We extend our gratitude to Wenjie Zhao for her invaluable assistance in sorting round spermatids through flow cytometry and to Fang Wang for her expertise in mouse embryo transfer. This work received partial support from the Natural Science Foundation of Jilin Province (No. YDZJ202301ZYTS461). We thank Bullet Edits Limited for the linguistic editing and proofreading of the manuscript.

B6D2F1 (C57BL/6 x DBA/2), C57BL/6, and ICR mice used in this experiment were purchased from Beijing Vital River Laboratory Animal Technologies Co. Ltd. (Beijing, China). All animal treatments adhered to the experimental procedures and standards approved by the Experimental Animal Ethics Committee of the First Hospital of Jilin University (approval number: 20200435).
1. Preparation of relevant reagents
<ol>
	<li>Acquire some reagents commercially and self-prepare the remaining reagents.
	<ol>
		<li>Obtain M2, a buffer system for in vitro processing of round spermatids (RS), spermatozoa, and oocytes from the supplier.</li>
		<li>Prepare the Ca2+-free C.L. Chatot, C.A. Ziomek, and B.D. Bavister (CZB) medium for AOA following the previously published protocol20.</li>
		<li>Obtain the potassium simplex optimized medium supplemented with amino acids (KSOMaa) solution for embryo culture from the supplier.</li>
		<li>Prepare hyaluronidase, with a working concentration of 0.1% (1 g/L), by dissolving 0.1 g of hyaluronidase in 1 mL of embryo water. Take 10 &#181;L of concentrated storage solution and dissolve in 990 mL of M2, which could be stored at 4 &#176;C for 1 week.</li>
		<li>Prepare polyvinylpyrrolidone (PVP), with an average molecular weight of 360,000 and a mass concentration of 12%, by dissolving 1.2 g of PVP in 10 mL of M2. The resulting solution could be stored at 4 &#176;C for 1 week.</li>
		<li>Store the cytochalasin B (CB) stock solution (2 &#181;L), soluble in 18 &#181;L of DMSO at -20 &#176;C. Take 5 &#181;L of the concentrated storage solution and dissolve in 995 mL of M2. The resulting solution could be stored at 4 &#176;C for 1 week.</li>
	</ol>
	</li>
</ol>
2. Preparation of oocytes
<ol>
	<li>Acclimate female B6D2F1 mice, aged 6-8 weeks, to their new environment for at least 1 week. Inject intraperitoneally with 7.5 IU of pregnant mare serum gonadotropin (PMSG) at 5 pm, followed by 7.5 IU of human chorionic gonadotropin (HCG) 48 h later. Ensure no contact with male mice occurs after the injection. 
	NOTE: The specified time (5 pm) is not mandatory. It is just for work convenience. The 14 hours after 5 pm is 7 am, when the oocytes are ready, and the injection is performed, so the injection time is scheduled for the morning. The start time of the injection can be adjusted according to the laboratory&#39;s daily schedule.</li>
	<li>After 14 h of HCG injection, euthanize the mice using the cervical dislocation method. Open the abdominal cavity and locate the uterus. Identify the ovaries along the uterus and sever the fallopian tubes near the ovaries using scissors. Repeat the same procedure on the other side and place the fallopian tubes in M2, which is preheated at 37 &#176;C in advance.</li>
	<li>Under an upright microscope (10x), locate the enlarged part of the fallopian tube, showing a transparent and swollen appearance. Use a 1 mL syringe to secure the fallopian tube while cutting the enlarged part, collecting oocyte coronal cumulus complexes (OCCCs).</li>
	<li>Place OCCCs into the preheated M2 operating solution containing 0.1% hyaluronidase and remove granulosa cells by gently blowing through an oral pipette. 
	NOTE: Keep the OCCCs in 0.1% hyaluronidase for a short duration, approximately 2 min, if needed.</li>
	<li>After removing the granulosa cells, transfer the oocyte to KSOMaa, rinse 3x, and place in a carbon dioxide incubator for later use (37 &#176;C, 5% CO2).</li>
</ol>
3. Preparation of round spermatids and spermatozoa
<ol>
	<li>Collect round spermatids and spermatozoa from the testis and epididymis of male C57BL/6 mice (8-10 weeks), respectively.
	<ol>
		<li>For spermatozoa, euthanize the mice by cervical dislocation. Open the abdominal cavity, locate the epididymis near the testis, and gently excise with scissors.</li>
		<li>Place the epididymis in M2 medium and gently excise using a 1 mL syringe to allow spermatozoa to flow out under an upright microscope (10x).</li>
		<li>Siphon the flowing spermatozoa and M2 suspension into the bottom of a tube containing 0.5 mL of M2, allowing the spermatozoa to flow upstream naturally. Set them aside for later use.</li>
		<li>For round spermatids, remove the testicles from the abdominal cavity and place them in M2. Gently cut the white membrane through using a 1 mL syringe and squeeze the convoluted seminiferous tubules out under an upright microscope (10x).</li>
		<li>Excise the seminiferous tubules carefully using two 1 mL syringes. Extract the suspension and filter with a 400-mesh sieve (38 &#181;m).</li>
	</ol>
	</li>
	<li>Centrifuge the filtered cells, discard the supernatant, and add 200 &#181;L of Hoechst 33342 for staining, followed by incubation at 37 &#176;C for 10 min.</li>
	<li>Perform fluorescence-activated cell sorting (FACS) to distinguish round spermatids from other cell types. Place the stained cells into a flow cytometry tube and perform a round spermatid screening.
	<ol>
		<li>Different models of flow cytometry machines have different regulatory parameters and can be communicated with technical support during operation. As the first step, adjust the voltage and select the cell population according to forward scatter (FSS) and side scatter (SSC; see Supplementary Table 1).</li>
		<li>For the second step, remove cell adhesions according to FSC and trigger pulse width.</li>
		<li>As the third step, select the target haploid round spermatids. Hoechst 33342 binds to DNA, and cells with different ploidy exhibit different fluorescence intensities. Set the excitation wavelength to 355. Select two detection channels, 460/50 and 670/30, under 355 wavelength laser to sort round spermatids. Subsequently, store the sorted round spermatids in a 4 &#176;C refrigerator for later use. 
		NOTE: It is also possible to directly select round spermatids based on their morphology under a microscope, but training is required to ensure selection accuracy.</li>
	</ol>
	</li>
</ol>
4. Round spermatid injection (ROSI)
<ol>
	<li>Carry out the ROSI procedure using an inverted microscope with a micromanipulation system.</li>
	<li>In the initial preparation step, make holding and injection needles according to the previously described procedure21. Keep the inner diameter of the injection needle between 6-7 &#181;m, designed for the extraction of round spermatids, and for the holding needle, have an inner diameter of 20 &#181;m for securing the oocyte.</li>
	<li>In the subsequent preparation step, create an injection vessel by placing 10 &#181;L of M2 droplets containing CB to soften oocytes and 10 &#181;L of M2 droplets for storing round spermatids. Use PVP droplets to moisten and cleanse the injection needles. 
	NOTE: Adding CB here is not a necessary step, but adding CB can greatly improve the survival rate after injection.</li>
	<li>Extract round spermatids using an injection needle.</li>
	<li>Rotate the oocyte with a holding needle, positioning the oocyte&#39;s polar body at either the 12 o&#39;clock or 6 o&#39;clock position.</li>
	<li>Place the injection needle carefully at the 3 o&#39;clock position on the oocyte, tightly adhering to the zona pellucida. Use a PiezoXpert device to create a hole in the zona pellucida. Adjust the strength from small to large, which is approximately intensity = 5 and speed = 15 here.</li>
	<li>Gently advance the injection needle through the zona pellucida and enter the oocyte horizontally. When it passes the center of the oocyte, apply a piezo (intensity = 1 and speed = 1), resulting in the rupture of the oocyte membrane and the gradual injection of the round spermatid into the oocyte&#39;s cytoplasm.</li>
	<li>Withdraw the injection needle and aspirate a small amount of the oocyte&#39;s membrane near the opening to seal it. This step is crucial, as failure to seal would result in cytoplasmic leakage and oocyte death. 
	NOTE: It is worth noting that ROSI required AOA either before or after the injection of round spermatids, which we have discussed in subsequent sections.</li>
</ol>
5. Intracytoplasmic sperm injection (ICSI)
<ol>
	<li>Carry out the same steps for ICSI as for ROSI, with the only difference being the diameter of the ICSI injection needle, which measures 9-10 &#181;m.</li>
	<li>Place spermatozoa on a PVP runway to reduce their swimming speed and facilitate sperm extraction.</li>
	<li>Perform separation of the spermatozoa head and tail for ICSI. Aspirate the spermatozoa from the tail and position the spermatozoa&#39;s neck precisely at the mouth of the injection needle. Apply a piezo (multiple piezo could be used as needed, intensity = 5 and speed = 15) to separate the head and tail of the spermatozoa.</li>
</ol>
6. Assisted oocyte activation (AOA)
<ol>
	<li>According to the protocol, prepare a CZB medium without calcium and magnesium.</li>
	<li>Use strontium chloride hexahydrate (SrCl26H2O), with a molecular weight of 266.62, at a concentration of 10 mM. To prepare it, dissolve 0.26662 g of SrCl26H2O in 1 mL of embryo water. At the time of use, add 10 &#181;L of concentrated storage solution to 990 &#181;L of Ca2+-free CZB medium and use immediately.</li>
	<li>Place the oocytes in the Ca2+-free CZB medium containing SrCl26H2O for activation; each droplet has a volume of 20 &#181;L, and 20 oocytes are placed each time. Incubate for 20 min.</li>
	<li>Simultaneously, establish a group for parthenogenesis activation without injecting round spermatids or spermatozoa. It should be noted that the parthenogenesis activation group is shown without statistical analysis, and the control group is ICSI.</li>
</ol>
7. Embryo transfer
<ol>
	<li>House the ligated ICR male and female mice in a ratio of 1:2. The following morning, examine the female mice for the presence of a vaginal plug and select those with a vaginal plug for transplantation.</li>
	<li>Inject the transferred embryos consisting of 2-cell embryos the previous morning and transfer them the next afternoon.</li>
	<li>Make an incision on the back of the female mouse, expose the white fat and ovaries, and locate the enlarged part of the fallopian tube under an upright microscope (10x).</li>
	<li>Use a 1 mL syringe to create an opening in the enlarged part of the fallopian tube and aspirate the embryos through an oral pipette. Gently introduce the embryos in the direction of the uterus. Suture the wound, using 4 x 10 models of curved needles with thread, with a thickness of 5-0 (0.09-0.11 mm). The suture is made in two layers, with one layer for the peritoneum and one layer for the outer skin.</li>
	<li>For surrogate mice, perform cesarean sections based on the experimental schedule. Euthanize the mice by the cervical dislocation method, disinfect with alcohol, and open the abdominal cavity. Gently and quickly cut open the uterus and open the amniotic cavity to show the fetus and placenta. Remove the umbilical cord and cut from the side close to the fetus; the fetus was fed by the same period of breastfeeding mice. 
	NOTE: Due to the need to calculate birth rates, cesarean section is more accurate regarding production capacity. In natural childbirth, the offspring may be eaten by the mother, resulting in inaccurate statistics. The cesarean section is performed on surrogate mice transplanted in the early stage, and euthanasia cesarean section is performed before delivery.</li>
</ol>
8. Statistical analysis
<ol>
	<li>Use GraphPad Prism 5 to analyze. Data are expressed in mean &#177; SD; P &#60; 0.05 indicates a statistical difference, and P &#60; 0.01 indicates a significant statistical difference.</li>
</ol>

Murine Round Spermatid Injection Procedure to Study Embryonic Development

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The authors declare no conflicts of financial or other interests.

Assisted oocyte activation 
A critical prerequisite for ROSI is AOA since round spermatids alone cannot initiate oocyte activation. Currently, the most established method in mice involves the use of strontium chloride23,24, while the most advanced human application employs electrical activation13,14. The timing of oocyte activation is also of great significance. As reported in the li...

The final stage of spermatogenesis involves the transformation of a round spermatid into a fully developed spermatozoon, characterized by distinct head, neck, and elongated tail structures1. This transformation encompasses significant changes in cell morphology, such as the condensation of chromatin in the nucleus, replacement of histones by protamine, acrosome formation, mitochondrial sheath development, centriole migration and loss, tail structure formation, and the removal of cellular residues2.
In 1992, the first human fetus was successfully born through intracytoplasmic sperm injection (ICSI) technology3. Since then, researchers have been exploring the potential of utilizing round spermatids, which share the same haploid genetic composition as mature spermatozoa, to fertilize oocytes and sustain viable pregnancies2,4. Subsequently, in 1996, the first human fetus conceived via round spermatid injection (ROSI) technology was delivered5,6. It is worth noting that studies involving ICSI and ROSI in mice lagged behind those in humans due to the susceptibility of the mouse oocyte membrane to damage during the injection process. This issue was successfully resolved with the introduction of the Piezo membrane-breaking device. Consequently, in 1995, the first mouse conceived through ROSI technology was born. Additionally, research on ROSI in various other animals is also underway7,8.
Currently, research on ROSI primarily centers around the following aspects: clinical application, mechanism elucidation, and strategies to enhance developmental efficiency, along with broader applications of ROSI technology. In the context of clinical applications, despite the birth of the first ROSI human fetus through ROSI in 1996, progress has been marked by a series of successes and failures9,10,11,12. To date, ROSI technology has not achieved widespread clinical implementation, largely due to its low efficiency and the need for further validation regarding the safety of fetuses conceived through ROSI technology. Incomplete statistics indicate that globally, fewer than 200 human ROSI-conceived fetuses have been delivered. A turning point in understanding the potential of ROSI technology occurred in 2015 when Tanaka and colleagues reported on the successful birth of 14 fetuses through ROSI technology, instilling renewed confidence in its clinical application and feasibility13,14. ROSI technology holds substantial promise for addressing reproductive biology challenges, particularly in non-obstructive azoospermia patients. In addition to its clinical applications, ROSI serves as a valuable tool for studying the intricate mechanisms of embryonic development15,16,17.
Numerous animal studies have been conducted to investigate the underlying factors contributing to the low efficiency of ROSI in achieving full embryonic development. These factors encompass the choice of assisted oocyte activation (AOA) methods and their timings, abnormalities in genomic stability, and, particularly, abnormalities in epigenetic modifications. It is important to recognize that round spermatids are immature germ cells, differing significantly from mature spermatozoa in various physiological aspects. Mizuki Sakamoto and colleagues indicated that H3K27me3, derived from round spermatids, is associated with chromatin that is less accessible and leads to impaired gene expression in ROSI embryos18. In a related study by Jing Wang and colleagues, reprogramming defects in ROSI embryos at the pronuclear stages were predominantly associated with the misexpression of a cohort of the genes responsible for minor zygotic genome activation19. They also found that introducing a selective euchromatic histone lysine methyltransferase 2 inhibitor, A366, could potentially enhance the overall developmental rate by approximately twofold.
The mouse stands as one of the most valuable model animals for studying embryonic development. This article elaborates on how to perform ROSI on mice. This comprehensive protocol encompasses the selection of suitable mice, detailed ovulation induction procedures, AOA techniques, injection techniques, and the preparation of surrogate mice. Furthermore, we present a comparative analysis of the effects of two injection regimens on birth efficiency: AOA followed by ROSI (A-ROSI; first regimen) and ROSI followed by AOA (ROSI-A; second regimen). We aim to encourage researchers to conduct mouse ROSI experiments with greater precision, offering more robust support for their clinical application and the fundamental research of embryonic development mechanisms.

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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.

We initiated our investigation by examining AOA&#39;s effect on embryos&#39; developmental capability. A schematic illustration of the experimental design is shown in Figure 1A. Before the spermatozoon injection, the oocytes underwent either AOA (A-ICSI) or remained untreated (ICSI). Detailed data on embryonic development is presented in Table 1. The results revealed no significant differences in cleavage, blastocyst, or birth rates between the A-ICSI and ICSI groups (P<...

Read this Scientific Article Protocol about Mouse Round Spermatid Injection at JoVE.com

Mouse Round Spermatid Injection

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 ...

Research

JoVE Journal

Developmental Biology

554 Views.  First Hospital of Jilin University. 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.

Video: Mouse Round Spermatid Injection

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the cephalochordate amphioxus, Branchiostoma lanceolatum. We use constructs for membrane and nuclear targeted mCherry and eGFP that have been modified to accommodate amphioxus codon usage and Kozak consensus sequences. We describe the type of injection needles to be used, the immobilization protocol for the unfertilized oocytes, and the overall injection set-up. This technique generates fluorescently labeled embryos, in which the dynamics of cell behaviors during early development can be analyzed using the latest in vivo imaging strategies. The development of a microinjection technique in this amphioxus species will allow live imaging analyses of cell behaviors in the embryo as well as gene-specific manipulations, including gene overexpression and knockdown. Altogether, this protocol will further consolidate the basal chordate amphioxus as an animal model for addressing questions related to the mechanisms of embryonic development and, more importantly, to their evolution.

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here the robust and efficient expression of fluorescent proteins after mRNA injection into unfertilized oocytes of Branchiostoma lanceolatum. The development of the microinjection technique in this basal chordate will pave the way for far-reaching technical innovations in this emerging model system, including in vivo imaging and gene-specific manipulations.

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the ...

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

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during oogenesis. In order to study roles of such maternal factors in early embryonic development, it is desirable to manipulate their functions from the very beginning of embryonic development. Conventional ways of gene interference are achieved by injection of antisense oligonucleotides (oligos) or mRNA into fertilized eggs, enabling under- or over-expression of specific proteins, respectively. However, these methods normally require more than several hours until protein expression is affected, and, hence, the interference of gene functions is not effective during early embryonic stages. Here, we introduce an experimental system in which expression levels of maternal proteins can be altered before fertilization. Xenopus laevis oocytes obtained from ovaries are defolliculated by incubating with enzymes. Antisense oligos or mRNAs are injected into defolliculated oocytes at the germinal vesicle (GV) stage. These oocytes are in vitro matured to eggs at the metaphase II (MII) stage, followed by intracytoplasmic sperm injection (ICSI). By this way, up to 10% of ICSI embryos can reach the swimming tadpole stage, thus allowing functional tests of specific gene knockdown or overexpression. This approach can be a useful way to study roles of maternally stored factors in early embryonic development.

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

We describe methods of manipulating Xenopus laevis immature oocytes, in vitro maturation of oocytes to eggs, and intracytoplasmic sperm injection. This protocol allows degradation of some maternal proteins and overexpression of genes of interest at fertilization, and hence is valuable to study roles of specific factors in early embryonic development.

Amphibian eggs have been widely used to study embryonic development. Early embryonic development is driven by maternally stored factors accumulated during ...

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

All developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, fluid and nutrient supply). Consequently both the nervous and vascular systems develop alongside each other and share striking similarities in their branching architecture. Here we report embryonic manipulations that allow us to study the simultaneous development of neural crest-derived nervous tissue (in this case the enteric nervous system), and the vascular system. This is achieved by generating chicken chimeras via transplantation of discrete segments of the neural tube, and associated neural crest, combined with vascular DiI injection in the same embryo. Our method uses transgenic chickGFP embryos for intraspecies grafting, making the transplant technique more powerful than the classical quail-chick interspecies grafting protocol used with great effect since the 1970s. ChickGFP-chick intraspecies grafting facilitates imaging of transplanted cells and their projections in intact tissues, and eliminates any potential bias in cell development linked to species differences. This method takes full advantage of the ease of access of the avian embryo (compared with other vertebrate embryos) to study the co-development of the enteric nervous system and the vascular system.

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

Here we report dual labeling of neural crest cells and blood vessels using chickGFP neural tube intraspecies grafting combined with intra-vascular DiI injection. This experimental technique allows us to simultaneously visualize and study development of the NCC-derived (enteric) nervous system and the vascular system, during organogenesis.

All developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, ...

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced skeletal muscle regeneration is a widely used, powerful model system to study the events involved in muscle regeneration as well as the mechanisms and different players. Indeed, a detailed knowledge of this process is essential for a better understanding of the pathological conditions that lead to skeletal muscle degeneration, and it aids in identifying new targeted therapeutic strategies. The present work describes a detailed and reproducible protocol to induce acute skeletal muscle regeneration in mice through a single intramuscular injection of cardiotoxin (CTX). CTX belongs to the family of snake venom toxins and causes myolysis of myofibers, which eventually triggers the regeneration events. The dynamics of skeletal muscle regeneration is evaluated by histological analysis of muscle sections. The protocol also illustrates the experimental procedures for dissecting, freezing, and cutting the Tibialis Anterior muscle, as well as the routine Hematoxylin &#38; Eosin staining that is widely used for subsequent morphological and morphometric analysis.

This manuscript describes a detailed protocol to induce acute skeletal muscle regeneration in adult mice and subsequent manipulations of the muscles, such as dissection, freezing, cutting, routine staining, and myofiber cross-sectional area analysis.

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced ...

Dissection and Culture of Mouse Embryonic Kidney

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney development, the two progenitor tissues, the ureteric bud and the metanephric mesenchyme, communicate and reciprocally induce cellular mechanisms to eventually form the collecting system and the nephrons of the kidney. As mammalian embryos grow intrauterine and therefore are inaccessible to the observer, an organ culture has been developed. With this method, it is possible to study epithelial-mesenchymal interactions and cellular behavior during kidney organogenesis. Furthermore, the origin of congenital kidney and urogenital tract malformations can be investigated. After careful dissection, the metanephric rudiments are transferred onto a filter that floats on culture medium and can be kept in a cell culture incubator for several days. However, one must be aware that the conditions are artificial and could influence the metabolism in the tissue. Also, the penetration of test substances could be limited due to the extracellular matrix and basal membrane present in the explant.
One main advantage of organ culture is that the experimenter can gain direct access to the organ. This technology is cheap, simple, and allows a large number of modifications, such as the addition of biologically active substances, the study of genetic variants, and the application of advanced imaging techniques.

This protocol describes a method for isolating and culturing metanephric rudiments from mouse embryos.

The goal of this protocol is to describe a method for the dissection, isolation, and culture of mouse metanephric rudiments. 
During mammalian kidney ...

Subcutaneous Neurotrophin 4 Infusion Using Osmotic Pumps or Direct Muscular Injection Enhances Aging Rat Laryngeal Muscles

Laryngeal dysfunction in the elderly is a major cause of disability, from voice disorders to dysphagia and loss of airway protective reflexes. Few, if any, therapies exist that target age-related laryngeal muscle dysfunction. Neurotrophins are involved in muscle innervation and differentiation of neuromuscular junctions (NMJs). It is thought that neurotrophins enhance neuromuscular transmission by increasing neurotransmitter release. The neuromuscular junctions (NMJs) become smaller and less abundant in aging rat laryngeal muscles, with evidence of functional denervation. We explored the effects of NTF4 for future clinical use as a therapeutic to improve function in aging human laryngeal muscles. Here, we provide the detailed protocol for systemic application and direct injection of NTF4 to investigate the ability of aging rat laryngeal muscle to remodel in response to NTF4 application. In this method, rats either received NTF4 either systemically via osmotic pump or by direct injection through the vocal folds. Laryngeal muscles were then dissected and used for histological examination of morphology and age-related denervation.

Here, we present a protocol to describe the use of neurotrophin 4 (NTF4) systemically and directly to remodel rat aging laryngeal muscles.

Laryngeal dysfunction in the elderly is a major cause of disability, from voice disorders to dysphagia and loss of airway protective reflexes. Few, if ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

A Novel Use of Three-dimensional High-frequency Ultrasonography for Early Pregnancy Characterization in the Mouse

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is routinely used as an in vivo model to study embryo implantation and pregnancy progression. Unfortunately, such murine studies require pregnancy interruption to enable follow-up phenotypic analysis. To address this issue, we used three-dimensional (3-D) reconstruction of HFUS imaging data for early detection and characterization of murine embryo implantation sites and their individual developmental progression in utero. Combining HFUS imaging with 3-D reconstruction and modeling, we were able to accurately quantify embryo implantation site number as well as monitor developmental progression in pregnant C57BL6J/129S mice from 5.5 days post coitus (d.p.c.) through to 9.5 d.p.c. with the use of a transducer. Measurements included: number, location, and volume of implantation sites as well as inter-implantation site spacing; embryo viability was assessed by cardiac activity monitoring. In the immediate post-implantation period (5.5 to 8.5 d.p.c.), 3-D reconstruction of the gravid uterus in both mesh and solid overlay format enabled visual representation of the developing pregnancies within each uterine horn. As genetically engineered mice continue to be used to characterize female reproductive phenotypes derived from uterine dysfunction, this method offers a new approach to detect, quantify, and characterize early implantation events in vivo. This novel use of 3-D HFUS imaging demonstrates the ability to successfully detect, visualize, and characterize embryo-implantation sites during early murine pregnancy in a non-invasive manner. The technology offers a significant improvement over current methods, which rely on the interruption of pregnancies for gross tissue and histopathologic characterization. Here we use a video and text format to describe how to successfully perform ultrasounds of early murine pregnancy to generate reliable and reproducible data with reconstruction of the uterine form in mesh and solid 3-D images.

Mice are widely used to study gestational biology. However, pregnancy termination is required for such studies which precludes longitudinal investigations and necessitates the use of large numbers of animals. Therefore, we describe a non-invasive technique of high-frequency ultrasonography for early detection and monitoring of post-implantation events in the pregnant mouse.

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is ...

A Mouse Distraction Osteogenesis Model

Distraction osteogenesis (DO) is a surgical procedure that involves skeletal tissue regeneration without cell transplantation. A DO model consists of the following three phases: the latency phase after osteotomy and placement of the external distractor; the distraction phase, wherein the separated bone ends are gradually and continuously distracted; and the consolidation phase. This custom-made distractor used for DO is comprised of two incomplete acrylic resin rings and an expansion screw. The process was initiated by making a mold with silicone impression material and then creating the custom-made distractor. Dental resin was poured into the formwork made of silicone impression material, and it was allowed to polymerize to create the incomplete resin rings required for the custom-made distractor. These rings were fixed with an expansion screw using transparent resin. The custom-made distractor created via this approach was attached to the tibia of mice. The tibia was fixed to the device using one pair of 25 G needles proximally, one pair of 27 G needles distally, and acrylic resin. After a latency period of 5 days, distraction was initiated at a rate of 0.2 mm/12 h. The lengthening was continued for 8 days, resulting in a total gap of 3.2 mm. The mice were sacrificed 4 weeks after distraction. Bone formation in the distraction gap was confirmed using both radiography and histology.

We present a mouse tibial distraction osteogenesis model developed using a custom-made distractor. The use of a mouse as an analysis target is advantageous for advancing research.

Distraction osteogenesis (DO) is a surgical procedure that involves skeletal tissue regeneration without cell transplantation. A DO model consists of the ...

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and behave by observing the surface of a tissue. This method provides a means to visualize the complex 3-dimensional vascular architecture of the lung.
To accomplish this, a catheter is inserted into the pulmonary artery and the vasculature is simultaneously flushed of blood and chemically dilated to limit resistance. Lungs are then inflated through the trachea at a standard pressure and the polymer compound is infused into the vascular bed at a standard flow rate. Once the entire arterial network is filled and allowed to cure, the lung vasculature may be visualized directly or imaged on a micro-CT (&#181;CT) scanner.
When performed successfully, one can appreciate the pulmonary arterial network in mice ranging from early postnatal ages to adults. Additionally, while demonstrated in the pulmonary arterial bed, this method can be applied to any vascular bed with optimized catheter placement and endpoints.

The aim of this technique is ex vivo visualization of pulmonary arterial networks of early postnatal and adult mice through lung inflation and injection of a radio-opaque polymer-based compound via the pulmonary artery. Potential applications for casted tissues are also discussed.

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and ...

Mouse Round Spermatid Injection

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Chapters in this video

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

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

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using Chick^GFPNeural Tube Grafting and Carbocyanine Dye DiI Injection

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

Dissection and Culture of Mouse Embryonic Kidney

Subcutaneous Neurotrophin 4 Infusion Using Osmotic Pumps or Direct Muscular Injection Enhances Aging Rat Laryngeal Muscles

Live Imaging of Mouse Secondary Palate Fusion

A Novel Use of Three-dimensional High-frequency Ultrasonography for Early Pregnancy Characterization in the Mouse

A Mouse Distraction Osteogenesis Model

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging