Name: reproductive techniques for ovarian monitoring and control in amphibians
Uploaded: 2019-05-12T12:30:00.000+00:00
Duration: 4 min 37 s
Description: Watch this Scientific Journal Video about Reproductive Techniques for Ovarian Monitoring and Control in Amphibians at JoVE.com

Natalie Calatayud would like to thank Dr. Barbara Durrant for training and assistance with ultrasound and to Exploradora de Immuebles, S.A. (EISA) for awarding financial aid to my research associate position at SDZG. Thanks to Dr. Kylie Cane for comments on the manuscript as well as to the official reviewers (whoever they may be). Thanks to Jonathan Dain our 2018 Summer Fellow, San Diego Zoo Institute for Conservation Research for providing photos (Figure 1A,B). Monica Stoops extends appreciation to the Association of Zoos and Aquariums Conservation Endowment Fund and the Disney Worldwide Conservation Fund for providing financial support to establish the captive Necturus population. Additionally, support was also received through private donations from amphibian advocate Ms. Iris de la Motte. Thanks, is given to Mr. Christopher DeChant and Dr. Mark Campbell for their significant contribution to the research.

Salamander procedures were approved by Cincinnati Zoo &#38; Botanical Garden (CZBG) Institutional Care and Use Committee (IACUC) protocols 11-106, 13-110, 14-133, and 15-138. All frog and toad procedures were approved by the San Diego Zoo Global (SDZG), Institutional Care and Use Committee (IACUC) protocols: 15-001, 16-005 and 18-003.
The care and treatment of animals was approved by the Ethical Committee of the National Museum of Natural History (Paris) (Museum National d&#39;Histoire Naturelle-M&#233;nagerie du Jardin des Plantes (MNHN)), in accordance with the Institutional and National Guidelines (Commission de G&#233;nie G&#233;n&#233;tique, Direction D&#233;partementale des Services V&#233;t&#233;rinaires, European Union Directive 2010/63/EU, agreement decision No. C75-05-01-2 for the European Convention for vertebrate animals used for experimental and other scientific purposes. All protocols used in this study were approved under the reference number 68-037.
1. Non-invasive ovarian monitoring techniques
<ol>
	<li>Visual inspection and palpation
	<ol>
		<li>Hold the female Anuran in one of the three ways described below.
		<ol>
			<li>Secure the frog or toad&#39;s legs with the ring and little finger, supporting the dorsal side (abdomen) of the frog&#8217;s body with the index and middle finger and the ventral side with the thumb (Figure 1A).</li>
			<li>Hold the frog or toad in the dominant hand with a thumb on its abdomen and the rest of the fingers securing the back of the animal. Using the non-dominant hand to palpate the animal&#8217;s abdomen, feel if there is subdermal bumpiness (Figure 1B).</li>
			<li>Rest the frog or toad&#39;s abdomen on the palm of the hand, front arms draped over the index finger and a thumb on its upper back.</li>
		</ol>
		</li>
		<li>Since Caudates are fully aquatic in nature, perform visual inspection by one of two methods described below.
		<ol>
			<li>Move animal to a separate 4 L container containing tank water. Hold the container (lid secured) up and shine flashlight on the underside to visualize the presence/absence of eggs.</li>
			<li>Anesthetize in MS222 (0.5 g/L; Tricaine methanesulfonate, buffered with 0.5 M NaHCO3). Following induction, rotate animal onto its back and examine abdomen.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Morphometric assessments
	<ol>
		<li>Anurans 
		NOTE: Anesthesia is not required.
		<ol>
			<li>Using calipers, measure the animal from the tip of the mouth, along the center of the body to the tip of the tail to obtain SUL and SVL (Figure 2A,B).</li>
			<li>Tare a plastic container on a digital precision scale. Place the animal in a tared container and weigh (Figure 2C).</li>
			<li>For larger animals, like bullfrogs, or when obtaining weights in the field, use a hanging scale (Figure 2D).</li>
			<li>As in many anuran species, distinguish R. muscosa adult females from males by their larger size and the lack of nuptial (thumb) pads on the hands (Figure 3).</li>
			<li>Calculate the body condition, as a basic assessment of overall health by the following formula: 
			Fulton&#39;s index: K = mass &#247; length3 
			NOTE: Fulton&#39;s Index uses a dimensional balance of volume related to mass and length where 3 is the scaling exponent that relates mass and length isometrically.</li>
		</ol>
		</li>
		<li>Caudates
		<ol>
			<li>Tare scale with an empty bag before placing the non-anesthetized animal inside. Take care to not introduce excess water (Figure 2D) and act quickly as animals secrete mucus as a stress response to being restrained.</li>
			<li>Obtain adult measures by immobilizing individuals in a straight position at the bottom of a re-sealable plastic bag or in a separate plastic container that can accommodate expanded calipers.</li>
			<li>Measure the body length with calipers (Figure 2E).</li>
			<li>Measure caudate from tip of snout to tip of tail (SVL) to monitor growth.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Behavioral observations
	<ol>
		<li>Physically observe animals in real-time or use a video camera to record behavior.</li>
		<li>Record observations of animals categorize behaviors and construct an ethogram (Figure 4).</li>
		<li>Classify reproductive behaviors 
		NOTE: Figure 4 exemplifies one type of reproductive behavior observed in anurans.</li>
	</ol>
	</li>
	<li>Ultrasound 
	NOTE: The ultrasound transducer of choice, in this instance, 7.5 mHz linear or a multi-frequency (10-6 mHz) micro-convex, is recommend for Necturus and a 10 MHz probe and water soluble, salt-free gel for R. muscosa. Performing ultrasound on salamanders may require anesthesia (see section 1.5 for instructions).
	<ol>
		<li>Anurans
		<ol>
			<li>Perform ultrasound on R. muscosa using two people (Figure 5A).</li>
			<li>First person: Hold the animal with the dominant hand and apply the water soluble, salt free gel to the animal&#8217;s abdomen.</li>
			<li>Second person (ultra-sonographer): Take the 10 MHz probe in the dominant hand and apply it to the abdomen making sure to make good contact between the probe and gel.</li>
			<li>Slide inwards from just below the arm pit towards the center of the animal&#8217;s abdominal midline to visualize the whole ovary.</li>
			<li>Ultra-sonographer: Use non-dominant hand to freeze frame and capture the desired images on the ultrasound.</li>
			<li>Categorize the stage of the ovarian cycle by the grading system established for the genus4 (Table 2, Figure 5B-F).</li>
			<li>Rinse any gel off animal at the end of the procedure.</li>
		</ol>
		</li>
		<li>Caudates
		<ol>
			<li>Transfer non-anesthetized Necturus to 4 L rectangular container filled with 2 L of tank water.</li>
			<li>Minimize animal movement by turning the room lights off and/or cup one hand over the head of the animal.</li>
			<li>Position the transducer at a distance of 1-2 cm from body wall.</li>
			<li>Locate the heart at the ventral midline level to the forelimbs and then move transducer distally and examine the ovarian tissue7.</li>
			<li>Categorize females according to grading system established for the genus4 (Figure 6A,B,C).</li>
			<li>Obtain accurate measures of eggs at mid-to late-gravid stage by capturing images when the animal&#8217;s body is at an angle to the transducer (i.e., not linear, but slight arc; Figure 6B). Otherwise, overlapping follicles make it difficult to differentiate individual egg size.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Anesthetic induction and recovery
	<ol>
		<li>Anurans
		<ol>
			<li>Anesthetize in MS222 (0.5 g/L; Tricaine methanesulfonate buffered (0.5 M NaHCO3) as previously described.</li>
			<li>Use the righting reflex as a primary indicator of the degree to which the animal has become anesthetized. Complete loss of reflex demonstrates a state of deep anesthesia.</li>
			<li>Remove the animal from the water-bath&#8211;based anesthesia (MS-222) once the righting reflex is lost.</li>
			<li>Place the animal on a wet (with anesthetic-free de-chlorinated water) towel.</li>
			<li>Make sure to keep the animal moist during the entire surgical procedure.</li>
			<li>Intubate small amphibians with red rubber catheters, uncuffed tubes or classical cuffed endotracheal tubes without inflating the cuff.</li>
			<li>Provide a low flow of oxygen (0.5-0.75 L/min) with 0.5-1% isoflurane.</li>
			<li>Stop the isoflurane after the procedure but keep the flow of oxygen for 1 minute.</li>
			<li>Extubate the animal and rinse the animal thoroughly with anesthetic-free dechlorinated water for 2 minutes.</li>
			<li>Put the animal in a shallow amount of de-chlorinated water or on a wet towel.</li>
			<li>Evaluate the animal&#39;s recovery by gently pulling on a hind limb to elongate. Any responding contraction of the limb indicates withdrawal reflex.</li>
			<li>Monitor other indicators of recovery such as gular respirations (throat movement) and the righting reflex.</li>
			<li>Consider the amphibian recovered when all of the reflexes have returned, and heart and respiration rates have returned to pre-anesthetic values.</li>
		</ol>
		</li>
		<li>Caudates
		<ol>
			<li>Anesthetize Necturus and Ambystoma in MS222 (0.5 g/L of Tricaine methanesulfonate, buffered with 0.5 M NaHCO3, (MS222) in a 4 L rectangular tank.</li>
			<li>Place an air stone (1 inch) and air pump into the tank and turn it on to a constant flow to provide adequate oxygenation.</li>
			<li>When the limb function and the righting reflex are lost, remove the animal from the water-bath&#8211;based anesthesia (MS-222) and put the animal on a wet (with anesthetic-free de-chlorinated water) towel.</li>
			<li>Maintain skin and gill moisture with a squeeze bottle of tank water.</li>
			<li>To recover the animal, carefully place it ventral side down in a 4 L plastic container filled with 2 L of tank water with an air stone. 
			NOTE: Recovery starts with gill flashing, followed by the ability to move its tail and propel forward and finally functional movement of limbs.</li>
			<li>Return the animal to its original housing tank and monitor closely over the next 24 h. 
			NOTE: Other methods of anesthesia for amphibians exist and these are described in Wright and Whitaker8.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Invasive ovarian monitor and control techniques
NOTE: This procedure has been adapted from Forz&#225;n et al.10.
<ol>
	<li>Hold the frog in the dominant hand, and dab dry the venipuncture side of the frog&#39;s face with a sterile wipe or gauze.</li>
	<li>Dry the face to avoid the blood from dispersing across the skin too much.</li>
	<li>Insert the needle (26 G 1/2&#34; and 27 G 1/2&#34;), with the bevel facing upwards, through the skin where the raised skin around the eye and the upper jaw ridge meet to form the point of a triangle (yellow outline) (Figure 7A) accessing the vena facialis near the vena orbitalis posterior.</li>
	<li>Puncture the facial vein below the right eye and above the upper jaw ridge, starting between 1-2 mm back from the midline of the eye (Figure 7A). 
	NOTE: For smaller frogs (under 20 g), move the insertion point closer to a position directly below the midline of the eye.</li>
	<li>Angle the microhematocrit tube downward to enable gravity to help blood flow into the tube. Blood should flow immediately after puncture (Figure 7B,C).</li>
	<li>At the first sign of blood flow, place the tip of microhematocrit tube at the puncture site and collect 1-2 full microhematocrit tubes of blood and place tubes into suitable receptacles for collection (Figure 7B,C).</li>
	<li>If blood does not flow readily, or the volume is very low, shift the insertion of the needle slightly or insert the needle into the other side of the face.</li>
	<li>Stop bleeding by pressing gauze firmly to the puncture site for at least 20 s.</li>
	<li>Leave frog out of water for 10 minutes to confirm puncture site does not reopen.</li>
	<li>Use a new needle and new microhematocrit tubes for each frog sampled.</li>
</ol>
3. Hormone induction
<ol>
	<li>Hormone preparation
	<ol>
		<li>Prepare hormone injections immediately prior to use to ensure maximum effect.</li>
		<li>Select a hormone from the selection listed in Table 1.</li>
		<li>Determine the concentration of the hormone to be injected using a &#181;L or mL/g of body weight16.</li>
		<li>Dilute the hormone in one of the following: water, phosphate buffered saline (PBS), saline amphibian Ringer&#8217;s solution (SARS) or saline.</li>
		<li>Do not exceed an injection volume of 200 &#181;L for frogs weighing 30-70 g and 300 &#181;L for frogs weighing 80-110 g (personal observation)16.</li>
		<li>For correct holding of an animal during hormone administration of any animal ranging from 10-100 g, use any of the appropriate methods for holding described in section 1.1.</li>
	</ol>
	</li>
	<li>Anurans
	<ol>
		<li>Calculate the concentration required per individual using a gram per body weight calculation (g/body weight).</li>
		<li>Just prior to administration, reconstitute in a sterile diluent of choice.</li>
		<li>Ensure no bubbles are left in the syringe before injection.</li>
		<li>Hold the animal securely in the non-dominant hand and administer the injection with dominant hand.</li>
		<li>Administer injection according to hormone specifications. The most common injections in anurans are sub-cutaneous, intra peritoneal or intra muscular (Figure 8).</li>
		<li>Administer IP injections in the lower part of the abdomen or in the lower section of the dorsal side of the body near the back leg (Figure 9).</li>
		<li>Administer intra-muscular injections preferably into the hind legs.</li>
	</ol>
	</li>
	<li>Caudates (Necturus)
	<ol>
		<li>Reconstitute the hormone of choice in sterile water according to the gram per body weight method described above.</li>
		<li>In the case of Necturus, use doses of 1.7-2.3 &#181;g GnRH/g body weight.</li>
		<li>Remove Necturus from anesthetic chamber and place on a 45&#176; surface covered with surgical drape.</li>
		<li>Position the animal with head pointing down.</li>
		<li>Approach posterior quadrant of the abdomen (caudal of rear leg) at a 15-20&#176; angle. Be careful not to introduce air into the syringe.</li>
		<li>Inject (IP) using an insulin syringe and 27-30 G needle.</li>
		<li>Inject the hormone using an insulin syringe and 27-30 G needle.</li>
	</ol>
	</li>
</ol>
4. Surgery
<ol>
	<li>General surgical preparation and procedure
	<ol>
		<li>To maintain aseptic procedures, use clear sterile plastic drapes to isolate surgical site. Reduce evaporation by keeping the surrounding skin moist.</li>
		<li>Moisten any materials that will contact the animal&#39;s skin with sterile water. Make skin incision with a number 15 or number 11 scalpel blade. 
		NOTE: A combination of cold steel, radiosurgery or diode laser. Hemostasis in mild hemorrhagic procedure may be achieved by electrocauter or diode laser.</li>
		<li>Use cotton-tipped spears or applicators to allow the application of localized pressure to small vessels keeping track of blood loss.</li>
		<li>Use cotton-tipped spears or applicators to manage small confined spaces instead standard gauze squares.</li>
		<li>Use micro-instruments, like ophthalmologic instruments, with fine, small tips, when performing surgery on animals&#39; weighing less than 1 kg.</li>
		<li>Use plastic, self-retaining retractors (e.g., Lone Star retractor) to fit different sizes of incisions.</li>
		<li>Use eyelid retractors for retracting coelomic incisions.</li>
		<li>Use magnification instrumentation where necessary for performing surgery on smaller patients. 
		NOTE: Analgesia is required with any surgical procedure in amphibians. Failure to administer adequate analgesia during surgery has been associated with delayed return of normal functions. Moreover, analgesia potentiates the effects of anesthetic drugs (Table 3)34.</li>
	</ol>
	</li>
	<li>Anurans
	<ol>
		<li>Once X. laevis has been anesthetized as described in step 1.5.1, position the animal in dorsal recumbency (Figure 10A,C).</li>
		<li>Prepare the surgical field aseptically by wiping moist sterile gauze with dilute povidone-iodine solution (1/10) on the site for 10-15 s or 0.75% chlorhexidine solution on the surgical site for at least 10 min before the surgery35.</li>
		<li>Make a 3 mm paramedian skin incision in the mid coelom (between the shoulders and the cloaca) with one bold stroke leaving a clean incision using a no. 15 or no. 11 scalpel. 
		NOTE: One may use a diode laser also for skin incisions.</li>
		<li>Elevate the abdominal membrane, make and incision and dissect carefully using a no. 15 or no. 11 scalpel. (Figure 10B,D).</li>
		<li>Retract the coelomic incisions with eyelid retractors (or any appropriate equipment).</li>
		<li>Excise a portion of egg mass without ligating any blood vessels.</li>
		<li>For complete ovariectomy, cauterize surrounding blood vessels by electrocautery or laser diode (Figure 11).</li>
		<li>Using monofilament suture, close celiotomy incision with an interrupted, everting suture pattern.</li>
	</ol>
	</li>
	<li>Caudates
	<ol>
		<li>Once&#160;A. mexicanum has been anesthetized, place it in the right lateral recumbency, with the left pelvic limb simply placed against the tail base.</li>
		<li>Prepare the surgical field aseptically by placing moist sterile gauze with dilute povidone-iodine solution (1:10) on the site for 10-15 s. Alternatively, use sterile gauze soaked in 0.75% chlorhexidine solution and place on the surgical site for at least 10 min before surgery (Figure 12A)36, 37 .</li>
		<li>Draw a line between the shoulder and the hind limbs to divide the body into three equal parts (Figure 12B).</li>
		<li>Make the incision site between the second and third parts.</li>
		<li>Grasp the underlying muscle and elevate away from the coelomic viscera.</li>
		<li>Gently force small hemostats through the coelomic musculature and into the coelomic cavity.</li>
		<li>Retract the coelomic incisions with eyelid retractors (or any appropriate material) (Figure 12C).</li>
		<li>For complete ovariectomy, cauterize surrounding blood vessels by electrocautery or laser diode (Figure 12D).</li>
		<li>Using monofilament suture, close celiotomy incision with an interrupted, everting suture pattern.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Direct handling, visual observation and morphometric measures provide non-invasive techniques and are the first assessment criteria for determining female reproductive stage. However, this study shows that gravid ovaries cannot always be reliably identified by palpation. Depending on the species, gravid ovaries can sometimes be visually detected through semi-translucent (Figure 13A, B) or completely translucent skin on the ventral side of the animal (Figure 13C). Females that have completed oviposition can show obvious changes to their appearance compared to gravid females (e.g., loose skin and loss of up to 30% of their body mass, Figure 13D). During breeding, males and females will display certain behaviors that provide information about proximity to ovulation and oviposition. In the case of R. muscosa indications that a female is close to oviposition begin with the female entering handstands.
The application of ultrasound technology to&#160;Anurans and Caudates permits the diagnosis of presence or absence of eggs and whether oviposition was associated with complete or partial release of developed eggs. Thus, this method provides a more complete and accurate assessment of reproductive status without being limited to determining gravid/non-gravid status via a visualization technique that varies by abdominal skin transparency, or epidermal consistency among the different amphibian species. Ultrasound can be performed with relative ease and with little stress to the animals (Figure 5 and Figure 13) and can be used to characterize reproductive cycles and to determine&#160;reproductive status4. It is critical to become familiar with the species; however, this study showed that Necturus and R. muscosa share common developmental signs in their reproductive patterns allowing for similar classification of reproductive stage (Figure 5). Through this technology there is now evidence that egg development is high in captive Necturus and R. muscosa and that both these species follow a seasonal pattern. Although the reasons for these phenomena are unknown and require further inquiry, without the use of ultrasound, several areas of ovarian dysfunction, such as egg retention and partial oviposition, would have gone undetected. Future applications to this technique will be used to determine whether females should be selected for breeding in any given year and whether oviposition is complete.
A blood collection protocol, such as that presented in R. muscosa, which is both effective and causes minimal distress to the animal, is optimal to study hormone profiles in captive and wild-caught Anurans (Calatayud, unpublished). To date, no information exists regarding the annual hormone profiles of captive R. muscosa and, therefore, no knowledge on how hormones are influencing their health and reproduction. Moreover, with evidence that females of this species may not be annual breeders, hormonal profiling will be another method for tracking ovarian cycles. Together with ultrasound, hormone analysis may lead to better prediction of what females will be ready for oviposition. Furthermore, in the past year, two cases of intersex in the captive R. muscosa population have been documented. In addition, the development of thumb pads has been noted on some of the older founding females. Reasons for this are currently under investigation but initial results suggest it may relate to changes in testosterone levels (Calatayud, unpublished). Discerning hormonal cycles in females of different ages will help us understand why females may develop male-associated secondary sexual characteristics and whether this is to be expected in an aging population.
Exogenous hormone therapy has been used to overcome reproductive dysfunctions frequently encountered in captive amphibians. However, for both R. muscosa and Necturus populations in this study, no significant differences in oviposition between hormone-treated and control females were detected over a 2 and 5-year period of time, respectively. This may indicate that hormone administration protocol, doses, priming and hormone combination used was not adequate for the species. Closer analysis of individual female reproductive histories suggests R. muscosa may not experience annual breeding, which could also account for the lack of hormonal-effect observed in treated females. Because a certain percentage of females consistently skipped breeding every year, understanding the natural history of the species can help determine whether there is a need for exogenous hormones and when they may be most effective. The procedures outlined in this article can be applied to a number of species, (Table 1) and are for anurans ranging from 5 g to 150 g; larger animals may require different syringes and needle gauges. The location of the injection varies with some hormones requiring intra-muscular, intra-peritoneal, sub-cutaneous or intradermal injection (Figure 7).
Surgery for the purpose of ovariectomy is a common method used in various amphibian species to obtain oocytes for embryological studies. Ovariectomy may also be indicated for population control and medical issues such as egg retention. In the case of partial ovariectomies in which, oocyte harvesting is performed for research purposes, surgery must ensure that the animal remains reproductive. Administration of PGF2&#945; has shown some promise in resolving egg retention in female R. muscosa. In several individuals, PGF2&#945; elicited complete deposition of previously retained eggs but in others only partial deposition occurred requiring manual stripping to remove all the eggs. While PGF2&#945; may serve as an alternative to surgery for egg retention in R. muscosa, its ability to remedy similar pathologic conditions in other amphibians will require species-specific validation. When surgical intervention is mandated for the Anuran or Caudate patient, it is necessary to ensure an adequate plane of anesthesia before incisions are made. Astute observation skills are needed to assess and monitor the normative induction and recovery responses as outlined in this study for each of the taxa. Once one is familiar with the specific anatomy, an appropriate surgical approach, hemostasis, gentle tissue manipulation and adequate postoperative management, reproductive surgeries pose no overwhelming obstacles.&#160;

Amphibians have long been recognized as important biological and medical models by a wide range of research disciplines. Data obtained by studying particular species such as Xenopus laevis and X. tropicalis,&#160;the Leopard frog (Lithobates (formerly Rana) pipiens)&#160;and the axolotl (Ambystoma mexicanum) have&#160;been applied to a number of other vertebrate species, including humans. The veterinary, husbandry and assisted reproductive techniques that have emerged from studying these and other amphibians provide assistance to those tasked with developing successful care, maintenance and sustainability of rarer populations in captivity1,2,3,4.
Interest is gaining for the concurrent use of in and ex situ conservation-based approaches to reverse the tide of extinction for many at risk amphibian species1,2. This article provides the methodologies presently available to monitor and control the amphibian ovarian function in model species of Anurans&#160;and Caudates. Additionally, existing techniques to address a common reproductive pathology of egg retention is presented.
As in many taxonomic groups, amphibian ovarian control involves a series of tightly synchronized interactions between the environment and physiology. Temperature and the photoperiod (known as proximate signals) are decoded by the eye and brain where they are rapidly converted into genetic, hormonal and circadian processes (ultimate signals)3,4. The methods to monitor and control ovarian function covered in this article include invasive and non-invasive techniques. Institutional Animal Care and Use Committee (IACUC) research and teaching requirements define non-invasive techniques as those that will cause minimal to no physical pain or mental distress and require no pain-relieving drugs5. Here, non-invasive techniques include visual inspection and palpation, behavioral observations, morphometric assessments and ultrasound. On the contrary, the techniques of blood collection, hormone administration and surgery (ovariectomy and removal of retained eggs) are classified as invasive since they may result in some pain or discomfort and require anesthesia or post-procedural drug therapy.
Non-invasive ovarian monitoring techniques can be easily incorporated into the daily care routine for most captive amphibians. Depending on the species, ovarian gravidity can often be determined by simple visual inspection (glass frog). In other cases, palpation may indicate whether a female is gravid. Various body condition indices (BCI) such as weight, snout urostyle length (SUL), snout-vent length (SVL) and standard mass index (SMI) are available for predicting the presence or absence of eggs4,6,7,8,9. However, care should be taken with the interpretation of results as most do not consider age, body shape or composition (e.g., water retained versus ovarian mass or fat)6. Definitive reproductive diagnoses can be achieved via ultrasound with more in depth knowledge gained regarding egg development and staging of ovarian cycle4,7. Ultrasound also provides a means to confirm and monitor reproductive pathologies and associated physiological conditions4, 8.
In addition to providing information regarding health status, blood sampling can be used to measure reproductive hormones. If hormone profiling is the final goal, it is important to avoid stress-related influences that may confound systemic steroid data. While a potentially powerful monitoring tool, there is yet to be a study demonstrating innate endocrinological responses to exogenous hormone administration in any amphibian species. Blood can safely be taken from several sites; in frogs this includes the ventral abdominal vein, lingual plexus, femoral vein and heart9, 10. In Caudates, blood is collected from the ventral tail vein. The degree of invasiveness, the amount of restraint required, the need for anesthetic, the delicacy of the organ being targeted, and the size of the animal are factors to be considered when choosing a collection technique for the amphibian patient. This article will present the technique of blood collection from the facial maxillary or musculocutaneous vein of frogs as originally described by Forzan et al.9.
Ovarian control is species-specific and, as such, hormone protocols should be tested and optimized. Other than seasonality and the associated circulating hormone milieu, ovarian control may also be tightly linked to age, time spent in captivity and exposure to repeated hormone administration, for which there is little information in the literature11,12,13. The implementation of hormone therapies to elicit reproductive behaviors, gamete production, maturation and oviposition has become a widely reported approach to resolving common reproductive problems associated with captivity4,8,14,15,16. Because the mechanisms controlling reproduction in vertebrates are highly conserved, there are a number of hormones, neuropeptides and commercially available drugs used therapeutically in other taxonomic groups that can also be used reliably in a number of amphibian species (Table 1). Gonadotropin releasing hormone (GnRH) and Human chorionic gonadotropin (hCG) (or variations thereof, i.e., PMSG and eCG)17, 18, either individually or in combination, have been used extensively in amphibian captive breeding programs including: the Southern Rocky Mountain boreal (Anaxyrus boreas boreas)4,19,20; the toad, Dusky Gopher frog, Rana sevosa (Langhorne et al., unpublished)7; the Gulf Coast Waterdog, Necturus beyeri20; Wyoming toad, Anaxyrus baxteri18; the bullfrog, Rana catesbiana21; the American toad, Anaxyrus americanus22; the grass frog, Lymnodyaster tasmaniensis23; the Coqui, Eleutherodactylus coqui24; the Xenopus, Xenopus laevis25; the Gunther&#39;s toadlet, Pseduophryne guentheri26; the Northern Leopard frog, Lithobates pipiens; the Argentinian Horned-frog, Ceratophrys ornate; the Cranwell&#8217;s horned-frog, C. cranwelli; the American ground-frog, Odontophrynus americanus27; and the fire salamander (Salamandra)228. Steroid hormones, like progesterone (P4), are less commonly reported but have demonstrated good efficacy in eliciting ovulation and oviposition in some species of anurans16,18,29. Prostaglandins (especially Prostaglandin 2-alpha (PGF2&#945;)) are involved in ovulation together with corticosteroids30,31,32,34 and reach high levels during the ovulatory phase31.
In in vitro studies, PGF2&#945; is a potent inducer of ovulation31, while in vivo it can induce oviposition of retained eggs in Rana muscosa4,30,32. Pituitary extracts are also effective inducers of ovulation15,16,34; however, concerns surrounding biosecurity and the potential for disease transmission are often a deterrent for captive breeding colonies when considering this approach35.
The last section of this article details surgical procedures and provides alternate approaches to expand ovarian studies or aid with the resolution of reproductive pathologies. Ovariectomies are most commonly performed in amphibians to obtain oocytes for embryological research. However, it can also provide a remedy for retained eggs when another options fail. Although this procedure is invasive, requiring full anesthesia and incisions to expose the egg masses, it does not require euthanasia. Furthermore, after partial ovariectomy, animals can make a full recovery and continue to be reproductively active post-surgery8,36.
The protocols described below outline the invasive and non-invasive methods of ovarian control and monitoring in Anurans and Caudates. The specific species chosen to illustrate techniques in Anurans include R. mucosa and X. laevis. Necturus maculosus, N. beyeri, N. alabamensis, and A. mexicanum comprise the species used to similarly describe techniques in Caudates.

<table><tbody><tr><td>GE logiq Book XP and 8C-RS probe 4e10 MHz GE Medical Systems</td><td>GE medical systems</td><td>GE logiq Book XP</td><td>Ultrasound</td></tr><tr><td>Aloka 500 7.5mHz linear or IBEX multi-frequency (10-6mHz) micro-convex</td><td>GE medical systems</td><td>8C-RS (10 MHz)</td><td>Ultrasound probe</td></tr><tr><td>BD disposable U-100 insulin syringe (28-29 G needle)</td><td>Mettler Electronics Corp CA</td><td>Sonigel</td><td>Ultrasound gel (water soluble, salt-free)</td></tr><tr><td>&lt;strong&gt;Hormone&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Gonadotropin releasing hormone</td><td>BACHEM</td><td>4012028</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; [Des-Gly10, D-Ala6, Pro-NHEt9]-GnRH acetate&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; GnRH</td></tr><tr><td>Lutenizing hormone releasing hormone</td><td>BACHEM, Sigma-Aldrich</td><td>4033013;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; L1898</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Pyr-His-Trp-Ser-Tyr-Gly-Leu-Arg-Pro-Gly-NH&lt;sub&gt;2&lt;/sub&gt; acetate salt;&lt;br/&gt; [D-Ala6}-LHRH acetate salt hydrate&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; LHRH</td></tr><tr><td>Human chorionic gonadotropin</td><td>BACHEM, Sigma-Aldrich</td><td>4030270, 4018894; C1063, CG5, CG10</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Chorionic gonadotropin-&amp;beta; (109-145)(109-119); Choriogonin, HCG (2,500IU, 5,000IU, 10,000IU)&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; hCG</td></tr><tr><td>Prostaglandin 2&amp;alpha;</td><td>Sigma-Aldrich</td><td>P40424;</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; (5Z,9&amp;alpha;,11&amp;alpha;,13E,15S)-9,11,15-Trihydroxyprosta-5,13-dienoic acid tris salt, PGF2&amp;alpha;&amp;minus;Tris;&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; PGF2&amp;alpha;</td></tr><tr><td>Follicle-stimulating hormone</td><td>Sigma-Aldrich</td><td>F4021, F8174</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; porcine, sheep&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; FSH</td></tr><tr><td>Progesterone</td><td>Sigma-Aldrich</td><td>46665;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; P7556</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Vetranal; P4 water soluble&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; P4</td></tr><tr><td>Pituitary extract</td><td>na</td><td></td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Check papers for amphibian species derivation&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; PE</td></tr><tr><td>Pregnant Mare Serum Gonadotropin</td><td>Prospec;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; Lee Biosolutions; Sigma-Aldrich</td><td>HOR-272;&amp;nbsp;&amp;nbsp;&amp;nbsp; 493-10; 9002-70-4</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Pregnant Mare Serum Gonadotropin&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; PMSG</td></tr><tr><td>Metaclopromide</td><td>Sigma-Aldrich</td><td>M0763</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; 4-Amino-5-chloro-N-[2-(diethylamino)ethyl]-2-methoxybenzamide, Methoxychloroprocainamide&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; MET</td></tr><tr><td>Lucrin</td><td>BACHEM; Sigma-Aldrich</td><td>4033014;&amp;nbsp;&amp;nbsp; L0399</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Leuprorelin acetate&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; Lucrin</td></tr><tr><td>Lutalyse</td><td>Pfizer</td><td></td><td>&lt;strong&gt;synonym:&lt;/strong&gt; PGF2&amp;alpha; - Dinoprost tromethamine&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; Lut</td></tr><tr><td>Pimozide</td><td>Sigma-Aldrich</td><td>P1793</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; 1-[1-[4,4-bis(4-Fluorophenyl)butyl]-4-piperidinyl]-1,3-dihydro-2H-benzimidazol-2-one&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; PZ</td></tr><tr><td>Amphiplex</td><td>see above</td><td></td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Gonadotropin releasing hormone + metoclopramide&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; GnRH + MET</td></tr><tr><td>Ovopel</td><td>Ovopel</td><td>na</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; GnRHa + dopamine receptor antagonist (administered 1 pellet/ kg)&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; Ovo</td></tr><tr><td>Ovaprim</td><td>Pentair aquatic eco-systems</td><td>Ova10</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; Salmon gonadotropin + domperidone&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; Ova</td></tr><tr><td>Domperidone</td><td>Sigma-Aldrich</td><td>D122</td><td>&lt;strong&gt;synonym:&lt;/strong&gt; 4-(5-Chloro-2-oxo-1-benzimidazolinyl)-1-[3-(2-oxobenzimidazolinyl)propyl]piperidine&lt;br/&gt; &lt;strong&gt;abbreviation:&lt;/strong&gt; DOM</td></tr></tbody></table>

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.

Watch this Scientific Journal Video about Reproductive Techniques for Ovarian Monitoring and Control in Amphibians at JoVE.com

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

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 ...

reproductive-techniques-for-ovarian-monitoring-control

Research

JoVE Journal

Biology

15.5K Views.  San Diego Zoo Institute for Conservation Research. 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.

Video: Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

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 ...

Basic Methods for the Study of Reproductive Ecology of Fish in Aquaria

Captive-rearing observations are valuable for revealing aspects of fish behavior and ecology when continuous field investigations are impossible. Here, a series of basic techniques are described to enable observations of the reproductive behavior of a wild-caught gobiid fish, as a model, kept in an aquarium. The method focuses on three steps: collection, transport, and observations of reproductive ecology of a substrate spawner. Essential aspects of live fish collection and transport are (1) preventing injury to the fish, and (2) careful acclimation to the aquarium. Preventing harm through injuries such as scratches or a sudden change of water pressure is imperative when collecting live fish, as any physical damage is likely to negatively affect the survival and later behavior of the fish. Careful acclimation to aquaria decreases the incidence death and mitigates the shock of transport. Observations during captive rearing include (1) the identification of individual fish and (2) monitoring spawned eggs without negative effects to the fish or eggs, thereby enabling detailed investigation of the study species' reproductive ecology. The subcutaneous injection of a visible implant elastomer (VIE) tag is a precise method for the subsequent identification of individual fish, and it can be used with a wide size range of fish, with minimal influence on their survival and behavior. If the study species is a substrate spawner that deposits adhesive eggs, an artificial nest site constructed from polyvinyl chloride (PVC) pipe with the addition of a removable waterproof sheet will facilitate counting and monitoring the eggs, lessening the investigator's influence on the nest-holding and egg-guarding behavior of the fish. Although this basic method entails techniques that are seldom mentioned in detail in research articles, they are fundamental for undertaking experiments that require the captive rearing of a wild fish.

A series of basic methods to enable the study of the reproductive ecology of fish kept in aquaria are described. These are useful protocols for collecting fish using SCUBA, transporting live fish, and observing the reproductive behavior of wild-caught fish kept in aquaria.

Captive-rearing observations are valuable for revealing aspects of fish behavior and ecology when continuous field investigations are impossible. Here, a ...

Behavior

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The efficient in vivo visualization of blood vessels and the reliable quantification of their complexity are key to understanding the biology and disease of the vascular network. Here, we provide a detailed method to visualize blood vessels with a commercially available fluorescent dye, human plasma acetylated low density lipoprotein DiI complex (DiI-AcLDL), and to quantify their complexity in Xenopus tropicalis. Blood vessels can be labeled by a simple injection of DiI-AcLDL into the beating heart of an embryo, and blood vessels in the entire embryo can be imaged in live or fixed embryos. Combined with gene perturbation by the targeted microinjection of nucleic acids and/or the bath application of pharmacological reagents, the roles of a gene or of a signaling pathway on vascular development can be investigated within one week without resorting to sophisticated genetically engineered animals. Because of the well-defined venous system of Xenopus and its stereotypic angiogenesis, the sprouting of pre-existing vessels, vessel complexity can be quantified efficiently after perturbation experiments. This relatively simple protocol should serve as an easily accessible tool in diverse fields of cardiovascular research.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The ...

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

Neuroscience

Fertilization of Xenopus oocytes using the Host Transfer Method

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to generate maternal-effect mutant animals. Additionally, many of the techniques to overexpress or inhibit gene function in organisms such as Xenopus and zebrafish fail to sufficiently target critical maternal signaling pathways, such as Wnt signaling. In Xenopus, manipulating gene function in cultured oocytes and subsequently fertilizing them can ameliorate these problems to some extent. Oocytes are manually defolliculated from donor ovary tissue, injected or treated in culture as desired, and then stimulated with progesterone to induce maturation. Next, the oocytes are introduced into the body cavity of an ovulating host female frog, whereupon they will be translocated through the host's oviduct and acquire modifications and jelly coats necessary for fertilization. The resulting embryos can then be raised to the desired stage and analyzed for the effects of any experimental perturbations. This host-transfer method has been highly effective in uncovering basic mechanisms of early development and allows a wide range of experimental possibilities not available in any other vertebrate model organism.

Fertilization of Xenopus oocytes using the Host Transfer Method

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Studying the contribution of maternally inherited molecules to vertebrate early development is often hampered by the time and expense necessary to ...

Analysis of Transforming Growth Factor &#223; Family Cleavage Products Secreted Into the Blastocoele of Xenopus laevis Embryos

The two arms of the Transforming Growth Factor &#223; (Tgf&#223;) superfamily, represented by Tgf&#223;/Nodal or Bone morphogenetic protein (Bmp) ligands, respectively, play essential roles in embryonic development and adult homeostasis. Members of the Tgf&#223; family are made as inactive precursors that dimerize and fold within the endoplasmic reticulum. The precursor is subsequently cleaved into ligand and prodomain fragments. Although only the dimeric ligand can engage Tgf&#223; receptors and activate downstream signaling, there is growing recognition that the prodomain moiety contributes to ligand activity. This article describes a protocol that can be used to identify cleavage products generated during activation of Tgf&#223; precursor proteins. RNA encoding Tgf&#223; precursors are first microinjected into X. laevis embryos. The following day, cleavage products are collected from the blastocoele of&#160;gastrula stage embryos and analyzed on Western blots. This protocol can be completed relatively quickly, does not require expensive reagents and provides a source of concentrated Tgf&#223; cleavage products under physiologic conditions.

Analysis of Transforming Growth Factor &#223; Family Cleavage Products Secreted Into the Blastocoele of Xenopus laevis Embryos

When Transforming Growth Factor &#223; family precursor proteins are ectopically expressed in Xenopus laevis embryos, they dimerize, get cleaved and are secreted into the blastocoele, which begins at the late blastula to early gastrula stage. We describe a method for aspirating cleavage products from the blastocoele cavity for immunoblot analysis.

The two arms of the Transforming Growth Factor &#223; (Tgf&#223;) superfamily, represented by Tgf&#223;/Nodal or Bone morphogenetic protein (Bmp) ligands, ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating gastric H+,K+-ATPase is more difficult to investigate. Many transport assays utilize radioisotopes to achieve a sufficient signal-to-noise ratio, however, the necessary security measures impose severe restrictions regarding human exposure or assay design. Furthermore, ion transport across cell membranes is critically influenced by the membrane potential, which is not straightforwardly controlled in cell culture or in proteoliposome preparations. Here, we make use of the outstanding sensitivity of atomic absorption spectrophotometry (AAS) towards trace amounts of chemical elements to measure Rb+ or Li+ transport by Na+,K+- or gastric H+,K+-ATPase in single cells. Using Xenopus oocytes as expression system, we determine the amount of Rb+ (Li+) transported into the cells by measuring samples of single-oocyte homogenates in an AAS device equipped with a transversely heated graphite atomizer (THGA) furnace, which is loaded from an autosampler. Since the background of unspecific Rb+ uptake into control oocytes or during application of ATPase-specific inhibitors is very small, it is possible to implement complex kinetic assay schemes involving a large number of experimental conditions simultaneously, or to compare the transport capacity and kinetics of site-specifically mutated transporters with high precision. Furthermore, since cation uptake is determined on single cells, the flux experiments can be carried out in combination with two-electrode voltage-clamping (TEVC) to achieve accurate control of the membrane potential and current. This allowed e.g. to quantitatively determine the 3Na+/2K+ transport stoichiometry of the Na+,K+-ATPase and enabled for the first time to investigate the voltage dependence of cation transport by the electroneutrally operating gastric H+,K+-ATPase. In principle, the assay is not limited to K+-transporting membrane proteins, but it may work equally well to address the activity of heavy or transition metal transporters, or uptake of chemical elements by endocytotic processes.

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

We describe a method to quantify the activity of K+-countertransporting P-type ATPases by heterologous expression of the enzymes in Xenopus oocytes and measuring Rb+ or Li+ uptake into individual cells by atomic absorption spectrophotometry. The method is a sensitive and safe alternative to radioisotope flux experiments facilitating complex kinetic studies.

Whereas cation transport by the electrogenic membrane transporter Na+,K+-ATPase can be measured by electrophysiology, the electroneutrally operating ...

Xenopus laevis as a Model to Identify Translation Impairment

Protein synthesis is a fundamental process to gene expression impacting diverse biological processes notably adaptation to environmental conditions. The initiation step, which involves the assembly of the ribosomal subunits at the mRNA initiation codon, involved initiation factor including eIF4G1. Defects in this rate limiting step of translation are linked to diverse disorders. To study the potential consequences of such deregulations, Xenopus laevis oocytes constitute an attractive model with high degrees of conservation of essential cellular and molecular mechanisms with human. In addition, during meiotic maturation, oocytes are transcriptionally repressed and all necessary proteins are translated from preexisting, maternally derived mRNAs. This inexpensive model enables exogenous mRNA to become perfectly integrated with an effective translation. Here is described a protocol for assessing translation with a factor of interest (here eIF4G1) using stored maternal mRNA that are the first to be polyadenylated and translated during oocyte maturation as a physiological readout. At first, mRNA synthetized by in vitro transcription of plasmids of interest (here eIF4G1) are injected in oocytes and kinetics of oocyte maturation by Germinal Vesicle Breakdown detection is determined. The studied maternal mRNA target is the serine/threonine-protein-kinase mos. Its polyadenylation and its subsequent translation are investigated together with the expression and phosphorylation of proteins of the mos signaling cascade involved in oocyte maturation. Variations of the current protocol to put forward translational defects are also proposed to emphasize its general applicability. In light of emerging evidence that aberrant protein synthesis may be involved in the pathogenesis of neurological disorders, such a model provides the opportunity to easily assess this impairment and identify new targets.

Xenopus laevis as a Model to Identify Translation Impairment

Protein synthesis control occurs mainly at the translation initiation step, deficiencies in which are linked to diverse disorders. To better understand their etiology, we described here a protocol using Xenopus laevis oocytes assessing the translation of mos transcript in the presence of a mutant of translation initiation factor eIF4G1.

Protein synthesis is a fundamental process to gene expression impacting diverse biological processes notably adaptation to environmental conditions. The ...

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

Summary

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Chapters in this video

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

Functional Cloning Using a Xenopus Oocyte Expression System

Basic Methods for the Study of Reproductive Ecology of Fish in Aquaria

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Fertilization of Xenopus oocytes using the Host Transfer Method

Analysis of Transforming Growth Factor ß Family Cleavage Products Secreted Into the Blastocoele of Xenopus laevis Embryos

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays

Xenopus laevis as a Model to Identify Translation Impairment