This study was supported by the major scientific and technological innovation project (2020CXGC010802).

NOTE: Pay attention to safety measurements before trapping wild-type migratory insects, it is suggested to wear safety gear (gloves, long-sleeved shirts, and goggles). Also, turn off the trap when not in use to avoid other safety hazards and overheating the light. It is important to follow safety protocols before dissection, such as wearing gloves, goggles, and lab coat to prevent exposure to body fluids and chemicals.
1. Trapping of migrant insects
<ol>
	<li>Begin this protocol by trapping insects using the searchlight lamp. In this protocol, test insect source is Jiyang district, Jinan city, Shandong province, China (36.977088&#176; N, 116.982747&#176; E).</li>
	<li>Use the main body of the searchlight lamp, that is made of non-rusted steel, the box, which is a rectangular body, and the GT75 type halogen headlamp, with a power of 1000 W. Place the headlamp in the middle as the light source.</li>
	<li>Place a funnel-shaped insect-collecting channel inside and at the bottom of the light, place a box for insect-collection with a diameter of 5 cm, followed by a 60-order insect-collecting net bag (0.5 m x 0.5 m), which is used to collect insects trapped by light. The known projection of the light is about 500 m above the ground.</li>
	<li>This protocol emphasizes on the immigratory dynamics of wild-type females and ovary development; therefore, collect different species of insects (here collection was done from April to August, during 2021 to 2022). Avoid collecting small sized and injured insects, select similarly sized large pests for this experiment.</li>
</ol>
2. Preparation of insects
<ol>
	<li>Transfer all collected insects from net bag (0.5 m) to the net cage (30 cm x 30 cm), and then provide a Petri dish containing sterilized solution of 10% honey water (feeding is optional). Place the cage at 27 &#177; 2 &#176;C, 65% &#177; 12 % relative humidity, and a maintain in dark for 8-12 h.</li>
	<li>Select wild-type females that have flown inside the cage on the same day, carefully transfer them into the individual vial tubes, and close each tube with a cotton lid. Avoid direct handling, that could damage or injure the pest due to excessive pressure. 
	&#8203;NOTE: All wild-type females were captured in the night-time and dissection was performed in daytime. Thus, each experiment was performed within a day.</li>
</ol>
3. Preparation for insect paralyzing method (Figure 1)
<ol>
	<li>Place selected female pest individually into a fly vial tube in the middle and anesthetize female using CO2 gas by holding blow gun needle to cause mild paralysis. To confirm whether the pest is paralyzed or not, gently nudge or touch the pest using a soft brush. No response to soft stimuli and immobility indicates a successful paralysis. 
	&#8203;NOTE: Low temperature (-20 &#176;C) can also be used as an alternative technique to paralyze insects.</li>
</ol>
4. Dissection of insects
<ol>
	<li>Place freshly paralyzed female into the dissecting Petri dish containing absolute ethanol (10 mL). To avoid the influence of scale hairs and powder of wings during dissection, infiltrate the living or paralyzed insect with absolute alcohol and rinse in clean water.</li>
	<li>Separate the dorsal wings from the junction of the chest and abdomen body, using two pairs of forceps.</li>
	<li>Transfer the abdomen into a new disposable Petri dish, containing an appropriate amount of water (2-5 mm deep), and gently peel the abdominal exoskeleton along the dorsal ventral line from the pointed mouth to the tail, using dissecting forceps. Repeat the same steps on the other side, and then put it into clean water to disperse the intact tissues.</li>
	<li>Carefully peel off the epidermis fat tissues using forceps, and gently pull and release the ovaries.</li>
	<li>Use dissecting forceps to gently remove fat particles and other organs around the ovaries. Generally, the pest ovaries are mostly folded inward on both sides of the abdomen, try to operate in a liquid environment while unfolding the ovaries, and slowly peel off the mating sac from the middle, and pick out the fat particles attached to the ovarian tubes.</li>
	<li>Gently hold on to the ovary and mating sac from the vertical posterior end and unfold it carefully downwards. To avoid damage during unfolding, transfer the ovary into a new or clean Petri dish containing water. Hold the ovary tip and unfold the ovary inwardly; carefully perform this step to avoid damage of the ovarioles.</li>
</ol>
5. Analyzing data for ovarian tissues anatomy
<ol>
	<li>At this step assess the eggs development for each insect, following the color and size of the eggs, to judge its maturity. Then, judge the ovarian grade according to egg development. 
	NOTE: The division of various ovarian development levels is mainly divided, either before egg laying or egg development with yolk precipitation level. Insects egg grain development are further divided into stages for more clarity such as, yolk occurrence stage, yolk maturity stage, and yolk demise stage. The maturity of egg grain depends on the fullness, color, and size of the egg to judge its maturity.</li>
	<li>After dissection, ensure to separate the female mating sacs tissues from the intact ovaries, and observe the morphology to distinguish the species because most mating sacs anatomy varies from species to species. Therefore, use mating sacs to distinguish between species.</li>
	<li>At this step, evaluate the ovarian anatomy, and analyze the ovarian development grading. Divide the ovarian tissues into five grades (grade 1 to grade 5).
	<ol>
		<li>Look for the following changes and structure to organize the tissue: first grade (1) is an early stage of development (milky transparent), full abdomen, soft, fat body fluffy, light color, difficult to peel. Second grade (2) is the yolk deposition stage and grade the ovaries separately if needed after observing the longer and thicker ovarian canal. Third grade (3) has fewer fat bodies and only a few granules attached to the ovary. Fourth grade (4) appears less elastic and easy to break, and some eggs can be present in the middle of uterine tube. Fifth grade (5) is easy to identify with less or no fat bodies, atrophic and fragile ovaries.</li>
	</ol>
	</li>
	<li>Capture images using a digital camera as per experimental need.</li>
</ol>

Dissection for Identifying Migratory Insects via Ovarian Reproductive Analysis

The authors have no conflicts of interest to declare.

Ovarian analysis methods are routinely used in plant protection, to elucidate the movement of insect flight and population for forecasting19,20,21 and to elaborate on the physiological variations in insects. It has been noticed that the unique migration and rapid dispersion ability of common agricultural pests, such as bollworm, armyworm, taro caterpillar,&#160;and beet moth, make it difficult for prediction from other regions. Some studies considered Athetis lepigone as migratory pest22,23, but the migration dynamics and prediction of this pest is elusive. Recently, another light trapping method was introduced where 1000 W searchlight lamps were used for monitoring of migratory pests18,24, which has a good application effect on migratory pest monitoring. It is also speculated that light trapping methods are not sufficient to distinguish migratory insects. In this study, during migratory peak, the number of adult individuals caught by searchlight trap were majorly migratory pests, while the number of regional individuals was negligible. This highlights that the emergence period of migratory species is quite similar25. To avoid regional individual, it is important to consider the migratory peak period to trap abundance of migrant individuals. Therefore, the identification of migratory insects with searchlight lamps and the grading of ovaries are both fundamental aspects. However, there are just handful of methods to identify the ovarian anatomy of the migratory insects13,26,27,28,29. The feasible method presented here is innovative work that establishes the use of both ovarian tissues, mating sacs and ovarian grading, to distinguish various developmental stages and tried to discuss morphogenesis properties of ovarian tissues in more detail than previous studies11,21,30; specifically with respect to the most dangerous wild female pests, such as H. armigera31, M. separata32, and S. litura33 that were captured using searchlight lamps.
Most previous studies have classified ovarian development using distinguishing methods to evaluate the identification of migratory insects. The focus of these studies was color maturity of yolk, oviposition, birth developments, mating frequency, etc.22,30,34,35. The method used in this study introduced bidirectional comparative evidence, based on two ovarian tissues. For the bollworm, armyworm, taro caterpillar, beet moth, Agrotis ipsilon, Spaelotis valida, Athetis lepigone, and Mamestra brassicae, mating sacs were studied. Egg grain development stages were directly assessed in Helicoverpa armigera, Mythimna separata, Spodoptera litura, and Spodoptera exigua for the ovarian anatomy. According to the insect female adult ovaries anatomy, the development of the egg varies from species to species16,17,36,37. Therefore, it is useful to study the ovarian morphologies at both levels of the migratory insects by analyzing the ovarian development grades and mating sacs by using this dissection method.
Generally, long-term, or improper storage, such as fluctuation in humidity, temperature and food source of the insects trapped using searchlight lamps may affect body weight, egg health and ovary size5,38,39,40,41,42. This leads to difficulty in insect sorting and dissection in the later stage and leads to incomplete data. To minimize these errors, short term and proper storage is necessary. Additionally, after applying the protocol, similar sized females were further selected. This selection and collection of females is time saving, with quick and significant results in the present study. This method demonstrates the dissection of wild-type adult migratory insects from natural environment and its ovarian grading. To perform this protocol with effective results, some critical steps should be considered. For example, insect selection from the natural environment is a crucial step in preventing empty ovaries and infertile eggs; the living insect with intact tissues organs is preferable, avoiding dead, weak, broken or shrunk abdominal cavity insects.
Accordingly, the dissection of ovarian tissues could be estimated by comparing ovarian gradients such as in bollworm (H. armigera) ovarian grading (Figure 4), eggs mature from milky white and transparent oogonia in the ovaries to dark yellow. Scattered egg laying (Figure 5) occurs in armyworms where ovaries mature from milky white transparent oogonia and appear milky white or pale yellow. Taro caterpillar&#160;showed obliques development from milky white transparent oogonia cells that were pink and tender, and further matured with brown or yellow coloration (Figure 6). The continuation of this work decides the ovarian anatomy in different species at the same time. This practice will help in the field of plant protection to study the movement of insects and can guide pest forecasting.
At present, this study has some limitations. The application of this method depends on the detailed analysis of egg grains and ovarian development properties. A key ongoing future challenge is to further advance and improve field-based analysis to evaluate the migratory dynamics of female pests in the ecosystem. Therefore, it is necessary to combine field data for statistical analysis, because ovarian grading is affected by environmental factors such as annual quantification of migratory insects, temperature, humidity, wind direction of the site and food, etc. It was assumed that all species would have the same ovarian grading and mating capsule levels. Still, it needs to be verified with relatively similar species since the availability of insects in nature varies and may result in lack of prediction. Thus, it is still necessary to verify the dissection of similar species of high-altitude migrant females using two ovarian tissues in the future.

Migration of insects plays a vital role in population dynamics of global insect distribution for insects like Helicoverpa armigera - the cotton bollworm, Mythimna separate - the oriental armyworm, Spodoptera litura - the taro caterpillar, Spodoptera exigua - the beet armyworm, that have been reported as serious pests in China1,2,3,4. The long travel distances, seasonal movements, high fecundity of migratory pests, and ecological factors have brought great difficulties in the prediction, forecast, and control of these pests5. Pest migration monitoring is required to reveal the adaptability and behavioral changes that facilitate migratory pests according to climate changes or cycles6. To sustain their growth, reproduction, and survival, insects have acquired sequential adaptability during evolution; this series of adaptive life has generated many changes in the reproductive system, such as migratory strategy leading to control of ovarian development in the long migratory process.
Ovarian development is common in migratory pests, which affects the growth of their population7. Therefore, ovarian development has been a hot topic of migratory pest research for a long time. A series of studies have led to several ovarian development indicators and classification strategies. Until now, several methods have been used to analyze ovary development, e.g., Loxostege sticticalis - the meadow moth- ovary development which includes the initial feathering stage, the early spawning period, the spawning period, and the end of oviposition8. Some researchers divide ovarian levels on the bases of yolk color development in migratory Lepidopteran pests, such as S. exigua - the beet armyworm, Pseudaletia unipuncta - the true armyworm, and Cnaphalocrocis medinalis- the rice leaf-folder, etc.9,10,11,12. In previous studies, the ovarian development levels for pests, such as cotton bollworm and rice leaf roller, were divided into five stages: yolk deposition stage, egg grain maturity stage, mature waiting for birth, peak ovogenesis period, and end spawning stage13,14. The ovarian development of the European corn borer was divided into six developmental stages: yolk deposition stage, egg maturation, pre-eggs dispositioning, peak spawning stage, and end-spawning stage15.
Moreover, insects of the same genus have different stages of development, such as ovarian development levels of Spodoptera frugiperda - the fall armyworm - falls into four levels: yolk deposition stage, mature waiting for delivery, peak ovi-positivity, and end spawning stage16. On the other hand, ovarian development in Spodoptera exigua - the beet moth - has five levels: transparent, vitellogenesis, maturation of eggs, egg release, and late egg-laying levels17.
Former studies can only classify development from single to multiple ovarian development levels using color maturity of yolk, oviposition, and egg developments, but classification cannot be done based on anatomy of the reproductive system. The development of an ovary based on the morphogenesis anatomy is a less studied area. Here, the dissection method was designed to predict migratory females in the population using two ovarian tissue types, to elaborate their reproductive dynamics based on the anatomical morphogenesis of -ovarian development stage and mating sac- providing direct evidence to distinguish migratory wild-type females.
Some studies have found that, migratory Noctuidae insect species were frequently captured by searchlights18. The ovary of most migratory Noctuidae insect species is in the early stages of development during the initial stage of migration and the ovarian level increases with the migratory progress. In this study, the dissection method for ovarian development grades is described, to study the two reproductive tissues of different female population pests, captured by search light. This method not only advances the research to understand the migratory dynamics, but also facilities in insect classification, insect physiology study, pest prediction, and forecasting of female pest species.

<table><tbody><tr><td>Digital camera</td><td>Canon ( China ) co., LTD</td><td>EOS 800D</td><td></td></tr><tr><td>Dropper</td><td>Qingdao jindian biochemical equipment co., LTD</td><td></td><td></td></tr><tr><td>Ethanol absolute (99.7%)</td><td>Shanghai Hushi Laboratory Equipmentco., LTD</td><td></td><td></td></tr><tr><td>Forceps</td><td>&amp;nbsp;Vetus Tools co., LTD</td><td>ST-14</td><td></td></tr><tr><td>GT75 type halogen headlamp (1000 W)</td><td>Shanghai Yadeng Industry co., LTD</td><td></td><td></td></tr><tr><td>Helicoverpa armigera, Mythimna separate, Spodoptera litura, Spodoptera exigua</td><td>Jiyang district, Jinan city, Shandong province, China</td><td></td><td></td></tr><tr><td>Measuring cylinder, beaker, flask</td><td>Qingdao jindian biochemical equipment co., LTD</td><td></td><td></td></tr><tr><td>Net bag&amp;nbsp;</td><td>Qingdao jindian biochemical equipment co., LTD</td><td></td><td>0.5 m&amp;nbsp;</td></tr><tr><td>Net cages&amp;nbsp;</td><td>Qingdao jindian biochemical equipment co., LTD</td><td></td><td>30 cm x 30 cm</td></tr><tr><td>Petri dishes</td><td>Qingdao jindian biochemical equipment co., LTD</td><td></td><td>&amp;nbsp;60 mm diameter</td></tr></tbody></table>

dissection and grading of ovarian development in wild-type female insects

Migratory insect pests pose serious challenges to food production and security all over the world. The migratory pests can be monitored and captured using searchlight traps. One of the most important techniques for migratory pest forecasting is to identify the migratory species. However, in most cases, it is difficult to get the information just by appearance. Therefore, using knowledge acquired by systematic analysis of the female reproductive system can help to understand the combined anatomic morphology of the ovarian mating sac and ovary developmental grading of wild-type migratory insects captured with searchlight traps. To demonstrate the applicability of this method, ovarian development status and egg grain development stages were directly assessed in Helicoverpa armigera, Mythimna separata, Spodoptera litura, and Spodoptera exigua for the ovarian anatomy, and the ovarian mating sacs were studied in Agrotis ipsilon, Spaelotis valida, Helicoverpa armigera, Athetis lepigone, Mythimna separata, Spodoptera litura, Mamestra brassicae, and Spodoptera exigua, to explore their relationships. This work shows the specific dissection method to predict wild-type migratory insects, comparing the unique reproductive system of different migratory insects. Then, both tissues, namely, the ovary and mating sacs, were further investigated. This method helps to predict the dynamics and the structural development of reproductive systems in wild-type female migratory insects.

Development of the eggs 
The above protocol was applied to analyze the development of eggs in the ovary. For this purpose, firstly, eggs were classified generally into four stages to distinguish early and mature stage of egg development among all species e.g., bollworm, armyworm, taro caterpillar, and beet moth. Here, the early stage of feathering (milky white transparent stage) was observed. Figure 2A shows that ovaries have not yet begun to develop, the ovarian duct is filamentous with good elasticity. The formed egg grain cannot be visualized under daylight, due to eggs delicacy, and milky white spots can be seen with light transmission by stretching the ovarian tube.
Ovarian development (yolk deposition stage) contains large eggs, slightly immature and milky white in color, and the eggs are closely in contact (Figure 2B). Mature pending delivery (Ripen stage) can be observed in Figure 2C, the eggs are bead shaped and fully ripe with bright yellow color. Figure 2D demonstrates the yolk extinction period (end-spawning stage), fragility of an ovarian tube. Most of the active egg grains are released, only a few or no egg grains, and maybe inactive eggs are present.
Comparison of mating sac morphologies 
A total of eight wild-type female species were compared on the bases of mating sacs. Each species insect count was more than 100 in number. The most representative mating sac was selected as an example for display (Figure 3). The mating sac of Helicoverpa armigera was a columnar right helix with an inner lumen (Figure 3A). Mythimna separata mating sac was G-shaped/ fishhook - shaped with dark brown bands on the inside (Figure 3B). Agrotis ipsilon mating sac was linear, with large coils and outer cysts on the back (Figure 3C).
Spaelotis valida (Figure 3D) had a mating sac that is slightly thin and J-shaped after enlargement; it looks like a leg and is slightly brownish in color. Spodoptera exigua species shows white translucent dumbbell shaped sac with a bubble cyst in the middle, and distal end is milky white (Figure 3E). Spodoptera litura mating sac is red at the end of mouth, and the upper part can be observed as milky white transparent, and the internal cystic cavity is faintly visible (Figure 3F). Pseudoptera lepigone mating sac is small coiled, with outer cysts that seem like a closed flower, having white and brown color from top to bottom (Figure 3G), and Mamestra brassicae has a double coiled sac with outer cysts olive in color (Figure 3H).
Ovarian development grading in migratory wild-type females 
In the ovarian development grading, grade 1 to grade 5 were evaluated in 4 wild-type female species, bollworm, armyworm, taro caterpillar, and beet moth. Analysis results of bollworm (H. armigera) ovarian grading (Figure 4) showed that the ovarian duct is transparent and elastic with subtle visible oocytes at grade 1 (Figure 4A). The ovarian duct is transparent, the eggs are tender yellow with transparent wraps at grade 2 (Figure 4B). At grade 3 (Figure 4C), no large fat bodies were present, only a few fat particles were attached to the ovarian duct. The ovarian duct is the longest and thickest near the ovipositor, has a clear division of mature areas, growth areas, and proto-egg areas, the ovarian duct is yellow and full rosary. Grade 4 and 5 (Figure 4D,E) showed clusters and less eggs in the ovaries with pale yellow and greenish color.
Similarly, all five grades of observations were noticed in armyworm (M. separata), as shown in Figure 5. A fat body is a flocculent texture composed of multiple fat particles, fluffy, light in color, and milky white with delicate fine visible oocytes (Figure 5A). At grade 2, the ovarian tube structure is yellow and full of bead shaped eggs (Figure 5B). While grade 3 and 4 (Figure 5C,D) are fully matured, the ovarian duct is white and cream colored, and the egg grain is white with a transparent curl wrap on the outside, the eggs are full but crowded. Grade 5 shows (Figure 5E) a&#160;decline of coloration from yellow to dark brown; it emphasizes the final stage of the developed ovary.
Taro caterpillar (S. litura) and beet moth (S. exigua) ovarian grading were also observed using the same ovarian grading levels as shown in Figure 6 and Figure 7. Ovaries at grade 1 and 2 (Figure 6A,B) of the taro caterpillar&#160;were thin enlarged and pinkish in color as compared to the ovaries of the beet moth (Figure 7A,B), that are white, short, fluffy, and wide. Both ovaries have similar grades of ovarian duct morphogenesis, with transparent, elastic, and subtle visible oocytes. Results of grade 3 and 4 of both species show that the ovaries are at the same stage of development, they are dark yellow with full eggs (Figure 6C,D). On the other hand, the same change in color can be visualized from sky blue to light greenish (Figure 7C,D); this change in color indicates the mature stage of ovary development. Late oviposition was evaluated with brownish and few eggs clusters in the oviduct (Figure 6E). Figure 7E shows an ovary with thin oviduct and fluffy eggs clusters that depict the grade 5 ovarian development for eggs, with ovaries gradually becoming larger and darker in color, whitish-bluish color.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65644/65644fig01.jpg" /> 
Figure 1: Tools used for experimentation. The basic tools required for the dissection of ovaries and mating sac from migratory insects. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig01large.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/65644/65644fig02.jpg" /> 
Figure 2:&#160;Egg development stages. Egg development stages in wild-type migratory insects (bollworm, armyworm, taro caterpillar, and beet moth), (A) Milky white transparent stage, (B) Yolk deposition stage, (C) Ripe stage, and (D) End-spawning stage or yolk extinction period. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig02large.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/65644/65644fig03.jpg" /> 
Figure 3:&#160;Mating sacs of different insects. (A) Mating sac of Helicoverpa armigera;&#160;(B) Mating sac of Mythimna separate; (C) Mating sac of Agrotis ipsilon; (D) Mating sac of Spaelotis valida; (E) Mating sac of Spodoptera exigua;&#160;(F) Mating sac of Spodoptera litura;&#160;(G)&#160;Mating sac of Athetis lepigone; (H) Mating sac of Mamestra brassicae. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65644/65644fig04.jpg" /> 
Figure 4: Ovarian development grading of Helicoverpa armigera.&#160;(A) Grade 1 (milky transparent period); (B) Grade 2 represents ovarian development period (yolk deposition period); (C) Grade 3 shows mature period; (D) Grade 4 indicates spawning grade; and (E) Grade 5 represents late oviposition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65644/65644fig05.jpg" /> 
Figure 5: Ovarian development grading of Mythimna separata. (A) Grade 1 (milky transparent period); (B) Grade 2 represents ovarian development period (yolk deposition period); (C) Grade 3 shows mature period; (D) Grade 4 indicates spawning level; (E) Grade 5 represents late oviposition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65644/65644fig06.jpg" /> 
Figure 6: Ovarian development grading of Spodoptera litura. (A) Grade 1 (milky transparent period); (B) Grade 2 represents ovarian development period (yolk deposition period); (C) Grade 3 shows mature period; (D) Grade 4 indicates spawning level; (E)&#160;Grade 5 represents late oviposition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/65644/65644fig07.jpg" /> 
Figure 7: Ovarian development grading of Spodoptera exigua. (A) Grade 1 (milky transparent period); (B) Grade 2 represents ovarian development period (yolk deposition period); (C) Grade 3 shows mature period; (D) Grade 4 indicates spawning level; (E) Grade 5 represents late oviposition. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65644/65644fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Analysis of Ovarian Anatomy in Migratory Insects to Overcome Experimental Challenges at JoVE.com

Author Spotlight: Analysis of Ovarian Anatomy in Migratory Insects to Overcome Experimental Challenges 

The protocol demonstrates a simple and easy dissection method, suitable for wild-type migratory female insects captured with searchlight traps. This technique can significantly clarify the same species by comparing both reproductive tissues, namely the mating sac and ovarian development of wild-type female insects.

Dissection and Grading of Ovarian Development in Wild-Type Female Insects

Migratory insect pests pose serious challenges to food production and security all over the world. The migratory pests can be monitored and captured using ...

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Nanyang Technological University

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Dissection of Drosophila Ovaries

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Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

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The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells ...

Microdissection and Dissociation of the Murine Oviduct: Individual Segment Identification and Single Cell Isolation

Mouse model systems are unmatched for the analysis of disease processes because of their genetic manipulability and the low cost of experimental treatments. However, because of their small body size, some structures, such as the oviduct with a diameter of 200-400 &#956;m, have proven to be relatively difficult to study except by immunohistochemistry. Recently, immunohistochemical studies have uncovered more complex differences in oviduct segments than were previously recognized; thus, the oviduct is divided into four functional segments with different ratios of seven distinct epithelial cell types. The different embryological origins and ratios of the epithelial cell types likely make the four functional regions differentially susceptible to disease. For example, precursor lesions to serous intraepithelial carcinomas arise from the infundibulum in mouse models and from the corresponding fimbrial region in the human fallopian tube. The protocol described here details a method for microdissection to subdivide the oviduct in such a way to yield a sufficient amount and purity of RNA necessary for downstream analysis such as reverse transcription-quantitative PCR (RT-qPCR) and RNA sequencing (RNAseq). Also described is a mostly non-enzymatic tissue dissociation method appropriate for flow cytometry or single cell RNAseq analysis of fully differentiated oviductal cells. The methods described will facilitate further research utilizing the murine oviduct in the field of reproduction, fertility, cancer, and immunology.

A method for microdissection of the mouse oviduct that allows collection of the individual segments while maintaining RNA integrity is presented. In addition, non-enzymatic oviductal cell dissociation procedure is described. The methods are appropriate for subsequent gene and protein analysis of the functionally different oviductal segments and dissociated oviductal cells.

Mouse model systems are unmatched for the analysis of disease processes because of their genetic manipulability and the low cost of experimental ...

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

The Utility of Stage-specific Mid-to-late Drosophila Follicle Isolation

Drosophila oogenesis or follicle development has been widely used to advance the understanding of complex developmental and cell biologic processes. This methods paper describes how to isolate mid-to-late stage follicles (Stage 10B-14) and utilize them to provide new insights into the molecular and morphologic events occurring during tight windows of developmental time. Isolated follicles can be used for a variety of experimental techniques, including in vitro development assays, live imaging, mRNA expression analysis and western blot analysis of proteins. Follicles at Stage 10B (S10B) or later will complete development in culture; this allows one to combine genetic or pharmacologic perturbations with in vitro development to define the effects of such manipulations on the processes occurring during specific periods of development. Additionally, because these follicles develop in culture, they are ideally suited for live imaging studies, which often reveal new mechanisms that mediate morphological events. Isolated follicles can also be used for molecular analyses. For example, changes in gene expression that result from genetic perturbations can be defined for specific developmental windows. Additionally, protein level, stability, and/or posttranslational modification state during a particular stage of follicle development can be examined through western blot analyses. Thus, stage-specific isolation of Drosophila follicles provides a rich source of information into widely conserved processes of development and morphogenesis.

The Utility of Stage-specific Mid-to-late Drosophila Follicle Isolation

Stage-specific isolation of mid-to-late Drosophila follicles is useful for a variety of purposes. Such follicles develop in culture, which allows for genetic and/or pharmacologic manipulations to be coupled with in vitro development assays and live imaging. Additionally, follicles can be used for molecular studies, such as isolating mRNA and protein.

Drosophila oogenesis or follicle development has been widely used to advance the understanding of complex developmental and cell biologic processes. This ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining

Accessibility to germ cells allows the study of germ cell development, meiosis, and recombination. The sexual biotype of the freshwater planarian, Schmidtea mediterranea, is a powerful invertebrate model to study the epigenetic specification of germ cells. Unlike the large number of testis and male germ cells, planarian oocytes are relatively difficult to locate and examine, as there are only two ovaries, each with 5-20 oocytes. Deeper localization within the planarian body and lack of protective epithelial tissues also make it challenging to dissect planarian ovaries directly. 
This protocol uses a brief fixation step to facilitate the localization and dissection of planarian ovaries for downstream analysis to overcome these difficulties. The dissected ovary is compatible for ultrastructural examination by transmission electron microscopy (TEM) and antibody immunostaining. The dissection technique outlined in this protocol also allows for gene perturbation experiments, in which the ovaries are examined under different RNA interference (RNAi) conditions. Direct access to the intact germ cells in the ovary achieved by this protocol will greatly improve the imaging depth and quality and allow cellular and subcellular interrogation of oocyte biology.

This protocol presents steps taken to dissect ovaries in the freshwater planarians, Schmidtea mediterranea. The dissected ovaries are compatible for antibody immunostaining and ultrastructural analysis with transmission electron microscopy to study the cell biology of the oocytes and somatic cells, providing an imaging depth and quality that were previously inaccessible.

Dissection and Grading of Ovarian Development in Wild-Type Female Insects

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