Work in the Lee lab was supported by the National Institute of General Medical Sciences under award number 1R15GM144861 and the National Institute for Child and Human Development under award number 1R15HD104115, both of the National Institutes of Health. DA was supported by the UML Kennedy College of Sciences Science Scholars program. NW was supported by a UML Honors College Fellowship and a UMLSAMP fellowship (funded by the National Science Foundation under grant number HRD-1712771). We thank A. Gartner for RAD-51 antibody. All C. elegans strains were provided by the Caenorhabditis Genetics Center, which is funded by the Office of Research Infrastructure Programs of the National Institutes of Health under award numberP40 OD010440.

<ol>
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The authors have no conflicts of interest to disclose. The content in this manuscript is solely the responsibility of the authors. It does not necessarily represent the official views of the National Institutes of Health or the National Science Foundation.

Sexual reproduction requires the creation of haploid gametes, which are produced via the specialized cell division of meiosis. C. elegans has become a popular model for the cytological study of meiosis due to its optical transparency, convenient germline anatomy, and powerful genetics28. Simple organismal experiments assessing fertility and embryonic lethality can be combined with molecular genetics to address many questions in the lab or the classroom. For example, because crossovers are essential for proper chromosome segregation, processes that disrupt their formation or resolution will generate aneuploid gametes. In turn, aneuploidy leads to inviable progeny, which can easily be assessed by counting progeny, or through the slightly more complicated method of determining embryonic lethality. Cytologically, a lack of crossovers will affect the numbers of DAPI-staining bodies observed during diakinesis. The robustness of DAPI staining and the ease of scoring makes this an ideal experiment to teach cytological techniques. The temporal-spatial layout of nuclei in the hermaphrodite germline provides a snapshot of each stage of meiosis at a single moment in time. Nuclei take about 54 hours to proceed from the distal mitotic end of the germline to the proximal end (for examples, see Jaramillo-Lambert et al.33, Stamper et al.34 and Libuda et al.35). This well-established timing of meiotic progress allows for pulse-chase labeling or DNA damaging experiments.
Because DAPI staining nearly always works, it serves as a useful technical control for fluorescence microscopy (and can provide satisfaction for microscopists-in-training). Antibody staining can be more variable and is a good measure of reproducibility between replicates. C.elegans gonads can be difficult to fix consistently, as this step requires a balance between preserving chromosomal structure while also allowing enough antibody diffusion. Fixing with formaldehyde best preserves chromosomal morphology, and preparing formaldehyde fresh from paraformaldehyde ampules has provided the most reproducible results. It is also possible to use formaldehyde prepared from formalin (37% aqueous formaldehyde and methanol). Fixation strength and timing may need to be empirically determined for each antibody. Alternative methods involve skipping the formaldehyde fixation in step 2.4 and, following freeze-crack, immersion in 100% ethanol at -20 &#176;C, or immersion in 100% methanol for 10 min followed by immersion in 100% acetone for 10 min at -20 &#176;C. These fix conditions allow better antibody penetration but will negatively affect tissue morphology (chromosomes will look blown-out and puffier).
Timing is very important for the dissection and fix steps outlined in step 2. Because the volume of dissection solution is so small, evaporation can impact the final fix concentration, which will cause variable antibody staining. An experienced C. elegans handler can prepare a slide in less than 2 min from the start (step 2.3) to fix (step 2.4). The main technical hurdle is the ability to pick animals into the drop of dissection solution and quickly dissect them. It can be easier to learn how to dissect in larger volumes. New trainees first start by picking animals into 50-200 &#181;L of dissection solution in a glass embryo staining dish (Figure 2D); the wider surface provides more room to maneuver when identifying an ideal needle position for good gonad extrusions, while the larger volume of liquid makes evaporation less of an issue. Once comfortable with the motions of dissecting, trainees start dissecting using only 50 &#181;L in the dish, then switch to dissecting in 20 &#181;L on a slide. Once dissecting on slides, trainees can quickly start reducing the amount of solution until they are quick enough at working in 4 &#181;L to make evaporation negligible. 
If the timing of dissection remains an issue, trainees can dissect in 8 &#181;L of the dissection solution during step 2.3. Then, during step 2.4, they can add 8 &#181;L of the fix solution, gently pipetting to mix thoroughly (watching closely to ensure that carcasses are not disrupted or pipetted away), and removing 8 &#181;L from the mixed solution. This alternative approach will result in the same final volume of 8 &#181;L and the same final fix concentration; this volume is important to ensure that carcasses contact both the coverslip and slide surface during the freeze-crack in step 2.5. If the timing of preparing each slide remains an issue even in larger dissection volumes, a suspension method for germline immunofluorescence protocol is described by Gervaise and Arur26.
The major limitation of using immunofluorescence to visualize proteins in situ is the availability of primary antibodies targeting a particular protein of interest. However, if a tagged version of the protein has been engineered, this protocol can be adapted to visualize the protein tag. Antibodies targeting common tags, like FLAG, HA, or GFP, are commonly available. Fluorescent tags like GFP can often be quenched by fixation steps, so it is recommended to use a primary antibody targeting GFP, rather than relying on the native fluorescence signal itself. Another limitation of this protocol is that it captures the germline within a single timeframe (although all stages of oogenesis will be represented within the germline). Therefore, this technique can miss dynamic changes that may occur as an oocyte progresses through oogenesis. However, the timing of gametogenesis has been well-studied in C. elegans; in a wild-type young adult, an oocyte takes about 60 h to progress from the distal tip of the germline (the mitotic zone) to diakinesis33. Therefore, a pulse chase or specific intervention like irradiation followed by a time course would allow for the observation of effects at different stages of meiosis.
In conclusion, this protocol describes C. elegans gonad dissection followed by DAPI and antibody staining for fluorescence microscopy. Dissection and fixing (step 2) takes 60-90 min, depending on the number of slides generated. Antibody staining (step 3) is mostly hands-off and can range from 7 h at minimum to 1.5 days, depending on antibody incubation times. Mounting (steps 4.1-4.7) takes about 15 min. This general approach to visualizing germline chromosomes and the gonad can be used for cytological studies of any protein if an antibody or fluorescent tag reagent exists. In its simplest form, DAPI staining of diakinesis nuclei can be used to screen for factors affecting meiotic recombination. When paired with organismal analyses of progeny number, the incidence of males (Him phenotype), and embryonic lethality, this cytological approach provides a single-cell counterpoint to population-based analyses.

Meiosis is the specialized cell division used to create gametes (eggs and sperm/pollen) in all sexually-reproducing organisms1,2. Crossover recombination is the reciprocal exchange of DNA between homologous chromosomes; it is essential for meiosis, both providing an important source of genetic diversity and promoting genome stability through generations. Chromosomes that fail to form at least one crossover during meiosis will segregate randomly, which can result in chromosome nondisjunction, creating gametes with the incorrect number of chromosomes &#8211; a condition that is usually fatal for resulting progeny3. During meiosis, crossovers are induced by programmed double-strand DNA breaks4. A subset of these breaks will be repaired as crossovers that provide physical linkages of DNA, called chiasmata, that help orient homologous chromosomes in preparation for cell division5. Meiotic stages are highly conserved across all eukaryotes, and their chromosomal conformation allows them to be easily identified.
As a fundamental concept in biology, meiosis is a topic that students encounter multiple times in different biology courses. They are often introduced to the mechanics of meiotic chromosome segregation in high school, while college-level courses focus on the cell biology of segregation and the genetic impact of crossover recombination. However, meiosis is a notoriously tricky concept for many students1. A failure to understand the relationship between genes, DNA, chromosomes, and meiosis can generate student misconceptions and gaps in understanding that impede a full understanding of genetic inheritance6,7. One way to improve student understanding of abstract topics is to provide concrete, hands-on activities. For example, when teaching meiosis, instructors can choose from activities that emulate molecular analysis8, 3D models that allow students to manipulate molecules9, or role-play where students themselves act out the molecular choreography1. Incorporating research with unknown outcomes is a particularly effective way of improving student understanding. This practice is known as a course-based undergraduate research experience (CURE) and has the added benefit of strengthening student attitudes and agency, especially for those belonging to groups that remain underrepresented in STEM10,11. The nematode worm Caenorhabditis elegans is particularly amenable for classroom studies of behavior, fertility, and genetic crosses, and is an effective model to introduce students to biological research12.
C. elegans makes a powerful model organism for cell biology by combining molecular genetics with simple cytological analysis. It is also particularly well-suited for use in a biology classroom 13,14,15. They are easy and economical to maintain in a lab, producing hundreds of progeny every 3 days, both at standard room temperature or at 20 &#176;C, the most common incubation temperature. Importantly, they can be frozen as glycerol stocks and kept in a -80 &#176;C freezer, which means that any husbandry mistakes made by novice researchers can be easily corrected16. Furthermore, its well-annotated genome allows for forward and reverse genetic techniques17,18, allowing C. elegans to be used to address biological questions ranging from the molecular to evolutionary. Finally, C. elegans researchers have created a supportive community that is often willing to provide help and advice for budding scientists19. These advantages have led to C. elegans being incorporated into a number of CUREs at various types of institutions12,19,20,21,22,23.
In addition to its benefits for research and teaching, C. elegans has become a popular model for studies of meiosis and germline development24,25,26. The optical clarity of these animals simplifies cytological approaches27, and in adults, gonads represent nearly half of the animal, providing hundreds of meiotic cells to study. In gonads, meiotic germline nuclei are arranged like an assembly line (Figure 1); mitotic replication occurs at the distal tip of the gonad, with nuclei progressing through meiotic stages as they migrate toward the proximal end of the gonad, where fertilized embryos emerge from the vulva. Because the stereotyped spatial organization also represents a temporal progression through meiosis, different stages can be easily identified based on their chromosomal organization and location in the gonad. Finally, processes that disrupt meiosis and cause aneuploidy create phenotypes that are straightforward to characterize, even for novices: sterility, embryonic lethality, or a high incidence of males (Him phenotype)28.
This is a simple protocol for visualizing meiotic chromosomes in C. elegans. Mounting, dissection, fixing, and antibody staining are all performed on the same microscope slide, which simplifies the protocol and allows near-perfect sample recovery. This method works for simple DAPI staining to visualize chromosomes and can be used for immunofluorescence to visualize the localization of proteins in the gonad. Students dissect gonads using basic dissecting microscopes, generate whole-mount preparations for visualization of DNA or immunofluorescence, and image them on a compound fluorescent microscope. This protocol has been taught to high-school students and undergraduates working in a C. elegans research lab and incorporated into a CURE at a liberal arts college12. Although the CURE had a relatively small class size, this protocol would be amenable for classes at a range of institutions due to the relatively low cost of worm strains and reagents. Instructors would only be limited by the number of dissecting microscopes available for use. The previous implementation had students working in groups of three to share a single microscope and took place over three 90-minute sessions: the first to practice dissection, the second to implement dissection and DAPI staining, and the third to image slides on a widefield fluorescence microscope. Participation in undergraduate research provides many benefits for students11,29, both academic and personal. Embedding research in courses via CUREs allows students to participate in research during normal class time11,30,31, which makes exposure to these benefits more accessible and equitable.

<table><tbody><tr><td>Alexa Fluor 488 Donkey Anti-Rabbit IgG Secondary Antibody</td><td>Molecular Probes</td><td>711-545-152</td><td></td></tr><tr><td>Aluminum heating/cooling block</td><td>Millipore Sigma</td><td>Z743497</td><td></td></tr><tr><td>Bacto Peptone</td><td>Gibco</td><td>DF0118 17 0</td><td></td></tr><tr><td>Borosilicate Glass Pasteur Pipets, Disposable, 5.75 inches</td><td>Fisherbrand</td><td>13 678 20B</td><td>Used to make worm picks</td></tr><tr><td>BSA, Bovine Serum Albumin</td><td>VWR</td><td>97061 420</td><td></td></tr><tr><td>&lt;em&gt;C. elegans&lt;/em&gt; wild type (ancestral)</td><td>&lt;em&gt;Caenorhabditis&lt;/em&gt; Genetics Center</td><td>N2 (ancestral)</td><td></td></tr><tr><td>&lt;em&gt;C. elegans kle-2 (ok1151)&lt;/em&gt; mutants</td><td>&lt;em&gt;Caenorhabditis&lt;/em&gt; Genetics Center</td><td>VC768</td><td>The full genotype of this strain is: &lt;em&gt;kle-2(ok1151) III/hT2 [bli-4(e937) let-?(q782) qIs48] (I;III)&lt;/em&gt;. It has &lt;em&gt;kle-2 &lt;/em&gt;balanced with a balancer chromosome marked with pharangeal GFP. Pick non-green animals to identify homozygous &lt;em&gt;kle-2 mutants&lt;/em&gt;.</td></tr><tr><td>&lt;em&gt;C. elegans spo-11 (me44)&lt;/em&gt; mutants</td><td>&lt;em&gt;Caenorhabditis&lt;/em&gt; Genetics Center</td><td>TY4342</td><td>The full genotype of this strains is: &lt;em&gt;spo-11(me44)&amp;nbsp; IV / nT1[qIs51] (IV:V)&lt;/em&gt;. It has spo-11 balanced with a balancer chromosome marked with pharangeal GFP. Pick non-green animals to identify homozygous &lt;em&gt;spo-11&lt;/em&gt; mutants.</td></tr><tr><td>Calcium Chloride Anhydrous</td><td>VWR</td><td>97062 590</td><td></td></tr><tr><td>Cholesterol</td><td>Sigma Aldrich</td><td>501848300</td><td></td></tr><tr><td>DAPI</td><td>Life Technologies</td><td>D1306</td><td></td></tr><tr><td>EDTA, Disodium Dihydrate Salt</td><td>Apex Bioresearch Products</td><td>20 147</td><td></td></tr><tr><td>Embryo Dish, glass, 30mm cavity</td><td>Electron Microscopy Services</td><td>100492-980</td><td>Used to practice dissecting; glass is preferred because dissected animals can stick to plastic</td></tr><tr><td>&lt;em&gt;Escherichia coli&lt;/em&gt;</td><td>&lt;em&gt;Caenorhabditis&lt;/em&gt; Genetics Center</td><td>OP50</td><td></td></tr><tr><td>Food storage container, plastic</td><td>various</td><td>n/a</td><td>Plastic containers with (1) relatively flat bottoms, to allow slides to rest level; (2) lids that are air-tight, but can still be easily removed without disturbing contents of container, and (3) are about 9 inches long by 6 inches wide are preferred</td></tr><tr><td>Glass Coplin Jar</td><td>DWK Life Sciences&amp;nbsp;Wheaton</td><td>08-813E</td><td></td></tr><tr><td>Hydrophobic Barrier PAP Pen</td><td>ImmEdge</td><td>NC9545623</td><td></td></tr><tr><td>Kimwipes</td><td>Kimtech Science</td><td>06 666A</td><td></td></tr><tr><td>Levamisole Hydrochloride</td><td>Sigma Aldrich</td><td>L0380000</td><td></td></tr><tr><td>Magnesium Sulfate Anhydrous</td><td>Fisher Chemical</td><td>M65 500</td><td></td></tr><tr><td>Needles, Single-use, 25 G</td><td>BD PrecisionGlide</td><td>14 821 13D</td><td>blue, 1 inch</td></tr><tr><td>NGM Agar, Granulated</td><td>Apex Bioresearch Products</td><td>20 249NGM</td><td></td></tr><tr><td>Parafilm</td><td>Bemis</td><td>16 101</td><td></td></tr><tr><td>Paraformaldehyde (16% w/v) Aqueous Solution</td><td>Electron Microscopy Sciences</td><td>50 980 487</td><td></td></tr><tr><td>PBS, Phosphate Buffered Saline (10x Solution)</td><td>Fisher BioReagents</td><td>BP399500</td><td></td></tr><tr><td>Petri Dishes, 60-mm</td><td>Tritech Research</td><td>NC9321999</td><td>Non-vented, sharp edge</td></tr><tr><td>Platinum wire (90% platinum, 10% iridium)</td><td>Tritech Research</td><td>PT 9010</td><td>Used to make worm picks</td></tr><tr><td>Potassium Chloride</td><td>Fisher</td><td>BP366-500</td><td></td></tr><tr><td>Potassium Phosphate Dibasic</td><td>VWR BDH Chemicals</td><td>BDH9266 500G</td><td></td></tr><tr><td>Potassium Phosphate Monobasic</td><td>VWR BDH Chemicals</td><td>BDH9268 500G</td><td></td></tr><tr><td>Premium Cover Glasses 18 mm x 18 mm</td><td>Fisherbrand</td><td>12-548-AP</td><td>0.13&amp;ndash;0.17 mm thickness</td></tr><tr><td>Razor Blades, Single Edge</td><td>VWR</td><td>55411 055</td><td></td></tr><tr><td>SlowFade Gold Antifade Mountant</td><td>Molecular Probes</td><td>S36937</td><td>Alternatives: VectaShield Antifade Mounting Medium (Vector Laboratories, H-1000-10) or ProLong Diamond Antifade (Thermo Fisher Scientific, P36970). If using ProLong Diamond, no nail polish is required for sealing, but slides must cure for 24 hours before imaging.</td></tr><tr><td>Sodium Azide</td><td>Fisher BioReagents</td><td>BP922I 500</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fisher Chemical</td><td>S271 500</td><td></td></tr><tr><td>Sodium Phosphate Dibasic Anhydrous</td><td>Sigma Aldrich</td><td>7558 79 4</td><td></td></tr><tr><td>Sodium Phosphate Dibasic Heptahydrate</td><td>Sigma Aldrich</td><td>S9390</td><td></td></tr><tr><td>Styrofoam box</td><td>various</td><td>n/a</td><td>Approximate dimensions of interior: 12 inches long, 9 inches wide, 8 inches deep</td></tr><tr><td>Superfrost Plus Microscope Slides</td><td>Fisherbrand</td><td>22 037 246</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher BioReagents</td><td>BP151 500</td><td></td></tr><tr><td>Tryptone</td><td>Apex Bioresearch Products</td><td>20 251</td><td></td></tr><tr><td>Tween 20</td><td>Fisher Chemical</td><td>BP337 500</td><td></td></tr><tr><td>Yeast Extract</td><td>Apex Bioresearch Products</td><td>20 254</td><td></td></tr></tbody></table>

c. elegans gonad dissection and freeze crack for immunofluorescence and dapi staining

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. Whole mount preparations preserve the structure of meiotic nuclei, and importantly, each gonad arm contains all stages of meiosis, organized in a temporal-spatial progression that makes it easy to identify nuclei at different stages. Adult hermaphrodites have two gonad arms, each organized as a closed tube with proliferating germline stem cells at the distal closed end and cellularized oocytes at the proximal open end, which join in the center at the uterus. Dissection releases one or both gonad arms from the body cavity, allowing the entirety of meiosis to be visualized. Here, a common protocol for immunofluorescence against a protein of interest is presented, followed by DAPI staining to mark all chromosomes. Young adults are immobilized in levamisole and quickly dissected using two syringe needles. After germline extrusion, the sample is fixed before undergoing a freeze crack in liquid nitrogen, which helps permeabilize the cuticle and other tissues. The sample can then be dehydrated in ethanol, rehydrated, and incubated with primary and secondary antibodies. DAPI is added to the sample in the mounting medium, which allows reliable visualization of DNA and makes it easy to find animals to image under a fluorescent microscope. This technique is readily adopted by those familiar with handling C. elegans after a few hours spent practicing the dissection method itself. This protocol has been taught to high-schoolers and undergraduates working in a research lab and incorporated into a course-based undergraduate research experience at a liberal arts college.

DAPI binds strongly to DNA, and its staining is robust even under a wide variety of conditions (Figure 3A,B). It should be present in all nuclei and therefore makes an effective positive control for the presence of any worm tissue on the slide and the ability to detect fluorescence on the microscope. Staining is effective when the antibody is present within meiotic nuclei (Figure 3A,B). For example, Figure 3A,B shows mid-pachytene nuclei stained with DAPI and an antibody targeting RAD-51, a marker for double-strand breaks. KLE-2 is a component of condensin, a highly conserved protein complex that structures chromosomes in preparation for mitosis and meiosis. kle-2/+ mutants have minor defects in chromosome structure, as shown by slightly disordered DAPI staining and an increase in double-strand break number reflected in the higher number of RAD-51 foci (Figure 3B). A common mistake for immunofluorescence is leaving the sample in the fix solution for too long. Over-fixing can lead to unsuccessful staining because it usually prevents antibodies from diffusing into nuclei. This mistake can be identified when the antibody is diffused in the cytoplasmic regions of the gonad, but appears to be excluded from nuclei.
In the germline (Figure 1), nuclei mitotically proliferate at the distal tip, enter meiosis during the transition zone, and progress through pachytene (which is often divided into three equal stages by a number of nuclei rows) before entering diplotene and finally diakinesis. Oocytes in diakinesis are numbered based on their proximity to the spermatheca, with the -1 oocyte being most proximal to the spermatheca, the -2 oocyte being the next distal, and so on. C. elegans have six chromosomes, which manifest as compact ovals in diakinesis nuclei. Each of these represents a homologous pair of two sister chromatids held together by a chiasma, also called a bivalent (Figure 3C). Mutants, like spo-11, that disrupt crossover formation will fail to form chiasmata; therefore, diakinesis nuclei will have 12 separate DAPI bodies (also called univalents), one for each sister chromatid (Figure 3D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64204/64204fig01.jpg" /> 
Figure 1: Germline nuclei are arrayed in a spatial-temporal manner that represents their progression through meiosis. (A)&#160;Whole-mount young adult hermaphrodite stained with DAPI. Gonad and meiotic stages are outlined, from the distal tip (asterisk) to the proximal region (the -1 oocyte, indicated with an arrow), which contains the spermatheca (triangle). Scale bar represents 100 &#181;m. (B)&#160;Projected fluorescence image of a dissected hermaphrodite gonad stained with DAPI. Meiotic stages are denoted, with representative nuclei shown in insets (all insets are shown to scale with one another). Nuclei can be easily staged based on their location in the germline and characteristic chromosome morphology, progressing from the mitotic zone at the distal tip (asterisk), to the transition zone, through pachytene, diplotene, and diakinesis. The spermatheca is marked by a triangle. This figure has been adapted from Hillers et al. under license CC BY 3.028. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64204/64204fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64204/64204fig02.jpg" /> 
Figure 2: Demonstration of dissection setup. (A) Microscope stage during dissection. Animals are dissected in 4 &#181;L of dissection solution on a coverslip placed on the holding slide, which is used to move the coverslip into view. The diagram shows ideal needle placement. Needles are held with a beveled edge facing down, crossed over the pharyngeal region. Pink arrows demonstrate a scissor-like motion for needles. The pink dashed line shows the ideal cut location. (B) Images of dissected animals, with an example of a full extrusion and an incomplete extrusion. Sections of extruded gonads and guts are labeled. (C) Image demonstrating the freeze-crack step. The slide should be held firmly in one hand, with the coverslip side facing away from the body. The opposite edge is braced against the bench. The razor should be held in the other hand. The pink arrow indicates the direction of movement for the razor to flick the coverslip off the slide, with a slight turn such that the coverslip flicks away from the body. Note that the coverslip has been placed such that one corner hangs over the long edge of the slide. (D) Image of the setup for dissection practice in a glass embryo staining dish. Animals are placed in 50-200 &#181;L of dissection solution, which negates the problem of evaporation and extends the time for dissection. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64204/64204fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64204/64204fig03.jpg" /> 
Figure 3: Representative fluorescence results of germline nuclei. (A,B) Z-stack projections of late pachytene nuclei stained with RAD-51 antibody (green) and DAPI (red) in (A) wild-type and (B) kle-2/+ heterozygotes. (C,D) Z-stack projections of a single diakinesis nucleus stained with DAPI in (C) wild-type (with six DAPI staining bodies) and (D) spo-11 mutants (with 12 DAPI staining bodies). Scale bars represent 5 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64204/64204fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining at JoVE.com

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

This is a commonly used method for C. elegans gonad dissection followed by freeze crack, which produces germline samples for immunofluorescence via antibody staining, or for simple DAPI staining to visualize DNA. This protocol has been successful for undergraduates in a research lab and in a course-based undergraduate research experience.

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. ...

c-elegans-gonad-dissection-freeze-crack-for-immunofluorescence-dapi

Nanyang Technological University

Research

JoVE Journal

Genetics

5.0K Views.  University of Massachusetts Lowell. This is a commonly used method for C. elegans gonad dissection followed by freeze crack, which produces germline samples for immunofluorescence via antibody staining, or for simple DAPI staining to visualize DNA. This protocol has been successful for undergraduates in a research lab and in a course-based undergraduate research experience.

Video: C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple cellular and developmental processes principally via activation of ERK, the terminal kinase of the pathway. Tight regulation of ERK activity is essential for normal development and homeostasis; overly active ERK results in excessive cellular proliferation, while underactive ERK causes cell death. C. elegans is a powerful model system that has helped characterize the function and regulation of RTK-RAS-ERK signaling pathway during development. In particular, the RTK-RAS-ERK pathway is essential for C. elegans germline development, which is the focus of this method. Using antibodies specific to the active, diphosphorylated form of ERK (dpERK), the stereotypical localization pattern can be visualized within the germline. Because this pattern is both spatially and temporally controlled, the ability to reproducibly assay dpERK is useful to identify regulators of the pathway that affect dpERK signal duration and amplitude and thus germline development. Here we demonstrate how to successfully dissect, stain, and image dpERK within the C. elegans gonad. This method can be adapted for spatial localization of any signaling or structural protein in the C. elegans gonad, provided an antibody compatible with immunofluorescence is available.

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

We present an immunofluorescence&#160;imaging-based method for spatial and temporal localization of active ERK in the dissected C. elegans gonad. The protocol described here can be adapted for visualization of any signaling or structural protein in the C. elegans gonad, provided a suitable antibody reagent is available.

The evolutionarily conserved&#160;extracellular signal transducing RTK-RAS-ERK pathway is an important kinase-signaling cascade that controls multiple ...

Developmental Biology

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging

Multicellular tubes, fundamental units of all internal organs, are composed of polarized epithelial or endothelial cells, with apical membranes lining the lumen and basolateral membranes contacting each other and/or the extracellular matrix. How this distinctive membrane asymmetry is established and maintained during organ morphogenesis is still an unresolved question of cell biology. This protocol describes the C. elegans intestine as a&#160;model for the analysis of polarized membrane biogenesis during tube morphogenesis, with emphasis on apical membrane and lumen biogenesis. The C. elegans twenty-cell single-layered intestinal epithelium is arranged into a simple bilaterally symmetrical tube, permitting analysis on a single-cell level. Membrane polarization occurs concomitantly with polarized cell division and migration during early embryogenesis, but de novo polarized membrane biogenesis continues throughout larval growth, when cells no longer proliferate and move. The latter setting allows one to separate subcellular changes that simultaneously mediate these different polarizing processes, difficult to distinguish in most polarity models. Apical-, basolateral membrane-, junctional-, cytoskeletal- and endomembrane components can be labeled and tracked throughout development by GFP fusion proteins, or assessed by in situ antibody staining. Together with the organism&#39;s genetic versatility, the C. elegans intestine thus provides a unique in vivo model for the visual, developmental, and molecular genetic analysis of polarized membrane and tube biogenesis. The specific methods (all standard) described here include how to: label intestinal subcellular components by antibody staining; analyze genes involved in polarized membrane biogenesis by loss-of-function studies adapted to the typically essential tubulogenesis genes; assess polarity defects during different developmental stages; interpret phenotypes by epifluorescence, differential interference contrast (DIC) and confocal microscopy; quantify visual defects. This protocol can be adapted to analyze any of the often highly conserved molecules involved in epithelial polarity, membrane biogenesis, tube and lumen morphogenesis.

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analy

The transparent C. elegans intestine can serve as an "in vivo tissue chamber" for studying apicobasal membrane and lumen biogenesis at the single-cell and subcellular level during multicellular tubulogenesis. This protocol describes how to combine standard labeling, loss-of-function genetic/RNAi and microscopic approaches to dissect these processes on a molecular level.

Multicellular tubes, fundamental units of all internal organs, are composed of polarized epithelial or endothelial cells, with apical membranes lining the ...

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

Neuroscience

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development&#160;and relies on external cues from neighboring tissues. C. elegans is an excellent model organism to study tissue morphogenesis in vivo due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this ...

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, germ cell development, piRNA biology, and niche formation. Here, we present a dissection technique to live-image the gonad ex vivo during a period when in vivo live-imaging is highly ineffective. This protocol outlines how to transfer embryos to an imaging dish, choose appropriately-staged male embryos, and dissect the gonad from its surrounding tissue while still maintaining its structural integrity. Following dissection, gonads can be imaged using a confocal microscope to visualize dynamic cellular processes. The dissection procedure requires precise timing and dexterity, but we provide insight on how to prevent common mistakes and how to overcome these challenges. To our knowledge this is the first dissection protocol for the Drosophila embryonic gonad, and will permit live-imaging during an otherwise inaccessible window of time. This technique can be combined with pharmacological or cell-type specific transgenic manipulations to study any dynamic processes occurring within or between the&#160;cells in their natural gonadal environment.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, ...

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using "Freeze-Cracking"

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

Biology

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein subcellular localization changes; in particular, a nuclear translocation under a heat stress condition. Proteins react correspondingly to external and internal signals. A common mechanism is to change its subcellular localization. This article describes a protocol to track protein localization that does not require an antibody, radioactive labeling, or a confocal microscope. In this article, GFP is used to tag the target protein EXL-1 in C. elegans, a member of the chloride intracellular channel proteins (CLICs) family, including mammalian CLIC4. An integrated translational exl-1::gfp transgenic line (with a promoter and a full gene sequence) was created by transformation and &#947;-radiation, and stably expresses the gene and gfp. Recent research showed that upon heat stress, not oxidative stress, EXL-1::GFP accumulates in the nucleus. Overlapping the GFP signal with both the nuclei structure and the DAPI signals confirms the EXL-1 subcellular localization changes under stress. This protocol presents two different fixation methods for DAPI staining: ethanol fixation and acetone fixation. The DAPI staining protocol presented in this article is fast and efficient and preserves both the GFP signal and the protein subcellular localization changes. This method only requires a fluorescence microscope with Nomarski, a FITC filter, and a DAPI filter. It is suitable for a small laboratory setting, undergraduate student research, high school student research, and biotechnology classrooms.

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

This protocol demonstrates how to track a protein nuclear translocation under heat stress by using a green fluorescence protein (GFP) fusion protein as a marker and 4',6-diamidino-2-phenylindole (DAPI) staining. The DAPI staining protocol is fast and preserves the GFP and protein subcellular localization signals.

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein ...

C. elegans Gonad Extrusion: A Rapid Dissection Technique

Source: Gervaise, A. L. and Arur, S. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/54901/spatial-and-temporal-analysis-of-active-erk-in-the-c-elegans-germline">Spatial and Temporal Analysis of Active ERK in the C. elegans Germline</a>. J. Vis. Exp. (2016).
This video describes a method to dissect gonads from adult C. elegans hermaphrodites, resulting in tissues that are suitable for immunostaining and fluorescent imaging.

C. elegans (C. elegans) Wytłaczanie gonad: technika szybkiej sekcji

Source: Gervaise, A. L. and Arur, S. Spatial and Temporal Analysis of Active ERK in the C. elegans Germline. J. Vis. Exp. (2016).
This video describes a ...

JoVE Encyclopedia of Experiments

Caenorhabditis elegans (worm)

Anatomy and Physiology

Freeze-Cracking of Nematodes: A Method to Expose Interior Worm Tissues for Staining

Source: Zhang, N., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/56100/the-c-elegans-intestine-as-model-for-intercellular-lumen">The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging.</a> J. Vis. Exp. (2017).
This video describes freeze-cracking, a method to make C. elegans tissues accessible for antibody staining by disrupting the cuticle.

Kraking zamrażalny nicieni: metoda wystawiania wewnętrznych tkanek robaków do barwienia

Source: Zhang, N., et al. The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the ...

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

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Rozdziały w tym wideo

Spatial and Temporal Analysis of Active ERK in the C. elegans Germline

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging

Automated Quantification of Synaptic Fluorescence in C. elegans

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Antibody Staining in C. Elegans Using "Freeze-Cracking"

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

C. elegans (C. elegans) Wytłaczanie gonad: technika szybkiej sekcji

Kraking zamrażalny nicieni: metoda wystawiania wewnętrznych tkanek robaków do barwienia