This work was supported by grant R01 EY032056 (to CC) from the National Eye Institute. The authors thank Dr. Theresa Fassel and Kimberly Vanderpool at the Scripps Research Core Microscopy Facility for their assistance with the electron microscope images.

Mice have been cared for based on an animal protocol approved by the Institutional Animal Care and Use Committee at Indiana University Bloomington. The mice used to generate representative data were control (wild-type) animals in the C57BL6/J background, female, and 8-12 weeks old. Both male and female mice can be used for this experiment, since the sex of the mice is very unlikely to affect the experiment&#39;s outcome.
1. Lens dissection and decapsulation
<ol>
	<li>Euthanize mice following the National Institutes of Health&#39;s &#34;Guide for the Care and Use of Laboratory Animals&#34; as well as animal use protocols approved by the institution. 
	NOTE: For the present study, mice were euthanized by CO2 overdose followed by cervical dislocation in accordance with an approved animal protocol (Indiana University).</li>
	<li>Enucleate the eyes from the mice using curved forceps by depressing the tissue around the eyes with one side of the forceps to displace the eye out of the socket. Next, close the forceps underneath the eye and lift to remove the eye from the socket. Transfer the eyes to fresh 1x phosphate-buffered saline (PBS) in a dissection tray.</li>
	<li>Cut the optic nerve with ultra-fine scissors as close as possible to the eyeball. Carefully insert fine-tip, straight tweezers into the eyeball through the optic nerve&#39;s exit at the posterior of the eye.</li>
	<li>Carefully insert scissors at the same location as the tweezers in step 1.3 and begin cutting an incision from the posterior toward the corneal-scleral junction. 
	NOTE: Rodent lenses occupy ~30% of their eyes. Accidental damage will occur if the tweezers or scissors are inserted too deep into the eye.</li>
	<li>Continue cutting along the corneal-scleral junction until at least half of the junction has been separated.</li>
	<li>Use tweezers to gently push on the cornea so that the lens can exit through the incision made with steps 1.4 and 1.5.</li>
	<li>Carefully remove any large pieces of tissue from the lens using fine-tip, straight tweezers. Inspect the lens to find the equatorial region.</li>
	<li>Shallowly pierce the lens using fine-tip, straight tweezers, and then remove the lens capsule. The mass of lens fiber cells will remain intact, and the lens epithelial monolayer will remain attached to the lens capsule. Discard the lens capsule.</li>
</ol>
2. Lens single fiber cell staining
<ol>
	<li>Transfer the lens fiber cell mass to a well plate with 500 &#181;L of freshly made 1% paraformaldehyde (PFA; in 1x PBS) solution and incubate the samples at 4 &#176;C overnight with gentle nutation (Figure 2A). 
	NOTE: The volumes for the solutions given are optimized for 48-well plates. If another size well-plate or tubes are used, adjust the volumes of the solutions accordingly. Fixation, blocking, and washing (1x PTX, 0.1% Triton X-100 in 1x PBS) solutions were made with 10x PBS and double distilled water (ddH2O) to a final concentration of 1x PBS.</li>
	<li>Transfer the lens fiber cells to a 60 mm dish with 1% PFA. Use a sharp scalpel to split the ball of fiber cells in half along its anterior-posterior axis (Figure 2B). Cut the halves in half again along the same axis to produce quarters. 
	NOTE: The anterior-posterior axis is easily recognized by the direction of the fiber cells in the tissue mass.</li>
	<li>Use straight tweezers to remove the nucleus region from the lens fiber cell quarters (Figure 2C). 
	NOTE: The lens nucleus region is rigid in rodent lenses, and the center of the lens will separate from the softer cortical fibers easily.</li>
	<li>Post-fix the quarters of the lens cortex region in 200 &#181;L of 1% PFA for 15 min at room temperature (RT) with gentle shaking (300 rpm on a plate shaker).</li>
	<li>Wash the tissue quarters twice in 750 &#181;L of 1x PBS for 5 min each with gentle shaking at RT.</li>
	<li>Block the samples using 200 &#181;L of blocking solution (5% serum, 0.3% Triton X-100, 1x PBS) for 1 h at RT with gentle shaking. 
	NOTE: For the representative data in this protocol, the samples were not stained with primary or secondary antibodies. After the blocking step, the samples were incubated with wheat germ agglutinin (WGA; 1:100) and phalloidin (1:100) (see Table of Materials) for 3 h at RT with gentle shaking and protection from light. Primary antibody staining has been demonstrated in previous publications22,23.</li>
	<li>Incubate with 100 &#181;L of primary antibody solution overnight at 4 &#176;C with gentle shaking. 
	NOTE: The antibodies are diluted in blocking solution. Compared to slides with tissue sections, there are more cells in this type of preparation. Primary antibody concentrations should be increased to provide ample antibodies to stain more cells. Doubling the antibody concentration from what is used on tissue sections is recommended. The same applies to secondary antibodies.</li>
	<li>Wash the tissue quarters three times with 1x PTX (0.1% Triton X-100, 1x PBS) for 5 min each at RT with gentle shaking.</li>
	<li>Incubate the fiber cells with 100 &#181;L of secondary antibody/dye solution for 3 h at RT with gentle shaking. Protect the samples from light during this and subsequent steps. 
	NOTE: WGA, phalloidin, and other fluorescent dyes can be added to the secondary antibody solution for simultaneous labeling of the cell membrane, cytoskeleton, or other organelles while also labeling the primary antibody.</li>
	<li>Wash the fiber cells four times with 1x PTX for 5 min each at RT with gentle shaking.</li>
	<li>Add one drop or 50 &#181;L of mounting media onto a plus-charged microscope slide before transferring the tissue quarters to the slide.</li>
	<li>Use tweezers to gently separate the fiber cells from each other and try to limit overlapping of the cell bundles.</li>
	<li>Gently apply a #1.5 coverslip on top of the sample in mounting media. The mounting media should spread to the edge of the coverslip; if this does not occur, add some additional mounting media to the edge of the coverslip. Aspirate away any excess mounting media around the edge of the coverslip and use nail polish to seal the edges of the coverslip on the slide. 
	&#8203;NOTE: Any type of mounting medium that is formulated for confocal microscopy can be used for these experiments.</li>
</ol>
3. Lens nucleus single fiber cell staining
<ol>
	<li>Complete the dissection outlined in section 1.</li>
	<li>Mechanically remove the cortical fibers from the ball of the lens fiber cells, leaving the lens nucleus, by gently transferring the fiber cell mass to wet, gloved fingertips and gently rolling the tissue mass. 
	NOTE: In mouse lenses, rolling the ball of lens fiber cells between gloved fingertips is an effective method for removal of the cortical fiber cells from the hard lens nucleus28,30. For lenses where this mechanical method is not effective, careful dissection or a vortexing method29 may be used to remove the cortical fiber cells.</li>
	<li>Transfer the lens nucleus to freshly made 1% PFA solution in a well-plate and incubate overnight at 4 &#176;C with gentle shaking.</li>
	<li>Transfer the sample to a 60 mm dish with 1% PFA and use a sharp scalpel to split the lens nucleus along the anterior-posterior axis. Halve the tissue samples again to produce nucleus quarters.</li>
	<li>Follow the process outlined in steps 2.4-2.13 for immunostaining and sample mounting.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65638/65638fig02v2.jpg" /> 
Figure 2: Graphical summary detailing the preparation and immunostaining of lens fiber cells. (A) This 48-well plate has been color-coded by column to demonstrate a sample plate setup for the described methods, allowing easy transfer of samples between the various immunostaining steps by gentle handling using forceps. While the representative data for this protocol is not incubated with a primary antibody, the diagram includes a column for primary antibody incubation, and the wells for washing can be reused after removing used wash buffers by aspiration. (B) After fixation of the lens fiber cell mass, the tissue is split along the anterior-posterior axis (red dashed lines) to preserve the original structure of the cells. Once the tissue mass has been halved, the samples are rotated and the halves split into quarters along the anterior-posterior axis (red dashed lines). (C) Removing the lens nucleus region (in pink) is easily done using tweezers to dig out the dense central tissue from the cortical fiber cells (in blue). Cartoon diagrams were partially created using BioRender.com and not drawn to scale. <a href="https://www.jove.com/files/ftp_upload/65638/65638fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Immunofluorescence Staining of Membrane Interdigitations in Lens Fiber Cells

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The authors have nothing to disclose.

This protocol has demonstrated the fixation, preservation, and immunostaining methods that faithfully preserve the 3D membrane morphology of bundles or singular lens fiber cells from various depths in the lens. The stained lens fibers are compared with SEM preparations that have long been used to study lens fiber cell morphology. The results show comparable membrane structures between both preparations. EM remains the gold standard for studying cell morphology, but immunolabeling is more challenging in SEM samples for lo...

The lens is a clear and ovoid tissue in the anterior chamber of the eye that is made up of two cell types, epithelial and fiber cells1 (Figure 1). There is a monolayer of epithelial cells that covers the anterior hemisphere of the lens. Fiber cells are differentiated from epithelial cells and make up the bulk of the lens. The highly specialized fiber cells undergo an elongation, differentiation, and maturation programming, marked by distinct changes in cell membrane morphology from the lens periphery to the lens center2,3,4,5,6,7,8,9,10,11,12, also known as the lens nucleus. The function of the lens to fine-focus light coming from various distances onto the retina depends on its biomechanical properties, including stiffness and elasticity13,14,15,16,17,18,19. The complex interdigitations of lens fibers have been hypothesized20,21 and recently shown to be important for lens stiffness22,23.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65638/65638fig01.jpg" /> 
Figure 1: Lens anatomy diagrams and representative scanning electron microscopy (SEM) images from lens fibers. The cartoon shows a longitudinal (anterior to posterior from top to bottom) view of the anterior monolayer of epithelial cells (shaded in light blue) and a bulk mass of lens fiber cells (white). The center of the lens (shaded in pink) is known as the nucleus and comprises highly compacted fiber cells. On the right, a cross-section cartoon reveals the elongated hexagon cell shape of lens fibers that are packed into a honeycomb pattern. Fiber cells have two broad sides and four short sides. Representative SEM images along the bottom show the complex membrane interdigitations between lens fiber cells at different depths of the lens. From the right, newly formed lens fibers at the lens periphery have small protrusions along the short sides and balls-and-sockets along the broad side (red boxes). During maturation, lens fibers develop large paddle domains that are decorated by small protrusions along the short sides (blue boxes). Mature fiber cells possess large paddle domains illustrated by small protrusions. These interlocking domains are important for lens biomechanical properties. Fiber cells in the lens nucleus have fewer small protrusions along their short sides and have complex tongue-and-groove interdigitations (purple boxes). The broad sides of the cell display a globular membrane morphology. The cartoon was modified from22,32 and not drawn to scale. Scale bar = 3 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65638/65638fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The lens grows by adding shells of new fiber cells overlaid on top of previous generations of fibers24,25. Fiber cells have an elongated, hexagonal cross section shape with two broad sides and four short sides. These cells extend from the anterior to the posterior pole of the lens, and depending on the species, the lens fibers can be several millimeters in length. To support the structure of these elongated and skinny cells, specialized interdigitations along the broad and short sides create interlocking structures to maintain the lens shape and biomechanical properties. Changes in cell membrane shape during fiber cell differentiation and maturation have been extensively documented by electron microscopy (EM) studies2,3,4,5,6,7,8,9,10,20,26,27,28,29. Newly formed fiber cells have balls-and-sockets along their broad sides with very small protrusions along their short sides, while mature fibers have interlocking protrusions and paddles along their short sides. Nuclear fibers display tongue-and-groove interdigitations and globular membrane morphology. Little is known about the proteins that are required for these complex interlocking membranes. Previous studies on protein localization in fiber cells have relied on lens tissue sections, which do not allow clear visualization of the complex cell architecture.
This work has created and perfected a novel method to fix single and bundles of lens fiber cells to preserve the complex morphology and to allow immunostaining for proteins at the cell membrane and within the cytoplasm. This method faithfully preserves cell membrane architecture, comparable to data from EM studies, and allows staining with primary antibodies for specific proteins. We have previously immunostained cortical lens fibers undergoing differentiation and maturation22,23. In this protocol, there is also a new method to stain fiber cells from the lens nucleus. This protocol opens the door to understanding the mechanisms for formation and changes in membrane interdigitations during fiber cell maturation and lens nucleus compaction.

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preparation and immunofluorescence staining of bundles and single fiber cells from the cortex and nucleus of the eye lens

The lens is a transparent and ellipsoid organ in the anterior chamber of the eye that changes shape to finely focus light onto the retina to form a clear image. The bulk of this tissue comprises specialized, differentiated fiber cells that have a hexagonal cross section and extend from the anterior to the posterior poles of the lens. These long and skinny cells are tightly opposed to neighboring cells and have complex interdigitations along the length of the cell. The specialized interlocking structures are required for normal biomechanical properties of the lens and have been extensively described using electron microscopy techniques. This protocol demonstrates the first method to preserve and immunostain singular as well as bundles of mouse lens fiber cells to allow the detailed localization of proteins within these complexly shaped cells. The representative data show staining of the peripheral, differentiating, mature, and nuclear fiber cells across all regions of the lens. This method can potentially be used on fiber cells isolated from lenses of other species.

Lens fiber cells are prepared from the lens cortex (differentiating fibers and mature fibers) and the nucleus, and the cells are stained with phalloidin for F-actin and WGA for the cell membrane. A mixture of bundles of cells or single lens fibers (Figure 3) are observed and imaged. From the lens cortex, two types of cells (Figure 3A) are found. Differentiating fiber cells in the lens periphery are straight, with very small protrusions along their short sides. A...

Watch this Scientific Journal Video about Advancing Lens Biomechanics Research Through a Novel Protocol for Imaging Complex Interdigitations and Protein Staining at JoVE.com

Author Spotlight: Advancing Lens Biomechanics Research Through a Novel Protocol for Imaging Complex Interdigitations and Protein Staining 

This protocol describes methods to prepare peripheral, mature, and nuclear eye lens fiber cells for immunofluorescence staining to study complex cell-to-cell interdigitations and the membrane architecture.

Preparation and Immunofluorescence Staining of Bundles and Single Fiber Cells from the Cortex and Nucleus of the Eye Lens

The lens is a transparent and ellipsoid organ in the anterior chamber of the eye that changes shape to finely focus light onto the retina to form a clear ...

preparation-immunofluorescence-staining-bundles-single-fiber-cells

Research

JoVE Journal

Developmental Biology

1.3K Views.  Indiana University. This protocol describes methods to prepare peripheral, mature, and nuclear eye lens fiber cells for immunofluorescence staining to study complex cell-to-cell interdigitations and the membrane architecture.

Video: Author Spotlight: Advancing Lens Biomechanics Research Through a Novel Protocol for Imaging Complex Interdigitations and Protein Staining 

Efficient Derivation of Retinal Pigment Epithelium Cells from Stem Cells

No cure has been discovered for age-related macular degeneration (AMD), the leading cause of vision loss in people over the age of 55. AMD is complex multifactorial disease with an unknown etiology, although it is largely thought to occur due to death or dysfunction of the retinal pigment epithelium (RPE), a monolayer of cells that underlies the retina and provides critical support for photoreceptors. RPE cell replacement strategies may hold great promise for providing therapeutic relief for a large subset of AMD patients, and RPE cells that strongly resemble primary human cells (hRPE) have been generated in multiple independent labs, including our own. In addition, the uses for iPS-RPE are not limited to cell-based therapies, but also have been used to model RPE diseases. These types of studies may not only elucidate the molecular bases of the diseases, but also serve as invaluable tools for developing and testing novel drugs. We present here an optimized protocol for directed differentiation of RPE from stem cells. Adding nicotinamide and either Activin A or IDE-1, a small molecule that mimics its effects, at specific time points, greatly enhances the yield of RPE cells. Using this technique we can derive large numbers of low passage RPE in as early as three months.

Stem cell-derived retinal pigment epithelium (RPE) cells may be used for multiple applications including cell-based therapies for retinal degeneration, disease modeling, and drug studies. Here we present a simple protocol for reproducibly deriving RPE from stem cells.

No cure has been discovered for age-related macular degeneration  (AMD), the leading cause of vision loss in people over the age of 55. AMD is complex ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Organ Culture and Whole Mount Immunofluorescence Staining of Mouse Wolffian Ducts

Tubal morphogenesis is a fundamental requirement for the development of most mammalian organs, including the male reproductive system. The epididymis, an integral part of the male reproductive tract, is responsible for sperm storage, maturation, and transport. The adult epididymis is a highly coiled tube that develops from a simple and straight embryonic precursor known as Wolffian duct (WD). Proper coiling of the epididymis is essential for male fertility, as sperm in the testis are unable to fertilize an oocyte. However, the mechanism responsible for epididymal development and coiling remains unclear, partially due to the lack of whole organ culture and imaging methods. In this study, we describe an in vitro culture system and whole mount immunofluorescence protocol to better visualize the process of WD coiling and development, which may also be applied to study other tubular organs.

We present a method for isolation and culture of the mouse Wolffian duct (WD). We also demonstrate a detailed procedure for whole mount immunostaining of cultured/freshly isolated WDs with fluorescently tagged antibodies. Together, these techniques enable the study of WD development, coiling, and differentiation.

Tubal morphogenesis is a fundamental requirement for the development of most mammalian organs, including the male reproductive system. The epididymis, an ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Identification of Skeletal Muscle Satellite Cells by Immunofluorescence with Pax7 and Laminin Antibodies

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, antibodies for specific cell markers are applied on tissue sections. In adult skeletal muscle, satellite cells (SCs) are stem cells that contribute to muscle repair and regeneration. Therefore, it is important to visualize and trace the satellite cell population under different physiological conditions. In resting skeletal muscle, SCs reside between the basal lamina and myofiber plasma membrane. A commonly used marker for identifying SCs on the myofibers or in cell culture is the paired box protein Pax7. In this article, an optimized Pax7 immunofluorescence protocol on skeletal muscle sections is presented that minimizes non-specific staining and background. Another antibody that recognizes a protein (laminin) of the basal lamina was also added to help identify SCs. Similar protocols can also be used to perform double or triple labeling with Pax7 and antibodies for additional proteins of interest.

The precise identification of satellite cells is essential for studying their functions under various physiological and pathological conditions. This article presents a protocol to identify satellite cells on adult skeletal muscle sections by immunofluorescence-based staining.

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, ...

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and chromosome dynamics. While the germline is an excellent model, the analysis is often two dimensional due to the time and labor required for three-dimensional analysis. Major readouts in such studies are the number/position of nuclei and protein distribution within the germline. Here, we present a method to perform automated analysis of the germline using confocal microscopy and computational approaches to determine the number and position of nuclei in each region of the germline. Our method also analyzes germline protein distribution that enables the three-dimensional examination of protein expression in different genetic backgrounds. Further, our study shows variations in cytoskeletal architecture in distinct regions of the germline that may accommodate specific spatial developmental requirements. Finally, our method enables automated counting of the sperm in the spermatheca of each germline. Taken together, our method enables rapid and reproducible phenotypic analysis of the C. elegans germline.

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

We present an automated method for three-dimensional reconstruction of the Caenorhabditis elegans germline. Our method determines the number and position of each nucleus within the germline and analyses germline protein distribution and cytoskeletal structure.

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and ...

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and simple protocol to detect protein expression during craniofacial morphogenesis/pathogenesis using mouse craniofacial tissues as examples. The protocol consists of preparation and cryosectioning of tissues, indirect immunofluorescence, image acquisition, and quantification. In addition, a method for preparation and cryosectioning of undecalcified hard tissues for immunostaining is described, using craniofacial tissues and long bones as examples. Those methods are key to determine the protein expression and morphological/anatomical changes in various tissues during craniofacial morphogenesis/pathogenesis. They are also applicable to other tissues with appropriate modifications. Knowledge of the histology and high quality of sections are critical to draw scientific conclusions from experimental outcomes. Potential limitations of this methodology include but are not limited to specificity of antibodies and difficulties of quantification, which are also discussed here.

Here, we present a detailed protocol to detect and quantify protein levels during craniofacial morphogenesis/pathogenesis by immunostaining using mouse craniofacial tissues as examples. In addition, we describe a method for preparation and cryosectioning of undecalcified hard tissues from young mice for immunostaining.

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and ...

In Vitro Generation of Somite Derivatives from Human Induced Pluripotent Stem Cells

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give rise to multiple cell types, including the myotome (MYO), sclerotome (SCL), dermatome (D), and syndetome (SYN), which in turn develop into skeletal muscle, axial skeleton, dorsal dermis, and axial tendon/ligament, respectively. Therefore, the generation of SMs and their derivatives from human induced pluripotent stem cells (iPSCs) is critical to obtain pluripotent stem cells (PSCs) for application in regenerative medicine and for disease research in the field of orthopedic surgery. Although the induction protocols for MYO and SCL from PSCs have been previously reported by several researchers, no study has yet demonstrated the induction of SYN and D from iPSCs. Therefore, efficient induction of fully competent SMs remains a major challenge. Here, we recapitulate human SM patterning with human iPSCs in vitro by mimicking the signaling environment during chick/mouse SM development, and report on methods of systematic induction of SM derivatives (MYO, SCL, D, and SYN) from human iPSCs under chemically defined conditions through the presomitic mesoderm (PSM) and SM states. Knowledge regarding chick/mouse&#160;SM development was successfully applied to the induction of SMs with human iPSCs. This method could be a novel tool for studying human somitogenesis and patterning without the use of embryos and for cell-based therapy and disease modeling.

We present here a protocol for the differentiation of human induced pluripotent stem cells into each somite derivative (myotome, sclerotome, dermatome, and syndetome) in chemically defined conditions, which has applications in future disease modeling and cell-based therapies in orthopedic surgery.

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give ...

Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs during early development are technically challenging due to the limiting cell number. Moreover, many approaches require a miRNA of interest to be defined in assays such as northern blotting, microarray, and qPCR. Therefore, the expression of many miRNAs and their isoforms have not been studied during early development. Here, we demonstrate a protocol for small RNA sequencing of sorted cells from early mouse embryos to enable relatively unbiased profiling of miRNAs in early populations of neural crest cells. We overcome the challenges of low cell input and size selection during library preparation using amplification and gel-based purification. We identify embryonic age as a variable accounting for variation between replicates and stage-matched mouse embryos must be used to accurately profile miRNAs in biological replicates. Our results suggest that this method can be broadly applied to profile the expression of miRNAs from other lineages of cells. In summary, this protocol can be used to study how miRNAs regulate developmental programs in different cell lineages of the early mouse embryo.

We describe a technique for profiling microRNAs in early mouse embryos. This protocol overcomes the challenge of low cell input and small RNA enrichment. This assay can be used to analyze changes in miRNA expression over time in different cell lineages of the early mouse embryo.

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs ...