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&#176;66&#8242;62&#8242;&#8242;N, 102&#176;72&#8242;33&#8242;&#8242;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). 
	&#8203;NOTE: Pre-mature female crabs have triangular and light-colored abdomen flaps along with a CW range of 80&#8722;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="Figure 1" class="xfigimg" src="/files/ftp_upload/65039/65039fig01.jpg" /> 
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%&#8722;5% of the crab&#39;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%&#8722;7% of the crab&#39;s body weight.</li>
	<li>Rear the crabs until sexually mature through molting (35 days). 
	&#8203;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 &#62;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&#160;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&#8722;15 s until the eyestalk turns an orange or reddish-orange color. Be careful when conducting this step to avoid damage to adjacent structures.&#160; 
		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&#160;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 &#176;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. 
	&#8203;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 &#8722;20 &#176;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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Aeration tube&#160;</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&#160;</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.&#160;</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Cotton hand gloves (thick)&#160;</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&#160;Instruments&#160;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&#160;</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&#160;</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&#160; 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&#160;</td>
			<td>Watsons Malaysia</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Perforated plastic baskets&#160;</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&#160;</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&#160;</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Stainless surgical scissors&#160;</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Submersible water pump&#160;</td>
			<td>AS</td>
			<td>N/A</td>
			<td>model: Astro 4000</td>
		</tr>
		<tr>
			<td>Tincture of iodine solution&#160; (Povidone Iodine)</td>
			<td>Farmasi Fajr Sdn Bhd</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Tissue paper&#160;</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&#160;</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 c...

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

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1.6K Views.  Universiti Malaysia Terengganu. Two eyestalk ablation protocols (i.e., cauterization and surgery approaches) were performed on anesthetized female crabs. The eyestalk ablation of mud crabs hastened the ripening of ovaries without decreasing the survival rate.

Video: Eyestalk Ablation to Increase Ovarian Maturation in Mud Crabs

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In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Laser Ablation of the Zebrafish Pronephros to Study Renal Epithelial Regeneration

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

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

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

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

Endothelial Cell Co-culture Mediates Maturation of Human Embryonic Stem Cell to Pancreatic Insulin Producing Cells in a Directed Differentiation Approach

Embryonic stem cells (ESC) have two main characteristics: they can be indefinitely propagated in vitro in an undifferentiated state and they are pluripotent, thus having the potential to differentiate into multiple lineages. Such properties make ESCs extremely attractive for cell based therapy and regenerative treatment applications 1. However for its full potential to be realized the cells have to be differentiated into mature and functional phenotypes, which is a daunting task. A promising approach in inducing cellular differentiation is to closely mimic the path of organogenesis in the in vitro setting. Pancreatic development is known to occur in specific stages 2, starting with endoderm, which can develop into several organs, including liver and pancreas. Endoderm induction can be achieved by modulation of the nodal pathway through addition of Activin A 3 in combination with several growth factors 4-7. Definitive endoderm cells then undergo pancreatic commitment by inhibition of sonic hedgehog inhibition, which can be achieved in vitro by addition of cyclopamine 8. Pancreatic maturation is mediated by several parallel events including inhibition of notch signaling; aggregation of pancreatic progenitors into 3-dimentional clusters; induction of vascularization; to name a few. By far the most successful in vitro maturation of ESC derived pancreatic progenitor cells have been achieved through inhibition of notch signaling by DAPT supplementation 9. Although successful, this results in low yield of the mature phenotype with reduced functionality. A less studied area is the effect of endothelial cell signaling in pancreatic maturation, which is increasingly being appreciated as an important contributing factor in in-vivo pancreatic islet maturation 10,11.
The current study explores such effect of endothelial cell signaling in maturation of human ESC derived pancreatic progenitor cells into insulin producing islet-like cells. We report a multi-stage directed differentiation protocol where the human ESCs are first induced towards endoderm by Activin A along with inhibition of PI3K pathway. Pancreatic specification of endoderm cells is achieved by inhibition of sonic hedgehog signaling by Cyclopamine along with retinoid induction by addition of Retinoic Acid. The final stage of maturation is induced by endothelial cell signaling achieved by a co-culture configuration. While several endothelial cells have been tested in the co-culture, herein we present our data with rat heart microvascular endothelial Cells (RHMVEC), primarily for the ease of analysis.

The current study describes a directed differentiation approach in inducing pancreatic differentiation of human embryonic stem cells. Of great significance is the finding that endothelial cell co-culture mediates maturation of human embryonic stem cell derived pancreatic progenitors into insulin expressing cells.

Embryonic stem cells (ESC) have two main characteristics: they can be indefinitely propagated in vitro in an undifferentiated state and they are ...

Validation of a Mouse Model to Disrupt LINC Complexes in a Cell-specific Manner

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the Nucleoskeleton and Cytoskeleton (LINC complexes). These protein scaffolds span the nuclear envelope and connect the interior of the nucleus to components of the surrounding cytoplasmic cytoskeleton. LINC complexes consist of two evolutionary-conserved protein families, Sun proteins and Nesprins that harbor C-terminal molecular signature motifs called the SUN and KASH domains, respectively. Sun proteins are transmembrane proteins of the inner nuclear membrane whose N-terminal nucleoplasmic domain interacts with the nuclear lamina while their C-terminal SUN domains protrudes into the perinuclear space and interacts with the KASH domain of Nesprins. Canonical Nesprin isoforms have a variable sized N-terminus that projects into the cytoplasm and interacts with components of the cytoskeleton. This protocol describes the validation of a dominant-negative transgenic mouse strategy that disrupts endogenous SUN/KASH interactions in a cell-type specific manner. Our approach is based on the Cre/Lox system that bypasses many drawbacks such as perinatal lethality and cell nonautonomous phenotypes that are associated with germline models of LINC complex inactivation. For this reason, this model provides a useful tool to understand the role of LINC complexes during development and homeostasis in a wide array of tissues.

Nuclear envelope proteins play a central role in many basic biological processes and have been implicated in a variety of human diseases. This protocol describes a new Cre/Lox-based mouse model that allows for the spatiotemporal control of LINC complexes disruption.

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the ...

Benefits of Cardiac Resynchronization Therapy in an Asynchronous Heart Failure Model Induced by Left Bundle Branch Ablation and Rapid Pacing

It is now well recognized that heart failure (HF) patients with left bundle branch block (LBBB) derive substantial clinical benefits from cardiac resynchronization therapy (CRT), and LBBB has become one of the important predictors for CRT response. The conventional tachypacing-induced HF model has several major limitations, including absence of stable LBBB and rapid reversal of left ventricular (LV) dysfunction after cessation of pacing. Hence, it is essential to establish an optimal model of chronic HF with isolated LBBB for studying CRT benefits. In the present study, a canine model of asynchronous HF induced by left bundle branch (LBB) ablation and 4 weeks of rapid right ventricular (RV) pacing is established. The RV and right atrial (RA) pacing electrodes via the jugular vein approach, together with an epicardial LV pacing electrode, were implanted for CRT performance. Presented here are the detailed protocols of radiofrequency (RF) catheter ablation, pacing leads implantation, and rapid pacing strategy. Intracardiac and surface electrograms during operation were also provided for a better understanding of LBB ablation. Two-dimensional speckle tracking imaging and aortic velocity time integral (aVTI) were acquired to validate the chronic stable HF model with LV asynchrony and CRT benefits. By coordinating ventricular activation and contraction, CRT uniformed the LV mechanical work and restored LV pump function, which was followed by reversal of LV dilation. Moreover, the histopathological study revealed a significant restoration of cardiomyocyte diameter and collagen volume fraction (CVF) after CRT performance, indicating a histologic and cellular reverse remodeling elicited by CRT. In this report, we described a feasible and valid method to develop a chronic asynchronous HF model, which was suitable for studying structural and biologic reverse remodeling following CRT.

The establishment of a chronic asynchronous heart failure (HF) model by rapid pacing combined with left bundle branch ablation is presented. Two-dimensional speckle tracking imaging and aortic velocity time integral are applied to validate this stable HF model with left ventricular asynchrony and the benefits of cardiac resynchronization therapy.

It is now well recognized that heart failure (HF) patients with left bundle branch block (LBBB) derive substantial clinical benefits from cardiac ...

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

Ovarian control and monitoring in amphibians require a multi-faceted approach. There are several applications that can successfully induce reproductive behaviors and the acquisition of gametes and embryos for physiological or molecular research. Amphibians contribute to one-quarter to one-third of vertebrate research, and of interest in this context is their contribution to the scientific community&#39;s knowledge of reproductive processes and embryological development. However, most of this knowledge is derived from a small number of species. In recent times, the decimation of amphibians across the globe has required increasing intervention by conservationists. The captive recovery and assurance colonies that continue to emerge in response to the extinction risk make existing research and clinical applications invaluable to the survival and reproduction of amphibians held under human care. The success of any captive population is founded on its health and reproduction and the ability to develop viable offspring that carry forward the most diverse genetic representation of their species. For researchers and veterinarians, the ability to monitor and control ovarian development and health is, therefore, imperative. The focus of this article is to highlight the different assisted reproductive techniques that can be used to monitor and, where appropriate or necessary, control ovarian function in amphibians. Ideally, any reproductive and health issues should be reduced through proper captive husbandry, but, as with any animal, issues of health and reproductive pathologies are inevitable. Non-invasive techniques include behavioral assessments, visual inspection and palpation and morphometric measurements for the calculation of body condition indices and ultrasound. Invasive techniques include hormonal injections, blood sampling, and surgery. Ovarian control can be exercised in a number of ways depending on the application required and species of interest.

The study of amphibian biology provides valuable information on the reproductive, physiological, embryological and developmental processes that drive organisms of many taxonomic groups. Here, we present a comprehensive guide on different methodologies that can be used to study ovarian control and monitoring in amphibians.

Ovarian control and monitoring in amphibians require a multi-faceted approach. There are several applications that can successfully induce reproductive ...

Bovine Ovarian Cortex Tissue Culture

Follicle development from the primordial to antral stage is a dynamic process within the ovarian cortex, which includes endocrine and paracrine factors from somatic cells and cumulus cell-oocyte communication. Little is known about the ovarian microenvironment and how the cytokines and steroids produced in the surrounding milieu affect follicle progression or arrest. In vitro culture of ovarian cortex enables follicles to develop in a normalized environment that remains supported by adjacent stroma. Our objective was to determine the effect of nutritional Stair-Step diet on the ovarian microenvironment (follicle development, steroid, and cytokine production) through in vitro culture of bovine ovarian cortex. To accomplish this, ovarian cortical pieces were removed from heifers undergoing two different nutritionally developed schemes prior to puberty: Control (traditional nutrition development) and Stair-Step (feeding and restriction during development) that were cut into approximately 0.5-1 mm3 pieces. These pieces were subsequently passed through a series of washes and positioned on a tissue culture insert that is set into a well containing Waymouth&#39;s culture medium. Ovarian cortex was cultured for 7 days with daily culture media changes. Histological sectioning was performed to determine follicle stage changes before and after the culture to determine effects of nutrition and impact of culture without additional treatment. Cortex culture medium was pooled over days to measure steroids, steroid metabolites, and cytokines. There were tendencies for increased steroid hormones in ovarian microenvironment that allowed for follicle progression in the Stair-Step versus Control ovarian cortex cultures. The ovarian cortex culture technique allows for a better understanding of the ovarian microenvironment, and how alterations in endocrine secretion may affect follicle progression and growth from both in vivo and in vitro treatments. This culture method may also prove beneficial for testing potential therapeutics that may improve follicle progression in women to promote fertility.

In vitro culture of bovine ovarian cortex and the effect of nutritional Stair-step diet on ovarian microenvironment is presented. Ovarian cortex pieces were cultured for seven days and steroids, cytokines, and follicle stages were evaluated. The Stair-Step diet treatment had increased steroidogenesis resulting in follicle progression in culture.

Follicle development from the primordial to antral stage is a dynamic process within the ovarian cortex, which includes endocrine and paracrine factors ...

Megakaryocyte Culture in 3D Methylcellulose-Based Hydrogel to Improve Cell Maturation and Study the Impact of Stiffness and Confinement

The 3D environment leading to both confinement and mechanical constraints is increasingly recognized as an important determinant of cell behavior. 3D culture has thus been developed to better approach the in vivo situation. Megakaryocytes differentiate from hematopoietic stem and progenitor cells (HSPCs) in the bone marrow (BM). The BM is one of the softest tissues of the body, confined inside the bone. The bone being poorly extensible at the cell scale, megakaryocytes are concomitantly subjected to a weak stiffness and high confinement. This protocol presents a method for the recovery of mouse lineage negative (Lin-) HSPCs by immuno-magnetic sorting and their differentiation into mature megakaryocytes in a 3D medium composed of methylcellulose. Methylcellulose is non-reactive towards megakaryocytes and its stiffness may be adjusted to that of normal bone marrow or increased to mimic a pathological fibrotic marrow. The process to recover the megakaryocytes for further cell analyses is also detailed in the protocol. Although proplatelet extension is prevented within the 3D milieu, it is described below how to resuspend the megakaryocytes in liquid medium and to quantify their capacity to extend proplatelets. Megakaryocytes grown in 3D hydrogel have a higher capacity to form proplatelets compared to those grown in a liquid milieu. This 3D culture allows i) to differentiate progenitors towards megakaryocytes reaching a higher maturation state, ii) to recapitulate phenotypes that may be observed in vivo but go unnoticed in classical liquid cultures, and iii) to study transduction pathways induced by the mechanical cues provided by a 3D environment.

It is now acknowledged that the three-dimensional environment of cells can play an important role in their behavior, maturation and/or differentiation. This protocol describes a three-dimensional cell culture model designed to study the impact of physical containment and mechanical constraints on megakaryocytes.

The 3D environment leading to both confinement and mechanical constraints is increasingly recognized as an important determinant of cell behavior. 3D ...