CSH Lab Manual 1997

Cold Spring Harbor 1997

EARLY

DEVELOPMENT OF

XENOPUS LAEVIS

COURSE MANUAL, COLD SPRING HARBOR FIFTH EDITION, 1997

EDITED BY:

HAZEL L. SIVE Whitehead Institute for Biomedical Research

ROBERT M. GRAINGER University of Virginia

RICHARD M. HARLAND University of California, Berkeley

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FOREWORD This manual arose from a course taught at Cold Spring Harbor Laboratory. We have made it available to the Xenopus community since it contains information that is not collected into a single work elsewhere. However, we would like to emphasize that this is a work in progress. It may contain innacuracies and mistakes, and for these we apologize. Only selected references to particular techniques and their uses have been cited, and we apologize for glaring omissions. We will be glad to remedy problems brought to our attention. Many of the techniques described here are illustrated in an accompanying set of videotapes which are cross-referenced to the appropriate manual section.

ACKNOWLEDGEMENTS The contribution of many people to entries in this manual cannot be overstated and we thank them. These include Enrique Amaya, Leila Bradley, Marietta Dunaway, Tabitha Doniach, Rick Elinson, Laura Gammill, John Gerhart, Jeremy Green, Ali Hemmati- Brivanlou, Ray Keller, Chris Kintner, Kris Kroll, Peggy Kolm, Mike Klymkowsky, Paul Krieg, Nancy Papalopulu, Charles Sagerstrom, Bill Smith, Dan Wainstock and Paul Wilson. We thank also students of the course and members of our laboratories for suggestions and criticism. Thanks for permission to use figures goes to Rick Elinson, Jonathan Slack...

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TABLE OF CONTENTS A. Introduction .................................................................................................................7 1. Where to obtain Xenopus ............................................................................7 2. How to house and feed them.......................................................................7 3. Raising Tadpoles and Frogs.........................................................................11 4. Diseases, preventions and cures.................................................................13 B. Equipment for embryo experiments .....................................................................15 1. Variable temperature incubator.....................................................................15 2. Microscopes...................................................................................................15 3. Microinjector, micromanipulator and needle puller.......................................15 4. Dissecting tools ..............................................................................................16 5. Cameras and taking pictures.........................................................................18 C. Getting embryos.......................................................................................................20 1. Handling Xenopus adults ..............................................................................20 2. Inducing ovulation...........................................................................................20 3. Isolating the testes .........................................................................................23 4. Collecting eggs ..............................................................................................23 5. In vitro fertilization............................................................................................24 6. Natural mating .................................................................................................25 7. Keeping track of frogs....................................................................................25 D. Preparing embryos for manipulation .......................................................................26 1 Dejellying embryos ........................................................................................26 2. Removing the vitelline membrane...............................................................26 E. Embryo perturbations...............................................................................................27 1. Ultraviolet light.................................................................................................28 2. LiCl ..................................................................................................................29 3. Retinoic acid....................................................................................................30 4. Inducing exogastrulation ................................................................................30 5. How to mark the future dorsal side by tipping and staining .......................30 F. Microinjection...............................................................................................................32 1. Injection checklist.............................................................................................32 2. Microinjection of oocytes ...............................................................................33 Frog surgery............................................................................................33 Preparation of oocytes...........................................................................33 References..............................................................................................34 Injection....................................................................................................34 3. Preparation of secreted proteins from oocytes...........................................36 4. Microinjection of embryos .............................................................................36 F. Lineage Labeling .......................................................................................................38 1. Finding the correct labeling site .....................................................................38 2. FDA and RDA................................................................................................38 3. β-galactosidase RNA Injections....................................................................39 4. X-gal Staining Xenopus Embryos ...............................................................40 5. Green Fluorescent Protein (GFP) RNA.......................................................40 G. Microdissection..........................................................................................................42 1. Animal cap isolation........................................................................................42 2. Ectodermal (animal cap) layer separations..................................................42 3. Dissociation/reaggregation of animal cap/ other cells..................................43 4. Animal cap/vegetal conjugations ..................................................................44 5. Animal cap/dorsal mesoderm conjugates....................................................45 3


6. Einstecks .........................................................................................................45 7. Keller sandwiches...........................................................................................46 8. Tissue transplantation.....................................................................................49 9. Dissection of tightly adhering tissues by trypsin treatment........................49 10. Fixing and dissecting Xenopus embryos .................................................50 11. Cortical isolation from oocytes and eggs...................................................51 H. Method for Generating Transgenic Frog Embryos...............................................52 1. Introduction.....................................................................................................53 2. Reagents and Equipment............................................................................57 3. Methods.........................................................................................................62 4. Notes..............................................................................................................69 5. References ...................................................................................................71 6. Figures and Legends.....................................................................................75 7. Transgenic frog embryos- the short method...............................................78 I. Immunocytochemistry.................................................................................................89 1. Solutions and Equipment..............................................................................89 2. Embryo preparation and fixation..................................................................90 3. Embryo rehydration.......................................................................................91 4. Antibody incubation and washes .................................................................91 5. Immunodetection and HRP staining.............................................................92 J. In situ hybridization .....................................................................................................93 1. The probe.......................................................................................................93 2. The embryos..................................................................................................94 3. Vials or Baskets? ...........................................................................................95 4. Hybridization procedure................................................................................97 5. Washing..........................................................................................................98 6. Antibody incubation.......................................................................................99 7. Mounting chambers .......................................................................................100 8. Observations..................................................................................................101 9. Short protocol.................................................................................................103 10. Double staining in situ protocol for whole mount in situs ..........................104 11. References...................................................................................................106 K. Histology ....................................................................................................................108 1. Paraffin Method ..............................................................................................108 2. Plastic Method................................................................................................110 APPENDIX I Nucleic acid and protein methods .....................................................................112 RNA Methods................................................................................................................112 1. RNA Isolation .................................................................................................112 Hot phenol/ LiCl RNA isolation.............................................................112 Proteinase K/ LiCl RNA preparation....................................................113 Acid Guanidinium/ Phenol RNA Isolation.............................................114 Oligo dT column......................................................................................115 2. RNA Detection...............................................................................................116 Northern Analysis ...................................................................................116 RNAse Protection ..................................................................................119 PCR-based Protocols...........................................................................121 Standard PCR Protocol.........................................................................125 3. Preparation of in vitro transcribed RNA........................................................126 4. RNA Solutions ...............................................................................................130 5. Siliconizing tubes, glassware ........................................................................132 DNA Methods................................................................................................................133 1. Extraction of DNA from single embryos......................................................133 2. Bulk DNA isolation from red blood cells ......................................................134 4


Isolation of frog Red Blood Cells .........................................................134 Preparation of genomic DNA from red blood cells.............................135 Western Blots.................................................................................................................136 APPENDIX II Culture media......................................................................................................137 1. General media................................................................................................137 Modified Barth's Saline..........................................................................137 Amphibian Ringers ...............................................................................138 NAM (normal amphibian medium).......................................................139 Holtfreter's solution .................................................................................139 2. Specialized media .........................................................................................140 Danilchik's Blastocoel Buffer ..................................................................140 Sater's modified blastocoel buffer........................................................140 LCMR (Low Calcium Magnesium Ringer's).......................................141 CMFM (Calcium Magnesium Free Medium)......................................141 PhoNaK...................................................................................................141 3. Oocyte Culture Media...................................................................................142 O-R2 Medium ........................................................................................142 Oocyte Culture Medium........................................................................142 Devitellinating Buffer...............................................................................143 APPENDIX III Fate maps of blastula stages............................................................................144 References..........................................................................................................151 APPENDIX IV Morphology of Xenopus embryos and adults ...............................................152 1. Early development........................................................................................152 2. Neural Development.....................................................................................155 3. Adult morphology..........................................................................................163 4. References .....................................................................................................164 APPENDIX V Timing of development and temperature dependence.................................166

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A. Introduction Hazel Sive

Welcome to the world of Xenopus! This manual contains enough information to get you started on developmental analysis of frogs. It is not, however, a comprehensive volume. An additional helpful reference is Methods in Cell Biology 1991. vol. 36. Editors Brian Kay and Benjamin Peng, Academic Press. You will also need the Normal Tables of Xenopus laevis, by P. D. Nieuwkoop and J. Faber, recently reprinted by Garland, 1994. Another useful reference is The Early Development of Xenopus laevis by P. Hausen and M. Riebesell, Springer-Verlag, 1991. This contains excellent histology of early embryos. Peter Vize (U. Texas, Austin) has set up a Xenopus Web Site, with lots of useful information. The address is: http://vize222.zo.utexas.edu Another good Web site is the University of Wisconsin Amphibian Development Tutorial. It contains a glossary, and movies of developmental events. The address is: http://www.library.wisc.edu/guides/Biology/demo/frog2/mainmenu.html Xenopus laevis is a gentle animal, that lives in fresh water and can repeatedly be induced to lay eggs by simple hormone injection. These features, coupled with the large size of the embryos, allowing their micromanipulation and microinjection, and their rapid development make Xenopus an excellent animal for analysing early vertebrate development. The chief disadvantages of Xenopus laevis are its long generation time (one to two years) and its tetraploidy. Another species, Xenopus tropicalis does not have these disadvantages and may be a useful future substitute. A note: In several instances throughout the manual, you will find two slightly different procedures for doing the same thing, reflecting differences in the methods use by our labs, and reinforcing the point that there is seldom only one "right" method. 1. Where to obtain Xenopus The two major US suppliers are Nasco (Wisconsin) and Xenopus I (Michigan). Adult frogs cost about $20 each, late juveniles will usually yield eggs, though in smaller numbers. Frogs are shipped in peat moss (or equivalent), and may not be shipped in extreme summer heat or in frigid winters. Wait at least two weeks after their arrival before using them. Females are larger than males, with a prominent cloaca, males have rough, dark pads on the inside wrist (used to clasp the female during mating or amplexus). Xenopus I animals are endemically infected with nematodes and you should treat them prophylactically when they arrive (see Nematodes, in the diseases section). Do not accept thin animals, they are probably sick and will not lay well (as opposed to small and plump, which are fine). 2. How to house and feed them IMPORTANT: The more carefully you look after your frogs, the healthier they will be and the better the egg quality will be. This can take some effort, but sloppy frog care will be reflected in poor eggs and embryos. (See “stress and problems”). A healthy frog is placid, with moderately slimy skin and a nice pear shape. Jumpy frogs, those with dry skin or excessively slimy skin, bloated frogs, frogs that look grey and thin, or reddish are not healthy. Do not try to collect eggs from sick frogs, they will get even sicker and anyway the eggs or oocytes will be no good. Containers:

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Xenopus never leave the water and it is therefore unnecessary to have containers with any dry area. Standing water tanks For fewer than fifty or so frogs, it is a simple matter to house the frogs in still tanks of waterfor a tank about 2.5 ft x 1.5 ft. - four females or up to six males in approximately ten gallons. Water should be 5 - 8 inches deep. Plastic tanks are best, and opaque sides may approximate a pond more than clear sides. Tanks should be covered with a heavy lideither plexiglass with half inch holes, or stainless steel mesh, since frogs are very talented at jumping out of tanks (up to eighteen inches high). An opaque 4 inch pipe gives frogs somewhere to hide, which they like. Water in standing tanks should be replaced at least three times per week. Drip through systems For large numbers of frogs, it is more cost effective to house animals in a system that is at least partially self-cleaning. One possibility is a continuous drip system, where fresh water drips in and drips out constantly, preventing accumulation of wastes. This is probably the optimal way to keep frogs, since levels of toxic wastes (ammonia) are kept low and solid waste can be drained continously. The disadvantage is that this uses a lot of water and one has to keep very close watch on the input water quality. The chlorine content, pH and salinity of tap water can vary widely with the season or the whim of the water authority. If you choose to drip water in, you will therefore need to monitor water quality very carefully, preferably with a continuous monitoring system. Alternatively, you can drip in distilled water supplemented to about 20mM NaCl with Instant Ocean (or equivalent). However, this can use more distilled water than you have available. The idea here is to drip the water in slowly, as a fast stream may be bad for the frogs and also wastes a lot of water. Recirculating systems We (HLS) and others use a continously recirculating system that incorporates a biological filter. The water in our system is collected into a common drain, with U-tubes between the tank and common drain to prevent mixing dirty water between tanks. Water then flows into a reservoir where it gross particulate matter is filtered by a cheap air-conditioner pad, that we change every day. From the reservoir, it is pumped into a biological (bacterial) filter to remove ammonia and nitrites. It then flows through a sand filter to remove finer particulates and then through a high capacity series of UV-lights to kill bacteria and some other potential pathogens, before being pumped back to the tanks. The sand filter is backwashed every day, with some fresh water entering the system, such that water in the entire system is replaced every five days. Water quality is monitored by analysing pH, and concentration of ammonia, nitrates and nitrites at least once per week. We also do bacterial counts on water going back to the tanks (they should be essentially zero). This type of system requires some care: in our system, which contains about 500gal. and has a capacity of about 400 frogs, care takes about an hour per day. However, the health of frogs can be excellent in the controlled environment. In a continuous flow or drip through system, the flow should be slowish. Frogs are not fish, and may respond to vigorous water flow by developing "gas bubble disease", although the flow can be quite vigorous without causing apparent problems. The choice of whether to use a recirculating or flow through system is largely dependent on the water quality to which you have access. A drip through system is easier to look after than a recirculating one and in my opinion the system of choice, if you have good water quality available. Poor or variable quality tap water may make you want to consider using distilled water plus salts, in which case you will probably need to conserve water by recirculating it. 7


Water: Frogs collected in South Africa have traditionally been caught in large numbers from agricultural runoff and cattle drinking ponds. However, natural ponds contain bacteria and other organisms capable of keeping water clean, so bear in mind that a dirty tank is not equivalent to a natural pond. Water quality is very important. Chlorine, high pH, and ammonia are very bad for frogs. Water should be declorinated before use- where chlorine is the only additive, it can be removed from tap water by exposure to the air for several days (in standing tubs). Many water authorities add chloramine to the water- this is very stable and cannot be removed by aging. Running the water through a carbon filter (obtainable as a cartridge-type from most aqua-dealers, for example Barnstead) will remove chloramine. The cartridge should be changed frequently- chlorine and chloramine levels can be monitored with easy-to-use kits (sold by Hach). Deaminating liquids (such as AmmoLock) also work and can be purchased through pet stores. In addition, filter tap water through a dirt/rust or particle filter, also obtainable through your local aqua-dealer. NaCl should be added to a final concentration of 20mM. Alternatively, (cheaper) rock salt can be added to 1g/l (approx. 20mM). Alternately, several labs maintain frogs in distilled or ROH2O supplemented with salts. This is a good idea if the tap water quality in your area is poor or vaiable. It is a very bad idea to keep frogs in distilled water (their skin will flake and they may develop stress-related diseases). However, you can use distilled water supplemented to 20mM NaCl equivalent with Instant Ocean salts and pHed with Seachem Neutralization Buffer or with soda lime. pH: pH 6 - 7 is optimal. High pH can be very bad since the level of free ammonia rises with pH- at low pH (below about 7) ammonia waste is ammonium ion which is not toxic, but at high pH free ammonia is rapidly formed. Even with no ammonia in the system we have found that a change in 1 or more pH units (for example from 6.5 to 7.5) stresses the frogs and can lead to susceptibility to fungal growth or other disease. One response of the frogs to high pH is loss of their protective mucus which makes them highly susceptible to pathogens.

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Light/ temperature: Frogs should be kept on some type of light/dark cycle- 12 hours light/12 hours dark seems fine, so does 14 hours light/10 hours dark. We use "daylight" spectrum fluorescent lighting. Some have the impression that egg quality improves when frogs are exposed to sunlight equivalent light levels. Despite normally living in dark ditches, the frogs will come up to sit under the light. If one uses bright light, however, it is also important to have dark areas, such as plastic pipes, where frogs can escape the light if they want to. Temperatures of 16 - 20°C are optimal. Food: Frogs should be fed at least three times per week- several hours before performing a water change. Purina trout chow pellets float, which frogs like (they hunt insects on the top of the water in the wild) and are cheap and not very messy. Alternatives include Tetrahydromin fish food, and Nasco frog brittle however in our opinion no “perfect” pellet food for amphibians exits. If frogs have previously been fed liver, it may take them a week or so to eat the pellets- this can be accelerated by including a frog that already eats pellets in with those that don't to intiate a "feeding frenzy". Seasonal variation: Many investigators report seasonal variation in the quality of embryos, even for animals that have been kept for years or even bred without seeing seasonal light/dark changes. This may be due to changes in water quality or more importantly temperature. High summer temperatures (above 26oC) are no good for egg production. Injection with 50 - 100 Units of pregnant mare serum a few weeks before inducing ovulation may help- although all that seems to be available is chrionic gonadotropin from PMS (Sigma). However, this less purified CG likely also contains FSH which may help maintain egg production.

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3. Raising Tadpoles and Frogs Enrique Amaya Given enough space and time, raising frogs is fairly easy. Successful rearing of tadpoles to metamorphosed frogs requires around one liter per tadpole. Rearing full sized frogs requires up to four liters per frog. Hundreds of tadpoles can be generated by fertilizing several dishes of eggs in vitro as described elsewhere in this manual. Alternatively, unused embryos from several days of experiments can be pooled. It is important to consider that dejellied embryos must not be crowded. This leads to abnormal development usually stemming from gastrulation defects. The cause of these defects is probably anoxia. Large numbers of dejellied embryos should therefore be separated from each other by at least one embryo diameter at all times. At the swimming tadpole stage the embryos should be transferred to a tank with an adequate volume of good quality water containing 20mM rock salt. The best method to generate thousands of tadpoles is to induce "natural" matings between several females and males. To do this, inject each male and female with 500-800 UI of human chorionic gonadotropin (HCG; Sigma) and place the frogs in a large tank (> or = 20 gal.) containing good quality water supplemented with 20mM rock salt. After 6-8 hours the males will clasp the females around the hips (amplexus) and will fertilize the eggs as they emerge from the cloaca. It is important not to disturb the frogs while mating. Doing so may result in the release of amplexus and a decreased fertilization efficiency. For this reason it is often preferable to inject the frogs in the afternoon so that the frogs will have most of the night to mate in peace. After 24 hours or so, the males release the females and mating ceases. At this time the frogs should be removed from the tank, since the energy depleted frogs get hungry and do not mind eating their own eggs. After 3-4 days the embryos hatch and start colonizing the sides and surface of the tank. The unfertilized eggs should be removed since they will allow bacteria and fungi to contaminate the water. If the fertilization frequency is low, bacterial contamination can kill all the embryos. Therefore, when the fertilization frequency is low, the healthy embryos should be screened soon after hatching and transferred to fresh water supplemented with penicillin and streptomycin or gentamycin. This procedure, though, is time consuming and very laborious. After a week the tadpoles should be free swimming and ready to begin feeding, as evidenced by the rhythmic opening of the tadpoles' mouths. The tadpoles are filter feeders and do best when fed very fine food. We routinely feed the tadpoles a combination of nettle powder, active dry yeast and powdered bone meal mixed at 7:2:1 respectively. These ingredients can be obtained at health food stores. Alternately, Nasco tadpole brittle can be used succesfully. Tadpoles can also be raised on crushed trout pellets, although this regimen may not be ideal. The food should be delivered to the tank as a water suspension through a pipet. This decreases the amount of surface scum that forms when the food is added directly to the tank. Care should be taken not to overfeed the tadpoles. Presumably overfed tadpoles become anoxic due to clogged gills. Death due to overfeeding is common especially when the tadpoles are young. Generally the tadpoles should clear the water within two hours after feeding. Ideally, the tadpoles should be fed daily. Once a week I add fresh whole milk to the tank until the water is slightly cloudy. The milk and bone meal add calcium and phosphate to the water which should eliminate the chance of developing skeletal deformities in the metamorphosing frogs. Although not essential, aerating the water with submerged bubblers may increase the growth rate of the tadpoles. The tadpole water does not need to be changed very often. Once or twice a month is sufficient. Even within siblings, the tadpoles grow very asynchronously and while some metamorphose after two months others require four to six months to metamorphose. 10


During metamorphosis the water level in the tank should be no more than 30 cm deep, since the newly metamorphosed froglets may have difficulty reaching the surface to breath. Froglets cannot eat the tadpole food and must be fed small size trout chow (Purina, Pellet size 4) or frog brittle (Nasco). The small size pellets sink to the bottom where the froglets prefer to eat. Generally there is no need to remove the froglets from the tank. Froglets and sibling tadpoles can co-exist for several months until all the tadpoles metamorphose. In fact, it can be rather convenient to have a mixture of tadpoles and froglets in the tank because the tadpoles keep the water clean. If separated, the daily fed froglets foul the water constantly. Once a third of the tadpoles have metamorphosed, it is no longer necessary to feed the remaining tadpoles the nettle powder mixture. This is because the froglets break up the trout chow sufficiently for the tadpoles to filter the scraps. This simplifies the feeding regimen to: -nettle powder mixture only up to the emergence of the first froglets -nettle powder mixture and small size trout chow pellets until a third of the froglets have emerged -trout chow pellets exclusively beyond this. Given enough space, the froglets will reach sexual maturity after a year and full size after three years.

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4. Diseases, preventions and cures Hazel Sive

Frogs do get sick. In general vetinarians are not familiar with Xenopus and not much help in their treatment. Watch your frogs carefully, catching a problem early is obviously a good idea. Most diseases flare up after the animals have been stressed, and the most common stress is induction of ovulation and egg collection. It is therefore imperative to try to minimize stress and to observe animals carefully after these procedures. We (HLS) keep our females out of the colony for 24 hours after egg collection, in 20mM NaCl plus gentamycin (0.1ml/l (ten times lower dose than is given to embryos (see “Getting Embryos”))). Keeping them overnight in oxytetracycline may also not be a bad idea (gentamycin and tetracycline target different bacteria). NEVER LEAVE A FROG IN A BUCKET OF DIRTY WATER, CONTAINING EXCESS FOOD OR ESPECIALLY, ROTTING EGGS. AFTER A FEW DAYS IT WILL GET SEPTACEMIC AND DIE A HORRIBLE DEATH. It is also worth noting that if you have a disease problem in your colony, treating the disease with drugs may not remove the precipitating cause of the disease. Bacterial, nematode or fungal infections often arise from pathogens endemic to the frogs, that become a problem only when frogs are stressed. Stresses could include a sudden increase in crowding, pH change or other change in water quality, bad frog handling procedures. It is therefore important to try to figure out whether there is a root cause that can be changed. There is no point in trying to get eggs from sick frogs, the frogs will get sicker and the egg quality poorer. Frogs should have slimy, but not excessively slimy skin- it should not be visibly flaking (though they do shed some skin normally), and the pigmentation should not be patchy (normal pigmentation is mottled, but the mottling should cover the entire body). Animals should not be bloated (as opposed to fat- you can easily tell the difference when you pick them up) and should not be excessively thin. The skin should not be red, which may indicate subcutaneous hemorrhaging. The two most common diseases are bacterial septicemia (the culprit bacterium varies widely) and nematode infestation. Fungal infections may also occur. a. Nematode infection (capillariasis) Symptoms: sloughing of skin, patchy pigmentation, skin becomes greyish and thin, weight loss. Precipitated by stress. Do not usually see any redness. Skin scrapings will reveal nematode presence. Treatment: best given as soon as possible. Infected animals should be isolated, since the eggs and adults are shed into the water. The drug used is Ivermectin, obtainable from PRO-VET, Illinois. 1-800-435-6902. Catalog # 08338A, Brand name IVOMEC (for pigs). The drug can be administered by oral gavage, but this is difficult to do without severely stressing the frogs, and so we administer the drug by injection into the dorsal lymph sac, giving two similar doses, two weeks apart. The dose we use is 0.2μg per g of body weight, or about 8μg per male; 18μg per female. We dilute the drug in sterile water, the undiluted drug is very stable at room temperature. This is a very effective treatment and frogs will recover, but slowly, so you want to catch this early. Once frogs are very thin with fragile skin, recovery can take several months. Note: Many frogs have this infection latently. Frogs obtained from Xenopus I are all infected, and we treat them prophylactically when they arrive. This apparently has no ill effects. References: Cromeens, D.M. et al. 1987. Lab. Animal Sciences 37: 58-59. Stephens, L.C. et al. 1987. Lab. Animal Science 37: 341-344. 12


b. Red Leg (bacterial septicemia) Symptoms: cutaneous hemorrhages, especially on flexor surfaces of thighs and foot webs, dull discoloration of skin, subcutaneous edema, neurologis disorders (trembling, intially of limbs). Etiologic asgents can be any of a number of gram-negative bacteris, primarily Enterobacteria including Aeromona, Pseudomaonas, Citrobacter. Precipitated by stress. Treatment: Possible, if you catch it early. Increasing NaCl concentration to 100mM may help, although this in itself can be stressful. Injected or orally administered antibiotics often work well, 100μg/ml of oxytetracycline final concentration added to the water over a week can be effective and is the easiest way to administer antibiotics. Change the water every day. For oral administration: tetracycline- 1mg/ 5g body weight or chloramphenicol 1mg/ 30mg body weight, given by gavage (mouth) repeated for 5 days. A less stressful, though less reliable method for oral administration involves pre-soaking food pellets in oxy tet. Separate infected frogs as disease can spread rapidly. c. Fungal Infections Symptoms: Generally, fungus starts to grow in the system on decaying food and may subsequently attack frogs. If you see thread like (generally whitish) bodies growing on tanks and pipes, and if frogs start to get infections at site of injection, you may have a fungal problem. Confirm by microscopy that this really is fungus. Treatment: “Mar-oxy” solution, for treating fish, seems to work quite well. We (HLS) have found that the permanganate treatment prescribed in Methods in Cell Biology, vol. 36 leads frogs to a horrible death. This is a case where the fungus involved may be nonpathogenic, but can cause oppotunistic infections. During one period in our lab., we found that an increase in pH and therefore ammonia in the system had stripped the protective mucus from the skin allowing fungus to kill frogs in a matter of hours, after even tiny abrasions. Although Mar-oxy helped, lowering the pH (and the overall ammonia content) was the long term solution.

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B. Equipment for embryo experiments Hazel Sive

1. Variable temperature incubator This is an extremely useful piece of equipment. LabLine sells a HiLo temp. incubator for about $1,400. A cheaper alternative is a small refrigerator ($100- $200) in which the thermostat is hooked up to a Honeywell temperature regulator. Xenopus embryos will develop normally from about 14°C to about 24°C, and their rate of development varies with temperature. This is very useful when you need different stages simultaneously, for example embryos incubated at 20°C develop 75% as quickly as those incubated at 22°C; at 16°C, development slows to about 50% and at 14°C to about 33% the 22°C rate. At 14°C, an additional significant slow-down occurs just before gastrulation. See Appendix V. 2. Microscopes You will need a good quality dissecting microscope that can give at least 50x magnification, for both microinjection and microdissection. This does not need to be placed on a vibrationfree table, since embryos are very large (until neurula, about 1mm diameter). It is very helpful to have a large flat base on the 'scope, so that one’s hands can rest comfortably during dissection. You will probably want a camera mount on one 'scope (see "taking pictures" section). You want a “beam splitter” attachment so that you can see the embryos through the scope while you are focussing for the picture. Many companies (Zeiss, Nikon, Wild etc) make acceptable microscopes, but before purchasing one test it- you want something that won't tire your eyes if you do an eight hour dissecting stint. 3. Microinjector, micromanipulator and needle puller These are likely to be essential in your experiments. Microinjectors vary from very inexpensive hand-cranked injectors ($5- $100), through fairly cheap models such as the Drummond Nanoject Variable (about $400), through the Picospritzer (about $1000) and the Narashige or Medical Systems models (about $4000). Even more expensive models that can inject femtoliters are not necessary for Xenopus. The cheapest models are okay if your budget is tight but it can be difficult to precisely regulate the volume delivered. You will need a micromanipulator to go with your injector. This holds the glass capillary needles with which you will inject. Narashige makes some excellent, basic models, as does Brinkman. You need something that can hold your needles firmly, and that can be angled at about 45o for injections. A micromanipulator bolted to a microscope (Zeiss sells some of these) that cannot be angled is no good. Needle pullers also vary widely, we use a Sutter Instruments puller which is top of the line (around $7000) but cheaper types will do. Narashige makes a cheaper version that we will test in the course ($4000). You can use a home made puller, but your needle tip size will not be as uniform as with a commercial puller. Although pullers are expensive, they can be shared by several labs, since one can pull a lot of needles quickly. Needle bevelers are used by some people (to grind needle tips smooth), but are not usually necessary.

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4. Dissecting tools see Hamburger, V. A Manual of Experimental Embryology University of Chicago Press. Now out of print. Richard Harland

All tools should be kept clean and rinsed with 70% ethanol to keep them sterile. Forceps Dumont number 5 or 5A forceps (Fine Science Tools) Stainless steel are adequate for most uses and do not rust easily, however they cannot be sharpened to such a fine point as carbon-steel forceps. Use a fine stone for sharpening, and watch under the dissecting microscope. Only slight pressure should be used. The points must meet cleanly at the end. Slight bends can be made or corrected by pressing the tips against the microscope stage. Label forceps for different uses, use the good ones only for fine work and keep old, battered ones (which can be bent and sharpened back to a blunt end) for all non-critical puroses. Needle nose pliers, or coarse forceps can be used to make adjustments to the forceps tip angles, but any excessive bend will break the tips. Many people use forceps for dissections (e.g. of animal caps). For such dissections it is important to have two sharp pairs of forceps. The same forceps are used for removal of the vitelline membrane and for cutting the cap. Hair loops. It is useful to have a variety of sizes. Small loops are resilient and useful for fine dissections. Larger loops are good for sorting or pushing embryos around. Start with a long-stem pasteur pipet. Heat it below the shoulders with a small bunsen flame and pull to about 15cm. Make a second, and finer pull. Break the end after scoring with a diamond pencil. The orifice should be big enough to thread the hair easily (even if you have to use the dissecting microscope and forceps to hold the hair) Careful flame polishing will help prevent the hair from getting cut on the glass, but is not essential. Start with a long hair, thread it through the end of the pipet until the other end emerges from the blunt end of the pipet. Now thread the free end into the tip far enough that it will not pop out. Now by pulling on the first end, the loop is pulled progressively tighter. Pull

Figure: Hair loop Now scrape a little beeswax into the end of another pasteur. Warm it over a flame until molten, then apply the tip to the small orifice of the hair loop. Wax will be drawn by capillarity into the tip. Final adjustments of the loop may be made and the wax remelted with a warm pipet tip. The set wax will hold the loop in place. You can also dip the hair loop into molten beeswax (melted on a hotplate). After removing the loop from the wax, blot excess wax from the loop by touching the loop to a hot kimwipe or tissue fragment (put the tissue on a warm hotplate or piece of metal that you’ve warmed in the bunsen. Eyebrow knives.

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These are made in the same way as hair loops, only more easily. Hairs vary enormously in their quality as cutting tools. Try different people as sources. You want a strong hair but not a brittle one. Simply insert the blunt end of the hair into the drawn out pipet, far enough that it leaves the desired amount of curvature/resilience. Set in place with beeswax. Eyebrow knives are not used to saw down into a tissue as is the case for a conventional knife, but to flick it from below (as in the case of an animal caps (see section G.1), or to flick down from above when the tissue has been removed from the embryo and needs trimming. In this latter case, trimming is done against the solid base of the dissecting dish (section G and see accompanying videotapes). Fine tungsten needles Tungsten needles and glass knives (below) are useful for older stages (neurula and beyond), when the embryonic tissues become tougher and resistant to eyebrow knives. Short pieces of fine tungsten wire (California fine wire company, Grover City CA 93433 size 0.001) can be mounted in pipet tips. The wire is cut, put in the drawn out tip, then the glass is fused carefully around the wire by heating in the edge of a flame. Overcooking makes the wire brittle. Tungsten needles with 1uM tips can be purchased from Fine Science Tools, as an alternative to eyebrow knives ($60 for 10, plus $30 for two needle holders). They are a poor substitute however. Electrosharpened tungsten needles can be very good. Glass needles Also good for dissecting older embryos, glass needles are very simply made from microinjection needles or hand pulled capillaries. Break off the tip so that you have something sturdy enough for dissection. The tips will be open. Mount needles in needle holders used for tungsten needles (available from Fine Science Tools or Carolina Biological). Throw needles away at the end of the day, since the tips are open it is difficult to sterilize them.

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5. Cameras and taking pictures

R. Harland

As I have learned to my embarrassment, if you supply a journal with a less than ideal print, it will look dreadful in the journal. It is worth taking pains to get good pictures. While it is certainly more convenient to have a high quality microscope camera, especially for the compound microscope, one can do quite well with a good quality single lens reflex. This can also be used for making slides, when fitted with a macro lens. We use a single lens reflex on the dissecting microscopes. It is a good idea to replace the focussing screen (usually fitted with a prism as a focusing aid, but very greedy for light and hard to use) with a plain focusing screen with lines as a focussing aid. For optical reasons beyond our understanding, it is extremely difficult to get low power pictures in focus. Therefore it is essential to use a telescope to magnify the image to be focused. The lines on the focusing screen and the image must be simultaneously in sharp focus. Focus on the screen first by adjusting the telescope (e.g. Zeiss 3x12B #52 20 12), then focus the specimen using the microscope focus. Unless you are old, your eyes quickly accomodate at low magnification, so it is important to keep rechecking that the lines on the focussing screen, or focussing reticle are in sharp focus. It is extremely easy to destroy a day's photography with a few simple errors. The checklist below is designed to avoid some of the errors we have made. Effects that are not very noticable by eye (such as uneven illumination) make a complete mess of a photograph. Film For non-fluorescent microscopy, where light is relatively plentiful. Ektachrome 64 slide film is a good choice since it gives fine grain and can usually be processed quickly at a local photography store. Ektar 25 is a high resolution print film which if exposed approximately correctly, can be printed perfectly, avoiding the need to bracket exposures. The local 1-hour print shop usually does a reasonable job of printing, but they need to be willing to adjust the computer exposure for the roll. If brightfield pictures of embryos ask to print 1 stop lighter, if darkfield ask for 1 stop darker. If you establish a good relationship with the printers, and get them to achieve the right background they can make great prints cheaply. However, in the end, its worth getting custom prints if you need to match exposures etc.

•Load film, Make sure its securely wound onto the spool. Better to waste an exposure than have the film loosely engaged and have it never wind on. With many cameras, you should see the film spool winding as the film is wound on (after the slack has been taken up). •Set

ASA

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Compound microscope •Blow dust away. •Check that no filter is engaged. What filters do you want to use? Daylight (light blue) in case of contrast with yellow and red, no filter or neutral density filter in case of contrast with blue. Use green filter if brown HRP signal and black and white film (don't use T-max, use tech pan) •Check condenser setting Field iris should be at edge of field and in focus. •Check condenser iris, too far closed can give bizarre effects •Check evenness of illumination. If uneven, check centering of field iris, check that all filters (e.g. Nomarski filters) are properly engaged and not stuck halfway. •Always use the telescope for low power photography. Specimen and crosshairrs must be in focus simultaneously. •Exposures. Is the field going to be mostly light or mostly dark? For brightfield pictures of embryos set auto exposure at +1. If darkfield, set at -1or -2 (depending how much bright embryo occupies the field). With films such as Ektar 25 there is no need to bracket exposures, any slight misexposure can be fixed during printing. For slide film its more important to bracket. SLR camera on dissecting microscope •Make sure that the spool turns when camera is winding. •Darkfield pictures. Dust shows up very badly in dark background pictures. Set a 90mm petri dish on top of stage instead of the normal black baseplate. Use a new clean petri dish for specimens.. Wash the embryos well to remove debris. Fill the space between the top and bottom dish with water or 70% ethanol. The background (under the microscope) will now be very distant and way out of focus. Use at least two light sources to get even illumination. Alternately, place the petri dish on a colored backgroundelectrical tape works well, most cardboards become grainy with magnification and are therefore no good as background.

Water Baseplate removed Figure: Darkfield photography of embryos •Be particularly careful to check for even lighting, shadows, reflections which show up more prominently on the film than in your eye.

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Making slides

•Set film speed Use telescope, focus on eyepiece cross hairs •set F stop 22 (for max depth of field) auto exposure •Exposures, Check that the camera is set on Automatic exposure for pictures with pale backgrounds, use up to +2 stops. for pictures with black backgrounds use up to -2 stops •For Ektachrome 64 set at ASA 25 for line drawings with white backgrounds, then bracket around 0. •Polaroid black and white. Polagraph 400 ASA high contrast use +12/3 stop •Turn off the overhead lights to avoid reflections look carefully for uneven illumination or glare C. Getting embryos This is done by inducing ovulation, collecting unfertlized eggs and fertilizing them in vitro. Alternatively, narural matings can be set up (see below). A really good female can lay many hundreds of eggs. However, since the actual number laid and ability to be fertilized is very variable, it is best to induce ovulation in at least two females if you need even a few eggs. Females can be induced to lay repeatedly for several years- however a rest period of two to three months between ovulations is required. See The testes from one male contain sufficient sperm to fertilize a few thousand eggs. 1. Handling Xenopus adults Pick up the frog by placing your hand over her back, with your forefinger between her legs and the rest of your hand wrapped around her middle. Use your other hand to cover her eyes which will calm her as well as physically prevent her from flying out of your hands. If a frog does land on the floor, pick it up immediately- Xenopus is very good at diappearing into non-existent spaces, and since the Genus is entirely aquatic, will dehydrate very quickly and die (two hours after leaving water, a frog will generally be in extremely bad shape). Frog skin is very thin and easily abraded- so pick up your frog and rinse her under the (distilled water) faucet right away (frogs generally enjoy this very much and will lie quietly in your hands). A note about gloves. Frogs can be picked up in clean, but soap-free hands (wash your hands when you’re done) or while you are wearing latex gloves. However, DO NOT wear POWDERED gloves (covered in either powder or cornstarch) or the TEXTURED latex gloves designed for better grip. Both types will abrade the frog’s skin. 2. Inducing ovulation This is done by injection of human chorionic gonadotropin (hCG) into the dorsal lymph sac of a female frog. In general frogs will become more kinetic after gonadotropin injection. Place the frog belly down on a clean surface (frog skin is very sensitive to abrasion)- we use benchcote turned upside down to expose the plastic side, and wrap the frog firmly in wet paper towels, completely covering her eyes, but leaving her hind legs and lower abdomen exposed. The frog should lie quietly during injection, but doesn't always do so- to prevent her kicking off the table, place your index and middle fingers over her thighs, so that you can press 19


down on these if she starts to move. In general, the frog will start to breathe very quickly before she attempts to move away so you have several seconds warning. For newly purchased females and those that have not been induced to ovulate for more than a year, we prime frogs with about 50 units of hCG at least five days before we induce ovulation. Do not use more hCG since higher doses sometimes induce partial ovulation, which you don't want- females will subsequently lay poor quality and few eggs when you try to induce them. The five day wait is necessary, since otherwise when you induce with higher hCG the night before you want eggs, egg laying will commence very quickly (likely in the middle of the night when you are not there!). The priming effect wears off after about a month. Alternately, treat with 50 units pregnant mare serum gonadotropin (calbiochem) which contains both hCG and other factors. PMSG can be used as little as one day before inducing ovulation, where its main effect is to make the time of layingmore predictable. For induction of ovulation, inject 500 - 800 units hCG, depending on frog size. hCG can be obtained from many suppliers, including Sigma and Pro-Vet, Illinois and costs between $15 and $30 for 10, 000 units. Use a FINE needle- 26 gauge needle attached to a 1ml syringe. Place the needle posteriorly about 20% of the distance between the hind and fore limbs, just outside the lateral line sense organs (see diagram). Slip the needle under the skin- push down intially to penetrate the skin and then when you are under the skin (you will be able to tell since it is quite loose and not attached to the underlying tissue) slip the needle laterally towards the dorsal midline across the lateral line "stitch" marks. The dorsal lymph sac lies on the back of the animal between the stitch marks. As you penetrate the wall of the sac you will feel some resistance and then relief thereof. Inject the liquid, wait five seconds and then slowly pull out the needle. Most important is that liquid injected does not run out of frog when the needle is removed. No bleeding should occur, if it does, the needle went in too deep, or the skin was torn by sideways shearing of the needle. It does not seem necessary (or possibly even desirable) to swab skin with alcohol before injection- frog skin naturally contains antibacterials. After induction of ovulation, egg laying begins about 9 - 10 hours later if frogs are kept at room temperature (about 23°C) or about 14 hours later if frogs are kept at 15°C.

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dorsal lymph sac

ne

ed

le

Needle placement for hormone injection into Xenopus

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3. Isolating the testes Males should be killed or deeply anesthetized in benzocaine (0.05% in water, a 10% stock can be kept in ethanol at 4o)- at least 30 min. before dissection. Several hours is okay. Make a fresh dilution of benzocaine in water every week and keep it at 4oC. When laid on its back, the frog should also not respond to ethanol in the nose. See section concerning Morphological Features for testes position. Use a tissue or sharp forceps to pick up the loose skin on the belly and make a cut with scissors. The skin can now be lifted with forceps. The testes are attached to the fat bodies, and the easiest way to remove them is to make a slit on either side of the dorsal midline, being careful not to cut across the midline where the large abdominal blood vessel is situated. Push aside the liver and pull out the fat body (yellowish, with many fingerlike lobes). Alternately, expose most of the belly, then similarly remove the muscle layer to expose the viscera. The testes each lie at the base of the fat bodies- they are whitish and covered with capillaries. Using scissors or forceps, free each testis from the fat body and surrounding connective tissue. If you are unsure whether you have the testis, crush a little on a microscope slide and view with phase contrast. The sperm are easy to see with a fine helical shape. Place in 80% serum (calf or chick- any type is probably fine) 20% 1xMBS + high salt; they also keep adequately in 1x MBS + high salt or 1x MMR. We add pen/strep or gentamycin (1μl per ml) and store the testes on ice or at 4°C. They are good for a couple of days, when kept intact (wholly or partially) after which sperm viability drops. 4. Collecting eggs Since septicemia is most often induced after ovulation, either by stress and/or by infection induced during handling, we keep females in 20mM NaCl + gentamycin (0.1ml/l- that is one tenth the amount embryos are incubated in, since gentamycin is nephrotoxic) during egg laying and for 12 hours after. In addition, we are fastidious about keeping the females in clean water after ovulation. Water in buckets should be changed every day- we guarantee that if you forget about a frog and leave her in dirty water over a weekend for example, she will become horribly septacemic and die. Manual collection Females should be laying in their containers (we use 4l plastic beakers filled to 3l mark, and kept covered with air holes) before attempting egg collection. The idea is to mimic the actions of a male frog and to encourage the female to lay- not to physically squeeze eggs out of the animal. The action is a gentle touch that has to be learned, the eggs collect in a sac near the cloaca, and gentle lateral and simultaneous vertical pressure should expel them. If performed gently, the females remain relaxed during the collection, except when vigorously pushing the eggs out. After induction of ovulation, the cloaca becomes red and swollen and is probably rather sensitive, so avoid touching it when picking up the frog. We hold the frog by hooking one hand around the top of each thigh, massaging her belly with one thumb, gently but firmly. After a minute (or sooner) she will begin to lay- and should be held over a sterile/clean petri dish (80mm) containing 1xMBS + high salt (see Appendix II). If she does not lay, you may need to massage her with more pressure. Eggs may be laid without movement from the frog, or she may push them out in a couple of vigorous bursts, at which time hold on to her tight (holding her against your shirt or clean lab coat really helps), let her settle down and then massage again- the eggs should now be easily laid. Do not continue collection for more than two to three minutes MAXIMUM. Collections can be made every hour (or less, if she seems to be laying in her container) for the first two or three hours after laying commences and then less frequently as the day progresses.

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Another method involves placing the frog belly down on clean aluminun foil or Saran wrap and keeping her lying down with one hand, including covering her eyes. The fingers of the other hand stroke firmly down either side of the animal, also mimicing amplexus movements, more or less. Eggs are collected dry and then washed off the foil with 1x MBS + high salt into a petri dish. Do not rub the back of a frog in attempting to collect eggs. You may get some, but there is very little subcutaneous fat on the back and she is much more likely to bruise and get sick. Collection into high salt solution Some researchers maintain the females in 1x MBS + high salt throughout the day and collect the eggs after they have been laid in the water. This is likely less stressful for the females than manual collection, though the number of eggs obtained may be lower. We (Sive lab) have found that this can work quite well, as long as the eggs are removed from the container soon after laying, and the MBS/high salt is maintained at pH7 (higher or lower pH irritates the frogs). If you try this method, be sure to check that the frogs are not moving around a lot in the high salt (relative to how they would move in low salt water, which should not be much, a happy frog is a placid one). A lot of movement indicates that the solution is irritating the frogs’ skins. If this occurs, move the frogs to low salt or check the pH. Females usually lay for about eight hours. Keep all layings in separate dishes and note the laying time thereon. Use the earliest layings first as competence for fertilization does decrease with age. Eggs should remain competent for fertilization up to 12 hours after laying when kept at 15°C, though viability does drop after 8 or so hours (seems to be frog dependent). During the time of laying, keep females separately so you can keep track of how each is doing. 5. In vitro fertilization Highest fertilization rates will be obtained if eggs are fertilized as soon as possible after collection. You can collect into 1 x MBS (or equivalent buffer) if you are going to fertilize right away (within a few minutes). collected into 1xMBS + high salt can be kept at 15oC for some time before fertilization, with the time they remain competent for fetilization largely dependent on the batch of eggs. A wonderful batch of eggs can wait 12 hours, however, most batches will not remain competent for more than a couple of hours. In general, you should plan your experiment around the eggs and fertilize them as soon as possible after laying. When ready to fertilize, remove all buffer from the eggs. Tease a piece of testes apart with forceps and rub it over the waiting eggs. It is important to touch every egg with the testis. At first it may help to do this under the dissecting microscope, to see that the follicles containing the sperm are being crushed. Alternately, the testis can be crushed and a sperm “slurry” mixed with the eggs. Make sure the eggs are dispersed in the dish. A pair of forceps can be used to break up clumps of eggs and spread them out . The eggs are very tough and will not break easily at this point. Flood with dilute buffer, 1/3 to 1/10 XMMR or 1/10x MBS. The first sign of fertilization is a contraction of the pigment so that the unpigmented territory is now maybe 2/3 to3/4 of the egg. The egg also gets firmer after fertilization, such that if you squeeze it gently with forceps it will be elastic and resist deformation if fertilization has occurred and will be much softer and deformable if fertilzation has been unsuccessful. After about 30 minutes the fertilized eggs will rotate within the vitelline membrane so that the animal hemisphere faces up. Always perform test fertilizations on a few eggs soon after laying, so that you can assess the quality of the eggs. If the first fertilization is poor, perform 23


another, on eggs laid at a different time to those used in the first test since the quality of eggs may vary during the day. Fertilization efficiencies range from 100% on down, with "good" frogs typically giving greater than 80% fertilization. 6. Natural mating Males and females can be allowed to mate naturally and the fertilized eggs collected by pipette. Alternately a plastic mesh bottom can be placed in the bucket through which the eggs can fall. Females should be injected with hCG as for the in vitro fertilization method, and males may need 50U a few days before mating, though if your frogs are healthy they should mate spontaneously (go into”amplexus” ) without having to inject the male (it is unlikely that the female will lay without stimulation). This is a convenient way to get many different stages of embryos, for in situ hybridization for example, with minimal stress to the frogs. 7. Keeping track of frogs A frog can repeatedly be induced to lay eggs, but must rest between ovulations. A resting time of at least two months between inducing ovulation is necessary. One therefore needs to record when a frog was last used. This can also be very useful to keep track of whether a particular frog laid good or bad eggs and is worth ovulating again. Most methods to mark individual frogs seem cruel (acid etched numbers, for example) or impermanent (for example, clipping toenails in specific patterns). The Slack lab. implants microchips subcutaneously (designed for mice) that can be detected by a hand-held computer. These chips are relatively expensive, but can be extracted from a frog and reused. After ovulation, the performance of a frog can be recorded and compared to performance in subsequent ovulations. A frog that repeatedly performs poorly (three times) is unlikely to improve. An alternative strategy is to keep batches of previously ovulated frogs together in “good”, “medium” or “poor” tanks designated according to their previous performance. Within these categories frogs are grouped according to when they were last ovulated. Subsequent to the next ovulation, frogs are re-rated. This is not as precise as the previous method, and one needs to get rid of frogs after two ovulations are poor, rather than three. However, it is very simple.and can be quite successful.

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D. Preparing embryos for manipulation The most important techniques here are removing the thick jelly coat, which is essential for both microinjection and micromanipulation, and removing the vitelline envelope, which is essential for micromanipulation. 1 Dejellying embryos Embryos are surrounded by a series of thick, protective jelly membranes that need to be removed before doing just about anything. Although this can easily be done by treating with cysteine, the embryos are very susceptible to damage during this process, and it is essential not to overdejelly. For microinjection, the jelly coat can be loosened, but not necessarily removed; while for dissection, the coat must be completely removed. Dejelly the eggs by removing buffer (MBS etc) and swirling gently in 2% cysteine in MMR or water, pH 8. For dilution in water, add 2g cysteine HCl and 3ml 5M NaOH per 100 ml water to get to pH 8. Always check the pH with pH paper before use. Dejellying takes about 2 - 4 minutes, but is frog dependent , and must be titrated for each frog. When the eggs just pack closely together and you can see fragments of jelly floating in the buffer, decant the cysteine and rinse the fertilized eggs 10 times in 1/3x MMR or 1/10x MBS, over a period of about 10 minutes. It is essential to rinse thoroughly. If it is important to remove vitelline membranes, the membrane will be easier to remove if the dejellying is done after the first signs of fertilization at about 5 minutes. The vitelline membrane firms up over about the first half hour after fertilization. If therefore, you aim to keep the embryos for a long time, its best not to dejelly until 30 minutes post-fertilization. Opinions differ, but we (Sive lab) generally dejelly close to the time embryos are needed, since post-dejellying mortality increases with time. However, embryos are particularly sensitive to dejellying during gastrula and neurula stages. After dejellying, keep embryos at low density (100 per 80 mM plate), and remove dead embryos promptly. 2. Removing the vitelline membrane This is an essential part of all dissection experiments, that becomes easier and easier with practice. For taking vitelline membranes off, it is useful to have a pair of forceps that are fairly sharp, even with the points slightly bent towards each other to give a pincer appearance. These are used to grab the vitelline membrane. For the other pair the tips can be slightly blunt, so that as the embryo is picked up, the blunt forceps can be used to take a second hold on the membrane. The bluntness helps to avoid poking through the membrane. For removal of vitelline membranes, different batches of embryos vary enormously. Floppy embryos, with a large space around the embryo are easiest to learn on. If the membrane is excessively snug, it can be loosened by a brief digestion with pronase or proteinase K (5μg/ml). As soon as the membranes are slightly elevated from the embryo, or if any of the membranes spontaneously breaks, wash the embryos well. The membrane will continue to elevate for a while. For all micromanipulation, however, embryos should be manually devitellinized since it is difficult not to damage the embryo with enzymatic treatment.

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Vitelline membrane

Figure: Removal of vitelline membrane Where embryo dissection will follow devitellinizing, start removing the membrane from a region of the embryos in which you are not interested, to minimize the effects of damage. For example, for animal cap dissections, remove the membrane from the marginal zone. Of whole intact embryos will be required, remove the membrane from the animal cap, since even if this is damaged it will heal very quickly. E. Embryo perturbations Although one cannot perform classical genetic experiments on Xenopus laevis, several reagents have been very useful in perturbing Xenopus development and acting as pseudogenetic tools. These include ultraviolet light, LICl and retinoic acid. Superficially (only!!), these treatments result at hatching stages in embryonic defects that fall onto a continuum of phenotypic defects- giving a "dorsoanterior index" (DAI) as defined by Kao and Elinson. 1988. Dev Biol 127:64-77. The “Index of Axial Deficiency” or IAD was a metric coined by Gerhat and colleagues (see Scharf and Gerhart (1983) Dev. Biol. 99:7587) in which normal is 0 and completely ventralized is 5. There was no scale for dorsalized embryos. The DAI scale is the one in use now, but you may see the IAD scale in older publications.

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1. Ultraviolet light Scharf and Gerhart (1983) Dev. Biol. 99:75-87. UV light will "ventralize" embryos when irradiated during the first 60% of the first cell cyclethat is before the time the cortico-cytoplasmic rotation is occurring. Since first cleavage occurs at 90 minutes after fertilization at room temperature (25°C), this leaves about 55 minutes in which irradiation will have an effect. Dejelly embryos as soon as they show pigment contraction in the animal hemisphere, or have turned. Place, in 0.1x MBS or equivalent, on a petri dish from which you have removed a strip of plastic (about 1 cm x 3 cm), and covered the strip with Saran wrap (NOT Parafilm). Place on a UVGL-25 lamp and irradiate batches of embryos for 30, 60 and 90 sec. It is necessary to perform a UV titration since UV sensitivity varies from frog to frog. Success of UV treatment can easily be scored at early gastrula, when formation of the blastopore should be delayed about an hour compared to controls. Irradiation can also be performed on an inverted Stratalinker. Dishes Quartz dishes are simple to make, convenient and long-lasting. Optical grade (distortionfree) quartz is not needed, and good quality quartz can be obtained for ~$10 for a square inch piece. If in doubt, check the quartz by putting it in the light path of the spectrophotometer. It should show good transmission down past 260nm. A window is made in 35mm dishes, about 1 cm wide, and slightly less than 1" long (for 1" square quartz). The corners of a 1" piece may need to be filed down to attach the square to a falcon 1008 dish. The quartz is attached with cement, or methylene chloride, or plexiglass glue to the bottom of the dish. The quartz should be cleaned of any cement residue with a Q-tip. The final dish has a rectangular well in the plastic, with a quartz floor. Alternately, the slit can be covered with Saran wrap that will stick to the dish by itself. The Saran can be replaced as needed. Embryos

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For effective UV treatment, the embryos must be treated prior to the initiation of rotation that occurs half way through the first cell cycle. The embryos are dejellied as soon as possible after fertilization. For pigmented embryos, the cortical contraction of animal pole pigment should be clear. The embryos do not need to have completed rotation. Following dejellying, the embryos are rinsed well, and good embryos transferred to a fresh dish. (Early dejellying increases the proportion of vitelline envelopes that rupture, and any embryos with ruptured membranes should not be transferred to the quartz dish). Transfer the embryos to the quartz-bottomed dish, and place over a UVLG lamp or the Stratalinker. Don't overcrowd the embryos. Dirty quartz or oozing protein will block the light, so keep things clean. A support for the embryos is made from corrugated cardboard. It should occupy less than 1/4 of the surface of the stratalinker. When the stratalinker is turned upside down, the cardboard is placed in the near left corner (away from the UV sensor). The cardboard has windows 10cm long by 1 cm wide that sit directly over the light tubes. The support for the embryos sits directly on the tubes. The optimal dose of UV will need to be determined empirically, but an energy reading of 1000 mJoules is a good starting point. This should be about a minute's irradiation. If the time needed for irradiation suddenly gets longer, the tubes may have burned out and should be replaced. Any embryos that have tipped will be rescued by the gravity-induced rotation; thus after irradiation embryos should be either left in place until after completion of first cleavage, or transferred gently to an agarose coated dish. Any embryos that stay tiipped after transfer should be removed. Anecdotal evidence suggests that the UV treatment is more effective if the embryos are left to develop at less than 20°C. Scoring the DAI (Dorso-anterior index). Bad embryos can be identified at the onset of gastruation. Any embryo that forms a lip at the same time as controls is likely to develop an axis. Good quality UV embryos should develop a lip circumferentially and at the time of ventral lip formation. The effectiveness of the uv can also be estimated easily during neurulation. Good treatment should result in no neural folds. For accurate scoring one needs to be able to identify the otocyst (the white refractile body in the ear of the animal). This becomes obvious as tadpoles mature, but the embryos have to be kept in good condition to develop this far. With experience, embryos can be scored earlier (in the late 20s to 30s) Hindbrain development is associated with a small nub of tissue at the anterior end. 2. LiCl Various regimens are available. The classic treatment is a brief immersion in 0.3M LiCl in 1/3 MMR at the 32-64 cell stage. Alternatively the embryos can be incubated in 0.1M LiCl from the 32 cell stage for 1 hour. Different batches vary in their tolerance to LiCl. Some batches survive after 1hour 15minutes, and are more extreme. Problems are usually manifest during gastrulation, when the yolk plug fails to be involuted. Incubation of the embryos in 5% ficoll can help gastrulation by putting additional pressure on the yolk plug. Kao, K. R. and Elinson, R. P. (1988) The entire mesodermal mantle behaves as Spemann's organizer in dorsoanterior enhanced Xenopus laevis embryos. Devl. Biol. 127, 64-77. Kao, K. R., Masui, Y. and Elinson, R. P. (1986) Lithium induced respecification of pattern in Xenopus laevis embryos. Nature 322, 371-373.

3. Retinoic acid Durston et al. 1989. Nature 340:140-144. Sive et al. 1990. Genes Dev. 4:932-942.

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Embryos are most retinoic acid-sensitive during late blastula and gastrula stages. Make a 0.1M retinoic acid stock in DMSO (all trans RA, Sigma). Store small aliquots at -80°C, avoid exposure to light. RA is very insoluble in water, and upon dilution will come out of solution- it is hard therefore to know what the real RA concentration in your final media is. A dilution of your stock to 1mM in 0.1xMBS or equivalent has powerful teratogenicity. Dejelly embryos. Immerse in a 1mM retinoic acid solution, for several hours, depending on the severity of defects you desire. Gross morpholocial defects can be observed after 5 minute treatment with 1mM, and subsequent incubation in buffer without RA (DAI 4). The most severe phenotypes resemble DAI 1 embryos. 4. Inducing exogastrulation Xenopus is reluctant to make good exogastrulae. To make exogastrulae take embryos and expose to high salt. Various concentrations can be tried, from 1x MMR to 1.3x MMR. They should be exposed from the early blastula stage. It is essential to take off the vitelline membrane (on an agarose bed) so that the yolk plug is free to push out. Frequently the involuting mesoderm still partially involutes, leading to a clearly recognizable miniature neural plate. Good exogastrulae will have a wrinkled-looking animal cap. During gastrulation, the embryos should be observed, since some transiently involute part of the mesoderm. These should be discarded. Animals with obvious neural folds or cement glands occur after mesoderm involution and cannot be considered complete exogastrulae. Because of various sloppy experiments exogastrulae are not considered a particularly clean or clear preparation, so if used, it is important to make them carefully. 5. How to mark the future dorsal side by tipping and staining Tabitha Doniach

The direction of the cortical-cytoplasmic (cc) rotation can be influenced by gravity. Thus, when embryos are tipped 90° with respect to their animal vegetal axis, before the cc rotation (3 days) probe so plan accordingly. Add the following ingredients at room temperature to prevent the DNA from precipitating. 2 μl 5x SP6 buffer (recipe below-also appropriate for T7 and T3 pol.) 1 μl 0.1 M DTT 1.5 μl ATP, GTP, UTP mix, 3.3 mM each. 4 μl 32P CTP 10 mCi/ml 400 Ci/mmol 1 μl 1 mg/ml DNA. water to 10 μl. 0.5 μl enzyme (5-10 units) In cases where a low specific activity probe is wanted (eg for EF1α), substitute the 4μl of 32P CTP with 4μl 200μM cold CTP and 0.5μl of 32P CTP. Incubate one hour at 37°C for SP6, T7 or T3 polymerase. To remove DNA add 0.5 μl RNAse free DNAse (BRL) 1 mg/ml and incubate for 10 min at 37°C. Add carrier RNA to 10 μg/ml. Precipitate with 1/4 volume 10M NH4OAc and 2 volumes ethanol in a dry ice/ethanol bath. Resuspend the RNA in 10 μl formamide + dyes. Heat to 65°C for 5-10 minutes or 80° for 3 minutes Load on preparative gel ~1cm width of well after xylene cyanol is 1/2 to 3/4 way down, dismantle gel, cover with saran wrap and put pieces of tape at edges. Put spots of fresh hot ink (month old 32P nucleotide + india ink) on the tape. After a 1-2 minute exposure cut windows in the film where the bands are. Do this on paper towels or cardboard, not on the gel, where the acrylamide will get smashed. Line up the film spots with the ink spots and cut the gel with a fresh razor blade. Lift the rectangle of gel into a tube of elution buffer (with or without saran wrap). If you are worried, reexpose to ensure that you hit the band, or Cerenkov count the elution tube. Elute at 37°C for 1-2 hours Extract with phenol (no need to remove the gel/saran) Transfer and precipitate the supernatant with 2 volumes of ethanol. Resuspend in 100μl DEP water. Count 1μl. Should be millions of cpm. Dilute probe so that 1μl contains 104 cpm (100μl/106 cpm). Use this immediately.

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If you must delay, add ethanol to 30%. This substantially stabilizes 32P labelled probes, markers etc. Store at -20°C RNA Hybridization and Digestion The RNA should previously have been extracted etc. RNA can be stored at -80° in 1mM EDTA, or as an ethanol precipitate. If the amount of sample RNA is small, add 10-20ug of torula RNA as carrier. Coprecipitate the target RNA with the probe. For rare RNAs 104 cpm is sufficient, and using less probe can reduce the background. For more abundant RNA use 105 cpm. Alternatively, the probe can be synthesized at low specific activity (see above) and 104 cpm used. To check that you have probe excess, do two samples of a positive control RNA with different amounts of probe. If probe is in excess the signal should be identical for each. Resuspend the RNA in 20 μl Formamide + hybridization salts 80% formamide + 20% 5X Salts (see below). Heat the mix to 65-75°C for 5 minutes then hybridize at 45°C overnight. Ideally, use a small waterbath, preheated to 75°. After putting the tubes in turn the bath down to 45° and go home. Next morning, add 300ul of the RNase Digestion Buffer containing 10 units/ml RNase T1 (Calbiochem 556785; its expensive, so it must be good), mix well, and incubate for 12hours at room temperature. Omitting RNAseA improves the stability of hybrids, with no serious background problem. Temperature is a factor in the sensitivity and lack of background. To optimize the assay in each case, try digesting at a range of temperature from 4°C to 37°C (I feel I have to suggest this, but room temp is usually best) Stop the digestion by adding 20ul of 10% SDS and 5ul of 10mg/ml proteinase K and incubating for 15 min. at 37°C. Add 20ug torula RNA as carrier. Make 2.5M in NH4 acetate and add 1 volume isopropanol. This moderately stringent precipitation avoids having to phenol extract. Wash and dry the pellet and resuspend in 4ul formamide and dyes. In the undigested probe sample, dilute the probe 1:100 or 1:200 and mix 1ul with 20ug torula RNA and formamide and dyes. Before loading, denature the samples by heating at 90°C for 3 min. or 65°C for 10 min. Analyze the samples on a 5% sequencing gel.

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PCR-based Protocols QUANTITATIVE RT-PCR

Charles Sagerstrom, Hazel Sive

RT-PCR analysis surpasses Northern blots and RNAse protection assays in its ability to detect low levels of gene expression. This technique has therefore found much use, particularly in applications where sample sizes are limiting. However, to fully compete with these other techniques RT-PCR must also be made quantitative. Researchers have primarily taken two approaches to this problem. First, relative quantitation can be achieved by amplifying an endogenously expressed gene, known to be expressed at identical levels in all samples, along with the gene of interest. As long as amplification of the two genes is linear, normalization to the internal control allows levels of the gene of interest to be compared directly between separate samples. With this technique it is therefore possible to conclude that, for instance, gene Y is expressed at 10X higher levels in sample A than in sample B. It is important to note that this protocol says nothing about the absolute levels of transcripts in either sample and it is therefore impossible to compare the expression levels of two different genes (i.e; Is gene Z expressed at higher levels than gene Y in sample A?). Second, absolute quantitation (aka competitive PCR) requires the inclusion of a known amount of control RNA or DNA in the PCR reaction. The control template should be amplified with the same primers as the gene of interest and should ideally have the same sequence, but still be distinguishable on a gel. (This can be achieved for instance by inserting a small piece of DNA or by including a restriction site). By amplifying serial dilutions of the control template with a constant amount of sample it is possible to estimate the absolute concentration of target template in the sample. If one does not need absolute information, relative quantitative PCR requires less work to set up. It does not require the construction of competitive templates, nor does it require that a titration curve be performed on each sample. We have found relative quantitation by RTPCR to be at least 100 times more sensitive than Northern blots. In addition, the RT-PCR gives results in a day or two whereas Northerns often require lengthy exposures. cDNA synthesis: The cDNA synthesis is performed on total RNA prepared by the method of Chomzcynski and Sacchi (Anal. Biochem. 162:156-159, 1987). (We routinely use 100-150 ng of total RNA for the RT reaction, or the equivalent of one small animal cap). Identical amounts of total RNA from each sample (resuspended in H2O) is added to parallell cDNA reactions and all reagents are aliquoted from a freshly prepared ‘master mix’. Using this protocol there appears to be minimal sample to sample variation in the effiency of the RT reaction. In addition, the internal control included in the PCR amplification should correct for such differences, should they arise.

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PCR amplification: The PCR reaction (see attached protocol) first has to be optimized with respect to the usual parameters, e.g. dNTP and MgCl2 concentration, for each primer pair. We then determine conditions for the coamplification of several primer pairs in one reaction tube. This may require varying the primer concentrations relative to one another and may also necessitate altering the MgCl2 and dNTP levels. This extra amount of work is worth it since combining several primer pairs in one reaction makes it possible to get more information from a small sample. At this stage we also establish conditions for coamplification of the internal control. The internal control is a gene that is known to be expressed at the same level in all samples and it is required for normalization between the samples. Next, since comparisons between samples have to be done whithin the linear range of the PCR amplification and as the linear range is dependent on template input (more template, i.e. higher expression of the gene of interest, gives linear amplification at lower cycle numbers and vice versa) it is necessary to find a range where all samples, regardless of expression level, are amplified linearly. We therefore test samples that we expect to have the highest and lowest signals and select one cycle number where both are in the linear range. We then perform subsequent experiments at that number of cycles. Every time a new treatment or a new sample type is to be analyzed we first verify that the amplification is linear at the cycle number to be used. In addition, we add the same volume of template from the cDNA synthesis to each PCR reaction to equalize any inhibitory effects of the RT reaction mixture and we set up the PCR reactions from a freshly made ‘master reaction’ mix. There is a practical limit to how little template can be detected, since the PCR reaction, when performed as in this protocol, appears to plateau after about 30 cycles regardless of the amount of template present, presumably as a result of exhaustion/inactivation of reaction components. We add 32P-dCTP as a tracer and use a sensitive detection system (a FUJI Phosphoimager) thereby detecting signals from less input template. We routinely use 2-4 ul of the cDNA synthesis (=cDNA from ~10-30 ng of RNA) for the PCR. This amount of template gives a linear range somewhere between 16 and 28 cycles for 12 different primer pairs that we have tested. Note that ‘linear amplification’ does not necessarily mean that the amount of product has to double with each additional cycle, only that it has to be a linear increase. We often find the increase to be around 1.5-1.7 fold. Post PCR work-up: We use Qiagens ‘QIAquick spin’ PCR purification columns to remove remaning 32PdCTP. This may not be necessary. 1/3 of the sample is subsequently run on a 5% acrylamide gel, dried and exposed to a Fuji phospoimager plate.

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Quantitation: We utilize the Fuji Phosphoimager software to quantitate the bands on the gel as well as to subtract the background (the signal from an area adjacent to the band). We then compare the signals from the internal controls in all the samples and calculate a correction factor for each sample. This factor is then used to correct the signal from the gene of interest. In addition, we usually express the results as percent of a control sample to permit comparisons between different experiments. Compare with independent assay: The acid test of any quantitative PCR is to compare the PCR results with an independent assay. We compared the PCR to Northern blots and found an excellent agreement between the two methods.

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PROTOCOLS: cDNA synthesis 10 ul 1 ul

10 ng-1 ug total RNA in H20. 20 uM random hexamers. Heat to 70˚C for 10 minutes.

4 ul 2 ul 1ul

5x RT buffer (GibcoBRL) 0.1 M DTT 10 mM each dNTP Incubate at 37˚C for 2 minutes Add 1 ul Superscript II RT (GibcoBRL). Incubate at 37˚C for 90 minutes. Incubate at 95˚C for 10 minutes.

PCR reaction. 5 ul 3 ul 1 ul 1 ul 1 ul 0.5 ul 0.5 ul X ul Y ul

10x PCR buffer 25 mM MgCl2 10mM each dNTP 20 uM 5’ Oligo 20 uM 3’ Oligo 5U/ul Taq DNA polymerase 10uCi/ul 32P dCTP template cDNA ddH20 to a total volume of 50 ul

PCR reaction cycles:

1 minute at 94˚C 1 minute at 55˚C. 2 minutes at 72˚C.

This is the standard PCR starting point. The PCR of course has to be optimized for each primer pair by titrating the concentration of MgCl2 , primers and dNTP as well as optimizing the annealing temperature etc. Standard PCR controls such as single primer controls and controls for DNA contamination of course have to be included. Analyse hot reactions by electrophoresing on a 5% acrylamide/TBE gel, dry down and analyse by autoradiography or phospho-imaging. It is important to include single primer and no primer controls, at least for the first time that one uses a new set of primers.

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Standard PCR Protocol 1. BUFFERS: 10x PCR buffer: 500mM KCl 100mM Tris pH 8.4 (at room temp.) 1mg/ml gelatin Magnesium: 25 mM MgCl2 2. For a 1x reaction (50 - 100ul total volume): 1x PCR buffer 1.5- 4.0 mM MgCl2 0.2mM each dNTP 0.1-0.2ug each non-degenerate primer (15-30mers) OR approx. 5ug each degenerate primer 2.5 units Taq polymerase template DNA (5pg plasmid, 50ng genomic DNA, 1st strand cDNA from reverse transcription of 1ug total RNA). 3. Overlay reaction with 50-100ul autoclaved paraffin oil. 4. Cycle settings (25 - 35 cycles): 94C, 45 - 60 sec. (denaturation) 50 - 65C, 45 - 60 sec. (annealing- we typically start with 55C) 72C, 60 - 120 sec. (elongation) Last cycle 72C, 10 min. to complete elongation. 5. You may need to optimize Mg, primer and template concentration, cycling conditions- ie whether you "hot start" the reaction or not, whether you do the first few cycles at low annealing temp. These optimizations are especially important when employing degenerate primers or non-plasmid templates.

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3. Preparation of in vitro transcribed RNA For introduction of sense RNAs into embryos. Can be used to test effects of overexpression, misexpression or expression of dominant negative constructs. The mMessage machine kit from Ambion is usually reliable, but for those who deride kits, the following works very well. Transcription to make capped RNA in bulk.

Paul Krieg, Richard Harland

(Adapted from Green, Melton and Maniatis (1983) Cell 32, 681-694.) This method assumes that the coding region of interest is already inserted behind an appropriate bacteriophage promoter. The method below differs from previously published procedures by increasing the concentration of nucleotides greatly and by decreasing the ratio of cap analog to GTP (to 3:1). Note: When preparing the reaction mix, be sure to warm the reaction mix, and add the 10X transcription buffer as one of the final ingredients, otherwise the spermidine and cap analog in the buffer will precipitate the DNA template. Reagents: Template DNA solution: Template plasmid DNA linearized at the 3’ end of the sequence to be transcribed.Concentration about 1 mg/ml in TE. 10X Transcription buffer:

120 mMMgCl2 800 mM HEPES-Cl pH 7.5 20 mM Spermidine-HCl

BSA must be clean. The BSA that comes with restriction enzymes etc. is good. 2X Nucleotide triphosphate mix: 6 mM ATP 6 mM CTP 6 mM UTP 3 mM GTP 9 mM m7(5’)Gppp(5’)G, cap analog (e.g. N.E. Biolabs #1404 orAmbion #8050). alternatively use unmethylated GpppG (e.g. N.E.B. #1407), which is cheaper. It is reported not to be as good as methylated analog in reticulocyte translation, but in oocytes and embryos this reaction yields RNA of similar potency to Ambion's mMessage Machine. Sephadex G-50 equilibrated in 0.3 M NaOAc, 0.1% SDS (DEPC treated and autoclaved). After autoclaving, replace the supernatant with fresh sterile solution, shake and settle. Repeat. This removes dextrans that may leach out of the sephadex (a problem with some batches). After making the column, prespin, wash with fresh buffer, respin and use. Method: 1. For a 20 μl reaction volume, assemble the following ingredients, into a warmed tube, in the order indicated. If the nucleotide crashes out of solution before synthesis, you will probably need to dilute and heat the reaction. A precipitate invariably forms during the reaction, but the yields and bioactivity are good. Linear template DNA (0.5 to 1 mg/ml) 2X Nucleotide triphosphate mix. 117

2 μl 10 μl


α32P

UTP (if available) DTT (200 mM) BSA (1mg/ml) 10X Transcription Buffer RNase Inhibitor (20 units/μl) Bacteriophage RNA polymerase (20 units/μl)

0.1μl (trace) 2 μl 2 μl 2 μl 0.5 μl 1.5 μl

2. Mix gently, and incubate the reaction at 37oC for 2 hours. 3. Add 2 units of RNase free DNase and incubate for a further10 minutes at 37oC. Cerenkov count the tube to obtain the total available cpm. 4. Dilute the reaction to 100 μl with a buffer such as 100 mM NaCl, 30 mM EDTA, 20 mM Tris pH 7.5 and 1% SDS. 5. Spin through sephadex G-50 equilibrated in 0.3 M NaOAc, 0.1% SDS (DEPC treated and autoclaved). The column has three purposes, first it allows you to tell that the transcription has worked, and it is easy to monitor precipitation of the RNA, since almost all the counts are now in RNA. Second, the exotic mix of nucleotides can cause RNA to aggregate at a phenol interface, thus requiring multiple reextractions of the organic phase for good recovery of RNA; the column removes most of the nucleotide. Finally, cap analog is extremely toxic to embryos, and its worth taking effort to remove it. 6. Extract with phenol/chloroform. Check the recovery of counts from the interface and if necessary extract again. This step is only for the obsessed,and can be omitted. 7. Add two volumes ethanol and precipitate. The RNA can be left at this point at -20° for as long as you like. Spin and resuspend in 100 μl DEP H2O. Take 1 μl for incorporated cpm. Count the total and incorporated (correcting for volume) and find the % cpm incorporated. 100% incorporation of UTP would be 80 μg mRNA, but the reaction is limited by the available GTP so the maximum theoretical yield is 50%. The usual experimental yield is 10-20% or 8-16μg mRNA. 8. Reprecipitate the RNA with DEP NaOAc (pH 5.5) and ethanol. Wash pellet in 70% ethanol. Resuspend at desired concentration. Remove 1 μl of the transcription mix and examine on a gel to assess the transcript yield and integrity. Check the resuspension of RNA by recovery of counts. Sometimes the RNA is reluctant to redissolve.

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Observations. Problems with yield or translatability While poor yield is usually due to poor template, the commercial reagents are sometimes at fault. Over the past years we have come across batches of GpppG that are poisonous to transcription, or worse, they allow RNA synthesis but the RNA is not translatable. It is therefore valuable to keep stocks of known quality reagents as positive controls, so that you can provide a basis for complaints to the manufacturer. Calculation of yield If the labeled nucleotide is UTP or CTP, then the reaction above contains 10 μl of 6mM UTP. In an average RNA, for every mole of UTP incorporated there will be one mole of each of the 4 nucleotides incorporated. We assume an average molecular weight of nucleotide monophosphate of 330 g/mol. So if all the UTP were incorporated (10x 10-6 liters)x(6x10-3mol/liter) x4x330g/mol = 79.2x10-6 g. Of course, there is half as much GTP as there is UTP, so incorporation should never be above 50%. Nevertheless, for the purposes of calculating mass each 1% of yield corresponds to 0.8μg RNA. For synthesis of large numbers of RNAs as in expression cloning, half-scale reactions yield sufficient RNA. For a half scale reaction each 1% incorporation of UTP corresponds to 0.4μg RNA. More highly radioactive mRNAs can be made without radiolytic degradation being problematic. 1μl of radioactive nucleotide can be added to the standard reaction. After storage at -80° in DEP H2O for 2 years the RNA is still active in translation (no more than 4fold worse than it started out). Removal of short transcripts For critical work you can remove very small RNA from premature termination by precipitation with LiCl. This requires that the concentration of RNA be more than 0.5 mg/ml. Add LiCl from a DEP treated 10M stock to 3M final. Leave on ice for > 1 hr (-20 overnight is also OK but thaw before spinning). The RNA may be invisible so monitor cpm, the pellet is also easily visible after 80% ethanol wash. You will lose a surprisingly large amount of counts, but be careful not to lose the real stuff. Follow the LiCl precipitation with a standard NaOAc Ethanol precipitation. Be patient when resuspending; vortex often, 1-2 min at 65° helps. Choice of vector All polymerases cap to the same efficiency (based on stability of RNA in oocytes). However some polylinkers contain sequence that is poisonous to translation. The Sive lab. (Kuo et al, Gene 176: 17-21, 1996) has shown that 5' homopolymeric deoxycytidine stretches can depress translation of adjacent mRNAs up to 10-fold in vitro and up to 100fold in Xenopus embryos. These 5' sequences would result from cDNA synthesis involving tailing with dG. This effect is observed when cDNAs containing 5' dC tracts are cloned downstream of T7 or T3 polymerase promoters, but not when cloned downstream of SP6 promoters. T3 and T7 promoters generally initiate transcription with 5'GGG as the first three bases, while SP6 transcripts initiate with 5'GAA (Fig.3). We postulate therefore that stem-loop structures may form between the 5' end of T3 and T7 transcripts and downstream dC tracts to prevent cap-binding protein from allowing ribosome assembly and translation. This suggests that in construction of cDNA libraries where eukaryotic

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expression is desired, avoiding priming second strand synthesis with oligodC, or avoiding cloning vectors with complementary dG tracts in translated regions may be prudent. Consistent with this, (Bob Brown unpublished) CAT in the Sma site of pSP65 is fine, but in the Bam site of bluescript and transcribed by T7 is dead, or T3 is weak. In general the less polylinker present the better. Many people routinely subclone DNAs for translation into pSP64T or derivatives, plasmids that contain globin 5' and 3' untranslated regions. Although no systematic analysis has been published these clearly boost translation many fold for many investigators and many mRNAs, while having no effect on others. This may be particularly relevant when one wants to overexpress maternal mRNAs. Since no transcription takes place before the midblastula transition, much regulation of maternal RNA expression is translational and regulatory regions often contained in the 3' and 5' untranslated stretches. Removing these regions and substituting the strong globin UTRs and even AUG Kozak consensus may help very significantly. The CS vectors (Turner and Weintraub, 1994) give very potent mRNAs, due to the incorporation of a polyadenylation signal in the primary transcript. This is cleaved and polyadenylated after injection into embryos, providing a boost of ten fold more protein per nanogram of injected mRNA over many equivalent transcripts made from pSP64T. However, for oocyte injections, pSP64T provides a ready made polyA tail that enhances translation over long term incubations, while CS2 transcripts would only be adenylated if they are injected into the nucleus. Vectors with a polyA stretch can be useful for selecting full length transcripts by oligo dT cellulose chromatography. However, watch out for instability of polydA tracts in plasmids, one can seldom maintain more than 50 As.

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4. RNA Solutions Note: Although protocols listed call for DEP treatment, one of us (HLS) has found that this is not necessary, when starting with good quality distilled water. DEP water 1ml DEP to 100ml water. Then autoclave. Since DEP attacks free amines, any amines should be added after autoclaving (e.g. Tris, EDTA) To avoid spontaneous appearance of nuclease in DEP water either use a fresh bottle, or add EDTA to 0.1mM after opening to stop fungi germinating and growing 5x SP6 buffer. 200 mM Tris, pH 7.5 30 mM MgCl2 20 mM spermidine. Autoclaved and stored frozen in aliquots. DNAse I is diluted from the concentrated stock into 20 mM NaOAc, pH 6.5, 5 mM CaCl2, 50% glycerol (dilution buffer should be DEP treated and autoclaved. Formamide+dyes: 98ml redistilled formamide (EM FX0421-3 or BRL 5515UB) 2ml 0.5M EDTA PH 8, 0.05% bromophenol blue and xylene cyanol Preparative gel for 100ml stock of 5% gel 12.5 ml 40% acrylamide 12.5 ml 2% bis acrylamide 10ml 10x TBE 42 g urea to 100ml with water 25ml is sufficient for 1 small gel (e.g. BRL SA32) add 50.μl TEMED and 50μl 20% ammonium persulphate to polymerize Gel Elution buffer 0.3M NaOAC pH 5.5 0.1% SDS 10mM EDTA 10μg/ml RNA carrier 5X Hybridization Salts 0.2M Pipes (pH 6.7) 2M NaCl 5mM EDTA

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RNase Digestion Buffer (1X) 10mM Tris-HCl (pH 7.5) 5mM EDTA 300mM NaCl 2ug/ml RNase T1 (Calbiochem 556785. Made up in water to 10,000 units per ml and frozen in 10μl aliquots). carrier RNA is from Sigma R3629 type IX Torula. Seems to be OK just redissolved in DEP water at 50mg/ml, but the purists will want to extract it first. Resuspend at 200mg (yes mg)/ml in 1%SDS 30mM EDTA 100mM NaCl 20mM TrispH7.5 or similar. Add proteinase K to 10μg/ml digest at room temp for a few days. Add NaOAC and isopropanol to precipitate, resuspend in DEP water, precipitate with 3M (final) LiCl. The pellet will contain longer RNA, the sup can be reprecipitated with NaOAc and isopropanol for shorter carrier. Measure OD for concentration. glycogen is Molecular Biology grade from Boehringer Mannheim. Comes as 20mg/ml solution, use 10 - 20 μg per ethanol precipitation in 1.5ml Eppendorf tube. Is an excellent carrier, inert in all reactions tested, comes through phenol/chloroform but not rhough butanol extractions.

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5. Siliconizing tubes, glassware Hazel Sive

Siliconozed tubes are a good idea when one is dealing with small amounts of any singlestranded nuleic acid, which can stick irrevesibly to plastic or glass tubes, especially after ethanol precipitation. Addition of glycogen as carrier is a big help (Boehringer, molecular biology grade, 10-20μg per ethanol precipitation in 1.5ml Eppendorf tube). In our experience, commercially siliconized tubes are a poor imitation of well prepared home siliconized tubes, however one can buy pre-siliconized tubes from Intermountain Scientific Corporation (1-800-999-2901). Put tubes etc. in dessicator in fume hood. Loosely cover tubes with kimwipes, paper towels, not foil. Put 10.20ml Dichlorodimethylsilane (SIGMA # D3879) in beaker in dessicator. BE CAREFUL - DCDMS IS VERY TOXIC. close dessicator, turn on vacuum. Wait - 2 min. Release vacuum briefly, to cause a "spray" of DCDMS. Turn on vacuum again. Keep under vacuum > 8h, until all DCDMS has vaporized. Rinse plastic etc. 4-5x w. good DH2O Dry Autoclave Dry NOTE:

Best to have a special dessicator for this procedure - since it gets very gunky. Line with paper before each use. Open only in hood (even when no DCDMS liquid present). CHECK TUBES FOR DROPLETS THAT WILL NOT DRY. DO NOT USE IF YOU FIND THESE - they are silica +? and will cause your precipitated nucleic acid to become insoluble FOR EVER. 2 things help above problem: a) Use fresh (relatively) DCDMS - don't use stuff that is years old. We buy 50ml bottles and use them within a couple of months. b) Make sure DCDMS has completely vaporized after vacuum treatment.

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DNA Methods Richard Harland

1. Extraction of DNA from single embryos When harvested the embryo should be homogenized in 0.5 ml of 1% SDS 10 mM EDTA 20 mM tris 7.5 100 mM NaCl or some similar buffer. Freeze until ready to process. Then add proteinase K to 100 μg/ml (final) This is conveniently done by adding 200μl of proteinase K solution to each vial. Incubate overnight at 55°. Extract with aqueous phenol at least twice. (if the DNA is viscous and seems to drag phenol with it it may pay to remove the lower phenol layer rather than retrieving the upper aqueous layer) Extract with 1:1 phenol chloroform and then once with chloroform Add Ammonium Acetate to 2M (can start with a 10M stock) To the total aqueous volume now add 0.6 vol isopropanol. Mix thoroughly, if the DNA is viscous this will take many inversions of the tube. If you can see a stringy DNA precipitate spin for a few seconds in the microfuge. If no DNA is visible, leave the tube on ice for 30 min (or overnight even at -20°). Spin for 5 min to recover DNA. Resuspend the DNA in 100 μl TE (10 mM tris pH 7.5, 1 mM EDTA) Digest with RNAse A (10 μg/ml) and RNAse T1 (10 units/ml) for 30' at room temp. Repeat the precipitation. Wash the pellet with 70% ethanol. Resuspend in 20 μl TE and proceed to digestion with restriction enzymes. Cautionary note There is something in eggs that is bad for restriction enzymes. It is therefore essential to extract the DNA thoroughly with the phenol to get good digestion.

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2. Bulk DNA isolation from red blood cells Isolation of frog Red Blood Cells Anesthetize the frog by immersion in ).05% benzocaine (from a 10% stock in ethanol). Open the ventral surface to expose the heart, preferably with a small hole so that the liver does not pop out (A hole is made ~1cm square in the upper ventral thorax. Cut the cartilaginous shield-shaped bone connecting the shoulders with heavy scissors and remove. The heart will be visible. Carefully lift the silvery pericardium away from the heart and cut open to expose the heart.) Inject 0.5 ml of 2mg/ml heparin in 0.85X SSC. Bleed the frog into 0.85X SSC by "draping" the frog over a beaker with the heart dipping into the SSC. The blood will be pumped into the beaker through the hole made by the heparin injection. If flow stops make another incision in the heart with scissors. If you can bring yourself to do it massaging and squeeezing the frog's legs will help the blood flow. Another source of RBCs is the liver. When using liver keep extra cold and avoid damaging the gall bladder. Mince the liver in 0.85x SSC and dounce with a B pestle. Pour the material through two layers of cheesecloth. Spin the cells down in a bench top centrifuge at about 2,000 rpm. Pour off the supernatant, but don't remove everything. Place pellets on ice. All further steps with ice cold buffers, etc. Partially resuspend the pellet in the residual supernatant. Lyse the cells in RSB + NP-40: 0.05% NP-40 10 mM NaCl 10 mM Tris-HCl, pH 8 5 mM MgCl2 Start with a small volume (5 ml), resuspend cells, and increase volume to approximately 50 ml. Leave on ice 1 minute and spin down nuclei. Nonidet 40 (NP-40) is a non-ionic detergent which breaks down cell membranes, but nuclear membranes are stable to this detergent. Pour off the supernatant gently. Resuspend loose pellet by vortexing in its juices. When resuspended add more NP-40 in RSB to 50 ml. Spin down nuclei. Repeat this step until nuclear pellet is white. If serious glopping occurs or if there are clots, dounce gently using the B pestle. Alternatively, pour the resuspended cells through two to four layers of cheesecloth to remove the clots. (optional) Before the last centrifugation, take a 10 ul aliquot of the nuclei and dilute to 1 ml in 1% SDS. Measure the A260. To store the nuclei, resuspend in nuclei freezing buffer at a concentration of 1 to 5 mg/ml: 50% glycerol 0.15 M NaCl 5 mM MgCl2 10 mM Tris-HCl, pH 8 0.1 mM EDTA 1 mM DTT Preparation of genomic DNA from red blood cells The preparation of genomic DNA without shearing requires gentle treatment. Procedures where shearing or nicking could be inadvertently introduced should be avoided. Therefore, it is desirable to use a procedure with a minimum of steps and without vortexing

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or phenol extraction. The following procedure will yield DNA of size greater than 200 kb if carefully done. This length DNA is suitable for cloning into cosmids and is certainly suitable for use in genomic Southern blots. Just remember not to vortex and not to get impatient. Dilute the nuclei to 100-200 ug/ml of DNA in RSB in a 50 ml Corning tube. Add proteinase K to 200 ug/ml (final concentration). Mix gently. Add an equal volume of 0.6 M NaCl, 20 mM Tris-HCl, pH 7.4, 20 mM EDTA, 1% SDS. Mix gently, but thoroughly. The solution will be come very viscous as the nuclei lyse. Incubate overnight at 37 - 50°C. Add 1/4 volume 10 M NH4 OAc. Mix thoroughly. Measure the total aqueous volume and add 0.6 volumes of isopropanol. Mix again. Fuse the end of a long pasteur pipet in a Bunsen burner flame. Use this rod to spool the DNA out of the mixture. When viscosity has disappeared and the DNA is completely spooled onto the pipet, discard the remaining solution and replace it with 20 ml of 70% ethanol. Using another pasteur pipet, scrape the DNA off into the 70% ethanol. Let wash until completely white and rather stringy looking. Carefully pipet off all the traces of the ethanol, avoiding the DNA. Suspend the DNA at an estimated concentration of 200 ug/ml in TE. Rock overnight to dissolve.

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Western Blots Peggy Kolm with help from Mike Klymkowsky

Lysis

For quick and dirty protein analysis, homogenize embryos in Laemmli buffer (20 μl /embryo). Pipet up and down with a yellow tip, then vortex. Centrifuge 5’ at 4° C. to separate out the yolk. Run one embryo equivalent of the supernatant on an SDS-PAGE gel. This is adequate for many proteins, however, the protein bands can be smeary due to the yolk. Additionally, proteins between 80 and 100 kDa are displaced by the yolk band. For cleaner gels, embryos can be lysed in ice-cold NP-40 buffer (10 μl/embryo homogenize thoroughly). Lysates are centrifuged 5-10 minutes at 4°C, then mixed with an equal volume of 2x Laemmli buffer. Solubility in this buffer is protein dependent, some proteins (e.g. desmoglein) are almost completely insoluble. NP-40 Buffer* 2X Laemmli buffer 0.5% NP-40 4% SDS 20% glycerol 20% glycerol 50 mM Tris-Cl, pH7.5 120 mM Tris pH6.8 150 mM NaCl 10% b-mercaptoethanol 1 mM DTT (add immediately before use) store at -20°C. store @ 4°C. * We have successfully used 0.5% NP40 in 1X PBS100x Protease Inhibitors (optional) (add immediately before use) 100 mM PMSF (dissolve in 100% EtOH) 1 mg/ml pepstatin (dissolve in 100% MeOH) 2 mg/ml aprotinin 1 mg/ml leupeptin store individual stocks @ -20°C. 25X Phosphatase Inhibitors (optional) (add immediately before use) 250 mM Na b-glycerophosphate 25mM NaF 2.5 mM Na orthovanadate store 25X mixture in aliquots @ -20°C. SDS-PAGE, Western Transfer, and Visualization A 10% SDS-PAGE gel is appropriate for separating proteins in the 40-60 kDa range. We recommend the Hoeffer Mighty Small gel running aparatus/Western transfer system. The 10-well comb allows the loading of 20μl of sample (one embryo equivalent). Prestained molecular weight markers (BioRad or BRL) allow you to see the progress of the gel as it is running, and easily orient the membranes after transfer. To visualize antibody binding the ECL chemoluminescence kit (Amersham) is simple and contains all the necessary reagents (blocking reagent, 2° antibody, detection reagents).

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APPENDIX II: Culture media from Gerhart, Harland, Sive labs and others

There are numerous, more or less interchangable media in which one can successfully culture Xenopus embryos. The chief differences are the monovalent cation concentration and the buffer. The original culture media devised by Holtfreter employed bicarbonate as buffer. This is a very poor buffer, but can be useful, where one wants, for example to alter the pH of the culture media. However, in general more efficient buffers are more reliable and used in our labs. Isotonic solutions are used for culture of isolated tissue, approximately 0.1 - 0.3 X isotonic solutions for culture of whole embryos. Specialized solutions include those that mimic blastocoel fluid, for Keller explant culture, for example, or solutions that prevent animal cap curling. 1. General media Modified Barth's Saline 1x=

2 solutions are made

88mM NaCl 1mM KCl 0.7mM CaCl2 1mM MgSO+ 5mM HEPES pH 7.8 2.5mM NaHCO3 -

CaCl2 (0.1M) NaCl/KCL/MgSO4HEPES (10 x stock) and autoclaved separately. Keep CaCl2 in aliquots / frozen or at 4°C

10 x MBS SALTS (1l) Stock solution. NaCl KCl MgSO 4 .7H2 O HEPES NaHCO 3 pH H2O AUTOCLAVE

51.3g 0.75g 2.0g 23.8g 2.0g 7.6 with NaOH to 1l

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1 x MBS (1l) For culture of explants or oocytes. For long term culture, use 0.5 X. 10 x MBS 100ml 0.1MCaCl2 7ml H2O 892ml Gentamycin (50mg/ml) 1ml (100u/ml penicillin and 100μg/ml streptomycin sulfate ((GIBCO), if desired) 1/10 x MBS (1l) Use for culture of whole embryos (+/- jelly or vitelline membrane, but must be intact) 1 x MBS H2O Gentamycin

100ml 900ml 1ml

1 x MBS + High Salt Eggs can be laid into this, in this solution they remain competent for fertilization for up to 12 hours. The better the batch of eggs, the longer they will remain competent. 0.1M CaCl2 10 x MBS 5M NaCl H2O

7ml 100ml 4ml 988ml

Amphibian Ringers General culture medium. Modified Frog Ringers - MR 0.1 M NaCl 1.8 mM KCl 2.0 mM CaCl2 1.0 mM MgCl2 5.0 mM Hepes-NaOH, pH 7.6 or 300 mg/l NaHCO3 MMR (Ubbels et al. (1983) JEEM 77:15-37) 0.1 M NaCl 2.0 mM KCl 1 mM MgSO4 2 mM CaCl2 5 mM HEPES, pH 7.8 0.1 mM EDTA

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NAM (normal amphibian medium) General culture medium JEEM 80 289, 283 JEEM 78 299 110 mM 2 mM 1 mM 1 mM 0.1 mM 1mM 2 mM

NaCl KCl Ca (NO3) Mg SO4 Na2EDTA NaHCO 3 Na Phosphate pH 7.4

27.5 ml 4M 1ml 2M 1 ml 1M 1ml 1M 0.5 ml 0.2M 1 ml 1M 2ml 1M

Holtfreter's solution General culture medium, poorly buffered, but can be useful. May be best buffer for progesterone-induced oocyte maturation. 10 x =

600mM 6mM 9mM 25mM

NaCl KCl CaCl2 NaHCO 3

Make up 10 x salts 10 x NaHCO3 separately Can autoclave salts, filter sterilize NaHCO3

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2. Specialized media Danilchik's Blastocoel Buffer For explant culture, especially Keller explants. Allows good convergent extension, likely because of high pH. For 500 ml use: 6.63 ml 4.0 M NaCl 9.38 ml 0.8 M NaHCO3 5.0 ml 0.25 ml 0.60 ml

0.45 M K-gluconate (Aldrich) or 0.527g 2 M CaCl2 or 0.074 g CaCl2-2H2O 0.83 M MgSO4 or 0.124 g MgSO 4 -7H 2 O

0.408 g bicine (Mann) ≈450 ml dH 2O Adjust pH to 8.3 with a measured volume of 1.0 M Na2CO3 (≈1.8 ml), calculate total Na and adjust the total Na to 95 mM with Na-isethionate (2-hydroxy ethanesulfonic acid from Sigma) (about 1.73 g). Volume is then brought to 500 ml. NOTES: 1) A small amount of precipitation occurs after 2-3 weeks so fresh stock should be made up weekly. 2) This medium depresses apical-surface healing responses. Mike D. interprets this as evidence that internal cells are content in this medium but he says it is a lousy rearing medium for intact embryos. However large volumes injected into the blastocoel seemed to have no detrimental effects. 3) Mike D. took the ion concentrations from the mean free intercellular activities measured by J. Gillespie for Xenopus (J. Physiol. 344: 359, 1983) 4) The ion concentrations in Danilchik's medium: Na+ 95 mM HCO 3 -/CO 3 = 18-19 mM + K 4.5 mM bicine 5.0 mM Ca + + 1.0 mM SO 4 = 1.0 mM Mg + + 1.0 mM isethionate 23-24 mM Cl 55.0 mM gluconate 4.5 mM Sater's modified blastocoel buffer Basically the same as Danilchick's. 49.5 mM NaCl 12.375 ml 4M NaCl 36.5 mM Gluconic acid, sodium salt 7.95 g 5 mM Na Carbonate (Na2CO3) 0.53 g (105.9 MWt) 4.5 mM KCl 2.25 ml 2M 1 mM CaCl2 1 ml 1M 1 mM Mg SO4 1 ml 1M ~6mM HEPES until pH 8.1 50 μg/ml gentamycin sulfate (powder from sigma) If desired, also add standard amounts of Penicillin and Streptomycin 1 mg/ml BSA optional Filter sterilize, freeze in 50ml aliquots. LCMR (Low Calcium Magnesium Ringer's)

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Slows healing of dissected embryos. Various recipes have been used, the first can be used for embryo dissection, but long term survival of explants is better in the second. Stewart, R. M., and Gerhart, J. C. (1990). The anterior extent of dorsal development of the Xenopus embryonic axis depends on the quantity of organizer in the late blastula. Development 109, 363-372. 66mM NaCl 1.33 mM KCl 0.33 mM CaCl2 0.17mM MgCl2 5mM HEPES pH 7.1 (adjust pH after making) 50μg/ml gentamycin For explant dissection and culture: Hemmati-Brivanlou, A., Stewart, R. M., and Harland, R. M. (1990). Region-specific neural induction of an engrailed protein by anterior notochord in Xenopus. Science 250, 800-802. 43 mM NaCl 0.85 mM KCl 0.37mM CaCl2 0.19 mM MgCl2 5 mM HEPES 50μg/ml gentacmycin 0.5%BSA if growth factors are added

CMFM (Calcium Magnesium Free Medium) For dissociating embryonic tissue. Will dissociate the inner, but not outer layer of an animal cap. (T. Sargent) for 1 L 500ml 10x NaCl 88 mM 22ml 4M 110ml 4M KCl 1 mM 0.5 ml 2M 2.5 ml 2M NaHCO 3 2.4 mM 2.4 ml 1M 12 ml 1M Tris pH7.6 7.5mM 3.75 ml 2M 18.75ml 2M

PhoNaK For dissociating embryos, much more vigorous than CMFM, will completely dissociate animal caps. (Slack - Dev. Biol. 134: 486 (1989)) 50mm Na Phosphate 5ml 1 M 35mm NaCl 0.7ml 5M 1mm KCl 0.1ml 1M ------- 100ml w. H2O

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3. Oocyte Culture Media O-R2 Medium (Wallace, et al. (1973). J. Exp. Zool. 184: 321-334)

NaCl KCl CaCl2-H2O MgCl 2 -6H 2 O Na2HPO4

mM 82.5 2.5 1.0 1.0 1.0

g/l 4.822 0.186 0.147 0.203 0.142

g/l 10x stock A 48.221 1.864 1.470 2.030 --- 1.420

10x stock B ---------

HEPES NaOH

5.0 3.8

1.192 0.152

11.915 1.520

-----

Use 1 volume of Stock A, 1 volume Stock B, and 8 volumes dH2O. This should yield a pH 7.8 solution. Oocyte Culture Medium (modified from Opresko, et al. (1980), Cell 22: 47-57) 50 % L-15 + glutamine (Gibco Leibovitz stock) 40% HEPES / insulin stock* 10% serum or VTG (vitellogenin)** 100 μg/ml gentamycin 0.5% GIBCO fungione/penicilin/streptomycin (100x stock) * 100 ml HEPES/insulin stock: 37.5 mM HEPES pH7.8 2.5 μg/ml Sigma insulin stock (22.5 IU/mg; 5 mg/ml insulin slurry in dH2O solubilized with 50μl 0.5 M EDTA pH7.7) **Fetal calf serum or frog serum with 50-200 mg/ml VTG

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Devitellinating Buffer (Horrell et al. 1987. Genes and Devel. 1: 433) Oocytes that have been manually removed from the ovarian tissue must first be treated for 30-60 min with 1 mg/ml collagenase (in CaMg free MR) before attempting to remove the vitellines. After rinsing out collagenase, incubate the oocytes in: 20 mM KCl 1 mM MgCl2 10 mM EGTA 10 mM HEPES, pH 7.2 After 5-10 min it should be possible to remove the vitelline envelope with forceps.

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APPENDIX III: Fate maps of blastula stages Rob Grainger

Since the early part of the century embryologists have used fate mapping techniques to be able to follow the developmental changes of particular regions in amphibian embryos. Such observations have yielded important insights, particularly about the remarkable intricacies of gastrulation. Early studies involved application of vital dyes to particular regions of the embryo, which could then be observed at later stages. In recent times these techniques have been extremely useful as well. For example the mechanisms of gastrulation in Xenopus, studied by Keller and coworkers, have utilized these methods (e.g. Keller, 1975 and 1976). Detailed descriptions of these methods are discussed by Rugh (1948) and Hamburger (1960). Shown below is a summary of gastrulation movements in Xenopus ascertained in part by such methods:

Legend . Xenopus development is shown at the early gastrula (top panel) late gastrula (middle panel), and late neurula (bottom panel) stages. Morphogenetic movements are

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shown from a surface viewpoint (far left column) and from a midsagital viewpoint (left center). Embryos are arbitrarily oriented to have the prospective dorsal midline on the left (in the visual plane of the page) and the prospective ventral midline on the right. Arrows indicate the direction of movement. (A, animal pole; AC, animal cap; AF, archenteron floor; AR, archenteron roof; BC, bottle cells; BLC, blastocoel; BPL, blastopore pigment line; DZ, deep zone; IMZ, involuting marginal zone, IMZ-D, IMZ deep layer; IMZ-S, IMZ superficial layer; LI, limit of involution; NIMZ, noninvoluting marginal zone; PBC, prospective bottle cells; RBC, respread bottle cells; VB, vegetal base; YP, yolk plug.) Figure and Legend from Gerhart and Keller, 1986. Several methods have been devised more recently that are very useful for fate mapping. The fate map of the Xenopus neural plate (summarized in the section on morphological development) by Eagleson and Harris (1989) utilizes two approaches. In one, a particular tissue is removed from the embryo, soaked in the fluorescent Hoechst DNA-binding dye (#33258) and then replaced back into the unlabeled host. After fixation at a later stage the labeled implant can be viewed by epi-fluorescence. Alternatively one can label particular regions of the embryo by application of small crystals of DiI or DiO, which can also be monitored at later stages by their fluorescence. A very useful modification permits observation of marked regions of the embryo after in situ hybridization (based on the method of Izpisua-Belmonte et al., 1993), thus permitting one to correlate a particular morphological region with a particular domain of gene expression. Either before or after fixation the region of the embryo to be marked is labeled by brief application of DiI crystals (Molecular Probes, In. #D-1125). At some point before beginning the in situ hybridization procedure a photoconversion process is used to create an insoluble product at the site of application of DiI (DiI itself lipid-soluble and would be lost during the hybridization procedure). The DiI-treated embryo is incubated in the dark with diaminobenzidine (1 mg/ml) in 0.1 M Tris-HCl (pH 7.4) for 30 min-1 hr. Embryos are then illuminated at 547 nm (the wavelength used to monitor rhodamine fluorescence) until red fluorescence begins to fade. Embryos are then washed 2-3 times over a 30 min period in 0.1 M Tris-HCl. In recent years a number of techniques have also been utilized to follow the fates of particular blastomeres. Injection of horseradish peroxidase (e.g. Dale and Slack, 1987; Moody, 1987) and fluoresceinated dextrans (Gimlich and Gerhart, 1984) has been particularly useful. From such studies the fates of every blastomere from early embryonic stages has been determined. Shown below are summaries of the fate maps of 16-cell and 32-cell embryos. Because it is often desirable to ascertain the effect of ectopically expressed genes on particular tissues in the embryo, injection of gene constructs or mRNAs into blastomeres that give rise to a restricted set of tissues can be very useful. The fate maps illustrated have been quite helpful for such studies. In addition to injecting the gene construct or mRNA being tested for biological activity in such experiments, investigators have also co-injected constructs/mRNA encoding beta-galactosidase, which is expressed in the embryo and can be used as a lineage label to identify definitively the tissues derived from the injected cell or cells (e.g. Smith and Harland, 1991). Fate Map of the 16-Cell Xenopus Embryo (Moody 1987a)

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Next page: Fate map of the 32-cell Xenopus embryo (after Moody, 1987b)

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THIS IS THE PAGE THAT THE COLOR PLATE GOES ON!!!!!! DO NOT INCLUDE THIS PAGE AS IS!!!!!!!

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Fate map of the thirty two cell embryo: Dale and Slack, 1987

Fig. 1. Nomenclature of blastomeres at the 32-cell stage. D: dorsal, V: ventral, AP: animal pole, VP: vegetal pole.

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Dale and Slack (1987) 32-cell fate map. Orientation of the embryos is as on previous page.

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References Bauer, D.V., Huang, S. and Moody, S.A. (1994) The cleavage stage origin of Spemann’s Organizer: analysis of the movements of blastomere clones before and during gastrulation in Xenopus. Development 120,1179-1189. Dale, L. and J.M.W. Slack (1987) Fate map for the 32-cell stage of Xenopus laevis. Development 99, 527-551. Eagleson, G.W. and Harris, W.A. (1989). Mapping of the Presumptive Brain Regions in the Neural Plate of Xenopus laevis. J. of Neurobiology 21, 427-440. Gerhart, J. and Keller, R. (1986). Region-specific Cell Activities in Amphibian Gastrulation. Ann. Rev. Cell Biol. 2, 201-229. Gimlich, R.L. and Gerhart, J.C. (1984). Early Cellular Interactions Promote Embryonic Axis Formation in Xenopus laevis. Dev. Biol. 104, 117-130. Hamburger, V. (1960). A Manual of Experimental Embryology. Chicago, University of Chicago Press. Izpisua-Belmonte, J.C., DeRobertis, E.M., Storey, K.G. and Stern, C.D. (1993). The Homeobox Gene goosecoid and the Origin of Organizer Cells in the Early Chick Blastoderm. Cell 74, 645-659. Keller, R. (1975). Vital Dye Mapping of the Gastrula and Neurula of Xenopus laevis I. Dev. Biol. 42, 222-241. Keller, R. (1976). Vital Dye Mapping of the Gastrula and Neurula of Xenopus laevis II. Dev. Biol. 51, 118-137. Moody, S.A. (1987a). Fates of the Blastomeres of the 16-cell Stage Xenopus Embryo. Dev. Biol. 119, 560-578. Moody, S.A. (1987b). Fates of the blastomeres of the 32-cell Xenopus embryos. Dev. Biol. 122, 300-319. Rugh, R. (1948). Experimental Embryology: A Manual of Techniques and Procedures. Minneapolis, Burgess Publishing Co. Smith, W.C. and Harland, R.M. (1991). Injected Xwnt-8 RNA Act Early in Xenopus embryos to promote formation of a vegetal dorsalizing center. Cell 67, 753-765.

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APPENDIX IV: Morphology of Xenopus embryos and adults 1. Early development The comprehensive treatises on the development of Xenopus laevis by Nieuwkoop and Faber (1994) and by Hausen and Riebesell (1991) (the latter focusing on early stages) are invaluable resources. All stage numbers mentioned here are according to the system of Nieuwkoop and Faber. In brief summary, after fertilization, cleavage proceeds rapidly and the blastula stage (st. 7-8) is reached by 4 to 5 hr (all times refer to development at 22oC). Gastrulation commences at 9 hr (st. 10) when the blastopore is first discernable and is complete by 13 hr (st. 12). Frog gastrulation is illustrated below, where one can see the late blastula stage (A) and the initial (B), middle (C) and late (D) stages of gastrulation.

From Balinsky, 1981

At st. 14 (16 hr) presence of the neural plate is quite clear (see section 2 below and Fate mapping section for further discussion). The closure of the neural plate to form the neural 143


tube occurs during the next several hr and is complete by st. 19 (21 hr). The process of frog neurulation is illustrated below:

.. A) Neural plate stage; B) mid-neurula; C; late neurula. From Balinsky, 1981.

Following neural tube closure the period of organogenesis commences, illustrated in the views of the early tailbud stage frog embryo below:

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. From Balinsky, 1981. .. Further organogenesis is seen in tailbud and tadpole stage frog embryos:

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. From Balinsky, 1981. 2. Neural Development Nancy Papalopulu

The vertebrate central nervous system can be divided into spinal cord, hindbrain (or rhombencephalon), midbrain (or mesencephalon) and forebrain (or prosencephalon). The forebrain is further subdivided into diencephalon and telencephalon. Diagramatic series of brain development

Legend: the positions of the boundaries between different brain regions is approximate.The asterisk shows the position on the neural tube of the original anterior end of the neural plate. C has been adapted from Keller, 1992a, B and D from Eagleson and Harris,1989. Abbreviations: cr, chiasmatic ridge; cf, cephalic flexure; di, diencephalon; ep:epiphysis; H: hindbrain; h: hypophysis; M: midbrain; ol, olfactory organ; op, olfactory placode; os, optic stalk; S, spinal cord; sl, sulcus limitans; tel, telencephalon.

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The Xenopus nervous system originates from the neural ectoderm which is induced on the dorsal side of the embryo during gastrulation, by signals originating from the dorsal mesoderm. Fate mapping in Xenopus (Keller, 1992b; see also Fate mapping section) have shown that in the early gastrula the prospective hindbrain and spinal cord extend well to the lateral sides of the embryo (panel a). During gastrulation and neurulation, cells in the posterior neural plate undergo dramatic convergence towards the midline and anteroposterior extension. Thus, material from the lateral sides of the gastrula is brought towards the midline and the neural plate transiently adopts a keyhole shape (panel b). This is the first stage where the neural ectoderm displays a clear medio-lateral and anteroposterior (or rostro-caudal) axis. The neural plate can be subdivided along the anteroposterior axis into two regions: the prechordal neural plate which gives rise to the forebrain and the epichordal neural plate which gives rise to the midbrain, hindbrain and spinal cord. The epichordal neural plate takes its name from the fact that it lies over the notochord, a structure that forms along the midline of the underlying dorsal mesodermal mantle during gastrulation (see section 1 above). During neurulation the midline of the neural plate deepens and the lateral edges rise towards each other and fuse creating the dorsal midline as seen in figures in section 1. The mediolateral axis of the neural plate becomes the dorso-ventral axis of the neural tube. Neurulation is more complicated in the anterior neural plate; In addition to lateral folding of the neural edges, there is also downward rotation of the A-P axis by approximately 90o at the cephalic flexure (panel c above). After st. 29/30 the forebrain gradually acquires a more straightened appearance (panel d). The key elements in the nervous system of a frog tadpole are shown below.

. From Nelson, 1953.

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The neural plate is surrounded by prospective neural crest and placodes. Cranial neural crest material segregates on the lateral edges of the anterior neural plate at st.15 and starts migrating after st 20. Cranial neural crest contributes to a variety of tissues, including the skeleton of the head and the branchial (gill) arches. X-twi .is a good marker for cranial neural crest at neural plate stages (Hopwood et al., 1989). Placodes develop as thickenings of the deep layer of the ectoderm, visible from about st. 22. The olfactory, adenohypophyseal, otic, epibrachial and dorsolateral placodes give rise to neuronal cells while the optic placode gives rise to the lens of the eye (see also eye development figure below). Neural crest at different stages:

Legend: Diagramatic series of neural crest development (taken form (Sadaghiani, 1987). Abbreviations: MCS, mandibular crest segment; HCS, hyoid crest segemnt.; aBCS, anterior branchial crest segment; pBCS, posterior branchial crest segment; VNC and TNC, vagal and trunk neural crest. .From Hausen and Riebesell, 1991. Ectodermal placodes at the neural plate stage:

Legend: The pattern of ectodermal placodes at the neural plate stage (taken from Knouff, 1935). Abbreviations: AHP, adenohypophyseal placode; AP, auditory (or otic) placode; LP, lens placode; CGA, cement gland anlage; OP, olfactory placode; NC, neural crest; NPL, neural plate; PPT, primitive placodal thickening. From Hausen and Reibesell, 1991. Both cranial neural crest and placodes contribute to the formation of the cranial ganglia. Cranial nerves can be visualised by hybridization with tannabin RNA (Hemmati-Brivanlou et al.1992, Neuron 9, 417-428) or immunostaining with anti-acetylated tubulin antibody. 148


The hypophysis originates from a very anterior placode at the neural late stage, but migrates posteriorly during neurulation and establishes a connection with the posterior diencephalon. Just outside the anterior neural plate, lies the primordium of the cement gland. The cement gland is a mucus-secreting ectodermal organ which helps the tadpole to suspend from the surface of the water, and it degenerates in later life. There are a number of genes that are expressed in the cement gland anlage from early neural stages onwards, such as XCG13 (Jamrich and Sato, 1989) XCG and XAG (Sive et al., 1989). A schematized version of tannabin expression in the tadpole (from Hemmati-Brivanlou et al, 1992) shown in the dark areas below, illustrates cranial nerve expression, and in other regions of the brain:

Legend: Summary of tannabin expression in st. 36 embryos; dorsal and lateral views. F, forebrain; M, midbrain, m/h midbrain-hindbrain boundary; r2-r8, rhombomeres 2-8; Cranial ganglia and nerves: V, VII, IX and X; E, Eye; OV, otic vesicle; s1-s5, somites; H, heart; LD, liver diverticulum. Morphological landmarks of the neural tube Unfortunately, relatively little is known about the anatomy of the embryonic Xenopus brain. It is only recently, with the discovery of genes which are expressed in the early neural tube in a regionalised manner, that the interest in the early neuroanatomy has been revived.

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Most classical neurobiology texts describe the anatomy of the adult brain which is shown below. From Nikundiwe, A. M. and Niuwenhuys, R. (1983)

Legend: Diagram of adult Xenopus brain with cranial nerves The nII, optic nerve, is not shown. Abbreviations: bolf, olfactory bulb; cereb, cerebellum; dienc, diencephalon; rhomb, rhombencephalon; tect, tectum; telenc, telencephalon. nI, olfactory nerve; nIII, oculomotor nerve; nIV, trochlear nerve; nV, trigeminal nerve; nVII, facial nerve; nVIII; acoustic nerve; nlX-X, glossopharyngeal and vagus nerves. In this section, some basic landmarks of the neural tube which are identifiable from early embryonic stages will be described. A diagram which illustrates some of the major features of later embryonic brain morphology in urodele embryos is shown in the figure which follows (taken from Slack and Tannehill, 1992). Several of the structures shown in this figure are described in the following sections. The hindbrain is relatively easy to identify because of its triangular shape and its thin roof (at stages late 20 onwards). Posteriorly, the hindbrain tapers into the spinal cord which is very long and thin. Anteriorly the hindbrain is separated from the midbrain by a prominent constriction, the isthmus. The hindbrain shows transiently a series of 7-8 bulges, which are called rhombomeres and are thought to be segmental units of neuronal development (Lumsden, 1990). Most of the work on hindbrain segmentation has been done in chickens and zebrafish and it is unclear how much can be applied to Xenopus (Hartenstein, 1993). Nevertheless, rhombomeres can be seen in sectioned or whole mount immunostained tadpoles, at about st.43, perhapseven earlier (e.g. see tannabin expression figure above). As in other species, the roots of some cranial nerves,show a two-segment relationship with the rhombomeres. Thus, the root of the trigeminal (Vth) root is located in rhombomere 2 (r2), that of the facial (VIIth) in r4 and the

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glossopharyngeal (IXth) in r6 (the full range of cranial nerves is shown in the previous figure showing adult brain morphology). Some prospective structures in this area of the brain can be identified based on gene expression at much earlier stages than it is possible based on morphology. For example, the Engrailed (En) gene is routinely used as a marker for the mib-hindbrain boundary (Hemmati Brivanlou, de la Torre et al., 1991), and Krox-20 as a marker for rhombomeres 3 and 5. Generally speaking however, one should be cautious in using gene expression as a stable cell marker, since expression may be modified between early and late stages. The otocyst, the prospective ear, provides another landmark, which is easy to identify under the dissecting microscope or with Nomarski optics. The otocyst develops from the otic placode and, at st. 27, appears as a small vesicle, adjacent to the middle of the hindbrain. By st. 43, the center of the otocyst lies opposite to rhombomere 4. The midbrain develops dorsally to the posterior aspect of the cephalic flexure and is the primary visual center in frogs; the dorsal part of the midbrain forms the tectum where optic fibers from the eyes synapse. The anterior border of the midbrain is difficult to identify accurately at early stages (st. 20-30). The tract of the posterior commissure (crossing fibers connecting right and left sides) classically defines the dorsal aspect of midbrain-forebrain boundary and can be identified by staining with anti-acetylated tubulin antibody. The epiphysis, or pineal gland, is a small evagination that develops on top of the diencephalon and can be identified by staining with HNK-1 antibody .Hartenstein, (1993) and X-lim3 (Taira, Hayes et al., 1993) expression, starting at st.24 .

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The eyes provide a useful, easily identifiable landmark of the diencephalon (eye development is summarized in the figure below). Around st.22, the future eyes develop as protrusions from the diencephalic wall, the optic vesicles. The optic vesicles invaginate to form

From Gilbert, 1994 double layered optic cups, connected with the ventral diencephalon by long thin tubular structures, the optic stalks. The outer layer of each optic cup forms the pigment epithelium and the inner layer forms the neural retina. Axons of the optic nerve, grow from the retina along the optic stalks, and enter the diencephalon in the chasmiatic ridge. The position where the optic stalks are connected to the brain, the optic recess, defines the ventral aspect of the telencephalic-diencephalic border.The dorsal aspect of this boundary is indicated by a slight depression at mid-20 stages. The telencephalon shows the greatest growth relative to other parts of the nervous system. The cerebral hemispheres begin to evaginate around st. 46 and the part of the telencephalon that receives input from the olfactory neurons forms the paired olfactory bulbs.

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3. Adult morphology Adult morphology and physiology is thoroughly described by Deuchar (1975). For the purposes of dissecting animals for the course, illustrations of the viscera of Xenopus and a diagram of the male urinogenital system are shown below: Xenopus viscera

From Deuchar, 1975 male urinogenital system

From Deuchar, 1975

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4. References Bradley, L.C., Snape, A., Bhatt, S. and Wilkinson, D.G. (1993). The structure and expression of the Xenopus Krox-20 gene: conserved and divergent patterns of expression in rhombomeres and neural crest. Mech. of Devel. 40, 73-84. Doniach, T. (1993). Planar and vertical induction of anteroposterior pattern during the development of the amphibian central nervous system. J. of Neurobiology 24, 12561275. Eagleson, G.W. and Harris, W.A. (1989). Mapping of the presumptive brain regions in the neural plate of Xenopus laevis. 21: 427-440. Hartenstein, V. (1993). Early pattern of neuronal differentiation in the Xenopus embryonic brainstem and spinal cord. J. Comp. Neurol. 328: 213-231. Hemmati-Brivanlou, A., et al. (1991). Cephalic expression and molecular characterization of Xenopus EN-2. Development 111: 715-724. Hopwood, N.D., Pluck, A. and Gurdon, J.B. (1989). A Xenopus mRNA related to Drosophila twist is expressed in response to induction in the mesoderm and the neural crest. Cell 59, 893-903. Jamrich, M. and S. Sato (1989). Differential gene expression in the anterior neural plate during gastrulation of Xenopus laevis. Development 105: 779-786. Keller, R., Shih, J., Sater, A.K., and Moreno, C. (1992a). Planar induction of convergence and extension of the neural plate by the organizer in Xenopus. 193: 218-234. Keller, R. S., J., and Sater, A. (1992b). The cellular basis of the convergence and extension of the Xenopus neural plate. Dev. Dynamics 193: 199-217. Knouff, R. A. (1935). The developmental pattern of ectodermal placodes in Rana Pipiens. J. Comp. Neurol. 62, 17-71. Lumsden, A. G. S. (1990). The development and significance of hindbrain segmentation. Seminars in Developmental Biology 1: 117-125. Nikundiwe, A. M. and Niuwenhuys, R. (1983). The cell masses in the brainstem of the South African clawed frog Xenopus laevis. J. Comp. Neurol. 213, 199-219. Sadaghiani, B., and Thiebaud, C.H. (1987). Neural crest development in the Xenopus laevis embryo, studied by interspecific transplantation and scanning electron microscopy. Dev. Biol. 124: 91-110. Saxen, L. (1989). Neural induction. Int. J. Dev. Biol. 33, 21-48. Sive, H. L., et al. (1989). Progressive determination during formation of the anteroposterior Axis in Xenopus Laevis. Cell 58: 171-180. Slack, , J.M.W. and Tannahill, D. (1992). Mechanism of anteroposterior axis specification in vertebrates: Lessons from the amphibians. Development 114: 285-302.

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Taira, M., et al. (1993). Expression of LIM class homeobox gene Xlim-3 in Xenopus development is limited to neural and neuroendocrine tissues. Dev. Biol. 159: 245-256. General reading Balinsky, B.I. (1981). An Introduction to Embryology, Fifth Edition. Philadelphia, Saunders. Deuchar, E. (1975). Xenopus: The South African Clawed Frog. London, J. Wiley and Sons. Gilbert, S.C. (1994). Developmental Biology, Fourth Edition. Sunderland, MA., Sinauer Hausen, P. and Riebesell, M. (1991). The Early Development of Xenopus Laevis. Berlin, Springer-Verlag. Kuhlenbeck, H. (1973). The central nervous system of vertebrates.Basel, Switzerland: S. Karger Nieuwkoop, P. D. and J. Faber (1994). Normal table of Xenopus laevis. New York, Garland Publishing, Inc. Romer, A.S. (1970). The Vertebrate Body. Philadelphia, Saunders.

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APPENDIX V: Timing of development and temperature dependence Stage 1 2 3 4 5 6 6.5 7

h@22-24ºC 0 1:15’-1:30’ 2h 2:15h 2:45h 3h 3:30h 4h

description 1 cell 2 cell 4 cell 8 cell 16 cell 32 64 tangential cleavage

Stg 24 25 26 27 28 29/30 31 32

h@22-24ºC description 26:15 27:30 29:30 31:15 32:30 tail bud forms 35 37:30 1d 16h

8 9 10

5h 7h 9h

mid blastula-MBT late blastula

33/34 35/36 37/38

1d 20:30h 2d 2h 2d 5:30h

10 1/4 10 1/2 11 11 1/2 12 12 1/2 13 13 1/2 14 15 16 17 18 19 20 21 22 23

10h 11h 11:45 12:30 13:15 14:15 14:45 15:30 16:15 17:30 18:15 18:45 19:45 20:45 21:45 22:30 24 24:45

39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55

2d 8h 2d 18h 3d 4h 3d 8h 3d 15h 3d 20h 4d 2h 4d 10h 5d 12h 7 1/2 d 12d 15d 17d 21d 24d 26d 32d

early gastrula (dorsal lip form)

late gastrula early neurula

neural tube closed

mouth opens

stages from Nieuwkoop and Faber,1967. Timing rules (from Rob Grainger): dependence of developmental rate on temperature 22ºC = 1x 20º = 3/4x 16º = 1/2x 14º = 1/3x

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