I thank the authors contributing to the initial work which is the basis of this publication. Thanks to Aitor Aguirre, Genevieve E. Ryan and Erica L. Schoeller for help preparing the manuscript. Thanks to Jessica Sora Lee and Austin Chin for technical assistance with the manuscript. H.M.H. was supported by Eunice Kennedy Shriver National Institute of Child Health &#38; Human Development of the National Institutes of Health under Award Number R00HD084759.

All methods described here have been approved by the Institutional Animal Care and Use Committee of Michigan State University and conducted in accordance with the Guide for the Care and Use of Laboratory Animals.
1. Determine Pubertal Onset
<ol>
	<li>Follow institutional guidelines for clothing, at a minimum, it is necessary to wear a clean lab coat and clean gloves. Always handle mice wearing clean gloves.</li>
	<li>Prepare the work area by placing a pad on the table.
	<ol>
		<li>Place a clean mouse cage top with the grid facing upwards on the pad.</li>
		<li>Set up a scale in the work area. Place a clean 500 &#8211; 1000 mL beaker on the scale and tare.</li>
		<li>Place a sheet next to the work area to record body weight and preputial separation, vaginal opening or first estrus.</li>
	</ol>
	</li>
	<li>Identify the mice for the study. Perform daily inspections for pubertal onset until puberty is reached. Perform the inspection at the same time of day throughout the study. Pubertal onset can occur as early as postnatal day 11-1219, however, in most experimental setups, pubertal onset monitoring starts at ~22 days.</li>
	<li>Place the mouse cage in the work area (Step 1.2). Open the cage and place the lid, water bottle and food holder on the table.</li>
	<li>Determining preputial separation in male mice.
	<ol>
		<li>While holding the mouse by the tail, place it on the clean mouse cage top from Step 1.2.1. While gently holding onto the tail, let the mouse explore the grid of the cage top for ~5 s.</li>
		<li>Gently pull the tail backward; this usually results in the animal moving forward while holding on to the grid with its forelegs. Approach the other hand to the skin close by the neck and ears.</li>
		<li>Grasp the loose skin between the shoulders and neck with the thumb and forefinger. It is important to get a good hold of the skin, as it will prevent the mouse from turning its head around to bite.</li>
		<li>Grab the skin on the back with the remaining fingers and hold it by pressing against the hand. Hold firmly without obstructing breathing, blood flow, or damaging bones, muscles or skin. Turn the hand to expose the belly of the mouse, while supporting the back of the mouse within the palm of the hand.</li>
		<li>With the free hand, gently push back the skin around the penis without forcing. At preputial separation, the preputial skin slides backward exposing the glans penis (Figure 2A, arrow showing preputial separation). Record the presence or absence of preputial separation.</li>
		<li>Weigh the mouse by gently placing it in the 500-1000 mL beaker from Step 1.2.2 and record the weight.</li>
		<li>Reintroduce the mouse in the housing cage by holding the beaker 0-2 cm over the cage floor and slowly tip over the beaker allowing the mouse to exit on its own. Clean the beaker with water and mild detergent and dry it off before returning it to the scale and taring the scale.</li>
	</ol>
	</li>
	<li>Determining vaginal opening in female mice.
	<ol>
		<li>Place a cotton ball, beaker and a bottle of sterile water in the work area (Step 1.2). Pour the water into the beaker. Humidify the cotton ball with the sterile water and place it next to the cage top from Step 1.2.1.</li>
		<li>Place the mouse cage in the work area (Step 1.2) and transfer the mouse from its home cage to the cage top (Step 1.2.1) by holding it at the base of the tail. Gently pull back the tail to encourage a forward movement of the mouse.</li>
		<li>Lift the tail while supporting the hips with the free fingers. Allow the hindlimbs to maintain contact with the top of the cage. If the mouse moves too much, hold it in the hand as described in Steps 1.5.1-1.5.4.</li>
		<li>Gently clean the vulva with the water humidified cotton ball from Step 1.6.1. Use a clean humidified cotton ball for each mouse. To determine vaginal opening, examine the vulva and determine whether the vagina is completely open (Figure 2A, female vaginal opening).</li>
		<li>Record if vaginal opening has occurred. Weigh the mouse and return it to the home cage as described in Steps 1.5.6-1.5.7.</li>
	</ol>
	</li>
	<li>First estrus an indicator of first ovulation.
	<ol>
		<li>Place a beaker with sterile water, a labeled glass slide and a 200 &#181;L pipette with a clean tip in the work area (Step 1.2).</li>
		<li>Hold the female as described in Step 1.6.2-1.6.3. Place the tip of the pipette containing ~50 &#181;L of sterile water at the vaginal opening.</li>
		<li>Gently flush the cells from the vaginal wall by introducing and reabsorbing ~50 &#181;L of water 2-4 times. Smear the content from the pipette tip onto a labeled glass microscope slide.</li>
		<li>Carefully observe the cellular morphology on a brightfield microscope using a 10X or 20X objective or let the smear airdry for ~1 h and then counterstain with a 0.1% methylene blue solution (dissolved in ddH2O) (Figure 2B). To counterstain, dip the dried slide into the 0.1% methylene blue solution for ~ 30 s. Let the slide air dry for 1 h.</li>
		<li>Observe the slides from Step 1.7.4 on a bright field microscope at 10X or 20X. Establish the stage of the estrous cycle (Figure 2B)20 by observing the cell morphology which allows to distinguish the four distinct stages of the estrous cycle (Figure 2B). 
		Note: Metestrus is characterized by a mixture of cornified squamous epithelial cells and leukocytes, diestrus by leukocytes, proestrus by a mixture of leukocytes and nucleated epithelial cells, and estrus by cornified epithelial cells (Figure 2B)20.</li>
		<li>Record the body weight and return the mouse to its home cage as described in Steps 1.5.6-1.5.7. Vaginal smears are terminated on the day when the mouse has its first estrus (Figure 2B).</li>
	</ol>
	</li>
</ol>
2. Desirable Breeding Room Conditions
<ol>
	<li>Ensure that the breeding room has a temperature of ~20-24 &#176;C, with 55 &#177; 10% humidity, and homogenous room light of an intensity of ~300-400 lux approximately 1 m above the floor21. This light intensity allows the desirable mid-cage light intensity of 25-80 lux. To test light intensity, use a light meter.</li>
	<li>Use an automated system to control the lighting in the breeding room to ensure appropriate timing of lighting throughout the experiment. Optimal day-length conditions to assess fertility in rodents range from a 12 h day (12 h light and 12 h darkness, LD12:12), to &#8220;summer-like&#8221; conditions with 14 h of light and 10 h darkness (LD14:10)21.</li>
	<li>Prepare breeding cages.
	<ol>
		<li>Prepare a clean mouse cage with lid, water, food, bedding and nesting per breeding pair.</li>
		<li>Fill out the cage cards with all the required information, including sex, date of birth, mouse strain, and the date when the mating is set up.</li>
	</ol>
	</li>
</ol>
3. Fertility Study
<ol>
	<li>Identify the animals to be used for the fertility study. Ensure that the animals are sexually mature (Step 1). A typically used age range to set up a fertility study in mice is 10-16 weeks of age, an age where sexual maturation has been completed in most experimental conditions.</li>
	<li>On a clean table, place the breeding cage prepared in Step 2.3.</li>
	<li>Catch and hold the mouse.
	<ol>
		<li>Slowly introduce a gloved hand into the mouse cage. Leave the hand in the cage for a short period of time (~5-30 s), allowing the mice to adapt to the smell of the glove and hand. Avoid hectic and jerky movements.</li>
		<li>Lower the cupped hand close to the bottom of the cage and slowly approach it to the mice. They will run away. When they run over the hand, trap the tail of one mouse between the thumb and index finger.</li>
		<li>Lift the mouse by the tail for 2-3 s. If the mouse needs to be held longer, hold onto the tail and place the mouse in the cupped hand maintaining the hold on the tail.</li>
	</ol>
	</li>
	<li>Transfer 1 male and thereafter 1 female into the breeding cage by holding them by the tail (Step 3.3). Ensure that the females are in the same estrous stage when setting up the fertility assay. Determine the estrous stage of the females by performing a vaginal smear as described in Step 1.7.</li>
	<li>Close the cage, and ensure that the mice have food, water and nesting material. Attach the cage card holder with the cage card to the cage.</li>
	<li>Place the breeding cage on the housing rack. Leave the breeding cage as undisturbed as possible for the full duration of the fertility study. Conduct the fertility study for 30 days. For each breeding pair, keep detailed records and note 1) the date the mating is set up, 2) the date litters are born and 3) the number of pups born, including dead and live pups. Perform litter checks daily throughout the study.</li>
	<li>Prolong the fertility study for an additional 30-60 days if no or a weak impact on fertility is established in Step 3.6.</li>
</ol>

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The author has nothing to disclose.

The overall wellbeing of the mice is critical for a successful fertility assay21. When performing a fertility assay, it is important to not physically check on the mice every day as this can cause stress. Further avoid frequent cage changes, as these are also stressful. Ideally cage changes will be done no more than 1-2 times per week. Light exposure during the dark phase negatively impacts breeding in nocturnal rodents. Do not turn on lights in the breeding room during the dark hours. If entry to...

The transition through puberty is required to attain sexual maturity and reproductive competence. The pubertal transition and the maintenance of fertility in adulthood is regulated by the reproductive axis, also termed the hypothalamic-pituitary-gonadal axis (Figure 1). The timing of pubertal onset and maintenance of fertility is tightly regulated by internal as well as environmental factors to increase the chances of survival of offspring and parents1,2. This protocol provides a non-invasive approach to determine pubertal onset in mice and rats to confirm sexual maturity prior to setting up a fertility study to assess reproductive competence.
A fertility study is performed in sexually mature animals and can be initiated after the animals have gone through puberty. Prior to pubertal onset, the reproductive axis is quiescent, and the key driver of sexual maturation, gonadotropin-releasing hormone (GnRH), is released onto the pituitary in insufficient amounts to initiate puberty (Figure 1). Pubertal onset is a complex process that results in increased GnRH release at the median eminence. GnRH promotes luteinizing hormone (LH) and follicle stimulating hormone (FSH) secretion from the pituitary, two hormones essential for gonadal maturation and reproductive function (Figure 1)3,4,5.
Insults to the reproductive axis result in reduced fertility and can also advance or delay pubertal onset. Conditions known to influence the timing of pubertal onset and reproductive competence include the exposure to endocrine disrupting chemicals6,7, increased/decreased body weight1,8, changes in day length2,9 and genetic mutations10,11,12,13,14,15.
The onset of sexual maturity is a critical step that needs to be completed prior to setting up a fertility assay. The advantages of determining pubertal onset through preputial separation, vaginal opening and first estrus, are the non-invasive characteristics of these procedures, as they do not require blood collection or sacrifice of the animal16,17.
After pubertal onset is determined, correctly setting up a fertility study will provide important information about the integrity of the reproductive axis, and usually has the second advantage of generating experimental animals for further studies (refinement)18. The fertility study setup described in this protocol can detect both minor and major deficits in reproductive competence in males and females. Key parameters evaluated include 1) time to the first litter, 2) number of litters generated in a given time frame and 3) litter size. Finally, recommendations for the type of follow up studies which can be conducted to identify the cause of fertility impairment are included.
The described protocol refers to mice and the representative data reflect work done in transgenic mice. However, all the included protocols are equally valid in rats.

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determination of reproductive competence by confirming pubertal onset and performing a fertility assay in mice and rats

Assessment of reproductive competence is critical for understanding the impact of a treatment or genetic manipulation on the reproductive axis, also termed the hypothalamic-pituitary-gonadal axis. The reproductive axis is a key integrator of environmental and internal input adapting fertility to favorable conditions for reproduction. Prior to embarking upon a fertility study in mice and rats, sexual maturity is evaluated to exclude the possibility that the observed reproductive phenotypes are caused by delayed or absent pubertal onset. This protocol describes a non-invasive approach to assess pubertal onset in males through the determination of preputial separation, and in females through vaginal opening and first estrus. After the confirmation of the completion of puberty and the achievement of sexual maturity, a fertility study can be initiated. The procedure describes the optimal breeding conditions for mice and rats, how to set up a fertility study, and what parameters to evaluate and determine if the treatment or gene deletion has an impact on fertility.

The presented results are from two different transgenic mouse models where the transcription factor Ventral anterior homeobox 1 (Vax1) has been deleted in the whole body on one allele, here referred to as heterozygote mice (HET)13, or Vax1 has been conditionally deleted within GnRH neurons22, here termed conditional KO (cKO). Prior to setting up the fertility study, it is important to confirm pubertal onset in all the mice....

Watch this Scientific Journal Video about Determination of Reproductive Competence by Confirming Pubertal Onset and Performing a Fertility Assay in Mice and Rats at JoVE.com

Determination of Reproductive Competence by Confirming Pubertal Onset and Performing a Fertility Assay in Mice and Rats

Many treatments and genetic mutations impact the timing of sexual maturity and fertility. This protocol describes a non-invasive method to evaluate pubertal onset in mice and rats prior to setting up a fertility study in sexually mature animals.

Assessment of reproductive competence is critical for understanding the impact of a treatment or genetic manipulation on the reproductive axis, also ...

determination-reproductive-competence-confirming-pubertal-onset

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

15.3K Views.  Michigan State University. Many treatments and genetic mutations impact the timing of sexual maturity and fertility. This protocol describes a non-invasive method to evaluate pubertal onset in mice and rats prior to setting up a fertility study in sexually mature animals.

Video: Determination of Reproductive Competence by Confirming Pubertal Onset and Performing a Fertility Assay in Mice and Rats

Collection of Post-mating Semen from the Female Reproductive Tract and Measurement of Semen Liquefaction in Mice

In mice, ejaculated semen is deposited in the uterus. After ejaculation, the semen changes consistency from gel-like to watery, a process called liquefaction. In this study, we show how to collect the post-ejaculated semen from the female reproductive tract in a mouse model. First, adult female mice in the estrus stage were housed in a male&#39;s cage overnight. The next morning, copulation was confirmed by the presence of copulatory plug at the vaginal opening. Female mice with copulatory plugs were euthanized, and each reproductive tract was collected as a whole (vagina, uterus, oviducts, ovaries), ensuring a closed system to contain the semen. The reproductive tract was placed in a 1.5 mL microcentrifuge tube, and the vagina was cut off to release the semen into the tube. To ensure maximum semen volume for analysis, toothless forceps were used to squeeze the uterine horns from ovarian end to vaginal end expelling remaining semen. The whole reproductive tract was then discarded. The semen-containing tube was briefly spun down. A 25 &#956;L capillary pipette was placed into the tube at a 180&#176; angle (parallel to the tube wall). The amount of time used to fill the capillary tube to the 25 &#956;L line was recorded. Semen from a proven male breeder usually takes approximately 60-180 s to fill a 25 &#956;L capillary tube. This semen collection technique can also be used in other downstream applications such as sperm imaging and motility analysis.

Semen liquefaction is required for sperm to be liberated from the seminal gel. This study provides the procedures for collecting semen from the female reproductive tract post-coitus, and measuring semen liquefaction time.

In mice, ejaculated semen is deposited in the uterus. After ejaculation, the semen changes consistency from gel-like to watery, a process called ...

Developmental Biology

Assessment of Sexual Behavior of Male Mice

Sexual behavior is highly species-specific. Although rodents have slightly different sexual behaviors, mice and rats have a similar sexual behavioral pattern. The purpose of this article is to describe the hormone-induced estrus ovariectomized female model and the experimental procedure for the assessment of sexual behavior of male mice. The most important sexual behavioral elements are demonstrated in the video and illustrations. The critical steps, advantages, and limitations of the sexual behavior test are explained as well. Finally, the behavior parameters are presented, and mounting, intromission, and ejaculation processes in mating are distinguished. Behavioral parameters are assessed in terms of the occurred duration and counts during the test period.

This article describes how to perform sexual behavior tests in male mice.

Sexual behavior is highly species-specific. Although rodents have slightly different sexual behaviors, mice and rats have a similar sexual behavioral ...

Behavior

A Hyperandrogenic Mouse Model to Study Polycystic Ovary Syndrome

Hyperandrogenemia plays a critical role in reproductive and metabolic function in females and is the hallmark of polycystic ovary syndrome. Developing a lean PCOS-like mouse model that mimics women with PCOS is clinically meaningful. In this protocol, we describe such a model. By inserting a 4 mm length of DHT (dihydrotestosterone) crystal powder pellet (total length of pellet is 8 mm), and replacing it monthly, we are able to produce a PCOS-like mouse model with serum DHT levels 2 fold higher than mice not implanted with DHT (no-DHT). We observed reproductive and metabolic dysfunction without changing body weight and body composition. While exhibiting a high degree of infertility, a small subset of these PCOS-like female mice can get pregnant and their offspring show delayed puberty and increased testosterone as adults. This PCOS-like lean mouse model is a useful tool to study the pathophysiology of PCOS and the offspring from these PCOS-like dams.

We describe the development of a lean PCOS-like mouse model with&#160;dihydrotestosterone pellet to study the pathophysiology of PCOS and the offspring from these PCOS-like dams.

Hyperandrogenemia plays a critical role in reproductive and metabolic function in females and is the hallmark of polycystic ovary syndrome. Developing a ...

Analysis of Congenital Heart Defects in Mouse Embryos Using Qualitative and Quantitative Histological Methods

Congenital heart defects (CHD) are the most common type of birth defect in humans, affecting up to 1% of all live births. However, the underlying causes for CHD are still poorly understood. The developing mouse constitutes a valuable model for the study of CHD, because cardiac developmental programs between mice and humans are highly conserved. The protocol describes in detail how to produce mouse embryos of the desired gestational stage, methods to isolate and preserve the heart for downstream processing, quantitative methods to identify common types of CHD by histology (e.g., ventricular septal defects, atrial septal defects, patent ductus arteriosus), and quantitative histomorphometry methods to measure common muscular compaction phenotypes. These methods articulate all the steps involved in sample preparation, collection, and analysis, allowing scientists to correctly and reproducibly measure CHD.

In this protocol, we describe procedures to qualitatively and quantitatively analyze developmental phenotypes in mice associated with congenital heart defects.

Congenital heart defects (CHD) are the most common type of birth defect in humans, affecting up to 1% of all live births. However, the underlying causes ...

Murine Round Spermatid Injection Procedure to Study Embryonic Development

Mouse Round Spermatid Injection

Round spermatids, characterized by their haploid genetic content, represent the precursor cells to mature spermatozoa. Through the innovative technique of round spermatid injection (ROSI), oocytes can be successfully fertilized and developed into viable fetuses. In a groundbreaking milestone achieved in 1995, the first mouse fetus was born through ROSI technology. ROSI has since emerged as a pivotal tool for unraveling the intricate mechanisms governing embryonic development and holds significant potential in various applications, including the acceleration of mouse generation and the production of genetically modified mice. In 1996, a milestone was reached when the first human fetus was born through ROSI technology. However, the clinical applications of this method have shown a fluctuating pattern of success and failure. To date, ROSI technology has not found widespread application in clinical practice, primarily due to its low birth efficiency and insufficient validation of fetal safety. This article provides a comprehensive account of the precise methods of performing ROSI in mice, aiming to shed new light on basic research and its potential clinical applications.

Here, we introduce a method for performing round spermatid injection (ROSI) in mice, a technique with promising clinical applications and utility for investigating the mechanisms underlying embryonic development.

Round spermatids, characterized by their haploid genetic content, represent the precursor cells to mature spermatozoa. Through the innovative technique of ...

Performing Vaginal Lavage, Crystal Violet Staining, and Vaginal Cytological Evaluation for Mouse Estrous Cycle Staging Identification

A rapid means of assessing reproductive status in rodents is useful not only in the study of reproductive dysfunction but is also required for the production of new mouse models of disease and investigations into the hormonal regulation of tissue degeneration (or regeneration) following pathological challenge. The murine reproductive (or estrous) cycle is divided into 4 stages: proestrus, estrus, metestrus, and diestrus. Defined fluctuations in circulating levels of the ovarian steroids 17-&beta;-estradiol and progesterone, the gonadotropins luteinizing and follicle stimulating hormones, and the luteotropic hormone prolactin signal transition through these reproductive stages. Changes in cell typology within the murine vaginal canal reflect these underlying endocrine events. Daily assessment of the relative ratio of nucleated epithelial cells, cornified squamous epithelial cells, and leukocytes present in vaginal smears can be used to identify murine estrous stages. The degree of invasiveness, however, employed in collecting these samples can alter reproductive status and elicit an inflammatory response that can confound cytological assessment of smears. Here, we describe a simple, non-invasive protocol that can be used to determine the stage of the estrous cycle of a female mouse without altering her reproductive cycle. We detail how to differentiate between the four stages of the estrous cycle by collection and analysis of predominant cell typology in vaginal smears and we show how these changes can be interpreted with respect to endocrine status.

Here, we describe how to identify the stage of the murine reproductive (proestrus, estrus, metestrus, or diestrus) by simple, non-invasive collection and cytological assessment of vaginal smear samples. We further describe how vaginal cytology reflects circulating hormonal levels underlying transition through the murine reproductive cycle.

A rapid means of assessing reproductive status in rodents is useful not only in the study of reproductive dysfunction but is also required for the ...

Ex Vivo Method for Assessing the Mouse Reproductive Tract Spontaneous Motility and a MATLAB-based Uterus Motion Tracking Algorithm for Data Analysis

Dysmenorrhea, or painful cramping, is the most common symptom associated with menses in females and its severity can hinder women&#39;s everyday lives. Here, we present an easy and inexpensive method that would be instrumental for testing new drugs decreasing uterine contractility. This method utilizes the unique ability of the entire mouse reproductive tract to exhibit spontaneous motility when maintained ex vivo in a Petri dish containing oxygenated Krebs buffer. This spontaneous motility resembles the wave-like myometrial activity of the human uterus, referred to as endometrial waves. To demonstrate the effectiveness of the method, we employed a well-known uterine relaxant drug, epinephrine. We demonstrate that the spontaneous motility of the entire mouse reproductive tract can be quickly and reversibly inhibited by 1 &#181;M epinephrine in this Petri dish model. Documenting the changes of uterine motility can be easily done using an ordinary smart phone or a sophisticated digital camera. We developed a MATLAB-based algorithm allowing motion tracking to quantify spontaneous uterine motility changes by measuring the rate of uterine horn movements. A major advantage of this ex vivo approach is that the reproductive tract remains intact throughout the entire experiment, preserving all intrinsic intrauterine cellular interactions. The major limitation of this approach is that up to 10-20% of uteri may exhibit no spontaneous motility. Thus far, this is the first quantitative ex vivo method for assessing spontaneous uterine motility in a Petri dish model.

Uterine contractions are important for the well-being of females. However, pathologically increased contractility may result in dysmenorrhea, especially in younger females. Here, we describe a simple ex vivo preparation allowing quick assessment of the efficacy of smooth muscle relaxants that may be used for treating dysmenorrhea.

Dysmenorrhea, or painful cramping, is the most common symptom associated with menses in females and its severity can hinder women&#39;s everyday lives. ...

Rodent Estrous Cycle Monitoring Utilizing Vaginal Lavage: No Such Thing As a Normal Cycle

The current methodology establishes a reproducible, standardized, and cost-effective approach to monitoring the estrous cycle of female Sprague Dawley (SD) adolescent rats. This study demonstrates the complexity of hormonal cycles and the broad spectrum of understanding required to construct a reliable and valid monitoring technique. Through an in-depth examination of principal experimental design and procedural elements,&#160;this description of the cycle and its fundamental principles provides a framework for further understanding and deconstructs misconceptions for future replication.
Along with an outline of the sample collection process employing vaginal lavage, the procedure describes the mechanism of data categorization into the four-stage model of proestrus, estrus, metestrus, and diestrus. These stages are characterized by a new proposed approach, utilizing the 4 categorizing determinants of vaginal fluid condition, cell type(s) present, cell arrangement, and cell quantity at the time of collection. Variations of each stage, favorable and unfavorable samples, the distinction between cyclicity and acyclicity, and graphic depictions of the collected categorizing components are presented alongside effective interpretive and organizational practices of the data. Overall, these tools allow for the publication of quantifiable data ranges for the first time, leading to the standardization of categorization factors upon replication.

This study details the crucial factors to consider in experimental designs involving female rats. In a larger sense, these data serve to decrease stigma and assist in the development of more inclusive diagnostic and intervention tools.

The current methodology establishes a reproducible, standardized, and cost-effective approach to monitoring the estrous cycle of female Sprague Dawley ...

Cervical Manipulation Induced Pseudopregnancy in Mice for Assisted Reproduction

Inducing Pseudopregnancy in Female Mice Without the Need for Vasectomized Males Prior to Non-Surgical Embryo Transfer or Artificial Insemination

For successfully maintaining pregnancy with embryo transfer or artificial insemination,&#160;female recipient mice must be induced into a pseudopregnant state. Female mice are traditionally paired overnight with vasectomized males, and the following morning, the presence of a copulation plug is assessed. To increase the efficiency of producing pseudopregnant females, a cervical manipulation technique has been standardized to be used in combination with non-surgical embryo transfer or artificial insemination techniques in mice. The blunt end of a small plastic rod is inserted vaginally to contact the cervix and is vibrated for 30 s by contact with a trimmer. The procedure is quick and does not require anesthesia or analgesia. This technique increases the reliability and predictability of producing pseudopregnant females and entirely eliminates the requirement for vasectomized males. For CD1 mice, the efficiency of pseudopregnancy induction using cervical manipulation was 83% for females in estrus (N = 76) but only 38% of females in estrus were plugged by vasectomized males (N = 24). Artificial insemination in CD1 mice was performed by estrus synchronization with hormones, cervical manipulation, and the uterine transfer of sperm. Artificial insemination recipients receiving cervical manipulation (N = 76) had a pregnancy rate of 72% and an average litter size of 8.3 pups. This method can also be used to produce pseudopregnant females for non-surgical embryo transfer. Therefore, inducing pseudopregnancy by cervical manipulation is a convenient and efficient alternative to mating with a vasectomized male when performing non-surgical assisted reproduction techniques. Using cervical manipulation provides 3Rs (replacement, reduction, and refinement) benefits for assisted reproduction techniques by reducing the number of animals required and eliminating the necessity for surgically altered males.

Author Spotlight: Inducing Pseudopregnancy in Female Mice Without the Need for Vasectomized Males Prior to Non-Surgical Embryo Transfer or Artificial Insemination  

The cervical manipulation method presented can induce pseudopregnancy in mice without the need for breeding females with vasectomized males. The induction of pseudopregnancy is required for the success of non-surgical embryo transfer and non-surgical artificial insemination, both of which are also presented.

For successfully maintaining pregnancy with embryo transfer or artificial insemination,&#160;female recipient mice must be induced into a pseudopregnant ...

Establishment of Rat Models Mimicking Gender-affirming Hormone Therapies

Transgender (TG) people are individuals whose gender identity and sex assigned at birth do not match. They often undergo gender-affirming hormone therapy (GAHT), a medical intervention that allows the acquisition of secondary sex characteristics more aligned with their individual gender identity, providing consistent results in the improvement of numerous socio-psychological variables. However, GAHT targets different body systems, and some side effects are recorded, although not yet fully identified and characterized. Therefore, TG people undergoing GAHT may be considered as a susceptible sub-group of population and specific attention should be paid in the frame of risk assessment, e.g., through the use of targeted animal models. The present work describes the procedures set to implement two rat models mimicking GAHT: the demasculinizing-feminizing model (dMF) mimicking the GAHT for TG women and the defeminizing-masculinizing model (dFM) mimicking the GAHT for TG men. The models have&#160;been implemented through the administration of the same hormones used for human GAHT, namely, &#946;-estradiol plus cyproterone acetate for dMF and testosterone for dFM, by the same routes of exposure for a 2 week period. Rats are checked daily during the treatment to evaluate health status and potentially aggressive behaviors. At sacrifice, blood and target tissues have been sampled and stored for biochemical, molecular, and histopathological analysis. Sex-specific parameters, namely, sperm count and clitoral dimensions, have also been evaluated. In addition, CYP450 isoforms, exclusively and/or preferentially expressed in male and female rat liver, are identified and characterized as novel biomarkers to verify the success of GAHT and to set the model. Thyroid involvement has also been explored as a key target in the endocrine system.

The protocol describes the development of two rodent models mimicking gender-affirming hormone therapies through subcutaneous administration of testosterone or estradiol plus cyproterone acetate (used in human therapies for transgender people): setting of the doses, identification of relevant biomarkers, and evaluation of the effects.

Transgender (TG) people are individuals whose gender identity and sex assigned at birth do not match. They often undergo gender-affirming hormone therapy ...