This study was supported by the Ministry of Education, Malaysia, under the Higher Institution Centre of Excellence (HICoE) program, Malaysia, accredited to the Institute of Tropical Aquaculture and Fisheries, Universiti Malaysia Terengganu (Vot No. 63933 &#38; Vot No. 56048). We acknowledge the support of Universiti Malaysia Terengganu and Sayap Jaya Sdn. Bhd. via the Private Partnership Research Grant (Vot. No. 55377). An adjunct Academic Fellow position from Universiti Sains Malaysia to Khor Waiho and Hanafiah Fazhan is also acknowledged.

This protocol follows the Malaysian Code of Practice for the Care and Use of Animals for Scientific Purposes outlined by the Laboratory Animal Science Association of Malaysia. The sacrifice of the experimental samples was done according to the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publications No. 8023, revised 1978). Sexually pre-mature female mud crabs (orange mud crab S. olivacea) were collected from the local market (5°66′62′′N, 102°72′33′′E) at the Setiu Wetlands in Malaysia. The mud crab species was identified based on morphological characteristics1.
1. Sample collection and disinfection
<ol>
	<li>Collect healthy, active, and pre-mature female mud crabs (Figure 1). 
	​NOTE: Pre-mature female crabs have triangular and light-colored abdomen flaps along with a CW range of 80−85 mm.</li>
	<li>Wash the crabs with chlorinated tap water (freshwater) to remove debris and osmophilic parasites.</li>
	<li>Soak the crabs in 150 ppm formaldehyde with 20 ppt salinity for 30 min.</li>
	<li>Maintain continuous and gentle aeration with airstones during the formaldehyde treatment. The aeration source can be from either a central aeration line or an aquarium aerator pump.</li>
	<li>Wash the crabs with flowing seawater to remove any residual formaldehyde.</li>
</ol>
<img alt="Crab maturity comparison: pre-mature vs. mature female crabs, dorsal view, scale 20 mm." class="xfigimg" src="/files/ftp_upload/65039/65039fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Abdominal morphology of female mud crabs used to identify the sexual maturation stages. <a href="https://www.jove.com/files/ftp_upload/65039/65039fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Acclimatization
<ol>
	<li>Transfer each disinfected female into a separate 32 L circular tank.</li>
	<li>Rear the females for 3 days in 20 ppt salinity and continue feeding twice per day (morning 09:00 am and evening 20:00 pm) with chopped marine fish at about 4%−5% of the crab's body weight.</li>
	<li>Remove excess and uneaten feed by siphoning before the morning feeding.</li>
	<li>Exchange 10% of the crab rearing seawater (20 ppt) daily.</li>
</ol>
3. Induced molting for sexual maturity
<ol>
	<li>Cut all the legs except the swimming legs using sterilized scissors.
	<ol>
		<li>Catch the crab with a scoop net, and hold the crab carefully. Cut both chelipeds first and then the walking legs at the second joint using scissors. The crab will autotomize the damaged appendages automatically. Anaesthesia is not required for limb autotomy.</li>
	</ol>
	</li>
	<li>Wash the crab in freshwater immediately after limb autotomy.</li>
	<li>Individually transfer the limb-autotomized crabs into perforated plastic baskets (28 cm L x 22 cm W x 7 cm H), and place them in a fiberglass tank (305 cm L x 120 cm W x 60 cm H). 
	NOTE: Two baskets can be tied and clipped together. The top basket is used as a cover so that the crab cannot escape from the basket.</li>
	<li>Use a recirculating aquaculture system (RAS) with 20 ppt salinity and a water depth of at least 10 cm to ensure that the whole plastic basket is submerged.</li>
	<li>Continue feeding the limb-autotomized female crab with chopped marine fish twice per day at 5%−7% of the crab's body weight.</li>
	<li>Rear the crabs until sexually mature through molting (35 days). 
	​NOTE: Induced molting can be skipped for commercial ovarian maturation and seed production with wild mature female mud crabs. Harvested mature females from the wild must be acclimatized and directly subjected to cold-shock anesthesia and subsequent eyestalk ablation.</li>
</ol>
4. Anesthesia
<ol>
	<li>Select sexually mature females with a dark-colored oval-shaped abdominal flap with a CW &gt;86 mm (Figure 1).</li>
	<li>Catch the crabs with a scoop net, and keep them individually in small aquariums for anesthesia.</li>
	<li>After 5 min of acclimatization period, add 2-phenoxyethanol (2-PE) at 2 mL/L into each aquarium and allow 15 min of anesthesia treatment.</li>
	<li>Ensure the crabs are fully anesthetized by the lack of spontaneous movement.</li>
</ol>
5. Eyestalk ablation
<ol>
	<li>Cauterization technique
	<ol>
		<li>Perform all procedures on top of a table and in an open area.</li>
		<li>Take a flat head nickel-steel metal rod (e.g., a screwdriver) with a wooden or plastic handle, and cover the handle with a wet cotton towel.</li>
		<li>Sterilize two stainless surgical forceps in an autoclave.</li>
		<li>Prepare 70% ethanol in a spray bottle and keep it away from any fire-related sources, such as blow torch and red hot screwdriver. Have tissue paper ready for use. 
		NOTE: Ethanol is highly flammable. Maintain a safe distance from fire sources.</li>
		<li>Connect a blowtorch to a gas cylinder (butane) securely. 
		CAUTION: Follow the instructions on the blowtorch and gas cylinder. Make sure that the blowtorch is switched off when connecting with the gas cylinder. Read and follow all the fire safety precautions mentioned on the gas cylinder.</li>
		<li>Wear thick cotton gloves to avoid injury from hot objects.</li>
		<li>Subject the tip of the metal rod to the fire of the blowtorch until the metal rod is bright red.</li>
		<li>Cover the anesthetized crab with a wet cotton towel. 
		NOTE: Cover the antennae of the crab to avoid unnecessary damage.</li>
		<li>Hold one eye of the crab with sterilized forceps. 
		NOTE: Sterilize the forceps in an autoclave for first-time use, and disinfect using 70% ethanol for subsequent use on other crabs.</li>
		<li>Hold the red-hot metal flat tip onto the eye of the crab and press slightly for about 10−15 s until the eyestalk turns an orange or reddish-orange color. Be careful when conducting this step to avoid damage to adjacent structures.  
		NOTE: Two people are needed to execute eyestalk ablation following the cauterization method: one to hold the crab and another to perform the ablation procedure.</li>
		<li>Disinfect the forceps with 70% ethanol spray to ensure no cross-contamination between crabs. 
		NOTE: Only perform this step at least waiting for 5 min after the eyestalk ablation procedure to ensure the forceps are cooled down before disinfection using 70% ethanol to prevent potential fire hazards.</li>
		<li>After performing the eyestalk ablation on all crabs, dip the hot nickel steel metal rod (screwdriver) into tap water.</li>
		<li>Disinfect the towel before reuse. Multiple towels can be used to save time. 
		NOTE: Wash the towel with tap water, and dip it into 30 ppm chlorinated water for 5 min. Then, wash the towel with tap water again, and dip it in a 1 g/L sodium thiosulphate solution.</li>
		<li>Keep the blowtorch in a safe place after turning it off, and wait until it returns to environmental temperature (about 30 min) before disconnecting.</li>
	</ol>
	</li>
	<li>Surgery technique
	<ol>
		<li>Perform the procedure in a well-ventilated area.</li>
		<li>Sterilize two surgical scissors and forceps in an autoclave.</li>
		<li>Pour 50 mL of 70% ethanol into a 100 mL glass beaker.</li>
		<li>Wear thick cotton gloves.</li>
		<li>Hold the anesthetized crab, and cover it with a wet cotton towel.</li>
		<li>Hold one eye of the crab with sterilized forceps.</li>
		<li>Swiftly cut off the eyestalk using sterilized surgical scissors. 
		NOTE: Hemolymph may be lost from the wounded part of the crab.</li>
		<li>Dip the scissors and forceps in 70% ethanol after every use, and dry them using tissue paper before reuse.</li>
	</ol>
	</li>
</ol>
6. Post-anesthesia care
<ol>
	<li>Prepare 20 ppt filtered seawater, and keep in an overhead tank with continuous aeration.</li>
	<li>Connect a flexible pipe with the overhead tank for gravitational water flow.</li>
	<li>Immediately after eyestalk ablation, place the crab into the basket, and subject the crab to flowing seawater (ambient water temperature: 28 °C) from the overhead tank.</li>
	<li>Keep the seawater flowing, and monitor the crab until it can move spontaneously, which indicates recovery from the anesthesia. 
	NOTE: Seawater can be prepared in a ground tank, and a submersible water pump can be used for the water flow.</li>
	<li>Keep the crabs individually in 20 ppt seawater with aeration in an aquarium for 30 min for further observation. 
	​NOTE: The recovered crabs will be cultured individually in the subsequent broodstock culture process.</li>
</ol>
7. Observation of ovarian maturation
<ol>
	<li>Broodstock rearing
	<ol>
		<li>Transfer the mature crabs to individual 32 L circular tanks.</li>
		<li>Continue feeding with chopped marine fish (frozen at −20 °C) twice per day (morning 09:00 am and evening 20:00 pm), and remove uneaten feed before the morning feeding.</li>
		<li>Rear the broodstock individually for 30 days in 20 ppt salinity.</li>
		<li>Remove feces, and exchange 10% of the seawater (20 ppt) daily.</li>
	</ol>
	</li>
	<li>Dissection
	<ol>
		<li>Clean a dissecting tray, scissors, and forceps with 70% ethanol.</li>
		<li>Anesthetize the females individually with the 2-PE immersion anesthesia method.</li>
		<li>Randomly select newly mature females (after the molting of pre-mature females) that have not gone through eyestalk ablation to confirm their gonadal stages.</li>
		<li>Sacrifice all the eyestalk-ablated experimental females individually, and identify the gonad maturation stages. Destroy the thoracic ganglia of the crab using a sharp sterile awl. Remove the top carapace first and then the hepatopancreas to make the ovary visible. Observe the ovary color, and identify the ovarian maturation stage (Figure 2).</li>
	</ol>
	</li>
	<li>Ovarian maturation stages identification
	<ol>
		<li>Observe the ovary color with the naked eye or under a stereomicroscope.</li>
		<li>Identify the ovarian maturation stages based on coloration30: the immature (stage-1) shows a translucent or creamy white color; the early maturing (stage-2) shows a pale to light yellowish color; (iii) the pre-maturing (stage-3) shows a yellow to light orange color; and (iv) the fully matured (stage-4) shows a dark orange to reddish color.</li>
	</ol>
	</li>
</ol>

None of the authors have any conflicts of interest.

This protocol was developed for the eyestalk ablation of the mud crab, Scylla spp., and can be applied as an efficient method to induce gonad maturation. This protocol can be easily replicated for the commercial ovary maturation of mud crabs and can be implemented to reduce the latent period (time from one spawning to another) in mud crab seed production.
The eyestalk ablation of crustaceans (i.e., freshwater prawn, marine shrimp) is typically done to induce gonad maturation and out-o...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/65039">Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs</a>. The Introduction, Protocol, Discussion and References were&#160;updated.

Erratum: Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs

All four mud crab species belonging to the genus Scylla are commercially important crustacean species in aquaculture1,2. The growth of crustaceans, including mud crabs, and their transformation from the pre-mature (sub-adult or pubertal) phase to the sexually mature (adult) phase occur through a molting process that involves the periodic shedding of older and smaller exoskeletons. Carapace width (CW), chelipeds, and abdominal flap morphologies are widely used to determine the sexual maturity of Scylla spp.3,4,5. The process of molting is regulated by the action of various hormones and requires a huge amount of energy6. In addition to the normal molting process, the loss of limbs, either voluntarily or induced by external factors, expedites the molting of crabs without affecting their survival rate7,8,9. Therefore, limb autotomy is commonly used for molt induction in the soft-shell mud crab farming industry7,9.
Unilateral or bilateral eyestalk ablation is mostly popular in freshwater prawn and marine shrimp for gonad maturation and seed production10,11,12,13. Common eyestalk ablation techniques in crustaceans include the following: (i) ligation at the base of the eyestalk using a string14,15; (ii) cauterization of the eyestalk using hot forceps or electrocautery devices16; (iii) removal or direct pinching of the eyestalk to leave an open wound12; and (iv) removal of the eyestalk contents through incision after slicing the distal portion of the eye with a razor17. The eyestalk X-organs are important endocrine organs in crustaceans as they regulate crustacean hyperglycemic hormones (CHH), molt-inhibiting hormones (MIH), and vitellogenesis-inhibiting hormones (VIH)6,18,19,20,21,22. Eyestalk X-organs (or the sinus gland complex) synthesize and release gonad-inhibiting hormones (GIH), also known as vitellogenesis-inhibiting hormones (VIH), belonging to the neuropeptide hormone family6. Unilateral or bilateral eyestalk ablation reduces GIH synthesis, resulting in the dominance of stimulating hormones (i.e., gonad stimulating hormones, GSH) and the acceleration of the ovarian maturation process in crustaceans23,24,25,26. Without the influence of GIH after eyestalk ablation, female crustaceans devote their energy to ovary development27. It has been found that unilateral eyestalk ablation is sufficient for the induction of ovarian maturation in crustaceans11 and that the ablated eyestalk of shrimps and crabs can regenerate after several moltings28. There are four ovarian development stages recorded in Scylla spp.: i) immature (stage-1), ii) early maturing (stage-2), iii) pre-maturing (stage-3), and iv) fully mature (stage-4)29,30. The immature ovarian stage is found in immature females. After pubertal molting and mating, the immature ovary starts developing and finally matures (stage-4) before spawning31.
An eyestalk ablation protocol is essential for mud crab broodstock development and seed production. In the global food market, mature mud crabs with fully mature ovaries (stage-4) rather than crabs with higher muscle content are preferred by consumers and, thus, have a higher commercial value, even higher than large males. There is no complete protocol for the eyestalk ablation of mud crabs. The eyestalk ablation protocol in this work minimizes stress by using fully anesthetized&#160;crabs and minimizes physical injury to personnel from crab bites. This protocol is easy and cost-effective. Here, we present a protocol for the eyestalk ablation of Scylla spp. that can induce the maturation of the gonad. Two techniques of eyestalk ablation (cauterization and surgery) were tested and their efficiencies compared based on the gonadal development rate of female mud crabs.

<table><tbody><tr><td>Aeration tube&amp;nbsp;</td><td>Ming Yu Three</td><td>N/A</td><td>aquarium and pet shop</td></tr><tr><td>Airstone</td><td>Ming Yu Three</td><td>N/A</td><td>aquarium and pet shop</td></tr><tr><td>Autoclave machine</td><td>HIRAYAMA MANUFACTURING CORPORATION</td><td>N/A</td><td>MADE IN JAPAN</td></tr><tr><td>Bleaching powder (Hi-Chlon 70%)</td><td>Nippon Soda Co.Ltd,Japan</td><td>N/A</td><td>N/A</td></tr><tr><td>Blow torch&amp;nbsp;</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Circular tank (32L)</td><td>BEST PLASTIC INDUSTRY SDN. BHD.&amp;nbsp;</td><td>N/A</td><td>N/A</td></tr><tr><td>Cotton hand gloves (thick)&amp;nbsp;</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Cotton towel</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Digital thermometer</td><td>Hanna Instrument</td><td>HI9814</td><td>Hanna&amp;nbsp;Instruments&amp;nbsp;GroLine Hydroponics Waterproof pH / EC / TDS / Temp. Portable Meter HI9814</td></tr><tr><td>Digital Vernier Caliper</td><td>INSIZE Co., Ltd.</td><td>N/A</td><td></td></tr><tr><td>Dissecting tray</td><td>Hatcheri AKUATROP&amp;nbsp;</td><td>N/A</td><td>Research Center of Universiti Malaysia Terengganu</td></tr><tr><td>Dropper bottle/Plastic Pipettes Dropper</td><td>Shopee Malaysia</td><td>N/A</td><td>N/A</td></tr><tr><td>Ethanol 70%</td><td>Thermo Scientific Chemicals</td><td>033361.M1</td><td>Diluted to 70% using double distilled water</td></tr><tr><td>Fiberglass tank (1 ton)</td><td>Hatcheri AKUATROP&amp;nbsp;</td><td>N/A</td><td>Research Center of Universiti Malaysia Terengganu</td></tr><tr><td>Fine sand</td><td>N/A</td><td>N/A</td><td>collected from Sea beach of Universiti Malaysia Terengganu</td></tr><tr><td>First Aid Kits</td><td>Watsons Malaysia</td><td>N/A</td><td>N/A</td></tr><tr><td>Flat head nickel steel metal rod (Screw driver)</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Formaldehyde</td><td>Thermo Scientific Chemicals</td><td>119690010</td><td></td></tr><tr><td>Gas cylinder (butane gas) for blow&amp;nbsp; torch</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Gas lighter gun (long head)</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Glass beaker (100 mL))</td><td>Corning Life Sciences</td><td>1000-100</td><td></td></tr><tr><td>Ice bag&amp;nbsp;</td><td>Watsons Malaysia</td><td>N/A</td><td>N/A</td></tr><tr><td>Perforated plastic baskets&amp;nbsp;</td><td>Eco-Shop Marketing Sdn. Bhd.</td><td>N/A</td><td>N/A</td></tr><tr><td>PVC pipe 15mm</td><td>Bina Plastic Industries Sdn Bhd (HQ)</td><td>N/A</td><td>N/A</td></tr><tr><td>Refractometer</td><td>ATAGO CO.,LTD.</td><td></td><td></td></tr><tr><td>Refrigerator</td><td>Sharp Corporation Japan</td><td>N/A</td><td>Chest Freezer SHARP 110L - SJC 118</td></tr><tr><td>Scoop net</td><td>MR D.I.Y. Group Berhad</td><td></td><td>N/A</td></tr><tr><td>Seawater</td><td>Hatcheri AKUATROP&amp;nbsp;</td><td>N/A</td><td>Research Center of Universiti Malaysia Terengganu</td></tr><tr><td>Siphoning pipe</td><td>MR D.I.Y. Group Berhad</td><td>N/A</td><td>N/A</td></tr><tr><td>Spray bottle</td><td>Mr. DIY Sdn Bhd</td><td>N/A</td><td>N/A</td></tr><tr><td>Stainless surgical forceps&amp;nbsp;</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>Stainless surgical scissors&amp;nbsp;</td><td>N/A</td><td>N/A</td><td>N/A</td></tr><tr><td>Submersible water pump&amp;nbsp;</td><td>AS</td><td>N/A</td><td>model: Astro 4000</td></tr><tr><td>Tincture of iodine solution&amp;nbsp; (Povidone Iodine)</td><td>Farmasi Fajr Sdn Bhd</td><td>N/A</td><td>N/A</td></tr><tr><td>Tissue paper&amp;nbsp;</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Transparent plastic aquarium</td><td>Ming Yu Three</td><td>N/A</td><td>aquarium and pet shop</td></tr><tr><td>Waterproof table</td><td>Hatcheri AKUATROP&amp;nbsp;</td><td>N/A</td><td>Research Center of Universiti Malaysia Terengganu</td></tr></tbody></table>

eyestalk ablation to increase ovarian maturation in mud crabs

Mud crabs (Scylla spp.) are commercially important crustacean species that can be found throughout the Indo-West Pacific region. During culture, the induction of ovarian maturation is important to meet the consumer demand for mature mud crabs and hasten seed production. Eyestalk ablation is an effective tool to enhance ovarian maturation in mud crabs. However, there is no standard protocol for the eyestalk ablation of mud crabs. In this study, two eyestalk ablation techniques are described: cauterization (the use of hot metal to ablate the eyestalk of an anesthetized crab) and surgery (the removal of the eyestalk using surgical scissors). Before eyestalk ablation, sexually mature females (CW &#62; 86 mm) were anesthetized using an ice bag (&#8722;20 &#176;C) with seawater. When the water temperature reached 4 &#176;C, the ice bag was removed from the water. Flowing seawater (ambient temperature: 28 &#176;C) was used for recovery from the anesthesia immediately after eyestalk ablation. Mortality did not occur during or after the process of eyestalk ablation. The eyestalk ablation protocol presented here accelerated the ovarian maturation of the mud crabs.

Gonad maturation 
Creamy white ovarian tissues (immature ovaries, stage-1) were found in 100% of the dissected females (n = 6) before performing the eyestalk ablation (Figure 2). The gonad maturation rate of the eyestalk-ablated female crabs (n = 63; 31 females with the cauterization technique and 32 females with the surgery technique) was higher compared to female crabs that were not subjected to eyestalk ablation (n = 31) after 30 days of individual rearing (<strong cla...

Watch this Scientific Journal Video about Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs at JoVE.com

Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs

Two eyestalk ablation protocols (i.e., cauterization and surgery&#160;approaches) were performed on anesthetized female crabs. The eyestalk ablation of mud crabs hastened the ripening of ovaries without decreasing the survival rate.

Mud crabs (Scylla spp.) are commercially important crustacean species that can be found throughout the Indo-West Pacific region. During culture, the ...

eyestalk-ablation-to-increase-ovarian-maturation-in-mud-crabs

Research

JoVE Journal

Biology

1.7K Views.  Universiti Malaysia Terengganu. Het beschreven protocol minimaliseert stress tijdens oogsteelablatie van modderkrabben, waardoor de sterfte als gevolg van oogsteelablatie wordt verminderd. Tijdsefficiëntie is de sleutel tot het minimaliseren van stress voor de dieren. Zorg ervoor dat alles is voorbereid voordat u de procedure uitvoert.Begin met het wassen van de gezonde, actieve en premature vrouwelijke modderkrabben met gechloreerd kraanwater om vuil en parasieten te verwijderen. Week de krabben 30 minuten in forma...

Video: Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during oogenesis. In order to study roles of such maternal factors in early embryonic development, it is desirable to manipulate their functions from the very beginning of embryonic development. Conventional ways of gene interference are achieved by injection of antisense oligonucleotides (oligos) or mRNA into fertilized eggs, enabling under- or over-expression of specific proteins, respectively. However, these methods normally require more than several hours until protein expression is affected, and, hence, the interference of gene functions is not effective during early embryonic stages. Here, we introduce an experimental system in which expression levels of maternal proteins can be altered before fertilization. Xenopus laevis oocytes obtained from ovaries are defolliculated by incubating with enzymes. Antisense oligos or mRNAs are injected into defolliculated oocytes at the germinal vesicle (GV) stage. These oocytes are in vitro matured to eggs at the metaphase II (MII) stage, followed by intracytoplasmic sperm injection (ICSI). By this way, up to 10% of ICSI embryos can reach the swimming tadpole stage, thus allowing functional tests of specific gene knockdown or overexpression. This approach can be a useful way to study roles of maternally stored factors in early embryonic development.

Manipulation and In Vitro Maturation of Xenopus laevis Oocytes, Followed by Intracytoplasmic Sperm Injection, to Study Embryonic Development

We describe methods of manipulating Xenopus laevis immature oocytes, in vitro maturation of oocytes to eggs, and intracytoplasmic sperm injection. This protocol allows degradation of some maternal proteins and overexpression of genes of interest at fertilization, and hence is valuable to study roles of specific factors in early embryonic development.

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during ...

Developmental Biology

Functional Cloning Using a Xenopus Oocyte Expression System

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in abundance, may not be secreted but tethered to the signaling cell(s), or may require the presence of binding partners or upstream regulatory molecules. Thus in a search for gene products capable of eliciting an early lens-inductive response in competent ectoderm, we utilized an expression cloning system that would allow identification of paracrine or juxtacrine factors as well as transcriptional or other regulatory proteins. Pools of mRNA were injected into Xenopus oocytes, and responding tissue placed directly on the oocytes and co-cultured. Following functional cloning of ldb1 from a neural plate stage cDNA library based on its ability to elicit the expression of the early lens placode marker foxe3 in lens-competent animal cap ectoderm, we characterized the mRNA expression pattern, and assayed developmental progression following overexpression or knockdown of ldb1. This system is suitable in a very wide variety of contexts where identification of an inducer or its upstream regulatory molecules is sought using a functional response in competent tissue.

Functional Cloning Using a Xenopus Oocyte Expression System

We describe a Xenopus oocyte and animal cap system for the expression cloning of genes capable of inducing a response in competent ectoderm, and discuss techniques for the subsequent analysis of such genes. This system is useful in the functional identification of a wide range of gene products.

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in ...

Dissection for Identifying Migratory Insects via Ovarian Reproductive Analysis

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.

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.

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

Behavior

Extraction and Quantification of Volatile Sex Pheromone Responses in C. elegans

Volatile Sex Pheromone Extraction and Chemoattraction Assay in Caenorhabditis elegans

Chemical communication is vital in organismal health, reproduction, and overall well-being. Understanding the molecular pathways, neural processes, and computations governing these signals remains an active area of research. The nematode Caenorhabditis elegans provides a powerful model for studying these processes as it produces a volatile sex pheromone. This pheromone is synthesized by virgin females or sperm-depleted hermaphrodites and serves as an attractant for males.
This protocol describes a detailed method for isolating the volatile sex pheromone from several C. elegans strains (WT strain N2, daf-22, and fog-2) and C. remanei. We also provide a protocol for quantifying the male chemotaxis response to the volatile sex pheromone. Our analysis utilizes measurements such as chemotaxis index (C.I.), arrival time (A.T.), and a trajectory plot to compare male responses under various conditions accurately. This method allows for robust comparisons between males of different genetic backgrounds or developmental stages. Furthermore, the experimental setup outlined here is adaptable to investigating other chemoattraction chemicals.

Author Spotlight: Examining Volatile Sex Pheromone Influence on Male C. elegans Behavior

This protocol establishes methods for extracting and quantifying responses to the volatile sex pheromone in C. elegans, providing tools to study chemical communication and navigation trajectory.

Chemical communication is vital in organismal health, reproduction, and overall well-being. Understanding the molecular pathways, neural processes, and ...

Multi-Photon Laser Ablation of Cytoplasmic Microtubule Organizing Centers in Mouse Oocytes

The fidelity of oocyte meiosis is critical for generating developmentally competent euploid eggs. In mammals, the oocyte undergoes a lengthy arrest at prophase I of the first meiotic division. After puberty and upon meiotic resumption, the nuclear membrane disassembles (nuclear envelope breakdown), and the spindle is assembled mainly at the oocyte center. Initial central spindle positioning is essential to protect against abnormal kinetochore-microtubule (MT) attachments and aneuploidy. The centrally positioned spindle migrates in a time-sensitive manner toward the cortex, and this is a necessary process to extrude a tiny polar body. In mitotic cells, spindle positioning relies on the interaction between centrosome-mediated astral MTs and the cell cortex. On the contrary, mouse oocytes lack classic centrosomes and, instead, contain numerous acentriolar MT organizing centers (MTOCs). At the metaphase I stage, mouse oocytes have two different sets of MTOCs: (1) MTOCs that are clustered and sorted to assemble spindle poles (polar MTOCs), and (2) metaphase cytoplasmic MTOCs (mcMTOCs) that remain in the cytoplasm and do not contribute directly to spindle formation but play a crucial role in regulating spindle positioning and timely spindle migration. Here, a multi-photon laser ablation method is described to selectively deplete endogenously labeled mcMTOCs in oocytes collected from Cep192-eGfp reporter mice. This method contributes to the understanding of the molecular mechanisms underlying spindle positioning and migration in mammalian oocytes.

An optimized protocol is presented that enables the depletion of cytoplasmic microtubule organizing centers in mouse oocytes during metaphase I using a near-infrared femtosecond laser.

The fidelity of oocyte meiosis is critical for generating developmentally competent euploid eggs. In mammals, the oocyte undergoes a lengthy arrest at ...

Tracking Calcium in the Early Developmental Phases of C. elegans

In Vivo Visualization of Calcium Transients during Fertilization and Early Development in C. elegans

Calcium is an important signaling molecule during the oocyte-to-embryo transition (OET) and early embryogenesis. The hermaphroditic nematode Caenorhabditis elegans provides several unique advantages for the study of the OET as it is transparent and has an ordered gonad that produces one mature oocyte every ~23 min at 20 &#176;C. We have modified the genetically encoded calcium indicator jGCaMP7s to fluorescently indicate the moment of fertilization within a living organism. We have termed this reporter &#34;CaFE&#34; for Calcium during Fertilization in C. elegans. The CaFE reporter was&#160;engineered into a safe harbor locus in single copy, has no significant impact on physiology or fecundity, and produces a robust signal upon fertilization. Here, a series of protocols is presented for utilizing the CaFE reporter as an in vivo tool for dissecting the OET and embryogenesis. We include methods to synchronize worms, examine the effects of RNAi knockdown, mount worms for imaging, and to visualize calcium in oocytes and embryos. Additionally, we present the generation of additional worm strains to aid in this type of analysis. Demonstrating the utility of the CaFE reporter to visualize the timing of fertilization, we report that double ovulation occurs when ipp-5 is targeted by RNAi and that only the first oocyte undergoes immediate fertilization. Furthermore, the discovery of single-cell calcium transients during early embryogenesis is reported here, demonstrating that the CaFE reporter persists into early development. Importantly, the CaFE reporter in worms is simple enough to use for incorporation into course-based undergraduate research (CURE) laboratory classes. The CaFE reporter, coupled with the ordered gonad and ease of RNAi in worms, facilitates inquiry into the cell-cell dynamics required to regulate internal fertilization and early embryogenesis.

Here, we present protocols and tools for visualizing calcium during fertilization and early embryogenesis using a genetically encoded calcium reporter expressed in the germline of the model nematode C. elegans.

Calcium is an important signaling molecule during the oocyte-to-embryo transition (OET) and early embryogenesis. The hermaphroditic nematode ...

Desensitization and Recovery of Crayfish Photoreceptors Upon Delivery of a Light Stimulus

A method to study desensitization and recovery of crayfish photoreceptors is presented. We performed intracellular electrical recordings of photoreceptor cells in isolated eyestalks using the discontinuous single electrode-switched voltage-clamp configuration. First, with a razor blade we made an opening in the dorsal cornea to get access to the retina. Thereafter, we inserted a glass electrode through the opening, and penetrated a cell as reported by the recording of a negative potential. Membrane potential was clamped at the photoreceptor's resting potential and a light-pulse was applied to activate currents. Finally, the two light-flash protocol was employed to measure current desensitization and recovery. The first light-flash triggers, after a lag period, the transduction ionic current, which after reaching a peak amplitude decays towards a desensitized state; the second flash, applied at varying time intervals, assesses the state of the light-activated conductance. To characterize the light-elicited current, three parameters were measured: 1) latency (the time elapsed between light flash delivery and the moment in which current achieves 10% of its maximum value); 2) peak current; and 3) desensitization time constant (exponential time constant of the current decay phase). All parameters are affected by the first pulse.
To quantify recovery from desensitization, the ratio p2/p1 was employed versus time between pulses. p1 is the peak current evoked by the first light-pulse, and p2 is the peak current evoked by the second pulse. These data were fitted to a sum of exponential functions. Finally, these measurements were carried out as function of circadian time.

A protocol for the study of desensitization and sensitivity recovery of crayfish photoreceptors as a function of circadian time is presented.

A method to study desensitization and recovery of crayfish photoreceptors is presented. We performed intracellular electrical recordings of photoreceptor ...

Neuroscience

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

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

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

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

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

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

Toxicity of misfolded proteins and mitochondrial dysfunction are pivotal factors that promote age-associated functional neuronal decline and neurodegenerative disease across species. Although these neurotoxic challenges have long been considered to be cell-intrinsic, considerable evidence now supports that misfolded human disease proteins originating in one neuron can appear in neighboring cells, a phenomenon proposed to promote pathology spread in human neurodegenerative disease.
C. elegans adult neurons that express aggregating proteins can extrude large (~4 &#181;m) membrane-surrounded vesicles that can include the aggregated protein, mitochondria, and lysosomes. These large vesicles are called &#8220;exophers&#8221; and are distinct from exosomes (which are about 100x smaller and have different biogenesis). Throwing out cellular debris in exophers may occur by a conserved mechanism that constitutes a fundamental, but formerly unrecognized, branch of neuronal proteostasis and mitochondrial quality control, relevant to processes by which aggregates spread in human neurodegenerative diseases.
While exophers have been mostly studied in animals that express high copy transgenic mCherry within touch neurons, these protocols are equally useful in the study of exophergenesis using fluorescently tagged organelles or other proteins of interest in various classes of neurons.
Described here are the physical features of C. elegans exophers, strategies for their detection, identification criteria, optimal timing for quantitation, and animal growth protocols that control for stresses that can modulate exopher production levels. Together, details of protocols outlined here should serve to establish a standard for quantitative analysis of exophers across laboratories. This document seeks to serve as a resource in the field for laboratories seeking to elaborate molecular mechanisms by which exophers are produced and by which exophers are reacted to by neighboring and distant cells.

Quantitative Approaches for Scoring in vivo Neuronal Aggregate and Organelle Extrusion in Large Exopher Vesicles in C. elegans

This protocol describes approaches for detection and quantitation of large aggregate and/or organelle extrusions (~4 &#181;m) produced by C. elegans cells in the form of membrane-bound exophers. We describe strains, growth conditions, scoring criteria, timing, and microscopy considerations needed to facilitate dissection of this debris expulsion mechanism.

Toxicity of misfolded proteins and mitochondrial dysfunction are pivotal factors that promote age-associated functional neuronal decline and ...

Unraveling the Role of Discrete Areas of the Rat Brain in the Regulation of Ovulation through Reversible Inactivation by Tetrodotoxin Microinjections

Many experimental approaches have been used for studying the role of the brain in the regulation of ovulation. Examples include the lesion and deafferentation of neuronal groups, which are both invasive methods that permanently impair the integrity of the target area. These methods are accompanied by collateral effects that can affect the analysis of acute and temporal regulatory mechanisms. The stereotaxic implantation of guide cannulas aimed at specific brain regions, followed by a recovery period, allows researchers to microinject different drugs after the disappearance of the undesired effects of the surgery. Tetrodotoxin has been used to determine the roles of several brain areas in diverse physiological processes because it transiently inhibits the sodium-dependent action potentials, thus blocking all neural activity in the target region. This protocol combines this method with strategies for the assessment of the estrous cycle and ovulation to reveal the role of discrete brain regions in the regulation of ovulation at particular times of any given stage of the estrous cycle. Awake and unrestrained rats (Rattus norvegicus) were used to avoid the blocking effects that anesthetics and stress hormones exert on ovulation. This protocol can be easily adapted to other species, brain targets and pharmacological agents to study different physiological processes. Future improvements to this method include the design of a microinjection system using glass capillaries of small diameter instead of guide cannulas. This will reduce the amount of tissue damaged during the implantation and decrease the spread of the infused drugs outside the target area.

This protocol describes the construction of a low-cost microinjection system, its stereotaxic implantation into deep-brain structures and the procedure for timed microinjections of tetrodotoxin in awake and unrestrained rats. The goal is to reveal the participation of hypothalamic structures in the regulation of ovulation by inhibiting their neural activity.

Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs

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