Name: Reproductive Techniques for Ovarian Monitoring and Control in Amphibians (Video) | JoVE Journal
Uploaded: 2019-05-12T12:30:00.000+00:00
Duration: 4 min 37 s
Description: Learn recoverable ovariectomy techniques for amphibian ovarian monitoring using female anurans. This method enables oocyte retrieval without euthanasia while preserving reproductive viability post-surgery.

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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

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 ...

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JoVE Journal

Biology

Procedure for fertilizing Xenopus oocytes by the host transfer method.

Fertilization of Xenopus oocytes using the Host Transfer Method

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.

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

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.

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

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.

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

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.

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

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.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

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.

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

When Transforming Growth Factor ß 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.

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

Here, we describe methods for tattooing adult Xenopus laevis (African clawed frog) with a rotary tattoo machine. Proper tattooing results in dark, easily legible numerals that last for several months and make animals easily distinguishable for research and recordkeeping purposes.

Methods for Tattooing Xenopus laevis with a Rotary Tattoo Machine

We present a protocol for the surgical removal of one testis from adult male X. laevis, which allows the use of the second testis at a later time. This protocol aims to reduce the number of male frogs required for in vitro fertilization.

Unilateral Gonadectomy in Adult Male Xenopus laevis

Learn recoverable ovariectomy techniques for amphibian ovarian monitoring using female anurans. This method enables oocyte retrieval without euthanasia while preserving reproductive viability post-surgery.

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

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