We thank Rayhan Arif and Hassaan Shahawy for camera work, Tripti Gupta and Frederike Alwes for assistance refining the cell ablation technique, and Extavour lab members for feedback on the data, video and manuscript. This work was partially supported by the Harvard Stem Cell Institute (Seed Grant number SG-0057-10-00) the Ellison Medical Foundation (New Scholar Award number AG-NS-07010-10) to CGE, and Harvard College Research Program awards to ARN.

Comments that may be helpful in executing certain steps are indicated in italics.
1. Day 1: Preparation of Materials
<ol>
	<li>Prepare the following materials (see Table of Materials and Equipment):
	<ul>
		<li>15 cm and 22 cm Pasteur pipettes</li>
		<li>Diamond scribe</li>
		<li>filtered artificial seawater (salinity between 0.0018-0.0020) containing 1 mg/ml Amphotericin B (1:100 of a 100 mg/ml stock solution), 100 units/ml penicillin and 100 mg/ml streptomycin (1:50 of a stock solution containing 5,000 units/ml of penicillin and 5,000 mg/ml of streptomycin)</li>
		<li>filtered artificial sea water (salinity between 0.0018-0.0020) without additives</li>
		<li>a large plastic container for housing couples (at least 15 cm x 15 cm x 5 cm)</li>
		<li>small (3 cm or 5 cm) Petri dishes lined with Sylgard</li>
		<li>small (3 cm or 5 cm) Petri dishes</li>
		<li>a watchglass or glass multi-well plate</li>
		<li>forceps (we use blunt forceps derived from old Dumont #5 forceps that were accidentally damaged in other laboratory procedures, which we have repurposed by using a forceps sharpening stone to ensure that the ends of the forceps are smooth, that both tines are the same length, and that they no longer have sharp points. Extremely fine forceps, such as new Dumont #5 forceps, are undesirable as they may injure embryos and/or females.)</li>
		<li>a Pasteur pipette with the end widened (see step 1.2 below)</li>
		<li>an embryo retrieval tool consisting of a thin, curved tungsten wire embedded into the thin end of a Pasteur pipette (could be replaced by an alternative implement; see 1.3 below)</li>
		<li>a mouth pipette with a small opening (see step 1.4 below)</li>
		<li>a glass needle made from a pulled glass capillary (see step 1.5 below)</li>
	</ul>
	No materials toxic to humans are used in this protocol. However, researchers should ensure that they wear gloves or other protective attire as required by the risk management guidelines of their institution.</li>
	<li>To make the widened Pasteur pipette, first score around a 15 cm Pasteur pipette at the point where the pipette begins to narrow with a diamond scribe, wrap the scored region of the pipette in a Kimwipe or paper towel to protect the fingers, and then carefully break off the tip at the score line. Hold the broken end of the pipette over a Bunsen burner flame for several seconds to smooth the edges. The opening should be slightly narrower than the maximum width of the pipette.</li>
	<li>To make the tungsten wire dissecting tool, insert a tungsten wire into the end of a 15 cm glass Pasteur pipette, and hold briefly over a Bunsen burner flame to melt the glass, fixing the wire in place. Use forceps to gently curve the end of the wire into a hooked shape. See Figure 3 for an example of an appropriate shape for this tool.</li>
	<li>To make a small opening for the mouth pipette, pull the thin end of a 22 cm Pasteur pipette into two parts over a flame, and break off the tip of the small end. Insert into the end of the mouth pipette holder.</li>
	<li>Make the glass needle that will be used to puncture the blastomeres of interest during the ablation procedure using a needle puller. A suitably shaped needle is shown in Figure 4. This needle was pulled on a Sutter P97 needle puller using a program with parameters 2x(heat = 566, pull = 100, velocity = 20, time = 250) + 1X(heat = 566, pull = 100, velocity = 100, time = 250). The specific program and instrument used to make the needle can vary, but the needle must have a fine point, yet also be sturdy enough not to break when pushed against the chorion of the embryo.</li>
</ol>
2. Day 1: Gathering Parhyale&#160;Mating Couples
<ol>
	<li>Use the widened Pasteur pipette (step 1.2 above) to collect mating P. hawaiensis couples from the large culture container, and transfer them to a separate container containing clean artificial seawater. It is not necessary that this seawater be filtered. It is best to collect mating pairs in the later afternoon and leave the couples at room temperature (20-23 &#186;C) overnight, so that the next morning, most embryos will have been recently fertilized and deposited, and thus at early cleavage stages suitable for ablation. Gather at least 20 mating pairs to ensure that some females will be carrying early cleavage stage embryos by the following morning.</li>
</ol>
3. Day 2: Gathering Parhyale&#160;Embryos
<ul>
	<li>The next morning, infuse filtered artificial seawater (FASW) with CO2. Some of the females will have swum free from their males during the night; collect these separated females and anesthetize them in FASW / CO2. The remaining unpaired males can be removed from the mating pair container and returned to the large culture. Remaining mating pairs can be saved for embryo collection later in the day or the next day.</li>
	<li>Transfer a single anaesthetized female carrying embryos to a watch glass in FASW / CO2. All separated females will likely have deposited eggs, which will be visible by eye as pale pink or purple spheroids through the transparent coxal plates of the females (Figure 1A). To determine if the embryos are at early cleavage stages and thus appropriate for blastomere ablation, the researcher will have to remove the embryos from the brood pouch and examine them under a microscope.</li>
	<li>Viewing the procedure under a dissecting microscope, grasp the coxal plates along one side of the body with forceps. Embryos will be visible in the ventral brood pouch between these coxal plates (Figure 1B).</li>
	<li>Insert the thin, curved wire tool (Figure 3; step 1.3 above) into the posterior end of the brood pouch, and gently lift the wire towards the anterior end of the brood pouch to separate the brood pouch coverings (these are modified ventral appendages called oostegites30), opening the pouch and loosening the embryos. Embryos that are within the first hour or so of being laid are all enclosed within a very thin, transparent membrane that will easily be disrupted by the wire tool; this membrane disappears by approximately 2-3 hr after egg laying, and embryos are not bound to each other or to the female in any way from this point onwards throughout embryogenesis. If researchers find it inconvenient or difficult to maneuver the curved wire tool to remove embryos from the brood pouch, an alternative implement may be used, as long as the movements are gentle and no harsh pressure is applied to the embryos or the brood pouch. Other tools appropriate for removing embryos from the brood pouch include blunted forceps or a glass capillary with a sealed and smoothed end.</li>
	<li>Use an unaltered 15 cm Pasteur pipette to flush water through the opened brood pouch, pushing out the embryos, which should settle to the bottom of the watch glass. Transfer the female to clean FASW (without CO2) to allow recovery before reintroducing her to the main culture. Multiple females can be placed into the same recovery dish, but allow females to recover fully before returning them to the main culture, as they may be eaten by other animals if they are still partially anesthetized.</li>
	<li>Use an unaltered 15 cm Pasteur pipette to transfer the embryos to a Petri dish with clean FASW, and view under the microscope to determine their embryonic stage. Separate embryos that are at the third cleavage (stage 4&#160;5), which are suitable for this procedure. It should also be possible to apply this protocol to any cleavage stage prior to germ disc formation (stages 7-8&#160;5).</li>
</ul>
4. Day 2: Ablating Single Cells
<ol>
	<li>Fill Sylgard-lined Petri dish with a shallow layer of FASW. Use an unaltered 15 cm Pasteur pipette to transfer a single embryo to the plate.</li>
	<li>Using blunt forceps, gently roll the embryo to identify the cell to be ablated (Figures 2A and&#160;2B). The germ line precursor g can be identified as the smallest micromere, which sits on top of the smallest macromere Mav. Additionally, g does not directly contact the cell en, which is the micromere directly across from it, as the mesoderm precursor micromeres ml and mr share a cell border in the middle of the embryo. mr is the blastomere to the right of g, and ml is the micromere to the left of g. The sister macromeres of mr, ml, and en are the ectoderm precursors Er, El, and Ep respectively.</li>
	<li>With forceps in the left hand if the researcher is right-handed (or in the right hand if the researcher is left-handed), orient the embryo with the cell of interest to the right (or to the left if the researcher is left-handed), and gently grasp the embryo between the forceps to stabilize it.</li>
	<li>Using a pulled glass needle (step 1.5 above; Figure 4), gently puncture the cell of interest. Do not insert the needle too far, so as not to push the needle into the cells neighboring or below the cell of interest. The needle only has to puncture the chorion and underlying cell membrane, and does not need to extend deeply into the cell to be ablated.</li>
	<li>Exchange the needle used to puncture the embryo for the glass end of a mouth pipette with a fine pulled Pasteur pipette tip (step 1.4 above). Hold the tip of the pipette close to the hole created in the cell of interest, and apply very gentle suction.</li>
	<li>Very gently squeeze the embryo with the forceps. As the content of the blastomere is pushed out of the chorion through the hole created in step 4.4, suck it into the mouth pipette to allow an unobstructed view of the embryo. If the hole is large enough, the nucleus of the punctured cell may be seen to emerge as well. This is a good check to make sure the entire cell contents have been removed and cannot be regenerated.</li>
	<li>Carefully observe the embryo as the blastomere of interest is removed. The border of the punctured cell will recede towards the hole in the chorion, making the intersections of the underlying cells visible. Continue applying pressure until all of the blastomere of interest has been removed. Use forceps to roll the embryo from side to side and visually confirm that all of the blastomere of interest is gone.</li>
	<li>Using an unaltered 15 cm Pasteur pipette, transfer the embryo to clean FASW+ (FASW + 1 mg/ml Amphotericin B, 100 units/ml penicillin and 100 mg/ml streptomycin).</li>
	<li>Raise blastomere-ablated embryos in individual wells of a clean 48-well plate in FASW+, changing 50% of antibiotic-supplemented FASW+ daily. In our hands embryos survive and develop well, even following ablation, at a range of temperatures from 18-28 &#186;C. Researchers should raise their embryos at a temperature for which they have a clean (but not necessarily sterile) surface on which to raise the embryos with minimal disturbance. To be able to compare the timing of developmental events in blastomere-ablated embryos with controls, we refer the reader to the detailed developmental staging information that is available for embryos raised at 18 &#186;C4, 25 &#186;C4, and 26 &#186;C5,6.</li>
</ol>

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The authors have nothing to disclose.

We describe a protocol for manual ablation and complete physical removal of single blastomeres from early cleavage stages of the amphipod P. hawaiensis. We demonstrate use of this protocol by removing the single germ line precursor cell g from an eight-cell stage embryo, and show that ablation has been successful by confirming absence of g&#8217;s daughter cells in later embryogenesis. This protocol can be used to remove any of the micromeres from the embryo at early cleavage st...

The amphipod crustacean Parhyale hawaiensis has emerged over the last decade as a promising model organism with great potential for use in evolutionary developmental biology research1. Among the arthropods, most model systems are insects, and the most extensively studied of these is the fruit fly Drosophila melanogaster. D. melanogaster is a member of the insect order Diptera, and as such displays many embryological features that are derived with respect to those of basally branching insects2. Moreover, insects are nested within the subphylum Pancrustacea3, meaning that insects have their closest relatives within the long-standing &#8220;natural&#8221; group called pancrustaceans, and that this group is paraphyletic. This suggests that in addition to basally branching insect models, studies of other crustaceans are required to gain a broader view of the evolutionary history of the developmental traits and molecular mechanisms that have been so well studied in D. melanogaster. However, very few crustaceans have been well established for experimental laboratory analysis of development. The amphipod P. hawaiensis is a highly tractable laboratory model system, amenable to a range of experimental techniques. Amphipods display many unique features within their parent superorder Peracarida (beach hoppers, scuds, and well shrimps), and are therefore thought to be relatively derived within this group of crustaceans. Nevertheless, the relative ease of embryological and functional genetic manipulation offered by Parhyale make this amphipod a valuable addition to the current inventory of model organisms.
As a laboratory animal, P. hawaiensis offers many advantages. Animals are tolerant to a wide range of temperatures and salinities, and survive well in large cultures of artificial seawater1. It is easy to distinguish between males and females based on clear morphological differences, most notably, the large, hooked, anterior trunk appendages that males use to grasp the females during mating. For embryological and developmental work, P. hawaiensis has several very appealing features. Embryogenesis lasts approximately 10 days and the time to sexual maturity is approximately six weeks at 28 &#186;C (but note that Parhyale survives well at temperatures ranging from approximately 20-30 &#186;C, and that detailed developmental staging information is available for embryos raised at 18 &#186;C4, 25 &#186;C4, and 26 &#186;C5,6). Adults mate all year round in the laboratory, so embryos are available at any time of year. Females lay 2-20 (depending on the age of the female) fertilized eggs into a ventral brood pouch located between the first several pairs of legs (Figures 1A and&#160;1B), and it is possible to gather these embryos very early in development without killing the female or damaging the embryos (Figure 1C). The embryos survive in filtered artificial sea water through to hatching, can be fixed for subsequent gene expression or histological analysis7, and a detailed staging table allows accurate identification of the progress through development5. Robust protocols have been used to perform gene expression analysis by in situ hybridization8-15 or immunostaining4,16,17, functional knockdown by RNA interference13,15 or morpholinos12, and stable germ line transgenesis18. Using the transgenesis system, inducible expression14 and enhancer trap19 methods can also be used to investigate gene function in P. hawaiensis. While a publicly available genome sequence is not currently available, a transcriptome containing transcripts produced during oogenesis and embryogenesis has been de novo assembled and annotated20, and deposited in a searchable database21, facilitating gene discovery. In sum, P. hawaiensis is a highly tractable model organism suitable for multiple experimental and genetic approaches to understanding development.
Unlike the early syncytial cleavages of D. melanogaster, P. hawaiensis embryos cleave holoblastically following fertilization (Figure 2A). Lineage tracing analysis has shown that by third cleavage, each of the third cleavage blastomeres is specifically fated to give rise to one of the three germ layers or the germ line6 (Figure 2B). These data, together with microarray data22, cell lineage analyses6,23, and blastomere isolation experiments4 have suggested that developmental potentials are segregated to at least some third cleavage blastomeres by asymmetric inheritance of cell fate determinants. Accordingly, in blastomere ablation experiments in which the germ line precursor (termed &#8220;g&#8221; in the Parhyale cell lineage nomenclature6) was removed at the eight cell stage, embryos lacked germ cells at later developmental stages4, as indicated by the absence of cells expressing the protein Vasa, which is a germ line marker in most metazoans24. In contrast, somatic blastomere ablation experiments showed that P. hawaiensis embryos also possess significant regulatory capabilities, such that the fates of mesoderm or ectoderm precursor blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres25. How regulative cell fate replacement can occur, and the extent of autonomous cell fate adoption by somatic blastomeres, remains unknown. Experimental embryological techniques such as blastomere ablation can be useful in understanding the relative autonomy and nonautonomy of cell fate decisions26,27&#160;and are therefore of interest in the study of P. hawaiensis embryogenesis.
In the experiments that demonstrated regulative replacement of ectodermal and mesodermal lineages, blastomere ablation was performed by injection28 and subsequent excitation of phototoxic dyes25. While this technique is effective at killing the injected blastomere(s), it does not completely remove the dead cell body from the embryo. In addition, differences have been observed between cell lineage data gathered through to gastrulation stages of embryogenesis by injecting blastomeres with fluorescent lineage tracers28,29, and data gathered by following unperturbed blastomeres through development of the same embryonic stages23. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications.
We previously published the results of cell lineage analyses of embryos in which single cells were manually ablated23. However, the delicate operations required to remove single blastomeres from early cleavage stage embryos have not yet been fully described. Here we present a protocol for collection of P. hawaiensis embryos and manual ablation of a single blastomere from an eight-cell stage embryo. The goal of this method is to achieve complete removal of the cell body from the embryo, allowing observation of the cellular behaviors and cell fate competencies of the remaining cells during embryogenesis and post-embryonic development. Our protocol shows removal of the germ line precursor g (Figure 2C), but can be applied to any cell at the eight-cell stage, or to blastomeres of earlier cleavage stages. In principle, this protocol could be applied to remove single cells from early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

<table border="0" cellpadding="0" cellspacing="0" width="1060">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">15 cm Pasteur pipette</td>
			<td style="width:164px;">Wheaton</td>
			<td style="width:123px;">53499-630</td>
			<td style="width:495px;">For transferring intact and blastomere-ablated embryos from dish to dish. 
			VWR</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:164px;">22 cm Pasteur pipette</td>
			<td style="width:164px;">Wheaton</td>
			<td style="width:123px;">53499-632</td>
			<td style="width:495px;">For making mouth pipette tips. 
			VWR</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">3 cm petri dishes</td>
			<td style="width:164px;">Thermo Scientific</td>
			<td style="width:123px;">25382-334</td>
			<td style="width:495px;">Use some unaltered to collect and store embryos; coat some with a layer of Sylgard (see below) to create a working surface for blastomere ablations. 
			VWR</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:164px;">48-well plates</td>
			<td style="width:164px;">Greiner Bio-One</td>
			<td style="width:123px;">82051-004</td>
			<td style="width:495px;">For culturing embryos following ablation. 
			VWR</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Bottle top filters 0.2 micron</td>
			<td style="width:164px;">Nalge Nunc International</td>
			<td style="width:123px;">28199-296</td>
			<td style="width:495px;">For creating FASW (See below) 
			VWR</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:164px;">Bunsen burner</td>
			<td style="width:164px;">VWR</td>
			<td style="width:123px;">89038-530</td>
			<td style="width:495px;">For creating tungsten wire tool and fine tip of mouth pipette. 
			VWR</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:164px;">Diamond scribe</td>
			<td style="width:164px;">Musco Sports Lighting</td>
			<td style="width:123px;">52865-005</td>
			<td style="width:495px;">For creating wide-mouthed Pasteur pipettes for collecting couples. 
			VWR</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:164px;">Forceps</td>
			<td style="width:164px;">Fine Science Tools</td>
			<td style="width:123px;">11050-10</td>
			<td style="width:495px;">For holding anaesthetized females during removal of embryos from brood pouch. These do not have to be fine-tipped Dumont forceps - the tips can be at least as blunt as that in the forceps for which the catalogue number is listed here. 
			Fine Science Tools</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Fungizone-Amphotericin B</td>
			<td style="width:164px;">Sigma</td>
			<td style="width:123px;">A2942</td>
			<td style="width:495px;">Add to FASW to a final concentration of 1mg/ml, and use this solution to culture blastomere-ablated embryos. 
			Sigma Aldrich</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">Glass capillaries 4 inches long, 1/0.58 OD/ID (mm)</td>
			<td style="width:164px;">World Precision Instruments</td>
			<td style="width:123px;">1B100-4</td>
			<td style="width:495px;">need to include glass capillary and needle puller machine? 
			World Precision Instruments</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">Glass watchglass 300 mm diameter</td>
			<td style="width:164px;">Electron Microscopy Sciences</td>
			<td style="width:123px;">70543-30</td>
			<td style="width:495px;">For use in removing embryos from the brood pouch of anaesthetized females. 
			Electron Microscopy Sciences</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Instant Ocean Artificial Sea Water (ASW)</td>
			<td style="width:164px;">Instant Ocean</td>
			<td style="width:123px;">IS160</td>
			<td style="width:495px;">Make ASW by dissolving salt in deionized water to a salinity of X. 
			Aquatic EcoSystems</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">Instant Ocean Filtered Artificial Sea Water (FASW)</td>
			<td style="width:164px;">Instant Ocean</td>
			<td style="width:123px;">IS160</td>
			<td style="width:495px;">Use 0.2 micron filters to sterilize ASW. Make up in small amounts (&#60; 250 ml) as needed. 
			Aquatic EcoSystems</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Kimwipes</td>
			<td style="width:164px;">Kimberly-Clark</td>
			<td style="width:123px;">21905-026</td>
			<td style="width:495px;">For safely breaking off the fine end of Pasteur pipettes modified for collecting couples. 
			VWR</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Mouth pipette adaptor</td>
			<td style="width:164px;">Drummond Labware</td>
			<td style="width:123px;">A5177-5EA</td>
			<td style="width:495px;">Insert the fine pulled pasteur pipette tip into this end, to be used for removing extruded cell contents during the ablation procedure. 
			Sigma Aldrich</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Mouth pipette tubing</td>
			<td style="width:164px;">Aldrich</td>
			<td style="width:123px;">Z280356</td>
			<td style="width:495px;">Cut this to be long enough to comfortably hang around your neck during the ablation procedure. 
			Sigma Aldrich</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">Needle Puller</td>
			<td style="width:164px;">Sutter Instrument Company</td>
			<td style="width:123px;">P97</td>
			<td style="width:495px;">For making needles from glass capillaries for puncturing the blastomere to be ablated. 
			Sutter Instrument Company</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">Penicillin-Streptomycin Solution</td>
			<td style="width:164px;">Mediatech</td>
			<td style="width:123px;">45000-650</td>
			<td style="width:495px;">Add to FASW to a final concentration of 100 units/ml penicillin and 100 mg/ml streptomycin, and use this solution to culture blastomere-ablated embryos. 
			VWR</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Rubber bulb</td>
			<td style="width:164px;">Electron Microscopy Sciences</td>
			<td style="width:123px;">100488-418</td>
			<td style="width:495px;">For using Pasteur pipettes to transfer embryos from dish to dish. 
			VWR</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:164px;">Sylgard 184</td>
			<td style="width:164px;">K. R. Anderson, Inc.</td>
			<td style="width:123px;">NC9659604</td>
			<td style="width:495px;">Make up according to manufacturer&#39;s instructions. Pour a layer 2-5 mm thick into a 3cm petri dish to create a working surface for blastomere ablations. 
			Fisher Scientific</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:164px;">Tungsten wire 0.004 inches diameter</td>
			<td style="width:164px;">A-M Systems</td>
			<td style="width:123px;">719000</td>
			<td style="width:495px;">Use this to make a tool for removing embryos from the brood pouch of anaesthetized females. 
			A-M Systems</td>
		</tr>
	</tbody>
</table>

ablation of a single cell from eight-cell embryos of the amphipod crustacean parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a promising model organism for laboratory studies of development, providing a useful outgroup comparison to the well studied arthropod model organism Drosophila melanogaster. In contrast to the syncytial cleavages of Drosophila, the early cleavages of Parhyale are holoblastic. Fate mapping using tracer dyes injected into early blastomeres have shown that all three germ layers and the germ line are established by the eight-cell stage. At this stage, three blastomeres are fated to give rise to the ectoderm, three are fated to give rise to the mesoderm, and the remaining two blastomeres are the precursors of the endoderm and germ line respectively. However, blastomere ablation experiments have shown that Parhyale embryos also possess significant regulatory capabilities, such that the fates of blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres. Blastomere ablation has previously been described by one of two methods: injection and subsequent activation of phototoxic dyes or manual ablation. However, photoablation kills blastomeres but does not remove the dead cell body from the embryo. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications. Here we present a protocol for manual removal of single blastomeres from the eight-cell stage of Parhyale embryos, illustrating the instruments and manual procedures necessary for complete removal of the cell body while keeping the remaining blastomeres alive and intact. This protocol can be applied to any Parhyale cell at the eight-cell stage, or to blastomeres of other early cleavage stages. In addition, in principle this protocol could be applicable to early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

Following successful ablation of single third cleavage P. hawaiensis micromeres as described in this protocol, the remaining micromeres gradually shift their positions slightly so as to partially occupy the space formerly occupied by the ablated blastomere. For example, when g is removed, the neighboring blastomeres mr and ml shift slightly and come to share the lateral cell borders that were formerly in direct contact with g (compare&#160;<stro...

Watch this Scientific Journal Video about Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis at JoVE.com

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a promising model organism for studies of crustacean embryology and comparative arthropod development and evolution. This protocol describes a method for manual removal of single blastomeres from early cleavage stage embryos of Parhyale.

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a ...

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12.5K Views.  Harvard University. The amphipod Parhyale hawaiensis is a promising model organism for studies of crustacean embryology and comparative arthropod development and evolution. This protocol describes a method for manual removal of single blastomeres from early cleavage stage embryos of Parhyale.

Video: Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

Dissection of Organizer and Animal Pole Explants from Xenopus laevis Embryos and Assembly of a Cell Adhesion Assay

This video demonstrates the technique used for preparation of organizer and animal pole explants from Xenopus laevis embryos, including the use of the eyebrow knife - a specialized dissection tool made of one's eyebrow. The protocol for assembling an adhesion assay is also given, which probes for the presence of key adhesion molecules present on the surface organizer or animal pole cells that are critical for proper development.

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Preparation of Single-Cell Suspensions from Mouse Spleen with the gentleMACS Dissociator

Single-cell suspensions are a prerequisite for experiments in cell separation, cell analysis and cell culture.  To avoid tedious and often painful manual dissociations the gentleMACS Dissociator allows one to dissociate tissue very efficiently under controlled and reproducible conditions.  The gentleMACS Dissociator can optimally dissociate mouse spleen, combining timesaving and standardization with user-safety.

This video describes how to dissociate mouse spleens using the gentleMACS Dissociator, an automated bench-top device that can mechanically disrupt tissues using special tubes to produce viable cell suspensions. Following dissociation, spleens are filtered, centrifuged, and resuspended for further applications.

This video describes a simple, time-saving technique for automated tissue dissociation using the gentleMACS Dissociator to prepare single-cell suspensions of mouse splenocytes.

Single-cell suspensions are a prerequisite for experiments in cell separation, cell analysis and cell culture.  To avoid tedious and often painful manual ...

Single Cell Transfection in Chick Embryos

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a thorough analysis of the fates of single cells under normal and experimental conditions. To this end, transfection methods that target single progenitors are a prerequisite. We describe here a technically straightforward method for transfecting single cells in chicken tissues in-ovo, allowing reliable lineage tracing as well as genetic manipulation. Specific tissue domains are targeted within the somite or neural tube, and DNA is injected directly into the epithelium of interest, resulting in sporadic transfection of single cells. Using reporters, clonal populations may consequently be traced for up to three days, and behavior of genetically manipulated clonal populations can be compared with that of controls. This method takes advantage of the accessibility of the chick embryo along with emerging tools for genetic manipulation. We compare and discuss its advantages over the widely-used electroporation method, and possible applications and use in additional in-vivo models are also suggested. We advocate the use of this method as a significant addition and complement for existing lineage tracing and genetic interference tools.

Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations.

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

Laser Ablation of the Zebrafish Pronephros to Study Renal Epithelial Regeneration

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in renal failure1. AKI can be caused by a number of factors including ischemia, drug-based toxicity, or obstructive injury1. This results in an inability to maintain fluid and electrolyte homeostasis. While AKI has been observed for decades, effective clinical therapies have yet to be developed. Intriguingly, some patients with AKI recover renal functions over time, a mysterious phenomenon that has been only rudimentally characterized1,2. Research using mammalian models of AKI has shown that ischemic or nephrotoxin-injured kidneys experience epithelial cell death in nephron tubules1,2, the functional units of the kidney that are made up of a series of specialized regions (segments) of epithelial cell types3. Within nephrons, epithelial cell death is highest in proximal tubule cells. There is evidence that suggests cell destruction is followed by dedifferentiation, proliferation, and migration of surrounding epithelial cells, which can regenerate the nephron entirely1,2. However, there are many unanswered questions about the mechanisms of renal epithelial regeneration, ranging from the signals that modulate these events to reasons for the wide variation of abilities among humans to regenerate injured kidneys.
The larval zebrafish provides an excellent model to study kidney epithelial regeneration as its pronephric kidney is comprised of nephrons that are conserved with higher vertebrates including mammals4,5. The nephrons of zebrafish larvae can be visualized with fluorescence techniques because of the relative transparency of the young zebrafish6. This provides a unique opportunity to image cell and molecular changes in real-time, in contrast to mammalian models where nephrons are inaccessible because the kidneys are structurally complex systems internalized within the animal. Recent studies have employed the aminoglycoside gentamicin as a toxic causative agent for study of AKI and subsequent renal failure: gentamicin and other antibiotics have been shown to cause AKI in humans, and researchers have formulated methods to use this agent to trigger kidney damage in zebrafish7,8. However, the effects of aminoglycoside toxicity in zebrafish larvae are catastrophic and lethal, which presents a difficulty when studying epithelial regeneration and function over time. Our method presents the use of targeted cell ablation as a novel tool for the study of epithelial injury in zebrafish. Laser ablation gives researchers the ability to induce cell death in a limited population of cells. Varying areas of cells can be targeted based on morphological location, function, or even expression of a particular cellular phenotype. Thus, laser ablation will increase the specificity of what researchers can study, and can be a powerful new approach to shed light on the mechanisms of renal epithelial regeneration. This protocol can be broadly applied to target cell populations in other organs in the zebrafish embryo to study injury and regeneration in any number of contexts of interest.

Acute kidney injury (AKI) in humans is a common clinical problem caused by damage to the epithelial cells that comprise kidney nephrons, and AKI is associated with high mortality rates of 50-70%1. Following epithelial cell destruction, nephrons have a limited ability to regenerate, though the mechanisms and limitations that guide this phenomenon remain poorly understood. In this video article, we describe our technique for targeted laser ablation of kidney nephron cells in the zebrafish embryo kidney, or pronephros. Our new method can be used to complement nephrotoxicity-induced models of AKI and gain a high-resolution understanding of the cell and molecular alterations that are associated with epithelial regeneration in the kidney nephron.

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Single-cell Microinjection for Cell Communication Analysis

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a hemichannel by each neighboring cell to form the gap junction. In mammalian cells, the hemichannel is formed by six connexins, monomers with four transmembrane domains and a C and N terminal within the cytoplasm. Gap junctions permit the exchange of ions, second messengers, and small metabolites. In addition, they have important roles in many forms of cellular communication within physiological processes such as synaptic transmission, heart contraction, cell growth and differentiation. We detail how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells. It is expected that in functional gap junctions, the dye will diffuse from the loaded cell to the connected cells. It is a very useful technique to study gap junctions since you can evaluate the diffusion of the fluorescence in real time. We discuss how to prepare the cells and the micropipette, how to use a micromanipulator and inject a low molecular weight fluorescent dye in an epithelial cell line.

We describe here how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells, and provide some useful tips. We expect that this paper will help everyone to evaluate the degree of cellular coupling due to functional gap junctions. Everything described here could be, in principle, adapted to other fluorescent dyes with molecular weight below 1,000 Daltons.

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a ...

Single Cell Collection of Trophoblast Cells in Peri-implantation Stage Human Embryos

Human implantation, the apposition and adhesion to the uterine surface epithelia and subsequent invasion of the blastocyst into the maternal decidua, is a critical yet enigmatic biological event that has been historically difficult to study due to technical and ethical limitations. Implantation is initiated by the development of the trophectoderm to early trophoblast and subsequent differentiation into distinct trophoblast sublineages. Aberrant early trophoblast differentiation may lead to implantation failure, placental pathologies, fetal abnormalities, and miscarriage. Recently, methods have been developed to allow human embryos to grow until day 13 post-fertilization in vitro in the absence of maternal tissues, a time-period that encompasses the implantation period in humans. This has given researchers the opportunity to investigate human implantation and recapitulate the dynamics of trophoblast differentiation during this critical period without confounding maternal influences and avoiding inherent obstacles to study early embryo differentiation events in vivo. To characterize different trophoblast sublineages during implantation, we have adopted existing two-dimensional (2D) extended culture methods and developed a procedure to enzymatically digest and isolate different types of trophoblast cells for downstream assays. Embryos cultured in 2D conditions have a relatively flattened morphology and may be suboptimal in modeling in vivo three-dimensional (3D) embryonic architectures. However, trophoblast differentiation seems to be less affected as demonstrated by anticipated morphology and gene expression changes over the course of extended culture. Different trophoblast sublineages, including cytotrophoblast, syncytiotrophoblast and migratory trophoblast can be separated by size, location, and temporal emergence, and used for further characterization or experimentation. Investigation of these early trophoblast cells may be instrumental in understanding human implantation, treating common placental pathologies, and mitigating the incidence of pregnancy loss.

Here, we describe a method for warming vitrified human blastocysts, culturing them through the implantation period in vitro, digesting them into single cells and collecting early trophoblast cells for further investigation.

Human implantation, the apposition and adhesion to the uterine surface epithelia and subsequent invasion of the blastocyst into the maternal decidua, is a ...

Real-Time Detection of Reactive Oxygen Species Production in Immune Response in Rice with a Chemiluminescence Assay

Reactive oxygen species (ROS) play vital roles in a variety of biological processes, including the sensing of abiotic and biotic stresses. Upon pathogen infection or challenge with pathogen-associated chemicals (pathogen-associated molecular patterns [PAMPs]), an array of immune responses, including a ROS burst, are quickly induced in plants, which is called PAMP-triggered immunity (PTI). A ROS burst is a hallmark PTI response, which is catalyzed by a group of plasma membrane-localized NADPH oxidases-the RBOH family proteins. The vast majority of ROS comprise hydrogen peroxide (H2O2), which can be easily and steadily detected by a luminol-based chemiluminescence method. Chemiluminescence is a photon-producing reaction in which luminol, or its derivative (such as L-012), undergoes a redox reaction with ROS under the action of a catalyst. This paper describes an optimized L-012-based chemiluminescence method to detect apoplast ROS production in real-time upon PAMP elicitation in rice tissues. The method is easy, steady, standardized, and highly reproducible under firmly controlled conditions.

Here, we describe a method for the real-time detection of apoplastic reactive oxygen species (ROS) production in rice tissues in pathogen-associated molecular pattern-triggered immune response. This method is simple, standardized, and generates highly reproducible results under controlled conditions.

Reactive oxygen species (ROS) play vital roles in a variety of biological processes, including the sensing of abiotic and biotic stresses. Upon pathogen ...