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>Hormone</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gonadotropin releasing hormone</td>
			<td>BACHEM</td>
			<td>4012028</td>
			<td>synonym: [Des-Gly10, D-Ala6, Pro-NHEt9]-GnRH acetate 
			abbreviation: GnRH</td>
		</tr>
		<tr>
			<td>Lutenizing hormone releasing hormone</td>
			<td>BACHEM, Sigma-Aldrich</td>
			<td>4033013;&#160;&#160;&#160;&#160; L1898</td>
			<td>synonym: Pyr-His-Trp-Ser-Tyr-Gly-Leu-Arg-Pro-Gly-NH2 acetate salt; 
			[D-Ala6}-LHRH acetate salt hydrate 
			abbreviation: LHRH</td>
		</tr>
		<tr>
			<td>Human chorionic gonadotropin</td>
			<td>BACHEM, Sigma-Aldrich</td>
			<td>4030270, 4018894; C1063, CG5, CG10</td>
			<td>synonym: Chorionic gonadotropin-&#946; (109-145)(109-119); Choriogonin, HCG (2,500IU, 5,000IU, 10,000IU) 
			abbreviation: hCG</td>
		</tr>
		<tr>
			<td>Prostaglandin 2&#945;</td>
			<td>Sigma-Aldrich</td>
			<td>P40424;</td>
			<td>synonym: (5Z,9&#945;,11&#945;,13E,15S)-9,11,15-Trihydroxyprosta-5,13-dienoic acid tris salt, PGF2&#945;&#8722;Tris; 
			abbreviation: PGF2&#945;</td>
		</tr>
		<tr>
			<td>Follicle-stimulating hormone</td>
			<td>Sigma-Aldrich</td>
			<td>F4021, F8174</td>
			<td>synonym: porcine, sheep 
			abbreviation: FSH</td>
		</tr>
		<tr>
			<td>Progesterone</td>
			<td>Sigma-Aldrich</td>
			<td>46665;&#160;&#160;&#160;&#160;&#160; P7556</td>
			<td>synonym: Vetranal; P4 water soluble 
			abbreviation: P4</td>
		</tr>
		<tr>
			<td>Pituitary extract</td>
			<td>na</td>
			<td></td>
			<td>synonym: Check papers for amphibian species derivation 
			abbreviation: PE</td>
		</tr>
		<tr>
			<td>Pregnant Mare Serum Gonadotropin</td>
			<td>Prospec;&#160;&#160;&#160;&#160;&#160;&#160; Lee Biosolutions; Sigma-Aldrich</td>
			<td>HOR-272;&#160;&#160;&#160; 493-10; 9002-70-4</td>
			<td>synonym: Pregnant Mare Serum Gonadotropin 
			abbreviation: PMSG</td>
		</tr>
		<tr>
			<td>Metaclopromide</td>
			<td>Sigma-Aldrich</td>
			<td>M0763</td>
			<td>synonym: 4-Amino-5-chloro-N-[2-(diethylamino)ethyl]-2-methoxybenzamide, Methoxychloroprocainamide 
			abbreviation: MET</td>
		</tr>
		<tr>
			<td>Lucrin</td>
			<td>BACHEM; Sigma-Aldrich</td>
			<td>4033014;&#160;&#160; L0399</td>
			<td>synonym: Leuprorelin acetate 
			abbreviation: Lucrin</td>
		</tr>
		<tr>
			<td>Lutalyse</td>
			<td>Pfizer</td>
			<td></td>
			<td>synonym: PGF2&#945; - Dinoprost tromethamine 
			abbreviation: Lut</td>
		</tr>
		<tr>
			<td>Pimozide</td>
			<td>Sigma-Aldrich</td>
			<td>P1793</td>
			<td>synonym: 1-[1-[4,4-bis(4-Fluorophenyl)butyl]-4-piperidinyl]-1,3-dihydro-2H-benzimidazol-2-one 
			abbreviation: PZ</td>
		</tr>
		<tr>
			<td>Amphiplex</td>
			<td>see above</td>
			<td></td>
			<td>synonym: Gonadotropin releasing hormone + metoclopramide 
			abbreviation: GnRH + MET</td>
		</tr>
		<tr>
			<td>Ovopel</td>
			<td>Ovopel</td>
			<td>na</td>
			<td>synonym: GnRHa + dopamine receptor antagonist (administered 1 pellet/ kg) 
			abbreviation: Ovo</td>
		</tr>
		<tr>
			<td>Ovaprim</td>
			<td>Pentair aquatic eco-systems</td>
			<td>Ova10</td>
			<td>synonym: Salmon gonadotropin + domperidone 
			abbreviation: Ova</td>
		</tr>
		<tr>
			<td>Domperidone</td>
			<td>Sigma-Aldrich</td>
			<td>D122</td>
			<td>synonym: 4-(5-Chloro-2-oxo-1-benzimidazolinyl)-1-[3-(2-oxobenzimidazolinyl)propyl]piperidine 
			abbreviation: 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

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

15.4K 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

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must be maintained to avoid deleterious consequences 1. External or internal factors that disrupt this balance can lead to protein aggregation, toxicity and cell death. In humans this is a major contributing factor to the symptoms associated with neurodegenerative disorders such as Huntington's, Parkinson's, and Alzheimer's diseases 10. It is therefore essential that the proteins involved in maintenance of proteostasis be identified in order to develop treatments for these debilitating diseases. This article describes techniques for monitoring in vivo protein folding at near-real time resolution using the model protein firefly luciferase fused to green fluorescent protein (FFL-GFP). FFL-GFP is a unique model chimeric protein as the FFL moiety is extremely sensitive to stress-induced misfolding and aggregation, which inactivates the enzyme 12. Luciferase activity is monitored using an enzymatic assay, and the GFP moiety provides a method of visualizing soluble or aggregated FFL using automated microscopy. These coupled methods incorporate two parallel and technically independent approaches to analyze both refolding and functional reactivation of an enzyme after stress. Activity recovery can be directly correlated with kinetics of disaggregation and re-solubilization to better understand how protein quality control factors such as protein chaperones collaborate to perform these functions. In addition, gene deletions or mutations can be used to test contributions of specific proteins or protein subunits to this process. In this article we examine the contributions of the protein disaggregase Hsp104 13, known to partner with the Hsp40/70/nucleotide exchange factor (NEF) refolding system 5, to protein refolding to validate this approach.

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

This article describes the use of a firefly luciferase-GFP fusion protein to investigate in vivo protein folding in Saccharomyces cerevisiae. Using this reagent, refolding of a model heat-denatured protein can be monitored simultaneously by fluorescence microscopy and an enzymatic assay to probe the roles of proteostasis network components in protein quality control.

Proteostasis, defined as the combined processes of protein folding/biogenesis, refolding/repair, and degradation, is a delicate cellular balance that must ...

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

Dried Blood Spots - Preparing and Processing for Use in Immunoassays and in Molecular Techniques

The idea of collecting blood on a paper card and subsequently using the dried blood spots (DBS) for diagnostic purposes originated a century ago. Since then, DBS testing for decades has remained predominantly focused on the diagnosis of infectious diseases especially in resource-limited settings or the systematic screening of newborns for inherited metabolic disorders and only recently have a variety of new and innovative DBS applications begun to emerge. For many years, pre-analytical variables were only inappropriately considered in the field of DBS testing and even today, with the exception of newborn screening, the entire pre-analytical phase, which comprises the preparation and processing of DBS for their final analysis has not been standardized. Given this background, a comprehensive step-by-step protocol, which covers al the essential phases, is proposed, i.e., collection of blood; preparation of blood spots; drying of blood spots; storage and transportation of DBS; elution of DBS, and finally analyses of DBS eluates. The effectiveness of this protocol was first evaluated with 1,762 coupled serum/DBS pairs for detecting markers of hepatitis B virus, hepatitis C virus, and human immunodeficiency virus infections on an automated analytical platform. In a second step, the protocol was utilized during a pilot study, which was conducted on active drug users in the German cities of Berlin and Essen.

The preparing and processing of dried blood spots (DBS) for their final analysis are still poorly standardized for most diagnostic applications. To overcome this shortcoming, a comprehensive step-by-step protocol is suggested and subsequently evaluated with regard to its effectiveness for detecting markers of viral infections.

The idea of collecting blood on a paper card and subsequently using the dried blood spots (DBS) for diagnostic purposes originated a century ago. Since ...

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

Micron-scale Phenotyping Techniques of Maize Vascular Bundles Based on X-ray Microcomputed Tomography

It is necessary to accurately quantify the anatomical structures of maize materials based on high-throughput image analysis techniques. Here, we provide a 'sample preparation protocol' for maize materials (i.e., stem, leaf, and root) suitable for ordinary microcomputed tomography (micro-CT) scanning. Based on high-resolution CT images of maize stem, leaf, and root, we describe two protocols for the phenotypic analysis of vascular bundles: (1) based on the CT image of maize stem and leaf, we developed a specific image analysis pipeline to automatically extract 31 and 33 phenotypic traits of vascular bundles; (2) based on the CT image series of maize root, we set up an image processing scheme for the three-dimensional (3-D) segmentation of metaxylem vessels, and extracted two-dimensional (2-D) and 3-D phenotypic traits, such as volume, surface area of metaxylem vessels, etc. Compared with traditional manual measurement of vascular bundles of maize materials, the proposed protocols significantly improve the efficiency and accuracy of micron-scale phenotypic quantification.

We provide a novel method to improve the X-ray absorption contrast of maize tissue suitable for ordinary microcomputed tomography scanning. Based on CT images, we introduce a set of image-processing workflows for different maize materials to effectively extract microscopic phenotypes of vascular bundles of maize.

It is necessary to accurately quantify the anatomical structures of maize materials based on high-throughput image analysis techniques. Here, we provide a ...

Single Cell Collection of Trophoblast Cells in Peri-implantation Stage Human Embryos

Human implantation, the apposition and adhesion to the uterine surface epithelia and subsequent invasion of the blastocyst into the maternal decidua, is a critical yet enigmatic biological event that has been historically difficult to study due to technical and ethical limitations. Implantation is initiated by the development of the trophectoderm to early trophoblast and subsequent differentiation into distinct trophoblast sublineages. Aberrant early trophoblast differentiation may lead to implantation failure, placental pathologies, fetal abnormalities, and miscarriage. Recently, methods have been developed to allow human embryos to grow until day 13 post-fertilization in vitro in the absence of maternal tissues, a time-period that encompasses the implantation period in humans. This has given researchers the opportunity to investigate human implantation and recapitulate the dynamics of trophoblast differentiation during this critical period without confounding maternal influences and avoiding inherent obstacles to study early embryo differentiation events in vivo. To characterize different trophoblast sublineages during implantation, we have adopted existing two-dimensional (2D) extended culture methods and developed a procedure to enzymatically digest and isolate different types of trophoblast cells for downstream assays. Embryos cultured in 2D conditions have a relatively flattened morphology and may be suboptimal in modeling in vivo three-dimensional (3D) embryonic architectures. However, trophoblast differentiation seems to be less affected as demonstrated by anticipated morphology and gene expression changes over the course of extended culture. Different trophoblast sublineages, including cytotrophoblast, syncytiotrophoblast and migratory trophoblast can be separated by size, location, and temporal emergence, and used for further characterization or experimentation. Investigation of these early trophoblast cells may be instrumental in understanding human implantation, treating common placental pathologies, and mitigating the incidence of pregnancy loss.

Here, we describe a method for warming vitrified human blastocysts, culturing them through the implantation period in vitro, digesting them into single cells and collecting early trophoblast cells for further investigation.

Human implantation, the apposition and adhesion to the uterine surface epithelia and subsequent invasion of the blastocyst into the maternal decidua, is a ...

Reproductive Techniques for Ovarian Monitoring and Control in Amphibians

Summary

Explore More Videos

Chapters in this video

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Techniques for Imaging Ca²⁺ Signaling in Human Sperm

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Coupled Assays for Monitoring Protein Refolding in Saccharomyces cerevisiae

In Vivo Microinjection and Electroporation of Mouse Testis

Dried Blood Spots - Preparing and Processing for Use in Immunoassays and in Molecular Techniques

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

Micron-scale Phenotyping Techniques of Maize Vascular Bundles Based on X-ray Microcomputed Tomography

Single Cell Collection of Trophoblast Cells in Peri-implantation Stage Human Embryos