PV is supported by the Inaugural Pedal the Cause Grant by the Alvin J. Siteman Cancer Center through The Foundation for Barnes-Jewish Hospital, Pilot Research Grant from Marsha Rivkin Center for Ovarian Cancer Research, Cancer Research Grant from Mary Kay Ash Foundation and V-Foundation. NR is supported by the NIH Cell and Molecular Biology training T32 grant to Washington University, St. Louis.

NOTE: The ovarian cancer cell line, OVCAR3, was used in these steps, but this protocol is broadly applicable to multiple other cell lines, including those derived from non-ovarian sources. A schematic of the protocol is shown in Figure 2.
1. Plating the cells
<ol>
	<li>Make poly-L-lysine coated coverslips.
	<ol>
		<li>Add autoclaved 12 mm diameter coverslips to a 50 mL conical tube with poly-L-lysine solution, and place on a rocker for 15 min.</li>
		<li>Aspirate the solution in a tissue culture hood. Wash the coverslips by adding sterile water, and place the tube containing the coverslips back on the rocker for 5 min. Repeat this wash step three times.</li>
		<li>Spread the coated coverslips on a sterile dish, and aspirate any remaining water. Let the coverslips dry in the tissue culture hood for 1 h or until no water droplets remain. Once dry, seal the dish with parafilm, and place at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Place one poly-L-lysine-coated coverslip in each well of a 24-well plate. The use of poly-lysine coverslips is critical to prevent the detachment of the cells from the coverslips during the pre-extraction step.</li>
	<li>Trypsinize OVCAR3 cells once they reach 70%-80% confluency.
	<ol>
		<li>To trypsinize a 10 cm culture dish that is 70%-80% confluent, aspirate the medium from the plate, and wash with 5-7 mL of PBS.</li>
		<li>Aspirate the PBS, add 1 mL of 0.25% trypsin, and place the dish in a 37 &#176;C incubator for 8-10 min, or until cells lift off from the bottom of the plate.</li>
		<li>Collect the cells with 5-10 mL of medium, and add to a conical tube.</li>
		<li>Count the cells manually with a hemocytometer using standard procedures.</li>
	</ol>
	</li>
	<li>Make a dilution of 25,000 cells/mL, and add 1 mL of the cells on poly-L-lysine coverslips placed in the wells of 24-well plates. For any other cell line, determine a number such that the cells are about 70%-80% confluent after three population doublings.</li>
	<li>Grow the cells in the culture medium under standard conditions.</li>
</ol>
2. Pulsing cells with IdU
<ol>
	<li>For the cells to properly spread onto the coverslips, one population doubling is enough. After one population doubling, pulse the cells with 10 &#181;M 5-iodo-2&#39;-deoxyuridine (IdU) (see Table of Materials on how to reconstitute IdU) for the two subsequent population doublings. 
	NOTE: We have tried different thymidine analogs, including bromodeoxyuridine (BrdU), 5-chloro-2&#8242;-deoxyuridine (CIdU), and IdU. Amongst the three analogs, IdU gives the best signal-to-noise ratio. Comparative images of cells pulsed with the three different analogs are shown in Figure 3. We recommend the use of two negative controls: (i) a no IdU pulsed sample, and (ii) a no primary antibody control. If the formation of ssDNA needs to be assessed due to a given treatment, we recommend adding the drug after the first round of IdU doubling.</li>
	<li>After pulsing with IdU for two population doublings, harvest the cells for imaging.
	<ol>
		<li>Replace the medium with ice-cold 0.5% PBSTx (PBS + 0.5% Triton X-100) on ice for 5 min. This pre-extraction step helps to release cytoplasmic and non-chromatin-bound proteins, leaving chromatin-bound proteins intact.&#160;Certain cell lines can easily peel off during pre-extraction. In such a scenario one can use 0.5% CSK buffer (10 mM PIPES (pH 6.8),100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 1 mM EGTA and 0.5% Triton X-100).</li>
	</ol>
	</li>
</ol>
3. Fixation
<ol>
	<li>Aspirate PBSTx, and incubate for 15 min with 3% PFA (Table of Materials) at room temperature, followed by three to four washes with 1x PBS. The fixed cells can be kept at 4 &#176;C until further steps. 
	&#8203;CAUTION: PFA is highly toxic and carcinogenic. Avoid contact with skin, eyes, and mucous membranes. Perform all the steps with PFA in a fume hood, and dispose of the materials properly.</li>
</ol>
4. Permeabilization and blocking
<ol>
	<li>After fixation, permeabilize the cells using 0.5% PBSTx on ice for 5 min. Use enough volume to cover the entire coverslip (typically between 500 &#181;L and 1 mL).</li>
	<li>Wash the cells three to four times with 0.2% PBST (1X PBS + 0.2% Tween-20) at room temperature. One mL of PBST is enough for washing each well. Perform the washes back to back without any incubations.</li>
	<li>Aspirate the PBST, and block the samples using 5% BSA (Table of Materials) made in 1x PBS (blocking buffer) for 30 min at room temperature.</li>
</ol>
5. Immunostaining with the IdU antibody
<ol>
	<li>For immunostaining, prepare a humidified chamber (wet paper towel on a flat-bottom Tupperware). Cover the lid of the 24-well plate with parafilm, place in the humidified chamber, and lay down the coverslips on the plate lid.</li>
	<li>Prepare the anti-BrdU primary mouse antibody (Table of Materials) by diluting it 1:200 in the blocking buffer from step 4.2.The anti-BrdU antibody has previously been shown to detect IdU.</li>
	<li>Add 60 &#181;L of IdU antibody dilution on the top of the coverslips. Incubate the coverslips for 1 h at 37 &#176;C.
	<ol>
		<li>Alternatively, use less antibody solution if the coverslips are flipped onto a drop (20-25 &#181;L) of 1:200 antibody dilution pipetted onto parafilm. This also decreases the likelihood of the solution drying out during the incubation.</li>
	</ol>
	</li>
	<li>After the 1 h incubation, aspirate the primary antibody. Return the coverslips back to a 24-well plate, and wash them four times with 0.2% PBST.</li>
	<li>For the secondary antibody, use the same humidified chamber described in step 5.1. Dilute anti-mouse conjugated secondary antibody (Table of Materials) in the blocking buffer (1:200). Add 60 &#181;L of secondary antibody to the coverslips, and incubate at room temperature in the dark for 1 h.This secondary antibody is light-sensitive.</li>
	<li>Aspirate the secondary antibody. Return the coverslips back to the 24-well plate, and wash four times with 0.2% PBST.</li>
	<li>Label microscope slides, and mount the coverslips on the slides with DAPI mounting medium (Table of Materials). Store slides in the dark at room temperature for 24 h. The 24 h incubation is recommended if the mounting medium needs to be cured or hardened. 
	NOTE: The slides can then be stored in 4 &#176;C before being imaged on a fluorescence microscope. A representative image is shown in Figure 4.</li>
</ol>
6. Automated quantification of IdU foci
NOTE: The power of this assay lies in the ability to automate the analysis for quick and efficient quantification. We present here an automated analysis pipeline that can be used to quantify IdU foci in a given image field. It is important that all the images within a given experiment are taken with the same exposure settings; otherwise, the quantification will not be reliable. It may also be valuable to include a non-stained control as a negative control, at least for the first time this experiment is run (Figure 5). The protocol below is specific for NIS General Analysis Software, but the same principles can be applied with other commercial software as well.
<ol>
	<li>In the View tab, go to Analysis Controls | Analysis Explorer. This opens a new side window called Analysis Explorer. Click on Create New | General Analysis 3. This opens the Analysis Explorer flowchart creation software.</li>
	<li>Go to the Sources tab, and drag the Channels command from the left menu into the working area. The two channels for the DAPI and IdU foci will automatically pop up as binary boxes (shown in Figure 5 as boxes with red outlines).</li>
	<li>Go to the Preprocessing tab, and drag the Local Contrast command from the left menu to the working area. Connect the Local Contrast with the IdU foci channel. From the Segmentation tab, drag the Threshold command (one for each channel).</li>
	<li>Connect each channel to a Threshold command. To eliminate any nuclei at the border of an image, go to the Binary Processing tab, drag the Touching Borders command, and connect to the DAPI threshold.</li>
	<li>Merge the data from the two channels by using the Aggregate Children command in the Measurement tab. Define the DAPI channel as Parent (A) and the foci channel as Child (B). In the dropdown menu in the Aggregate Children command, select the option child is inside the parent. This step helps to count the number of foci (child) per nucleus (parent).
	<ol>
		<li>To export the data in a tabular format, click on the Modify Columns command in the Data Management tab. In the dropdown menu from the Modify Columns command, select DAPI-ID (this helps to assign a unique number to each nucleus within a given image) and IdU count (this provides the number of IdU foci in each nucleus). To export the data in a CSV format, use the Table To CSV command in the Reference tab. The GA3 can now be saved with a descriptive title.</li>
	</ol>
	</li>
	<li>For the analysis, open an image in the software.
	<ol>
		<li>Open the Analysis Explorer tab as was done in step 6.1, and locate the newly made GA3. Double-click on the GA3 file, which opens a new window called GA3 Wizard where one can set the threshold for the DAPI and the IdU channels.</li>
		<li>Use the slider in the window to set the minimum and maximum threshold for each channel and, hence, define the signal. A good thresholding value is where the boundaries for each nucleus and IdU focus are clearly defined. Once satisfied with threshold values, retain these numbers for analyzing all the files in a given experiment.</li>
		<li>Click on the Run button to obtain the count of IdU foci/per nucleus. 
		NOTE: The software generates a table of foci per nucleus, which can thereafter be used for any graphing purposes (Figure 5).</li>
	</ol>
	</li>
</ol>

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As was mentioned in the protocol, it is valuable to include a few experimental controls to ensure that the assay is working. These include a no IdU treated sample as well as a no primary antibody treated sample. Both negative controls should yield cells that are stained by DAPI but contain no IdU signal.
Based on the experimental conditions and cell lines used, different antibody dilutions may be needed to obtain the best fluorescent signal. Too much signal may result in an inability to quanti...

Genomic DNA is frequently exposed to multiple assaults from various endogenous and exogenous sources1. The frequency of endogenous damage directly correlates with the levels of metabolic byproducts, such as reactive oxygen species or aldehydes, which are intrinsically higher in multiple cancer types, including ovarian cancers2,3. It is imperative that DNA damage is efficiently resolved; otherwise, it can foster genotoxic lesions and, consequently, mutagenesis. The ability of cells to repair genotoxic lesions is reliant on the functionality of error-free DNA repair pathways and the efficient regulation of cell cycle progression in response to DNA damage. Notably, many ovarian cancers bear functionally inactivating mutations in p53 and, thus, have a defective G1/S checkpoint, leading the cells to initiate DNA replication despite the presence of unrepaired genomic lesions4,5. The degree of DNA damage in ovarian cancers is further compounded by the observation that more than 50% of high-grade serous ovarian carcinoma (HGSOC) have defects in BRCA1- and BRCA2-mediated homologous recombination, the error-free DNA repair pathway, and around 20% have amplification in the gene CCNE1, which prematurely pushes G1 cells into the S-phase6. Together the high frequency of endogenous DNA damage, defective checkpoints, and malfunctioning repair pathways exponentially enhance the accumulation of genomic lesions in ovarian cancers. These lesions can serve as impediments to the progression of critical cellular processes such as DNA replication and transcription. As discussed below, such impediments catalyze the generation of single-stranded DNA (ssDNA) in cells.
The double helix of DNA is critical for safeguarding the genome from multiple mutagenic processes, such as spontaneous depurination and depyrimidination, the activity of cytosine deaminases, and oxidative DNA damage1,7. In contrast, ssDNA is highly vulnerable to these mutational events. Multiple processes in cells can result in the generation of ssDNA (Figure 1). These include the following:
(i) Stalling of the DNA replication machinery: This leads to an uncoupling of the DNA helicase and polymerase, leaving stretches of ssDNA8,9.
(ii) Stalling of the transcription machinery: Persistent stalling of RNA polymerase leads to the generation of three-stranded hybrid DNA/RNA structures called R-loops. R-loop formation exposes the displaced, non-transcribed DNA as a single strand10.
(iii) DNA end-resection: The initiation of homology-directed repair requires the generation of a 3&#39; ssDNA to catalyze the search for a homologous sequence11.
(iv) D-loop: Strand invasion during homologous recombination can result in the displacement of the non-template complementary strand, resulting in ssDNA12.
(v) Replication-coupled gaps: During DNA replication, lagging strand synthesis happens in a discontinuous fashion, whereby Okazaki fragments are first generated and then ligated. A delay or defect in processing the Okazaki fragments can also result in ssDNA formation. Finally, if the replication fork on a leading strand encounters a stalling lesion, DNA polymerase, and primase, PRIMPOL can reprime the synthesis downstream, leaving an ssDNA gap behind13,14.
Evidently, most of these events either happen when the DNA replication machinery faces genomic lesions or during replication-coupled repair, suggesting that higher DNA damage leads to increased levels of ssDNA. As many of these events are replication-associated, the formation of ssDNA is considered the marker of &#34;replication stress&#34; in cells15,16.
Here, we describe an assay that can be used to reliably quantify ssDNA in cells. The simplicity, reproducibility, and cost benefits of this approach make it amenable to be used for assessing the replication-stress response in cells. Emerging studies have revealed that the level of ssDNA can also be a predictor of responses to chemotherapy, such as inhibitors of PARP1/2 enzymes, ATR, and Wee1 kinase17,18,19,20,21. These inhibitors are being pursued in the treatment regimen of several HGSOCs22. Therefore, this assay can also be a useful tool to predict chemotherapeutic responses in ovarian cancer cells.

<table><tbody><tr><td>3% Paraformaldehyde (PFA)</td><td>Fisher Scientific</td><td>NC0179595</td><td>10 g sucrose + 100 mL 10X PBS + water to make volume to 925 mL. Add 75 mL 40% Methanol free PFA, mix, and make aliquots of 50 mL before storage&lt;br/&gt; Storage: Store in -20 &amp;deg;C</td></tr><tr><td>5-iodo-2&#039;-deoxyuridine (IdU)</td><td>Sigma Aldrich</td><td>I7125-5G</td><td>MW = 354.10 g/mol.For 10 mM stock: dissolve 3.541 mg IdU to 1 mL 1 N liquid ammonia&lt;br/&gt; Storage: Stored in -20 &amp;deg;C</td></tr><tr><td>Anti-BrdU antibody</td><td>BD Biosciences</td><td>347580</td><td>Storage: Store in 4 &amp;deg;C</td></tr><tr><td>Anti-mouse Alexa Fluor Plus 488 secondary antibody</td><td>Thermo Scientific</td><td>A32766</td><td>Light sensitive - keep in dark&lt;br/&gt; Storage: Store in 4 &amp;deg;C</td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma Aldrich</td><td>A7906-100G</td><td>Made by adding specific mass to volume of PBS&lt;br/&gt; Storage: Store in 4 &amp;deg;C</td></tr><tr><td>Circular Cover Glass&amp;nbsp;</td><td>Electron Microscopy Sciences</td><td>72230-01</td><td></td></tr><tr><td>NIS GA3 Software&amp;nbsp;</td><td>Nikon&amp;nbsp;</td><td>77010604</td><td></td></tr><tr><td>OVCAR3</td><td>ATCC</td><td>HTB-161</td><td>Growth Media: RPMI supplemented with L-glutamine, 0.01 mg/mL bovine insulin; fetal bovine serum to a final concentration of 20% and 1X Pen Strep&lt;br/&gt; Storage: Freezing Media: growth media + 5% DMSO and stored in -80 &amp;deg;C</td></tr><tr><td>Poly-L-Lysine solution</td><td>Sigma Aldrich</td><td>P4832-50ML</td><td>Storage: Store in 4 &amp;deg;C</td></tr><tr><td>ProLong Diamond Antifade Mountant with DAPI</td><td>Thermo Scientific</td><td>P36962</td><td>Storage: Store in 4 &amp;deg;C</td></tr><tr><td>Trypsin-EDTA, 0.25%</td><td>Genesee Scientific</td><td>25-510</td><td>Storage: Store in 4 &amp;deg;C</td></tr><tr><td>Water, sterile-filtered</td><td>Sigma Aldrich</td><td>W3500-6X500ML</td><td>Storage: Store in 4 &amp;deg;C</td></tr></tbody></table>

quantifying replication stress in ovarian cancer cells using single-stranded dna immunofluorescence

Replication stress is a hallmark of several ovarian cancers. Replication stress can emerge from multiple sources, including double-strand breaks, transcription-replication conflicts, or amplified oncogenes, inevitably resulting in the generation of single-stranded DNA (ssDNA). Quantifying ssDNA, therefore, presents an opportunity to assess the level of replication stress in different cell types and under various DNA-damaging conditions or treatments. Emerging evidence also suggests that ssDNA can be a predictor of responses to chemotherapeutic drugs that target DNA repair. Here, we describe a detailed immunofluorescence-based methodology to quantify ssDNA. This methodology involves labeling the genome with a thymidine analog, followed by the antibody-based detection of the analog at the chromatin under non-denaturing conditions. Stretches of ssDNA can be visualized as foci under a fluorescence microscope. The number and intensity of the foci directly co-relate with the level of ssDNA present in the nucleus. We also describe an automated pipeline to quantify the ssDNA signal. The method is rapid and reproducible. Furthermore, the simplicity of this methodology makes it amenable to high-throughput applications such as drug and genetic screens.

Representative images and the quantification of IdU foci from the nuclei derived from the untreated cells and cells treated with 0.5 mM hydroxyurea for 24 h are shown in Figure 4. Both nuclei are stained and identifiable in the DAPI channel. The analysis of these images consists of quantifying the number of foci in each nucleus. The number of foci is proportional to the degree of replication stress.
<img alt="Figure 1" cl...

Watch this Scientific Journal Video about Quantifying Replication Stress in Ovarian Cancer Cells Using Single-Stranded DNA Immunofluorescence at JoVE.com

Quantifying Replication Stress in Ovarian Cancer Cells Using Single-Stranded DNA Immunofluorescence

Here, we describe an immunofluorescence-based method to quantify the levels of single-stranded DNA in cells. This efficient and reproducible method can be utilized to examine replication stress, a common feature in several ovarian cancers. Additionally, this assay is compatible with an automated analysis pipeline, which further increases its efficiency.

Replication stress is a hallmark of several ovarian cancers. Replication stress can emerge from multiple sources, including double-strand breaks, ...

quantifying-replication-stress-ovarian-cancer-cells-using-single

Universidad de Cartagena UDEC

Research

JoVE Journal

Cancer Research

2.0K Views.  Washington University School of Medicine. Here, we describe an immunofluorescence-based method to quantify the levels of single-stranded DNA in cells. This efficient and reproducible method can be utilized to examine replication stress, a common feature in several ovarian cancers. Additionally, this assay is compatible with an automated analysis pipeline, which further increases its efficiency.

Video: Quantifying Replication Stress in Ovarian Cancer Cells Using Single-Stranded DNA Immunofluorescence

Immunofluorescence Imaging of DNA Damage and Repair Foci in Human Colon Cancer Cells

The purpose of the manuscript is to provide a step-by-step protocol for performing immunofluorescence microscopy to study the radiation-induced DNA damage response induced by neutron-gamma mixed-beam used in boron neutron capture therapy (BNCT). Specifically, the proposed methodology is applied for the detection of repair proteins activation which can be visualized as foci using antibodies specific to DNA double-strand breaks (DNA-DSBs). DNA repair foci were assessed by immunofluorescence in colon cancer cells (HCT-116) after irradiation with the neutron-mixed beam. DNA-DSBs are the most genotoxic lesions and are repaired in mammalian cells by two major pathways: non-homologous end-joining pathway (NHEJ) and homologous recombination repair (HRR). The frequencies of foci, immunochemically stained, for commonly used markers in radiobiology like &#947;-H2AX, 53BP1 are associated with DNA-DSB number and are considered as efficient and sensitive markers for monitoring the induction and repair of DNA-DSBs. It was established that &#947;-H2AX foci attract repair proteins, leading to a higher concentration of repair factors near a DSB. To monitor DNA damage at the cellular level, immunofluorescence analysis for the presence of DNA-PKcs representative repair protein foci from the NHEJ pathway and Rad52 from the HRR pathway was planned. We have developed and introduced a reliable immunofluorescence staining protocol for the detection of radiation-induced DNA damage response with antibodies specific for repair factors from NHEJ and HRR pathways and observed radiation-induced foci (RIF). The proposed methodology can be used for investigating repair protein that is highly activated in the case of neutron-mixed beam radiation, thereby indicating the dominance of the repair pathway.

Radiation-induced DNA damage response activated by neutron mixed-beam used in boron neutron capture therapy (BNCT) has not been fully established. This protocol provides a step-by-step procedure to detect radiation-induced foci (RIF) of repair proteins by immunofluorescence staining in human colon cancer cell lines after irradiation with the neutron-mixed beam.

The purpose of the manuscript is to provide a step-by-step protocol for performing immunofluorescence microscopy to study the radiation-induced DNA damage ...

Exploring X Chromosomal Aberrations in Ovarian Cells by Using Fluorescence In Situ Hybridization

Millions of people worldwide deal with issues concerning fertility. Reduced fertility, or even infertility, may be due to many different causes, including genetic disorders, of which chromosomal abnormalities are the most common. Fluorescence in situ hybridization (FISH) is a well-known and frequently used method to detect chromosomal aberrations in humans. FISH is mainly used for the analysis of chromosomal abnormalities in the spermatozoa of males with numerical or structural chromosomal aberrations. Furthermore, this technique is also frequently applied in females to detect X chromosomal aberrations that are known to cause ovarian dysgenesis. However, information on the X chromosomal content of ovarian cells from females with X chromosomal aberrations in lymphocytes and/or buccal cells is still lacking.
The aim of this study is to advance basic research regarding X chromosomal aberrations in females, by presenting two methods based on FISH to identify the X chromosomal content of ovarian cells. First, a method is described to determine the X chromosomal content of isolated ovarian cells (oocytes, granulosa cells, and stromal cells) in non-grafted ovarian cortex tissue from females with X chromosomal aberrations. The second method is directed at evaluating the effect of chromosomal aberrations on folliculogenesis by determining the X chromosomal content of ovarian cells of newly formed secondary and antral follicles in ovarian tissue, from females with X chromosomal aberrations after long-term grafting into immunocompromised mice. Both methods could be helpful in future research to gain insight into the reproductive potential of females with X chromosomal aberrations.

Exploring X Chromosomal Aberrations in Ovarian Cells by Using Fluorescence In Situ Hybridization

This article presents two methods based on fluorescence in situ hybridization to determine the X chromosomal content of ovarian cells in non-grafted and grafted ovarian cortex tissue from females with X chromosomal aberrations.

Millions of people worldwide deal with issues concerning fertility. Reduced fertility, or even infertility, may be due to many different causes, including ...

Genetics

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Biology

Visualizing DNA Damage Repair Proteins in Patient-Derived Ovarian Cancer Organoids via Immunofluorescence Assays

Immunofluorescence is one of the most widely used techniques to visualize target antigens with high sensitivity and specificity, allowing for the accurate identification and localization of proteins, glycans, and small molecules. While this technique is well-established in two-dimensional (2D) cell culture, less is known about its use in three-dimensional (3D) cell models. Ovarian cancer organoids are 3D tumor models that recapitulate tumor cell clonal heterogeneity, the tumor microenvironment, and cell-cell and cell-matrix interactions. Thus, they are superior to cell lines for the evaluation of drug sensitivity and functional biomarkers. Therefore, the ability to utilize immunofluorescence on primary ovarian cancer organoids is extremely beneficial in understanding the biology of this cancer. The current study describes the technique of immunofluorescence to detect DNA damage repair proteins in high-grade serous patient-derived ovarian cancer organoids (PDOs). After exposing the PDOs to ionizing radiation, immunofluorescence is performed on intact organoids to evaluate nuclear proteins as foci. Images are collected using z-stack imaging on confocal microscopy and analyzed using automated foci counting software. The described methods allow for the analysis of temporal and special recruitment of DNA damage repair proteins and colocalization of these proteins with cell-cycle markers.

Visualizing DNA Damage Repair Proteins in Patient-Derived Ovarian Cancer Organoids via Immunofluorescence Assays

The present protocol describes methods for evaluating DNA damage repair proteins in patient-derived ovarian cancer organoids. Included here are comprehensive plating and staining methods, as well as detailed, objective quantification procedures.

Immunofluorescence is one of the most widely used techniques to visualize target antigens with high sensitivity and specificity, allowing for the accurate ...

Identifying DNA Damage in Prophase-Arrested Oocytes of Mice

Detection of DNA Double-Stranded Breaks in Mouse Oocytes

Oocytes are amongst the biggest and most long-lived cells in the female body. They are formed in the ovaries during embryonic development and remain arrested at the prophase of meiosis I. The quiescent state may last for years until the oocytes receive a stimulus to grow and obtain the competency to resume meiosis. This protracted state of arrest makes them extremely susceptible to accumulating DNA-damaging insults, which affect the genetic integrity of the female gametes and, therefore, the genetic integrity of the future embryo. 
Consequently, the development of an accurate method to detect DNA damage, which is the first step for the establishment of DNA damage response mechanisms, is of vital importance. This paper describes a common protocol to test the presence and progress of DNA damage in prophase-arrested oocytes during a period of 20 h. Specifically, we dissect mouse ovaries, retrieve the cumulus-oocyte complexes (COCs), remove the cumulus cells from the COCs, and culture the oocytes in &#924;2 medium containing 3-isobutyl-1-methylxanthine to maintain the state of arrest. Thereafter, the oocytes are treated with the cytotoxic, antineoplasmic drug, etoposide, to engender double-strand breaks (DSBs). 
By using immunofluorescence and confocal microscopy, we detect and quantify the levels of the core protein &#947;H2AX, which is the phosphorylated form of the histone H2AX. H2AX becomes phosphorylated at the sites of DSBs after DNA damage. The inability to restore DNA integrity following DNA damage in oocytes can lead to infertility, birth defects, and increased rates of spontaneous abortions. Therefore, the understanding of DNA damage response mechanisms and, at the same time, the establishment of an intact method for studying these mechanisms are essential for reproductive biology research.

Author Spotlight: Understanding DNA Damage Response in Mammalian Oocytes and Preimplantation Embryos 

Maintaining oocyte genome integrity is necessary to ensure the genetic fidelity in the resulting embryo. Here, we present an accurate protocol for detecting DNA double-strand breaks in mammalian female germ cells.

Oocytes are amongst the biggest and most long-lived cells in the female body. They are formed in the ovaries during embryonic development and remain ...

Developmental Biology

Quantitation of Intra-peritoneal Ovarian Cancer Metastasis

Epithelial ovarian cancer (EOC) is the leading cause of death from gynecologic malignancy in the United States. Mortality is due to diagnosis of 75% of women with late stage disease, when metastasis is already present. EOC is characterized by diffuse and widely disseminated intra-peritoneal metastasis. Cells shed from the primary tumor anchor in the mesothelium that lines the peritoneal cavity as well as in the omentum, resulting in multi-focal metastasis, often in the presence of peritoneal ascites. Efforts in our laboratory are directed at a more detailed understanding of factors that regulate EOC metastatic success. However, quantifying metastatic tumor burden represents a significant technical challenge due to the large number, small size and broad distribution of lesions throughout the peritoneum. Herein we describe a method for analysis of EOC metastasis using cells labeled with red fluorescent protein (RFP) coupled with in vivo multispectral imaging. Following intra-peritoneal injection of RFP-labelled tumor cells, mice are imaged weekly until time of sacrifice. At this time, the peritoneal cavity is surgically exposed and organs are imaged in situ. Dissected organs are then placed on a labeled transparent template and imaged ex vivo. Removal of tissue auto-fluorescence during image processing using multispectral unmixing enables accurate quantitation of relative tumor burden. This method has utility in a variety of applications including therapeutic studies to evaluate compounds that may inhibit metastasis and thereby improve overall survival.

Ovarian cancer metastasis is characterized by numerous diffuse intra-peritoneal lesions, such that accurate visual quantitation of tumor burden is challenging. Herein we describe a method for in situ and ex vivo quantitation of metastatic tumor burden using red fluorescent protein (RFP)-labeled tumor cells and optical imaging.

Epithelial ovarian cancer (EOC) is the leading cause of death from gynecologic malignancy in the United States.  Mortality is due to diagnosis of 75% of ...

Medicine

Substructure Analyzer: A User-Friendly Workflow for Rapid Exploration and Accurate Analysis of Cellular Bodies in Fluorescence Microscopy Images

The last decade has been characterized by breakthroughs in fluorescence microscopy techniques illustrated by spatial resolution improvement but also in live-cell imaging and high-throughput microscopy techniques. This led to a constant increase in the amount and complexity of the microscopy data for a single experiment. Because manual analysis of microscopy data is very time consuming, subjective, and prohibits quantitative analyses, automation of bioimage analysis is becoming almost unavoidable. We built an informatics workflow called Substructure Analyzer to fully automate signal analysis in bioimages from fluorescent microscopy. This workflow is developed on the user-friendly open-source platform Icy and is completed by functionalities from ImageJ. It includes the pre-processing of images to improve the signal to noise ratio, the individual segmentation of cells (detection of cell boundaries) and the detection/quantification of cell bodies enriched in specific cell compartments. The main advantage of this workflow is to propose complex bio-imaging functionalities to users without image analysis expertise through a user-friendly interface. Moreover, it is highly modular and adapted to several issues from the characterization of nuclear/cytoplasmic translocation to the comparative analysis of different cell bodies in different cellular sub-structures. The functionality of this workflow is illustrated through the study of the Cajal (coiled) Bodies under oxidative stress (OS) conditions. Data from fluorescence microscopy show that their integrity in human cells is impacted a few hours after the induction of OS. This effect is characterized by a decrease of coilin nucleation into characteristic Cajal Bodies, associated with a nucleoplasmic redistribution of coilin into an increased number of smaller foci. The central role of coilin in the exchange between CB components and the surrounding nucleoplasm suggests that OS induced redistribution of coilin could affect the composition and the functionality of Cajal Bodies.

We present a freely available workflow built for rapid exploration and accurate analysis of cellular bodies in specific cell compartments in fluorescence microscopy images. This user-friendly workflow is designed on the open-source software Icy and also uses ImageJ functionalities. The pipeline is affordable without knowledge in image analysis.

The last decade has been characterized by breakthroughs in fluorescence microscopy techniques illustrated by spatial resolution improvement but also in ...

Bioengineering

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The specification of replication timing is thought to be a dynamic process regulated by tissue-specific and developmental cues that are responsive to epigenetic modifications. However, the mechanisms regulating where and when DNA replication initiates along chromosomes remains poorly understood. Homologous chromosomes usually replicate synchronously, however there are notable exceptions to this rule. For example, in female mammalian cells one of the two X chromosomes becomes late replicating through a process known as X inactivation1. Along with this delay in replication timing, estimated to be 2-3 hr, the majority of genes become transcriptionally silenced on one X chromosome. In addition, a discrete cis-acting locus, known as the X inactivation center, regulates this X inactivation process, including the induction of delayed replication timing on the entire inactive X chromosome. In addition, certain chromosome rearrangements found in cancer cells and in cells exposed to ionizing radiation display a significant delay in replication timing of &gt;3 hours that affects the entire chromosome2,3. Recent work from our lab indicates that disruption of discrete cis-acting autosomal loci result in an extremely late replicating phenotype that affects the entire chromosome4. Additional 'chromosome engineering' studies indicate that certain chromosome rearrangements affecting many different chromosomes result in this abnormal replication-timing phenotype, suggesting that all mammalian chromosomes contain discrete cis-acting loci that control proper replication timing of individual chromosomes5.
Here, we present a method for the quantitative analysis of chromosome replication timing combined with fluorescent in situ hybridization. This method allows for a direct comparison of replication timing between homologous chromosomes within the same cell, and was adapted from6. In addition, this method allows for the unambiguous identification of chromosomal rearrangements that correlate with changes in replication timing that affect the entire chromosome. This method has advantages over recently developed high throughput micro-array or sequencing protocols that cannot distinguish between homologous alleles present on rearranged and un-rearranged chromosomes. In addition, because the method described here evaluates single cells, it can detect changes in chromosome replication timing on chromosomal rearrangements that are present in only a fraction of the cells in a population.

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

A quantitative method for the analysis of chromosome replication timing is described. The method utilizes BrdU incorporation in combination with fluorescent in situ hybridization (FISH) to assess replication timing of mammalian chromosomes. This technique allows for the direct comparison of rearranged and un-rearranged chromosomes within the same cell.

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The ...

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA insults. Genotoxic agents can damage the DNA backbone, break it, or modify the chemical nature of individual bases. Following DNA insult, DNA damage response (DDR) pathways are activated and proteins involved in the repair are recruited. A plethora of factors are involved in sensing the type of damage and activating the appropriate repair response. Failure to correctly activate and recruit DDR factors can lead to genomic instability, which underlies many human pathologies including cancer. Studies of DDR proteins can provide insights into genotoxic drug response and cellular mechanisms of drug resistance.
There are two major ways of visualizing&#160;proteins in vivo: direct observation, by tagging the protein of interest with a fluorescent protein and following it by live imaging, or indirect immunofluorescence on fixed samples. While visualization of fluorescently tagged proteins allows precise monitoring over time, direct tagging in N- or C-terminus can interfere with the protein localization or function. Observation of proteins in their unmodified, endogenous version is preferred. When DNA repair proteins are recruited to the DNA insult, their concentration increases locally and they form groups, or &#8220;foci&#8221;, that can be visualized by indirect immunofluorescence using specific antibodies.
Although detection of protein foci does not provide a definitive proof of direct interaction, co-localization of proteins in cells indicates that they regroup to the site of damage and can inform of the sequence of events required for complex formation. Careful analysis of foci spatial overlap in cells expressing wild type or mutant versions of a protein can provide precious clues on functional domains important for DNA repair function. Last, co-localization of proteins indicates possible direct interactions that can be verified by co-immunoprecipitation in cells, or direct pulldown using purified proteins.

Following DNA damage, human cells activate essential repair pathways to restore the integrity of their genome. Here, we describe the method of indirect immunofluorescence as a means to detect DNA repair proteins, analyze their spatial and temporal recruitment, and help interrogate protein-protein interaction at the sites of DNA damage.

Mammalian cells are constantly exposed to chemicals, radiations, and naturally occurring metabolic by-products, which create specific types of DNA ...

Detection of G1 Phase Single-Stranded DNA Foci via RPA2 Immunofluorescent Staining

Visualizing Single-Stranded DNA Foci in the G1 Phase of the Cell Cycle

DNA has dedicated cellular repair pathways capable of coping with lesions that could arise from both endogenous and/or exogenous sources. DNA repair necessitates collaboration between numerous proteins, responsible for covering a broad range of tasks from recognizing and signaling the presence of a DNA lesion to physically repairing it. During this process, tracks of single-stranded DNA (ssDNA) are often created, which are eventually filled by DNA polymerases. The nature of these ssDNA tracks (in terms of both length and number), along with the polymerase recruited to fill these gaps, are repair pathway-specific. The visualization of these ssDNA tracks can help us understand the complicated dynamics of DNA repair mechanisms.
This protocol provides a detailed method for the preparation of G1 synchronized cells to measure ssDNA foci formation upon genotoxic stress. Using an easy-to-utilize immunofluorescence approach, we visualize ssDNA by staining for RPA2, a component of the heterotrimeric replication protein A complex (RPA). RPA2 binds to and stabilizes ssDNA intermediates that arise upon genotoxic stress or replication to control DNA repair and DNA damage checkpoint activation. 5-Ethynyl-2'-deoxyuridine (EdU) staining is used to visualize DNA replication to exclude any S phase cells. This protocol provides an alternative approach to the conventional, non-denaturing 5-bromo-2'-deoxyuridine (BrdU)-based assays and is better suited for the detection of ssDNA foci outside the S phase.

Author Spotlight: Visualizing Single-Stranded DNA During DNA Repair for Therapeutic Insights 

The following protocol presents the detection of single-stranded DNA foci in the G1 phase of the cell cycle utilizing cell cycle synchronization followed by RPA2 immunofluorescent staining.

DNA has dedicated cellular repair pathways capable of coping with lesions that could arise from both endogenous and/or exogenous sources. DNA repair ...

Quantifying Replication Stress in Ovarian Cancer Cells Using Single-Stranded DNA Immunofluorescence

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Immunofluorescence Imaging of DNA Damage and Repair Foci in Human Colon Cancer Cells

Exploring X Chromosomal Aberrations in Ovarian Cells by Using Fluorescence In Situ Hybridization

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

Visualizing DNA Damage Repair Proteins in Patient-Derived Ovarian Cancer Organoids via Immunofluorescence Assays

Detection of DNA Double-Stranded Breaks in Mouse Oocytes

Quantitation of Intra-peritoneal Ovarian Cancer Metastasis

Substructure Analyzer: A User-Friendly Workflow for Rapid Exploration and Accurate Analysis of Cellular Bodies in Fluorescence Microscopy Images

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Visualization of DNA Repair Proteins Interaction by Immunofluorescence

Visualizing Single-Stranded DNA Foci in the G1 Phase of the Cell Cycle