them to 2% Acetic Acid for 30 sec, which stops the action of the Developer (this may not be crucial). 6. Immerse the slides in Fixer for 5 min (CAUTION! See Hint #1). 7. Rinse the slides in distilled water for 15 min with two changes of water during the incubation. The slides can be exposed to light from this point on. K. Giemsa Staining of Sections 1. The degree of Giemsa staining desired varies with the intensity of the signal. If the signal from the probe is strong, stain for approximately 15 to 20 min. If the signal from the probe is weak, stain for only 30 sec to 1 min. A good rule of thumb is to stain the first batch of slides while assuming a weak probe signal and adjust for the next batches accordingly. 2. Immediately before staining the slides, make up a 5% Giemsa in 10 mM Sodium Phosphate buffer, pH 6.8. 3. Incubate the slides in the Giemsa solution for the desired amount of time. 4. To destain, gently pour ddH2O into the container until the film that forms at the surface of the Giemsa solution is poured away. This step helps to ensure that the film does not coat the slides. 5. Rinse the slides two or three times in water for about 30 sec each. While the slides are wet, they can be viewed at low magnification; and if they appear over-stained, they can be rinsed more thoroughly, although this is usually unnecessary. 13. Air-dry the slides and then mount them in DPX Mounting Medium under a coverslip. |
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| Solutions |
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| BioReagents and Chemicals |
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Hydrochloric Acid Formamide Acetic Acid Sodium Carbonate Sodium Bicarbonate Ammonium Acetate DNase, RNase-free [35S]-UTP DTT rGTP DPX Mounting Medium Fixer Developer rCTP Emulsion rATP Spermidine Magnesium Chloride Isoamyl Alcohol Chloroform Phenol Proteinase K Tris-HCl Poly-D-Lysine Oligonucleotide tRNA RNasin Paraplast Plus Embedding Medium Xylenes Heptane Giemsa RNase A Ethanol Sodium Acetate 2-Mercaptoethanol Methanol Paraformaldehyde Formaldehyde Sodium Phosphate, Monobasic Potassium Phosphate, Monobasic Triton X-100 Bleach, Household Sodium Hydroxide Bovine Serum Albumin (BSA), Fraction V Plolyvinylpyrrolidone Ficoll 400 Dextran Sulfate Triethanolamine Acetic Anhydride Glycine Protease, Type XIV: Bacterial Trichloroacetic Acid (TCA) EDTA Sodium Citrate Sodium Phosphate, Dibasic Potassium Chloride Sodium Chloride |
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1. CAUTION! This substance is a biohazard. Consult this agent's MSDS for proper handling instructions. 2. Make sure to use equilibrated phenol. 3. Paraplast Plus melts at 56°C and must not get hotter than 62°C. The solid wax takes quite a long time to melt and should be set up at least 1 hr in advance. Also, set up the water bath and a hot block for the test tubes of embryos far enough in advance so that the temperature has completely stabilized at 60 to 61°C. 4. The microscope slides are coated in poly-D-lysine to allow the sections to adhere better to the slide. The slides can be subbed in large batches and stored in a dust-free environment. 5. Use poly-D-lysine with a molecular weight range of 70,000 to 150,000. 6. The Stratagene transcription kit can be used. 7. For the example above, 75% incorporation of 200 μCi, use 96 μl of Formamide. The probe made here is 5X concentrated. Store at -20°C until use. 8. To minimize degradation of RNA by RNases, wear gloves when handling samples and reagents, and change gloves regularly while working. Treat water and solutions with DEPC (Diethyl Pyrocarbonate) to inactivate RNases and use solutions prepared with RNase-free water and equipment (see the BioTools section of this website). For more information about precautions when working with RNA, see Reference Pages under Working with RNA. 9. It is wise to run a small aliquot on a sequencing gel to check the size. Aim for no smaller than 50 bases and no larger than 200 bases (average sizes). 10. When making the 1:1 diluted emulsion, melt the emulsion at 45°C (takes about 45 min) and gently mix in water pre-warmed to 45°C. Do not allow bubbles to form; that will increase background. As the emulsion is very sensitive, make sure that no radioactivity is also stored in the refrigerators where the emulsion is stored. 11. While dipping the slides and drying them, they must be kept out of all light other than a weak safelight appropriate for the emulsion used. 12. It is convenient to have three or four slides for each probe used and to put one of each type into each box. This way, when you want to develop them, only one box needs to be opened. |
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| Citation and/or Web Resources |
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1. Cox KH, DeLeon DV, Angerer LM, Angerer RC. Detection of mRNAs in sea urchin embryos by in situ hybridization using asymmetric RNA probes. Dev. Biol. 1984;101:485-502. |
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