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[
Worm Breeder's Gazette,
1988]
My laboratory is interested in isolating maternal and early embryonic mRNAs which segregate with, or are produced in, the two germ line precursor cells of the early nematode embryo. I propose a final assay for putative messages to be in situ hybridization to Ascaris embryos. Because the vitelline membrane of Ascaris is highly impermeable, it has been necessary to use frozen 10 m sections of eggs (sliced eggs!). This has worked well for the probes tested to date. A 1.0 kb subclone from the 26 S subunit of an Ascaris ribosomal gene ( Bach, et al., (1984) NAR 12-3:1313.) has been put into the SP6 transcription vector in both orientations. With the antisense transcript all cells of the paraformaldehyde-fixed embryos hybridize strongly to the hydrolyzed [35S] RNA probe, while the sense strand shows no hybridization above background. In order to determine the resolution of our assay and see if the two germ cells can be selectively hybridized, I have also used an RNA probe from the Ascaris satellite sequence which is lost from all the somatic lineages and retained in the germ line during chromatic diminution (Muller, et al., (1982) NAR 10-23:7493.). With 60 cell embryos (after diminution), the satellite probe shows high grain accumulation in the two primordial germ cells. The pattern seen in Ascaris looks much like the antibody staining with the anti-P granule antibodies in C. elegans, the two germ cells being covered with silver grains in this case. The sensitivity of the hybridization has yet to be tested, as both the ribosomal and satellite probes are abundant in the RNA and germ line genomic DNA, respectively. Helpful, current resource material on in situ hybridization techniques includes: Jorgensen, E. M. and R. L. Garber. Pro Mega Notes 10:2-5 (1987).
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[
Worm Breeder's Gazette,
1993]
We plan to use frozen sections of synchronous Ascaris embryos to test whether cyclin B RNA is localized to the nematode P4 cell as it is to the germline precursor pole cells in Drosophila (see Kreutzer, et al., this volume). To establish whether we could detect a hybridization signal that is localized to a single cell, or a group of cells, we have used various 35S labeled riboprobes. Our first positive results were to the eliminated Ascaris satellite DNA. This DNA is lost from the somatic genome by the 28 cell stage, but retained in the germline. After hybridizing early Ascaris embryos with the satellite probe, only the P4 cell is positive; while at the two fold-stage, the Z2 and Z3 ,cells are detected. To detect RNAs, we first used an antisense probe transcribed from a small subclone of the Ascaris 26S ribosomal RNA gene cluster (Back, M ller, and Tobler 1984. NAR 12,no.3:1333-1346). This RNA probe gave an extremely strong signal in all cells, as expected. At the same time we tested the sense strand of this ribosomal probe, as a negative control. Upon long exposure (~ 1 4 days) this sense strand probe. which we call E3 ,is not negative, but surprisingly, hybridizes specifically to a single cell in the anterior of the 28 cell embryo. Following the localized hybridization through older embryonic stages, this cell appears to be following the lineage pattern of the
hyp-2 cells of C. elegans; therefore we have tentatively identified the one cell detected in the 28 cell embryo as the Ascaris equivalent of the Caenorhabditis ABalpha cell. (Based on the ~100 year old diagrams of the lineage of Ascaris embryogenesis of Boveri, zur Strassen and M ller, the early lineages look similar, if not identical, to those of Caenorhabditis). The signal is very specific and the result is reproducible (3 different experiments). Sequence gazing at the 26S C.elegans ribosomal gene (Ellis et al. 1986. NAR 14no.5:2345-2364) reveals at least three small ORFs of 60-137aa on the opposite strand, which start with a methionine. We will sequence our Ascaris subclone, which is 770bp, and will use it on Northerns of early embryonic RNA, looking for recognition of a small RNA. However because the signal is cell specific, even in young embryos, detecting an RNA that is rare may be difficult. We have yet to show the signal goes away with RNAsing, but the only way we detect DNA hybridization with the satellite probe is after denaturing the chromosomal DNA, which we don't do before hybridizing with E3 .And we haven't tried E3 on mixed stage, sectioned C. elegans embryos yet. The Kodacolor "baby pictures" of these embryos (combined DAPI stained and Nomarski) have been fun to look at, and have caused a stir at the local Photomat. I'd be glad to send copies, and share protocols with others doing in situs. The bottom line is we're now fairly convinced that we'll be able to detect the cyclins, which aren't rare messages, whether or not the RNA is localized.
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[
Worm Breeder's Gazette,
1989]
My major interest is to understand the network of regulation of gene expression in embryogenesis of C. elegans. To attack this problem, I am developing a new methodology for isolating genes which are specifically expressed in a stage and/or some cell(s) of the embryo because I think that such genes must play important roles in embryogenesis. For isolation of differentially expressed genes, cDNA- mRNA subtractive hybridization is very powerful. However, conventional subtraction methods need considerable amount of driver mRNA and the driver mRNA should be isolated from cells as closely related as possible to the cells from which cDNA is made. Therefore, I prefer to develop a new method to make a set of cDNA from single embryos of various stages, amplify them in vitro using PCR (polymerase chain reaction) and then perform subtractive hybridization, rather than to deal with a large amount of synchronized embryos. Outline of the method is as follows; (1) Each embryo that had just completed 1st cleavage was picked up, washed, transferred to an agarose pad and incubated at 22 C for an appropriate period. The developmental stage of the embryo was confirmed on a Nomarski microscope. (2) The embryo was transferred onto an inverted lid from a 0.5 ml eppendorf tube and squashed in GuEST (D.Pilgrim,WBG 10(2),154) buffer. (3) Total RNA was extracted twice by PC1 (ibid.) and precipitated by ethanol using glycogen as carrier. (4) cDNA was made on the RNA using 5'-
tag1(20mer)-oligo(dT)(12)-3' as primer. In this reaction [32P]-dCTP was used at high specific activity (400 Ci/mmole) and the resulting cDNA was stood for 2 days to shorten about 1 kb by [32P] disintegration. This is to ensure that the amplified cDNA represents the original cDNA, since the efficiency of PCR drops as the size of the target DNA increases. (5) After dG tailing (20-30 mer) by Terminal transferase, the cDNA was amplified by PCR using 5'-
tag2(20mer)-oligo(dC)(12)-3' and
tag1 as primers. (6) [32P]-labeled antisense-DNA and 10-fold amount of unlabeled sense-DNA were synthesized by further PCR reaction on thus amplified cDNA made from two different stages of embryos, respectively, using only
tag1 and
tag2 as primer, respectively. They were hybridized using PERT (phenol emulsion reassociation technique) (Kohne et al., Biochem. 16,5329(1977)) to ensure subtractive hybridization even at a low concentration of DNA. (7) The labeled DNA remaining single-stranded was isolated by hydroxyapatite chromotography and used to screen a cDNA library. In a preliminary experiment, I performed two subtractions, namely, 6. 5hr (after 1st cleavage) embryo-3hr embryo and 3hr embryo-2hr embryo. After screening some 5x10+E4 plaques of Ahringer's embryonic cDNA library, I got 5 and 2 very faint signals, respectively. There was no common signal between them, suggesting that the subtraction did work. But the problem is signal intensity. Since an embryo is a mixture of different types of cells, it is anticipated that, even if abundant expression of a gene occurs in some cell(s), the message is still rare in the whole mRNA. Thus, I am developing a method to boost the signal by amplifying the subtracted cDNA using two different sets of primers.
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[
Worm Breeder's Gazette,
1992]
As Dr. R. Horvitz's laboratory asked us for a good protocol for RNA extraction from Ascaris eggs and embryos, other labs might also be interested in this method. The problem with Ascaris is that it has a very rough egg shell; that's why it is rather difficult to obtain RNA from different developmental stages. With the method described below, one can isolate it efficiently. This RNA can be used to perform Northern blots, polyA+ isolation, c-DNA cloning, etc. Collection of eggs from Ascaris lumbricoides var. suum -Decomposition of the uteri and removal of the proteinatious layer of the eggs are done by incubating the uteri overnight at 4 C in 0.5 N NaOH with gentle agitation with a magnetic stirrer. -Let the eggs settle down (ca. 1 hour) or centrifuge them in a swing-out rotor at 7009 for 2 min. Rinse the eggs 5 times in 10 volumes of tap water. At this stage, the eggs are resuspended in 0.1 N H2S04 and can be stored at 4 C for days, months or even years! To obtain different embryonic stages, one incubates the eggs for the desired time with 10 volumes of 0.1 N H2 SO4at 30 C in a large erlenmeyer; the latter, closed by a paper towel, is gently agitated in a rotatory shaker. Removal of the chitinous layer -Resuspend a pellet of eggs in 5 vol. 3.8% NaClO during 1 min. in a 50 ml Falcon tube. -Centrifuge in a swing-out rotor at 7009 during 1 min. -Resuspend the eggs in 5 vol. 3.8% NaClO by inverting the tube 3-4 times. -Leave the tube in a vertical position for 10 min. The eggs should now swim on top of the solution. -Add 1 vol of water and mix. -Centrifuge in a swing-out rotor at 7009 during 1 min. -Rinse the eggs 5 times in 10 volumes of distilled water. Note: between the second and third rinse the non-fertilized eggs burst, giving a white color to the water. RNA extraction: This method is based on Chomczynski's protocol (Analytical Biochemistry 162 (1987), 156-159) -Resuspend the eggs (ca.0.7 ml of packed eggs) in 3 ml of solution D: 4 M guanidinium thiocyanate, 25 mM sodium citrate, pH 7, 0.5% sarcosyl, 0.1 M 2-mercaptoethanol. -Homogenize the eggs in a teflon-glass homogenizer with 1 mg of glass beads (diameter 0.1 mm); 3-4 strokes are sufficient. -Check whether the lysis of the eggs is complete with a microscope -Transfer the solution into a 15 ml Falcon tube. -Add 0.3 ml of 2 M NaAc (pH 4) and mix. -Add 3 ml of water-saturated phenol and mix. -Add 0.6 ml of CHCl3 /Isoamylalcohol 49:1. Mix vigourously during 20 sec. -Chill on ice for 15 min. -Centrifuge for 20 min. in a swing-out rotor at 10 0009 at 4 C. -Resuspend the pellet in 0.7 ml of solution D. Heating at 68 C and vortexing make the resuspension easier. -Transfer the solution into a sterile Eppendorf tube. Precipitate 1 hour at -20 C with 1 volume of isopropanol. -Centrifuge for 15 min. at 4 C. -Wash with 1 ml of 70% ethanol. -Dry the pellet at 68 C. -Resuspend the pellet in 0.1 ml RNAase-free water at 68 C. By using this method, one should obtain a total RNA concentration of about 1 mg/ml from 0.7 ml packed eggs.
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[
Worm Breeder's Gazette,
1987]
Ascaris embryos offer several advantages with regard to studying early nematode development, including the capability of obtaining large populations of synchronous embryos. The fertilized, but uncleaved eggs remain arrested and viable at 4 C for months. With the intention of producing an oocyte and an embryonic cDNA library to screen with our P granule antibodies, Dr. Bennett's new laboratory has recently isolated polyA+ RNAs from Ascaris oocytes and from Ascaris embryos at two stages, 4-8 cells and 16-32 cells. These RNAs have been tested with various probes, including our Ascaris actin gene. All contain actin message, but it had yet to be established whether young Ascaris embryos carry out new zygotic transcription, or rely entirely on maternal messages. We wanted to establish that the embryonic library will not be redundant. To address this question, we have isolated nuclei from Ascaris embryos using the protocol of Dixon et al.(WBG 9:3,73-74). The embryos used were slightly older than those used in the RNA preparations, in the 5-6th cleavage divisions, our reasoning being with more nuclei we'd have a better chance of success with the first isolations. The nuclei are intact, as assayed by diamidinophenylindole (DAPI) staining. We have used this preparation, along with nuclei from a mixed population of Caenorhabditis eggs, in run-on transcription assays, essentially as described by McKnight and Palmiter (JBC 1979, 254:9050). These Ascaris nuclei incorporate significant amounts of label (~35 fold over background). Preliminary results of testing the alpha- amanitin sensitivity of the run-on transcripts show these young Ascaris nuclei have a striking profile, with the transcripts being over 80% sensitive to 1 microgram/ml alpha-amanitin, implying high levels of Polymerase II transcription. After assuring ourselves that we are optimizing conditions for Ascaris (we're currently using those worked out in the Golomb lab for Caenorhabditis), we will assay transcription with specific probes, including actin. We plan to look at earlier embryos; the synchronous Ascaris populations offer the opportunity to profile transcription throughout nematode embryogenesis. And it's on for the embryonic library!
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[
Worm Breeder's Gazette,
1990]
Various protocols for permeabilization and staining of adult C. elegans in such a way as to avoid killing embryos in the uterus have been described (Sebastiano et al Genetics 112 459, AF WBG 9#2, Stone and Shaw WBG 10#3
p27, Candido MRC lore). I have been trying to optimize this procedure to obtain strong, well localized staining of lines carrying -gal fusion proteins, to allow for selection of altered staining patterns after mutagenesis. A current protocol is described below. The treatment is somewhat more drastic than previous methods, but allows excellent staining and reasonable preservation with survival of 3-5 embryos per stained adult. 1. Pellet animals from H20 and resuspend in 50% acetone for 10-15 min at room temp (the time for this step is critical...too long and embryos are inviable, too short and the adults fail to permeabilize and awaken in the staining solution.). 2. Wash animals 3x in PMB and resuspend in staining mix. Stain at room temperature. The hatched animals can survive within the adult in staining solution for several hours, but eventually die or free themselves, limiting staining time to about 12 hr. 3. Scoring with a dissecting microscope is best with epi-illumination over a white background PMB: 50mM NaPi pH 7.5, 1mM MgCl2 Staining Mix: PMB + 10mM KFerrocyanide + 10mM KFerricyanide + .004% SDS + 10 l/ml of 12.5% XGAL in DMF (XGAL stock should be stored at - 70 C) Optimal permeabilization so far has been around 95% of animals. Thus the technique could be used to screen for ectopic expression or expression of an otherwise inactive construct, but loss of expression would be difficult to screen for. N.B. Some permeabilization of pharyngeal muscle occurs without the acetone step (putting animals directly into staining solution).
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[
Worm Breeder's Gazette,
1982]
I have used the following procedure to stain C. elegans embryos histochemically for AChE activity. 1. On a microscope slide subbed with a 1% BSA solution, place a few young adults into a 20 l drop of 2% glutaraldehyde solution in 100 mM maleate buffer, pH 6.0. The animals are cut with a scalpel blade at the vulva to release the embryos. 2. Very gently, a 16 X 16 mm coverslip is placed on top of the buffer drop. The slide is then immersed into a liquid nitrogen container for about 15 seconds, and the coverslip is quickly pried off with a scalpel blade. 3. Another 16 X 16 mm coverslip with its two opposite edges coated with grease is placed over the embryos. The slide is incubated in a humid chamber at O C for two hours. 4. About 10 l of the staining solution ( containing 10 mg acetylthiocholinechloride, 8.8 ml 100 mM maleate buffer, pH 6.0. 1ml 50 mM sodium citrate, 100 l 300 mM copper sulfate, and 100 l potassium ferrocyanide, added in this order) is applied at the open edge of the coverslip, and is absorbed at the other open edge with an absorbant paper ( see fig.). This step is repeated 3-4 times to ensure that all embryos have been immersed in the staining solution. The slide is then incubated in a humid chamber at room temperature for about 6 hours before light microscopy. A reddish-brown stain appears in the regions showing AChE activity. [See Figure 1] The above procedure can also be used for larvae and the adult animals. Significantly, this method reveals the same areas of AChE activity in C. elegans as in the method reported by Culotti et al. ( Genetics: 97, 281-305,1981), in which the animals are treated with 95% acetone for 3 minutes. The prominent areas showing the stain for AChE activity are, the nerve ring, lateral ganglion, ventral nerve cord, dorsal nerve cord, pre-anal ganglion, and the pharryngeo-intestinal valve cells.
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[
Worm Breeder's Gazette,
1993]
SAMPLE PREPARATION-Egg Preps The Bennett laboratory uses the same protocol Ascaris or Caenorhabditis, although we currently have a lot more experience with Ascaris. We use the hypochlorite solution (40% Chlorox, 50% 1 N NaOH. 10% water) for gravid C. elegans adults or synchronous cultures of Ascaris eggs. When adults have dissolved and eggs have been stripped of their proteinaceous coat, rinse 3-4 times in M9 (or water). Transfer egg pellet to 1.5ml eppendorf tube. Add equal volume of 6:3:1 (methanol:acetic acid:chloroform) to permeabilize vitelline membrane. Invert. Immediately dilute in M9 or water and spin down eggs. Rinse several times. Fix in 4% paraformaldehyde in M9 or PBS (in DEPC water) on ice for 15 min. Rinse 2-3 times. A few embryos should be checked with DAPI at this point to assure they are intact and permeabilized. Mix fixed egg pellet with ~1/3 volume of O.C.T. embedding medium for frozen specimens (Tissue-Tek brand). Freeze the egg mixture on a microtome chuck (or a quarter) which has been pre-chilled on a block of dry ice. These egg blocks are stored at -80 C. until cutting. We have stored blocks for several weeks. SLICING EGGS We set the microtome for 12-16 m sections at -15 C. We fill a slide with sections and let the sections adhere to the slide on a 37 C hot plate for 30 min. After that the slides are fixed two more times in 4% paraformaldehyde (made fresh) in DEPC M9 or PBS + Mg. The first fixation is on ice. The second is at room temperature with the addition of 0.1% deoxycholate and 0.1% Triton-X100. All fixations are done in glass containers which have been baked in the "hot oven"(250 C for >4 hrs.) to remove RNases. Both fixations are for 15 min. The cellular RNA is acetylated in acetic anhydride/triethanolamine solution. (freshly made and used within 30 minutes) at room temp. for 5 min. 100mM Triethanolamine-HCl pH 7.5 250ml 250 ml DEPC-H(2)0 4.64g Triethanolamine-HCL (Sigma T-1502) 560 I 1ON NaOH add 625 l acetic anhydride immediately before adding to the slides Slides are then rinsed 3 times in 1xPBS+Mg or M9 .Slides are dehydrated in 30%,60%,80%,95% ETOH made in DEPC-H(2)0. We air dry them O/N. Store at -80 C until ready for hybridization. TRANSCRIPTION-ANTISENSE The transcription and hybridization protocols are from Dr. Etsuko Wada, courtesy of Dr. Sandra Petersen, UMC. Some comments may be useful. The lack of any cold nucleotide to supplement the hot label makes the reaction suboptimal in terms of molarity, and causes the probe to be very small (probably 50-75nt). However incorporation is often 95% or better and this small probe doesn't need hydrolysis. Dry down 250 uCi [35S]-CTP in speed-vac (CTP may also be used, cold UTP would then be added.) ADD: 1. 2ul 5X Transcription Buffer (we use that which comes with SP6 ,T7 or T3 polymerase) 2. 1 l 100mM DTT (Made fresh) 3. 0.5 l RNasin 40U/ l 4. 2 l 1OmM ATP 5. 2 l 1OmM UTP (or CTP when using hot UTP) 6. 2 l 1OmM GTP 7. 1 l Linearized DNA Template 1 g/ l 8. 0.5 l RNA Polymerase 20U/ l Gently mix. (Tap tube with finger.) Incubate 30min at 37 C. Add 0.5 l Polymerase and incubate another 30min. Add 89.5 l nuclease-free water. Check % incorporation with PEI chromatography paper. ([35S] count in fluor) Add: 1. 5 l 1 M Tris-HCL pH 8 2. 1 l 1M MgCl(2) 3. 0.5 l tRNA 25 g/ l 4. 0.5 l RNasin 40U/ l 5. 2ul DNase I 1U/ l Gently mix. (DNase is very labile. Do not vortex.) Incubate 30min at 37 C. Run Sephadex G-50 spin column to remove unincorporated counts.
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[
Worm Breeder's Gazette,
1988]
We have exploited some of the unique features of embryos of the parasitic roundworm, Ascaris des to observe transcriptional patterns in nematode early embryogenesis. In contrast to the free-living nematode Caenorhabditis ble to obtain large synchronous populations of Ascaris embryos at various stages of development. It has been reported in C. elegans that transcription is detectable at approximately the 100-cell stage. [Hecht, et al., (1981). Dev. Biol. 83:374-379]. More recent work in C. elegans suggests that transcription is occurring at or before the 30-cell stage. [Schauer, et al., WBG 10:1, 72-73.]. Our laboratory has recently reported that Ascaris embryos in the 5-6th cleavage stage are actively transcribing. [Dalley, et al., WBG 10:1, 76.]. From staged embryos we have isolated nuclei to determine run-on transcription characteristics. Initial studies performed on nuclei isolated by the method of Dixon et al. [WBG 9:3, 73-74.] resulted in contamination of the embryonic preparations with mitochondria, as seen by microscopic examination after diamidinophenylindole (DAPI) staining, as well as a strong hybridization signal using a mitochondrial probe. Subsequent purification of nuclei on 50% Percoll gradients has yielded about a 100-fold visual reduction in mitochondria, although a hybridization signal can still be detected. Run-on transcription assays were performed on nuclei isolated from staged embryos of 4-8 cell, 24-30 cell, ~60 cell, and 10 day embryos (about 600 cells). Parallel assays were carried out with and without alpha-amanitin present at 1 g/ml a level which inhibits RNA polymerase II activity. The results are shown on Fig. 1 as counts of [3H]-UTP incorporated per 10 l reaction. Total levels of incorporation were typically 10-35 fold over background, which has been subtracted from each of the values shown. The darkened portion of each bar represents the alpha-amanitin sensitive (RNA polymerase II) fraction of total transcription. We consistently detect RNA polymerase II activity as early as the 4-8 cell stage and by the 30 cell stage active mRNA transcription is occurring. When DNA content of preparations was measured by fluorimetric assay the highest level of incorporation per pg DNA was at the 30 cell stage. Similar transcription reactions were carried out with [32P]-UTP, with the labelled RNAs used to probe duplicate dot blots containing cold DNA probes. Results from the 60-cell stage showed no hybridization to the SP6 vector or the Ascaris sperm- specific MSP cDNA. Ascaris ribosomal and actin probes showed significant hybridization (Fig. 2). When the labelling was carried out in the presence of 1 I/ml alpha-amanitin, the actin signal was eliminated, while the ribosomal signal was retained as expected. This shows that we are able to detect specific mRNA transcription occurring in staged embryos. Similar studies are being carried out with each of the mentioned cell stages to establish if specific messages, i.e., actin for now, can be detected at the 4-8 cell stage. We propose that differential library screening using staged early vs. Iate RNA populations will yield transcripts unique to the early embryo. The following two references have been valuable for run-on transcriptions: J. R. Nevins, Methods in Enzymology, (1987), 172:235-241. M. E. Greenberg, Current Protocols in Molecular Biology, (1987), 4. 10.1-4.10. 7
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[
Worm Breeder's Gazette,
1980]
We have continued experiments on the cleavage patterns of partial embryos and isolated blastomeres obtained by crushing early N2 eggs in embryonic culture medium supplemented with Ascaris coelomic fluid (W.B. G. June 1978). Neither isolated somatic precursor cells (blast cells) nor isolated P cells give rise to the spatial patterns formed by their progeny in intact embryos. However, all our results so far are consistent with the following simple hypothesis. The blast cells, and their progeny, are internally determined to cleave in a simple helical pattern. The P cells are internally determined to cleave linearly, maintaining the spindle orientation of preceding P-cell divisions parallel to the anterior-posterior axis. In the intact embryo, interactions of cells with each other (and perhaps with the eggshell) combine with these two intrinsic cleavage patterns to dictate normal cellular organization, which therefore depends upon both extracellular and intracellular cues. Cleavage of an isolated P2 blastomere results in an approximately linear chain of cells in the order C, C, D, P4. This result, together with the approximately anterior-posterior orientation of the preceding cleavages that give rise to the AB, EMSt, E, and MSt cells suggests the possibility that in the zygote, determinants for blast-cell primary differentiation are arranged along the posterior-anterior axis in the order C (ectoderm), D (mesoderm), P4 (germ line), E (endoderm), MSt (mesoderm), AB (ectoderm). This arrangement corresponds to that of a fate map proposed for the Ascaris des zygote by Zur Strassen (1896). It differs from that of the fate map proposed for the C. elegans zygote by Schierenberg (1978), who postulated the order P4, D, C, E, MSt, AB based on observations of division orientation in intact embryos,cytoplasmic streaming,and cell movements during early cleavage.