This work was made possible through Victorian State Government Operational Infrastructure Support and Australian Government NHMRC IRIISS and supported by funding from National Health and Medical Research Council (ALW #1120300) and Australian Research Council (KJH #FT190100265). The authors would like to acknowledge the technical support of the Monash Animal Research Platform, Monash Histology Platform and Monash Micro Imaging facility.

Ovaries were collected from female C57BL6J mice. All animal procedures and experiments were performed in accordance with the NHMRC Australian Code of Practice for the Care and Use of Animals and approved by the Monash Animal Research Platform Animal Ethics Committee.
NOTE: A chemotherapy agent shown to deplete primordial follicle oocytes, as determined using stereology11 and direct counts12,13 was used in this report to compare the two counting methods in the same animal. Female, 8-week-old (young adult) mice were weighed prior to a single intraperitoneal injection of 75 mg/kg/bodyweight of cyclophosphamide, or saline vehicle control (n=5/group). This dose has been shown to cause an approximate 50% depletion of primordial follicles, but not reported to cause morbidity or mortality in mice14. Ovaries were harvested 48 hours after treatment. One ovary from each animal was fixed in 10% (v/v) neutral buffered formalin solution for 24 hours, and the other fixed in Bouin&#8217;s solution for 24 hours. Tissue was then embedded in either glycolmethacrylate resin and serially sectioned at 20 &#181;m, or in paraffin and serially sectioned at 5 &#181;m. All tissues were stained with periodic acid Schiff and haematoxylin.
1. Histological preparation: fixation, processing, embedding and sectioning mouse ovaries
<ol>
	<li>Dissect mouse ovaries by trimming the oviduct and all surrounding fat, without damaging or cutting the ovary itself. If necessary, use a dissecting microscope and fine blade for this step (Figure 1A).</li>
	<li>Fix tissues immediately by placing into a small labelled tube containing either Bouin&#8217;s fixative (stereology), or formalin fixative (direct counts) for 24 hours (Figure 1B), before transferring tissues into 70% ethanol. 
	NOTE: Follicle morphology is conserved best within Bouin&#8217;s fixed ovarian tissue, embedded into glycolmethacrylate resin (Figure 2).</li>
	<li>Process whole fixed ovaries and then embed in glycolmethacrylate resin for stereology (Supplementary File 1), or paraffin wax for direct counts using a standard histological protocol. 
	CAUTION: Resin is toxic, so ensure all tissue processing steps are performed in a fume hood and gloves are worn at all times.</li>
	<li>Use a specialized resin methacrylate microtome (Figure 1C) fitted with a glass knife (Figure 1D) to exhaustively cut thick glycolmethacrylate resin sections (e.g., 20 &#181;m). Collect the sections at a regular interval (e.g., every 3rd section) onto glass microscope slides for stereology.</li>
	<li>Use a standard microtome to cut thin paraffin sections (e.g., 4-6 &#181;m). Collect tissue sections at a regular interval (e.g., every 9th section) onto a glass microscope slide for direct follicle counts.</li>
	<li>Stain the slides with periodic acid Schiff and haematoxylin (Supplementary File 2).</li>
	<li>Coverslip with standard DPX for paraffin sections, or thick DPX for glycolmethacrylate resin sections (Figure 1E). 
	CAUTION: Glycolmethacrylate resin DPX is hazardous, so perform this step in to fume hood. 
	NOTE: Glycolmethacrylate resin DPX is extremely viscous. The glass coverslip must be adhered firmly by pressing it down with a spatula to ensure the DPX is evenly and thinly dispersed, and there are no air bubbles present under the coverslip (Figure 1F).</li>
</ol>
2. Stereological estimation of primordial follicle number using the optical fractionator
<ol>
	<li>Turn on the computer, the multi-control unit, the camera and the light source within the stereology setup and set the microscope objective to a low magnification (e.g., 10x).</li>
	<li>Open the stereology software.</li>
	<li>Put the first slide securely on the microscope stage.</li>
	<li>Adjust the light exposure by checking Automatic under Exposure in the Camera Settings panel (Supplementary Figure 1A). Alternatively, manually adjust the light exposure.</li>
	<li>Use the joystick to locate the first tissue sample and bring the sample into focus.</li>
	<li>Adjust the white balance, either by clicking on More Settings (located bottom right of the Camera Settings panel), and then click White Balance and click on Automatic (Supplementary Figure 1B). Alternatively, click the White Balance button adjacent to the More Settings button (or Select Area in More Settings), to set the white balance manually by selecting a white area on the section.</li>
	<li>Go to the Probes drop down menu and click on Optical Fractionator Workflow. Then click Start New Project and click OK.
	<ol>
		<li>If an existing sampling configuration has been saved, under Sampling Parameters click Yes | &#8230; and select the desired sampling configuration.</li>
		<li>If not, click No and manually enter the serial section information (Supplementary Figure 1C) and define the probe configuration at step 2.13.</li>
	</ol>
	</li>
	<li>Click on Next, set the microscope to Low Magnification and choose 10x magnification from the dropdown menu.</li>
	<li>Click on Next, and then trace around the entire ovarian section &#8211; start by left clicking around the section and at the end, right click and choose Close Contour.</li>
	<li>Click on Next, set the microscope to High Magnification and choose 100x Oil magnification from the dropdown menu.</li>
	<li>Place a drop of oil on the tissue section on the slide and move the microscope objective to 100x magnification.</li>
	<li>Adjust the light exposure (as in step 2.4) and click Next.</li>
	<li>Set up the Sampling Parameters to define the probe configuration. Here, the counting frame was set to 47.5 &#181;m x 47.5 &#181;m (2,256.25 &#181;m2) and the step length was set to 100 &#181;m x 100 &#181;m (10,000 &#181;m2) (Supplementary Figure 2). Once the sampling parameters are established, save the template and re-open during subsequent analysis sessions at step 2.7.</li>
	<li>Click on Start Counting (Supplementary Figure 1D). Focus to the top of the sample, click OK and begin counting.</li>
	<li>Use the focusing knob to move through the 10 &#181;m sampling depth and count any primordial follicles whose oocyte nucleus comes into focus. Click Next to move to the next area.
	<ol>
		<li>Classify follicles as a primordial if the oocyte is surrounded by squamous (flattened) granulosa cells, but no cuboidal granulosa cells (Figure 2A). 
		NOTE: Primordial follicles are distinct from intermediate/transitional follicles, which comprise a combination of cuboidal and squamous granulosa cells (Figure 2B), and primary follicles, which are surrounded predominantly by cuboidal granulosa cells (Figure 2C). These follicle classes should be quantified separately.</li>
		<li>Only count follicles in which the oocyte nucleus is visible. The oocyte nucleus must appear within the counting frame or be touching the green inclusion lines of the counting frame (Supplementary Figure 1E,F).</li>
		<li>If the oocyte nucleus falls outside the counting frame (Supplementary Figure 1G) or touches the red exclusion lines of the counting frame, do not count this follicle.</li>
		<li>When assessing primordial follicle depletion in response to an exogenous chemical (e.g., chemotherapy), ensure all follicles counted are healthy and thus have normal morphology (Figure 2). Count any abnormal or atretic follicles separately. Often, follicle death is induced by insults such as chemotherapy, and quantification of the atretic follicles should be obtained separately in order to distinguish between healthy and atretic follicles, as only healthy follicles comprise the ovarian reserve15.</li>
	</ol>
	</li>
	<li>Once counting is complete on that section, do one of the following:
	<ol>
		<li>Click Add New Section, and then return to step 2.3 to set up the next section for counting.</li>
		<li>Click I&#8217;ve Finished Counting to end the session. Return the objective to 10x, exit the stereology software and turn off the light source, camera, multi-control unit and computer.</li>
	</ol>
	</li>
	<li>Obtain the sum raw follicle number (Q) from each tissue section sampled from the entire ovary, then using a spreadsheet, use the equation below to obtain the final value from each replicate animal (N)4. 
	<img alt="Equation 1" src="/files/ftp_upload/61782/61782equ01.jpg" />, where: 
	N = Total estimated number of follicles within the ovary. 
	Q = Raw primordial follicle count. 
	f1 = Sampling interval. Every 3rd section was sampled thus <img alt="Equation 2" src="/files/ftp_upload/61782/61782equ02.jpg" />. 
	f2 = Relationship between the counting frame (sample area) and stepper, calculated as <img alt="Equation 3" src="/files/ftp_upload/61782/61782equ03.jpg" />. Since the sample area was 2256 &#181;m2 (47.5 &#181;m x 47.5 &#181;m) and the stepper area was 10000 &#181;m2 (100 &#181;m x 100 &#181;m), <img alt="Equation 4" src="/files/ftp_upload/61782/61782equ04.jpg" />. 
	f3 = Fraction of ovarian section sampled. Since 10 &#181;m of the 20 &#181;m section was sampled, <img alt="Equation 5" src="/files/ftp_upload/61782/61782equ05.jpg" />. 
	Therefore, <img alt="Equation 6" src="/files/ftp_upload/61782/61782equ06.jpg" />. 
	NOTE: This protocol describes how to apply these principles of stereological analyses using a widely cited stereology software (Table of Materials); however, other stereological software is available. The principles applied during stereological analyses of ovarian follicles are the same, regardless of the software used to set up the parameters. Stereology is most accurate when 100 or more objects are counted in an adult mouse ovary4, as this gives a coefficient of error for the estimate below 10%16. The sampling parameters outlined here have been optimized to ensure a minimum of approximately 100 objects (i.e., primordial, transitional and primary follicles) can be counted in control adult wild-type C57BL6J ovaries. A pilot study can be conducted including a small sample size to establish the optimal sampling parameters, such as the interval and number of sections to be analyzed and the number of optical dissectors within the sampled sections17.</li>
</ol>
3. Estimation of primordial follicle number by direct ovarian follicle counts
<ol>
	<li>Place the microscope slide securely under a standard light microscope and perform direct counts to obtain raw primordial follicle number.
	<ol>
		<li>Classify follicles as a primordial if the oocyte is clearly visible and is surrounded by squamous (flattened) granulosa cells, but no cuboidal granulosa cells (Figure 2D). 
		NOTE: Primordial follicles are distinct from intermediate/transitional follicles, which comprise a combination of cuboidal and squamous granulosa cells (Figure 2E), and primary follicles, which are surrounded predominantly by cuboidal granulosa cells (Figure 2F). These follicle classes should be quantified separately.</li>
		<li>Ensure all follicles counted are healthy and thus have normal morphology (Figure 2). Count any abnormal or atretic follicles separately, as only healthy follicles comprise the ovarian reserve.</li>
	</ol>
	</li>
	<li>Alternatively, take multiple, or stitched high-power (e.g., 20x) photomicrographs of the entire ovarian tissue section to perform counts by opening the image file(s). This can be done manually, or using an automated slide scanner.</li>
	<li>Obtain the sum raw follicle number (Q) from each tissue section sampled from the entire ovary at the predetermined interval. Multiply this number by the inverse of the sampling fraction to obtain the final value for each replicate animal (N), using the equation below. 
	<img alt="Equation 7" src="/files/ftp_upload/61782/61782equ07.jpg" />, where: 
	N = Total estimated number of follicles within the ovary. 
	Q = Raw follicle count (of each individual type, calculated separately). 
	f1 = Sampling interval. Every 9th section was sampled thus <img alt="Equation 8" src="/files/ftp_upload/61782/61782equ08.jpg" />. 
	Therefore, <img alt="Equation 9" src="/files/ftp_upload/61782/61782equ09.jpg" />.</li>
</ol>

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The authors have nothing to disclose.

This article provides a step-by-step protocol for the gold standard technique for enumerating mouse primordial follicles, stereology, and the more commonly employed method of direct follicle counting. Chemotherapy treatment was used to compare and contrast the results obtained from these two different methods within the left and right ovary from the same animal. Both methods revealed high inter-animal variability in primordial follicle numbers. A significant depletion of the ovarian reserve was recorded using stereology,...

The immature, meiotically-arrested oocytes stored within primordial follicles in the ovary are the storage unit for the female germline and comprise an individual&#8217;s lifetime ovarian reserve. Primordial follicle numbers decline naturally with age1, or alternatively, can be prematurely depleted following exposure to exogenous chemicals, including some pharmaceuticals and environmental toxicants in air, food and water2. Given that the primordial follicle number is finite, the quantity and quality of follicles present within the ovary largely determine female fertility and offspring health. Thus, accurate quantification of primordial follicle number in women is essential for evaluating the off-target impacts of exogenous insults on the ovarian reserve.
In women, analysis of the whole ovary is generally not possible, thus non-invasive surrogate measures of the ovarian reserve must be utilized in a clinical setting. Anti-M&#971;llerian hormone (AMH) is the most widely used surrogate biomarker clinically3. Serum AMH levels are often measured in women of advanced maternal age, or before and after cancer treatment, such as chemotherapy. However, AMH is produced by growing follicles and not by primordial follicles, and thus, serum levels do not inform on absolute primordial follicle number.
With the absence of methods to accurately determine primordial follicle number in women in situ, counting ovarian follicles in small animal models, such as rodents, remains an essential research tool to assess the degree by which exogenous insults impact on primordial follicles and thus, fertility. Unfortunately though, reports throughout the literature of primordial follicle numbers in rodent models are highly variable4. A major reason for this is widely reported technical differences in the counting method employed. Predominately, there are two different techniques described in the literature for enumerating primordial follicles in mice. These include stereology, which employs the fractionator optical dissector method, and direct follicle counts.
Stereology is widely regarded the gold standard as it uses systematic uniform random sampling5, making it the most accurate method of quantifying primordial follicle number in whole mouse, or rat ovaries4,6,7. Stereology is unbiased, as it accounts for the three-dimensional structure of the object of interest8. Using an optical dissector/fractionator method, three levels of sampling are applied to quantify primordial follicles using thick tissue sections (e.g., 20 &#181;m) within a known fraction of the total mouse ovary. Firstly, the sampling interval is chosen (e.g., every 3rd section) at a random start (sampling fraction 1, f1)4. Sections are then sampled in a systematic, uniform manner from this point through the whole ovary. Then, an unbiased counting frame is superimposed over the ovarian section and progressively moved along a defined, randomized counting grid (sampling fraction 2, f2)8. Lastly, a known fraction of the section thickness is optically sampled (e.g., 10 &#181;m) and follicles within this area are counted (sampling fraction 3, f3)4. The raw follicle number is multiplied by the inverse of these sampling fractions to obtain the final value. This method requires expert training and equipment, including a microscope with a motorized stage driven by stereological software. Tissues should be preserved in a specialized Bouin&#8217;s fixative, and embedded in glycolmethacrylate resin to allow for thick tissue sections to be cut using a glycolmethacrylate resin microtome with a glass knife. This method is designed to account for tissue shrinkage and deformation, to best preserve the three-dimensional morphological structure of the ovary and follicles9.
Direct follicle counting is the most frequently used method for counting follicles10. More common fixatives (i.e., formalin) can be used, followed by paraffin wax embedding and exhaustive serial sectioning using a standard microtome at a thickness of between 4-6 &#181;m. Follicles are systematically counted in the entire tissue section at a defined interval, and then multiplied by the inverse of the sampling interval to obtain the total follicle estimate. This method is quick, easy, can be performed using archived tissues, and prepared using standard histological techniques. It requires only a light microscope with standard imaging capabilities. However, despite these advantages, direct follicle counting lacks the accuracy and strict counting parameters of stereology, making it more prone to investigator bias. Additionally, tissues may undergo shrinkage and deformation during processing, disrupting the integrity and morphology of the ovary and thus making follicle classification and quantification difficult.
The aim of this article is to describe two commonly-used methods to quantitatively assess primordial follicle number in mouse ovaries: stereology and direct follicle counting. We will provide detailed protocols for these two methods and highlight some of their strengths and weaknesses, in order to enhance reproducibility in our field and allow researchers to make an informed decision of the most appropriate counting method for their studies.

<table>
	<tbody>
		<tr>
			<td>1-Butanol (HPLC)</td>
			<td>Fisher Chemical</td>
			<td>#A383-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Acid alcohol</td>
			<td>Amber Scientific</td>
			<td>#ACDL</td>
			<td></td>
		</tr>
		<tr>
			<td>Bouin&#8217;s fixative</td>
			<td>Sigma-Aldrich</td>
			<td>#HT10132</td>
			<td>Picric acid 0.9% (w/v), formaldehyde 9% (v/v), acetic acid 5% (w/v)</td>
		</tr>
		<tr>
			<td>Cyclophosphamide</td>
			<td>Sigma-Aldrich</td>
			<td>#C0768-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Dibutylphthalate Polystyrene Xylene (DPX)</td>
			<td>Sigma-Aldrich</td>
			<td>#06522</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Amber Scientific</td>
			<td>#ETH</td>
			<td>Ethanol 100%</td>
		</tr>
		<tr>
			<td>Micro Feather opthalmic scalpel with aluminium handle</td>
			<td>Designs for Vision</td>
			<td>#FEA-745-SR</td>
			<td>Feather blade for dissections (seen in Figure 1)</td>
		</tr>
		<tr>
			<td>Formalin fixative</td>
			<td>Australian Biostain</td>
			<td>#ANBFC</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslip</td>
			<td>Thermo Scientific</td>
			<td>#MENCS22501GP</td>
			<td>22 mm x 50 mm</td>
		</tr>
		<tr>
			<td>Glycomethacrylate resin RM2165 microtome</td>
			<td>Leica Microsystems</td>
			<td>#RM2165</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycolmethacrylate DPX</td>
			<td>*made in house</td>
			<td></td>
			<td>*Mix 1.5 L Xylene; 800 g polystyrene pellets; 100mL Dibutyl phthalate for 3 weeks</td>
		</tr>
		<tr>
			<td>Histolene</td>
			<td>Trajan</td>
			<td>#11031</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayer&#8217;s haematoxylin</td>
			<td>Amber Scientific</td>
			<td>#MH</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus BX50 microscope</td>
			<td>Olympus</td>
			<td>#BX50</td>
			<td>Brightfield microscope fitted with 10x dry &#38; 100x oil immersion objective (numerical aperture 1.3)</td>
		</tr>
		<tr>
			<td>Olympus immersion oil type-F</td>
			<td>Olympus</td>
			<td>#IMMOIL-F30CC</td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus TH4-200 light source</td>
			<td>Olympus</td>
			<td>#TH4-200</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin wax</td>
			<td>Sigma-Aldrich</td>
			<td>#03987</td>
			<td></td>
		</tr>
		<tr>
			<td>Periodic acid</td>
			<td>Trajan</td>
			<td>#PERI1%</td>
			<td>Periodic acid 1%</td>
		</tr>
		<tr>
			<td>Rotary Microtome CUT 4060</td>
			<td>MicroTec</td>
			<td>#4060R/F</td>
			<td>Used to cut paraffin sections</td>
		</tr>
		<tr>
			<td>Schiff&#8217;s reagent</td>
			<td>Trajan</td>
			<td>#SCHF</td>
			<td></td>
		</tr>
		<tr>
			<td>Scott&#39;s tap water</td>
			<td>Amber Scientific</td>
			<td>#SCOT</td>
			<td>Potassium carbonate, magnesium sulphate, water</td>
		</tr>
		<tr>
			<td>StereoInvestigator Stereological System</td>
			<td>MBF Bioscience</td>
			<td></td>
			<td>Includes StereoInvestigator software, multi-control unit, automatic stage and joystick</td>
		</tr>
		<tr>
			<td>Superfrost microscope slides</td>
			<td>Thermo Scientific</td>
			<td>#MENSF41296SP</td>
			<td>1 mm, 72 pcs</td>
		</tr>
		<tr>
			<td>Technovit 7100 Plastic embedding system</td>
			<td>Emgrid Australia</td>
			<td>#64709003</td>
			<td>500 mL/5 x 1 g/40 mL</td>
		</tr>
		<tr>
			<td>Technovit 3040 yellow</td>
			<td>Emgrid Australia</td>
			<td>#64708805</td>
			<td>100 g/80 mL</td>
		</tr>
	</tbody>
</table>

accurate follicle enumeration in adult mouse ovaries

Sexually reproducing female mammals are born with their entire lifetime supply of oocytes. Immature, quiescent oocytes are found within primordial follicles, the storage unit of the female germline. They are non-renewable, thus their number at birth and subsequent rate of loss largely dictates the female fertile lifespan. Accurate quantification of primordial follicle numbers in women and animals is essential for determining the impact of medicines and toxicants on the ovarian reserve. It is also necessary for evaluating the need for, and success of, existing and emerging fertility preservation techniques. Currently, no methods exist to accurately measure the number of primordial follicles comprising the ovarian reserve in women. Furthermore, obtaining ovarian tissue from large animals or endangered species for experimentation is often not feasible. Thus, mice have become an essential model for such studies, and the ability to evaluate primordial follicle numbers in whole mouse ovaries is a critical tool. However, reports of absolute follicle numbers in mouse ovaries in the literature are highly variable, making it difficult to compare and/or replicate data. This is due to a number of factors including strain, age, treatment groups, as well as technical differences in the methods of counting employed. In this article, we provide a step-by-step instructional guide for preparing histological sections and counting primordial follicles in mouse ovaries using two different methods: [1] stereology, which relies on the fractionator/optical dissector technique; and [2] the direct count technique. Some of the key advantages and drawbacks of each method will be highlighted so that reproducibility can be improved in the field and to enable researchers to select the most appropriate method for their studies.

A well-characterized model of follicle depletion was used, whereby young adult female mice were administered a single dose of cyclophosphamide chemotherapy, or saline vehicle control (n=5/group) and both ovaries were harvested from each animal after 48 hours. One ovary per animal was prepared as described in Step 1 for each of the two methods: stereology or direct counts. The left and right ovary from each animal was randomly assigned to each group. These data show that when using stereology, a significant depletion of m...

Watch this Scientific Journal Video about Accurate Follicle Enumeration in Adult Mouse Ovaries at JoVE.com

Accurate Follicle Enumeration in Adult Mouse Ovaries

Here, we describe, compare, and contrast two different techniques for accurate follicle counting of fixed mouse ovarian tissues.

Sexually reproducing female mammals are born with their entire lifetime supply of oocytes. Immature, quiescent oocytes are found within primordial ...

accurate-follicle-enumeration-in-adult-mouse-ovaries

Research

JoVE Journal

Developmental Biology

8.0K Views.  Monash University. Here, we describe, compare, and contrast two different techniques for accurate follicle counting of fixed mouse ovarian tissues.

Video: Accurate Follicle Enumeration in Adult Mouse Ovaries

Culture and Co-Culture of Mouse Ovaries and Ovarian Follicles

The mammalian ovary is composed of ovarian follicles, each follicle consisting of a single oocyte surrounded by somatic granulosa cells, enclosed together within a basement membrane. A finite pool of follicles is laid down during embryonic development, when oocytes in meiotic arrest form a close association with flattened granulosa cells, forming primordial follicles. By or shortly after birth, mammalian ovaries contain their lifetime&#8217;s supply of primordial follicles, from which point onwards there is a steady release of follicles into the growing follicular pool.
The ovary is particularly amenable to development in vitro, with follicles growing in a highly physiological manner in culture. This work describes the culture of whole neonatal ovaries containing primordial follicles, and the culture of individual ovarian follicles, a method which can support the development of follicles from an immature through to the preovulatory stage, after which their oocytes are able to undergo fertilization in vitro. The work outlined here uses culture systems to determine how the ovary is affected by exposure to external compounds. We also describe a co-culture system, which allows investigation of the interactions that occur between growing follicles and the non-growing pool of primordial follicles.

This protocol describes the primary culture/co-culture of mouse ovarian tissue, using ovaries from neonatal mice and individual ovarian follicles from prepubertal mice. The culture techniques support development in a highly physiological manner, allowing investigation of the effect of extrinsic agents on the ovary, and of interactions between ovarian follicles.

The mammalian ovary is composed of ovarian follicles, each follicle consisting of a single oocyte surrounded by somatic granulosa cells, enclosed together ...

Isolating Hair Follicle Stem Cells and Epidermal Keratinocytes from Dorsal Mouse Skin

The hair follicle (HF) is an ideal system for studying the biology and regulation of adult stem cells (SCs). This dynamic mini organ is replenished by distinct pools of SCs, which are located in the permanent portion of the HF, a region known as the bulge. These multipotent bulge SCs were initially identified as slow cycling label retaining cells; however, their isolation has been made feasible after identification of specific cell markers, such as CD34 and keratin 15 (K15). Here, we describe a robust method for isolating bulge SCs and epidermal keratinocytes from mouse HFs utilizing fluorescence activated cell-sorting (FACS) technology. Isolated hair follicle SCs (HFSCs) can be utilized in various in vivo grafting models and are a valuable in vitro model for studying the mechanisms that govern multipotency, quiescence and activation.

An ideal model for studying adult stem cell biology is the mouse hair follicle. Here we present a protocol for isolating different populations of hair follicles stem cells and epidermal keratinocytes, employing enzymatic digestion of mouse dorsal skin followed by FACS analysis.

The hair follicle (HF) is an ideal system for studying the biology and regulation of adult stem cells (SCs). This dynamic mini organ is replenished by ...

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 ...

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

Research in the field of mammalian reproductive biology often involves evaluating the overall health of ovaries and testes. Specifically, in females, ovarian fitness is often assessed by visualizing and quantifying follicles and oocytes. Because the ovary is an opaque three-dimensional tissue, traditional approaches require laboriously slicing the tissue into numerous serial sections in order to visualize cells throughout the entire organ. Furthermore, because quantification by this method typically entails scoring only a subset of the sections separated by the approximate diameter of an oocyte, it is prone to inaccuracy. Here, a protocol is described that instead utilizes whole organ tissue clearing and immunofluorescence staining of mouse ovaries to visualize follicles and oocytes. Compared to more traditional approaches, this protocol is advantageous for visualizing cells within the ovary for numerous reasons: 1) the ovary remains intact throughout sample preparation and processing; 2) small ovaries, which are difficult to section, can be examined with ease; 3) cellular quantification is more readily and accurately achieved; and 4) the whole organ imaged.

Here, we present a protocol to quantify follicles in cultured ovaries of young mice without serial sectioning. Using whole organ immunofluorescence and tissue clearing, physical sectioning is replaced with optical sectioning. This method of sample preparation and visualization maintains organ integrity and facilitates automated quantification of specific cells.

Research in the field of mammalian reproductive biology often involves evaluating the overall health of ovaries and testes. Specifically, in females, ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

Direct Lineage Reprogramming of Adult Mouse Fibroblast to Erythroid Progenitors

Erythroid cell commitment and differentiation proceed through activation of a lineage-restricted transcriptional network orchestrated by a group of cell fate determining and maturing factors. We previously set out to define the minimal set of factors necessary for instructing red blood cell development using direct lineage reprogramming of fibroblasts into induced erythroid progenitors/precursors (iEPs). We showed that overexpression of Gata1, Tal1, Lmo2, and c-Myc (GTLM) can rapidly convert murine and human fibroblasts directly to iEPs that resemble bona fide erythroid cells in terms of morphology, phenotype, and gene expression. We intend that iEPs will provide an invaluable tool to study erythropoiesis and cell fate regulation. Here we describe the stepwise process of converting murine tail tip fibroblasts into iEPs via transcription factor-driven direct lineage reprogramming (DLR). In this example, we perform the reprogramming in fibroblasts from erythroid lineage-tracing mice that express the yellow fluorescent protein (YFP) under the control of the erythropoietin receptor gene (EpoR) promoter, enabling visualization of erythroid cell fate induction upon reprogramming. Following this protocol, fibroblasts can be reprogrammed into iEPs within five to eight days. 
While improvements can still be made to the process, we show that GTLM-mediated reprogramming is a rapid and direct process, yielding cells with properties of bona fide erythroid progenitor and precursor cells.

Here we present our protocol for producing induced erythroid progenitors (iEPs) from mouse adult fibroblasts using transcription factor-driven direct lineage reprogramming (DLR).

Erythroid cell commitment and differentiation proceed through activation of a lineage-restricted transcriptional network orchestrated by a group of cell ...

Determining Bile Duct Density in the Mouse Liver

Mouse is broadly used as a model organism to study biliary diseases. To evaluate the development and function of the biliary system, various techniques are used, including serum chemistry, histological analysis, and immunostaining for specific markers. Although these techniques can provide important information about the biliary system, they often do not present a full picture of bile duct (BD) developmental defects across the whole liver. This is in part due to the robust ability of the mouse liver to drain the bile even in animals with significant impairment in biliary development. Here we present a simple method to calculate the average number of BDs associated with each portal vein (PV) in sections covering all lobes of mutant/transgenic mice. In this method, livers are mounted and sectioned in a stereotypic manner to facilitate comparison among various genotypes and experimental conditions. BDs are identified via light microscopy of cytokeratin-stained cholangiocytes, and then counted and divided by the total number of PVs present in liver section. As an example, we show how this method can clearly distinguish between wild-type mice and a mouse model of Alagille syndrome. The method presented here cannot substitute for techniques that visualize the three-dimensional structure of the biliary tree. However, it offers an easy and direct way to quantitatively assess BD development and the degree of ductular reaction formation in mice.

We present a rather simple and sensitive method for accurate quantification of bile duct density in the mouse liver. This method can aid in determining the effects of genetic and environmental modifiers and the effectiveness of potential therapies in mouse models of biliary diseases.

Mouse is broadly used as a model organism to study biliary diseases. To evaluate the development and function of the biliary system, various techniques ...

Adult Mouse Digit Amputation and Regeneration: A Simple Model to Investigate Mammalian Blastema Formation and Intramembranous Ossification

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic regeneration, which involves blastema formation and intramembranous ossification analyzed by fluorescence immunohistochemistry and sequential in-vivo microcomputed tomography (&#956;CT). Mammalian regeneration is restricted to amputations transecting the distal region of the terminal phalanx (P3); digits amputated at more proximal levels fail to regenerate and undergo fibrotic healing and scar formation. The regeneration response is mediated by the formation of a proliferative blastema, followed by bone regeneration via intramembranous ossification to restore the amputated skeletal length. P3 amputation is a preclinical model to investigate epimorphic regeneration in mammals, and is a powerful tool for the design of therapeutic strategies to replace fibrotic healing with a successful regenerative response. Our protocol uses fluorescence immunohistochemistry to 1) identify early-and-late blastema cell populations, 2) study revascularization in the context of regeneration, and 3) investigate intramembranous ossification without the need for complex bone stabilization devices. We also demonstrate the use of sequential in vivo &#956;CT to create high resolution images to examine morphological changes after amputation, as well as quantify volume and length changes in the same digit over the course of regeneration. We believe this protocol offers tremendous utility to investigate both epimorphic and tissue regenerative responses in mammals.

Here, we present a protocol of adult mouse terminal phalanx amputation to investigate mammalian blastema formation and intramembranous ossification, analyzed by fluorescent immunohistochemistry and sequential in-vivo microcomputed tomography.

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic ...