The work was supported, in part, by the National Institute of Health Center of Biological Research Excellence [grant number 1P20GM109005], the Arkansas Space Grant Consortium through National Aeronautics and Space Administration [grant number NNX15AK32A], and the National Institute for Occupational Safety and Health (NIOSH) [grant number 2T420H008436]. The authors would like to thank Christopher Fettes for proofreading and editing this manuscript.

1. Particle Collection and Preparation
<ol>
	<li>Calibration of the high volume total suspended particles sampler
	<ol>
		<li>Disconnect the sampler motor from the mass flow controller and connect the motor to a stable AC power source (Figure 1). 
		NOTE: A filter is not used during this procedure.</li>
		<li>Mount the calibrator orifice and top loading adapter plate to the sampler. Tighten the top loading adapter hold-down nuts securely to ensure that no air leaks are present. Allow the sampler motor to warm up to its normal operating temperature (approximately 10-15 min).</li>
		<li>Conduct a leak test by covering the holes on top of the orifice and pressure tap on the orifice with hands. Listen for a high-pitched squealing sound made by escaping air. If this sound is heard, re-tighten the top loading adapter hold-down nuts as a leak is present. If the sound is lower, the leak is near one of the other gaskets in the system.</li>
		<li>Connect one side of a water manometer to the pressure tap on the side of the orifice with a rubber vacuum tube. Leave the opposite side of the manometer open to the atmosphere. Hold a manometer vertically to ensure accurate readings. Tapping the backside of the continuous flow recorder will help to center the pen and provide accurate readings.</li>
		<li>Repeat step 1.1.2 using five flow rates representing operating flow rates from 30 to 60 ft/min (approximately every 5-6 ft3/min). Adjust the knob on the variable orifice to five different positions and take five different readings.</li>
		<li>Record the ambient air temperature, ambient barometric pressure, sampler serial number, orifice serial number, orifice slope and intercept with date last certified, the date, site location, and operator&#39;s initials on the calibration sheet.</li>
	</ol>
	</li>
	<li>Total suspended particles sample collection and gravimetric analysis
	<ol>
		<li>Wrap an 8&#34; x 10&#34; quartz fiber filter in two layers of aluminum foil. Place the wrapped filter in the furnace and set the temperature at 550 &#176;C. Bake the filter for at least 4 hr. Let the furnace cool and retrieve the filter. Place the filter in the desiccator with CaCO3.</li>
		<li>Weigh the filter in a microbalance with precision of 10 &#181;g. Repeat the measurement three times during a day. Individual measurements should be within 25 &#181;g. If this is not achieved, return the wrapped filter to the desiccator and repeat the process the following day.</li>
		<li>Open the shelter lid (Figure 2). Remove filter holder frame by loosening the four wing nuts, allowing the brass bolts and washers to swing down out of the way. Unfold the filter from the aluminum foil, and use cleaned forceps to carefully center it, rougher side up, on the supporting screen. Properly align the filter on the screen.</li>
		<li>Place the frame on the screen and secure it with the brass bolts and washers while applying sufficient pressure to avoid air leakage at the edges. Wipe any dust accumulation from around the filter holder with a clean cloth. Close shelter lid carefully and secure it.</li>
		<li>Make sure all cords are plugged into their appropriate receptacle sockets and the tubing between the blower motor pressure tap and the continuous flow recorder is connected.</li>
		<li>Prepare the flow recorder. Depress pen arm lifter to raise pen point and insert a new chart. Align the tab of the chart to the drive hub of the recorder and press with thumb to lower chart center onto hub. Set time by rotating the drive hub clock-wise until the correct time on chart is aligned with time index pointer.</li>
		<li>Manually switch on the sampler and start the collection. Monitor the recorder to be sure it is inking correctly and the elapsed time indicator to be sure it is working properly.</li>
		<li>At the end of the sampling period, switch off the sampler, record the elapsed time and retrieve the chart. Remove the frame to expose the filter. Remove the exposed filter from the supporting screen by holding it gently at the ends with the forceps. Fold the filter lengthwise so that sample touches sample twice and wrap it in the original aluminum foil.</li>
		<li>Store the filter in a desiccator with CaCO3 for at least 24 hr prior to weighing. Repeat step 1.2.2 to measure the weight of the filters after sampling. Place the filter in a zip plastic bag and store it in -80 &#176;C until analysis.</li>
	</ol>
	</li>
	<li>Analysis of collected total suspended particles sample
	<ol>
		<li>Retrieve the stored filter, unfold the aluminum foil and transfer the filter on a clean Teflon surface. Using forceps and scalpel, cut outer (white) part of the filter and dispose.
		<ol>
			<li>Next, cut the filter in 0.5&#34; x 0.5&#34; pieces and place pieces in a 100 ml flask. Add 50 ml of ultrapure water. Place the flask in an ultrasound bath and sonicate for 60 min at 33&#177;3 kHz.</li>
		</ol>
		</li>
		<li>Filter extract through a 0.45 &#181;m polypropylene syringe filter to a 100 ml round bottom flask. Evaporate the water using a rotavap instrument under pressure at 50 &#176;C until about 5 ml of extract remains in the flask.
		<ol>
			<li>Evaporate the remaining water in a 15 ml pear-shaped flask of known weight. Weigh the flask with the dry extract and record the mass of the dry extract.</li>
		</ol>
		</li>
		<li>Dissolve the dry sample with 400 &#181;l of D2O by sonicating for 15 min and transfer into a 5 mm standard NMR tube. Centrifuge for 5 min at 12,000 x g at room temperature to remove any undissolved matter.</li>
		<li>Repeat with 300 &#181;l of D2O. Add 10 &#181;l of NaN3 1.4% (final conc.: 0.02% w/v) in the NMR tube. Add 10 &#181;l (32 &#181;g) of total suspended particles (TSP)-d4 (3.2 mg/ml) to D2O (final concentration 0.258 mM) in the NMR tube.</li>
		<li>Acquire 1H NMR spectra using a NMR spectrometer operating at 600.17 MHz.
		<ol>
			<li>Insert NMR tube into the triple resonance cryoprobe. Calibrate the instrument in the following order: lock, tune, shim, 90&#176; pulse length calibration and receiver gain adjustment.</li>
			<li>Measure spectra at 25 &#176;C with a 1 D excitation sculpting sequence using 180 water-selective pulses in Digital Quad detection mode, 8,192 scans, 4 dummy scans, sweep width of 8012.57 Hz, frequency offset for water signal suppression set at 4.70 ppm, 1.70 sec acquisition time, 32,768 time domain data points (TD), 1 msec duration gradient (p16), 100 &#181;sec recovery delay after gradient (d16).</li>
		</ol>
		</li>
		<li>Process 1H NMR spectra by applying a line broadening of 1 Hz and zero filling by a factor of 2 (65,536 data points). Correct the phase and baseline manually and integrate. Set TSP-d4 peak at 0.0 ppm.</li>
	</ol>
	</li>
</ol>
2. Particle Exposure
<ol>
	<li>Culture set up
	<ol>
		<li>Warm complete growth media containing 10% fetal bovine serum (FBS) and penicillin-streptomycin solution (100 U/ml), 0.25% trypsin solution and calcium magnesium free phosphate buffer saline (PBS) in a water bath (at 37 &#176;C) for at least 30 min before culture initiation.</li>
		<li>Take out a T75 flask from 5% CO2 incubator containing cultured RAW264.7 cells and wash the cells with 2 ml of pre-warmed PBS two times.</li>
		<li>Dislodge adherent RAW264.7 cells from the bottom of tissue culture vessel using 1 ml of pre-warmed trypsin solution followed by scraping.</li>
		<li>Collect the cells from tissue culture vessel and make a single cell suspension by repeated pipetting. Count the number of cells with trypan blue and plate at a density of 8,000/cm2 (500,000 cells total) in a 100 mm tissue culture dish with pre-warmed complete media. Adjust media volume to 10 ml for a final concentration of 50,000 cells per ml. 
		NOTE: Double the number of plates if DNA methylation analysis is also performed.</li>
		<li>Place the newly plated RAW264.7 cells back in 5% CO2 incubator at 37 &#176;C and allow to attach for 24 hr.</li>
		<li>Also in a biosafety cabinet, resuspend filtered particles. For example, dilute particles at a concentration of 0.5 mg/ml in sterile PBS for a 5 &#181;g/ml treatment dose and at 5 mg/ml for a 50 &#181;g/ml treatment dose. Dilutions can be prepared up to month in advance and stored at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Treatment with particulate matter
	<ol>
		<li>Take out the cells from 5% CO2 incubator and check under 10x magnification using an inverted microscope for cell growth, cell morphology, culture condition, and contamination. One should expect to have at least 30% confluent cells at this stage.</li>
		<li>Aseptically, aspirate the media using sterilized pipette and wash the culture vessel with 2 ml 37 &#176;C PBS two times to completely remove the media and floating or dead cells.</li>
		<li>Add pre-determined dose of particulate matter with fresh media in the culture vessels and incubate for desired time at 37 &#176;C in 5% CO2 incubator. In this study, treat RAW264.7 cells with PM for 72 hr to parallel the exposure used for concurrent genotoxic and epigenotoxic analysis. 
		NOTE: Cytotoxicity may negatively affect the quality of the cytogenetic analysis. Therefore, additional assays such as cytotoxicity, DNA methylation, Western blotting, or gene expression analysis are typically performed on treated cells. Expose cells as described and proceed with the specific assay desired. Inexperienced users may also want to include a positive control such as methyl methanesulphonate to test their ability to detect CAs.</li>
	</ol>
	</li>
</ol>
3. Cytogenetic Assay
<ol>
	<li>Treatment to arrest cells at metaphase
	<ol>
		<li>Warm the colcemid solution (10 &#181;g/ml) in a 37 &#176;C water bath for 30 min. Wipe the lid with 70% alcohol under a biosafety cabinet to minimize the risk of contamination</li>
		<li>Completely remove the media aseptically and wash the culture vessels with 2 ml pre-warmed PBS two times.</li>
		<li>Add pre-warmed fresh media (2 ml media/106 cells) containing colcemid solution (5 &#181;l/ml) in the culture vessel and incubate for 2 hr.</li>
	</ol>
	</li>
	<li>Cell harvest
	<ol>
		<li>Warm trypsin solution and PBS in a water bath maintained at 37 &#176;C for 30 min. From this step onwards, there is no need to maintain aseptic condition.</li>
		<li>Completely remove media, wash the culture vessels with 2 ml 37 &#176;C PBS for two times, add required volume of pre-warmed trypsin solution (usually 2 ml for 60 mm culture dish or T25 flask and 4 ml for 100 mm dish or T75 flask), and place the culture vessel back in 5% CO2 incubator at 37 &#176;C for 1 min.</li>
		<li>Take out the culture vessel, detach the cells using a scraper, and add 10 ml of PBS to stop trypsin action.</li>
		<li>Pipette up and down at least ten times to make single cell suspension and transfer cells in a 15 ml conical bottom centrifuge tube.</li>
		<li>Centrifuge cell suspension at 400 x g for 5 min in a swing out centrifuge at room temperature.</li>
		<li>Remove the supernatant, break the cell pellet by gentle tapping, add 10 ml of PBS, and again centrifuge for five min.</li>
	</ol>
	</li>
	<li>Hypotonic treatment and fixation
	<ol>
		<li>Pre-warm 0.075 M potassium chloride solution in a 37 &#176;C water bath at least for 30 min.</li>
		<li>Remove supernatant without disturbing the cell pellet and leave approximately 0.5 ml of PBS.</li>
		<li>Gently break the cell pellet and make single cell suspension using a P200 pipette.</li>
		<li>Add 4 ml of pre-warmed potassium chloride solution drop by drop with gentle shaking.</li>
		<li>Incubate the cells in hypotonic potassium chloride solution in a 37 &#176;C water bath for 20 min.</li>
		<li>Make fresh acetomethanol solution by adding 3-parts methanol with 1-part glacial acetic acid. Keep this freshly prepared fixative at room temperature.</li>
		<li>After hypotonic treatment, add an equal volume (4 ml) of fixative in the tube and gently mix the solution by inverting the tube.</li>
		<li>Centrifuge at 400 x g for 5 min at room temperature, and remove the supernatant. At this stage cell pellet size increases, become whitish and more visible.</li>
		<li>Remove supernatant, add 4 ml fresh fixative and keep at room temperature for 30 min.</li>
		<li>Centrifuge at 400 x g for 5 min, remove supernatant, and give two more washes with 4 ml fixative.</li>
	</ol>
	</li>
	<li>Slide cleaning and metaphase spread preparation
	<ol>
		<li>Immerse glass slides completely in 10% liquid detergent for 30 min.</li>
		<li>Rinse the slides thoroughly in cold running tap water for 30 min.</li>
		<li>Rinse slides three times with distilled water to completely remove detergent, immerse in distilled water, and store in the refrigerator for future use. 
		NOTE: Cleaning slides before use facilitates uniformity of spreading and increases metaphase quality.</li>
		<li>Drop 10 &#181;l cell suspension on chilled wet slide and allow it to air dry overnight at room temperature.</li>
	</ol>
	</li>
	<li>Staining and mounting
	<ol>
		<li>Prepare 50 ml Giemsa staining solution by mixing one part of Giemsa stain solution with one part of PBS and pour in a Coplin jar.</li>
		<li>Immerse the slides in staining solution for 20 min.</li>
		<li>Rinse the slides in distilled water to remove excess stain and immediately dry the slides using a hair-dryer.</li>
		<li>Leave the slides at room temperature overnight.</li>
		<li>Apply a single drop of mounting medium on the slide, gently place a coverslip on the mounting medium, allow the medium to spread over the slide slowly, and remove the excess medium with a paper towel. Allow the mounted slide to dry for at least 1 hr prior to moving or viewing under a microscope to avoid movement of the coverslip. 
		Caution: It is important to place the coverslip carefully without forming any bubbles during this step. The use of excessive heat to remove bubbles is not recommended.</li>
	</ol>
	</li>
	<li>Microscopic observation and aberration types
	<ol>
		<li>Examine the structural CAs in metaphase spreads at 100X magnification using a good-quality bright-field microscope.</li>
		<li>Score at least 50 to 100 metaphase spreads for each treatment group for treatments that induce a large number of CAs. For smaller effect sizes, the analysis of up to 300 metaphase spreads is advised.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Cytogenetic study or microscopic analysis of the numbers or structures of chromosomes, primarily in metaphase spreads, provides information crucial for prognosis, risk assessment, and treatment for various diseases. It is now well-established that cytogenetic abnormalities are linked with the progression and development of several diseases, including cancer. To date, CAs have been found in all major tumor types. CAs may arise spontaneously or by either external or internal stimuli, such as ionizing radiation (IR) &#8212;...

It has been estimated that exposure to particulate matter (PM) causes over 3 million excess deaths annually, primarily from cardiopulmonary disease and lung cancer1. Indeed, PM was recognized as carcinogenic to humans by the International Agency for Research on Cancer (Group 1), as an increased risk of lung cancer with increasing levels of exposure to PM has been shown2. Interestingly, almost all the cancer cells harbor numerical and/or structural chromosomal abnormalities. Particulate organic carbon is a variant, complex, and heterogeneous mixture, whose composition and size distribution depends on emissions as well as physical and chemical transformations. It accounts from 35-55% of urban PM2.5 mass (PM 2.5 &#181;m in size and smaller) and more than 60% of rural and continental background PM2.5 mass3, 4. The water soluble fraction accounts for 30-90% of organic aerosol. A large number of organic compounds have been identified, including aliphatic hydrocarbons, polycyclic aromatic hydrocarbons, and their oxygenated and nitrated derivatives, aliphatic aldehydes and alcohols, free fatty acids and their salts, di-carboxylic acids, multifunctional compounds, proteins, and humic-like macromolecules (HULIS) using chromatographic methods coupled with mass spectrometric techniques5-8. These compounds represent less than 10-20% of particulate organic carbon, thus most of organic carbon is unknown9.
Experimental evidence suggests that cytotoxicity, oxidative stress, and inflammation are involved in the development of PM-associated pathological states. It has been recently shown, however, that exposure to PM also results in a number of epigenetic alterations, including alterations in DNA methylation of repetitive elements, in both experimental in vitro systems and in human subjects10-12. Of particular interest are the effects of PM on satellite DNA &#8212; major and minor satellites &#8212; which are found in the heterochromatic region around the centromere of chromosomes. It was shown that these effects may be persistent by nature, as they can be detected for at least 72 hr after exposure12. Alterations in DNA methylation, particularly around centromeres, may lead to accumulation of satellite DNA mRNA transcripts, compromise chromosomal integrity during cell division and, subsequently, result in the development of a variety of pathological states13.
Adaptation of cytogenetic approach for the analysis of CAs as an end-point of epigenetic alterations caused by PM, thus, is of high importance. Here, we report the approach for the ambient PM collection and preparation, in vitro exposure and analysis of CAs using the murine macrophage RAW264.7 model system. Macrophages comprise the first line of defense against the inhaled foreign objects and, therefore, this cell line serves as an established and the most frequently used model in particle toxicology11, 12, 14, 15.

<table><tbody><tr><td>Total suspended particulate sampler</td><td>Tisch Environmental</td><td>TE-5170</td><td></td></tr><tr><td>Bruker Avance III NMR spectrometer&amp;nbsp;</td><td>Bruker</td><td>NA</td><td></td></tr><tr><td>TopSpin 3.5/pl2 software</td><td>Bruker</td><td>NA</td><td></td></tr><tr><td>ACD/NMR Processor Academic Edition</td><td>ACD/Labs</td><td>NA</td><td></td></tr><tr><td>RAW264.7 murine macrophages</td><td>ATCC</td><td>ATCC TIB-71</td><td></td></tr><tr><td>High glucose DMEM GlutaMAX media</td><td>ThermoFisher</td><td>10569010</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>Fetal Bovine Serum</td><td>ThermoFisher</td><td>16000044</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>Penicillin-Streptomycin (10,000 U/ml)</td><td>ThermoFisher</td><td>15140163</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>Trypsin-EDTA (0.25%)</td><td>ThermoFisher</td><td>25200056</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>PBS, pH 7.4</td><td>ThermoFisher</td><td>10010049</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>KaryoMAX Colcemid Solution in PBS</td><td>ThermoFisher</td><td>15212012</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>KaryoMAX Potassium Chloride Solution</td><td>ThermoFisher</td><td>10575090</td><td>Warm in a 37 &amp;deg;C waterbath before use</td></tr><tr><td>Methanol (HPLC)</td><td>Fisher Scientific</td><td>A452N1-19</td><td></td></tr><tr><td>Acetic Acid, Glacial</td><td>Fisher Scientific</td><td>BP1185-500</td><td></td></tr><tr><td>Decon Contrad 70 Liquid Detergent</td><td>Fisher Scientific</td><td>04-355-1</td><td></td></tr><tr><td>Wright and Wright-Giemsa Stain Solutions</td><td>Fisher Scientific</td><td>23-200733</td><td></td></tr><tr><td>Permount Mounting Medium</td><td>Fisher Scientific</td><td>SP15-100</td><td></td></tr><tr><td>Axio Imager 2</td><td>Zeiss</td><td>NA</td><td></td></tr></tbody></table>

analysis of the ambient particulate matter-induced chromosomal aberrations using an in vitro system

Exposure to particulate matter (PM) is a major world health concern, which may damage various cellular components, including the nuclear genetic material. To assess the impact of PM on nuclear genetic integrity, structural chromosomal aberrations are scored in the metaphase spreads of mouse RAW264.7 macrophage cells. PM is collected from ambient air with a high volume total suspended particles sampler. The collected material is solubilized and filtered to retain the water-soluble, fine portion. The particles are characterized for chemical composition by nuclear magnetic resonance (NMR) spectroscopy. Different concentrations of particle suspension are added onto an in vitro culture of RAW264.7 mouse macrophages for a total exposure time of 72 hr, along with untreated control cells. At the end of exposure, the culture is treated with colcemid to arrest cells in metaphase. Cells are then harvested, treated with hypotonic solution, fixed in acetomethanol, dropped onto glass slides and finally stained with Giemsa solution. Slides are examined to assess the structural chromosomal aberrations (CAs) in metaphase spreads at 1,000X magnification using a bright-field microscope. 50 to 100 metaphase spread are scored for each treatment group. This technique is adapted for the detection of structural chromosomal aberrations (CAs), such as chromatid-type breaks, chromatid-type exchanges, acentric fragments, dicentric and ring chromosomes, double minutes, endoreduplication, and Robertsonian translocations in vitro after exposure to PM. It is a powerful method to associate a well-established cytogenetic endpoint to epigenetic alterations.

Great care should be taken in choosing the location of the TSP sampler, as well as the time of the year when collection is performed. The chemical composition as well as the size of particles may substantially influence the results. The material collected should be visible against the white filter. A normal mouse metaphase spread will have 40 acentric chromosomes. The goal of the technique is to demonstrate a change (or absence thereof) in the proportion of abnormal chromosomes in treated...

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Analysis of the Ambient Particulate Matter-induced Chromosomal Aberrations Using an In Vitro System

This protocol describes techniques for the quantification and characterization of chromosomal aberrations in vitro in RAW264.7 mouse macrophages after treatment with ambient air particulate matter.

Analysis of the Ambient Particulate Matter-induced Chromosomal Aberrations Using an In Vitro System

Exposure to particulate matter (PM) is a major world health concern, which may damage various cellular components, including the nuclear genetic material. ...

analysis-ambient-particulate-matter-induced-chromosomal-aberrations

Research

JoVE Journal

Genetics

8.6K Views.  University of Arkansas for Medical Sciences.  Dit protocol beschrijft technieken voor de kwantificatie en karakterisatie van chromosomale afwijkingen in vitro in RAW264.7 muis macrofagen na behandeling met de lucht zwevende deeltjes. 

Video: Analysis of the Ambient Particulate Matter-induced Chromosomal Aberrations Using an In Vitro System

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the field of environmental health. This is due in part to the paucity of model systems that can (1) accurately recapitulate keys features of reproductive processes and (2) do so in a medium- to high-throughput fashion, without the need for a high number of vertebrate animals.
We describe here an assay in the nematode C. elegans that allows the rapid identification of germline toxicants by monitoring the induction of aneuploid embryos. By making use of a GFP reporter line, errors in chromosome segregation resulting from germline disruption are easily visualized and quantified by automated fluorescence microscopy. Thus the screening of a particular set of compounds for its toxicity can be performed in a 96- to 384-well plate format in a matter of days. Secondary analysis of positive hits can be performed to determine whether the chromosome abnormalities originated from meiotic disruption or from early embryonic chromosome segregation errors. Altogether, this assay represents a fast first-pass strategy for the rapid assessment of germline dysfunction following chemical exposure.

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

We describe the detailed steps of a high-throughput chemical assay in the nematode Caenorhabditis elegans used to assess germline toxicity. In this assay, disruption of germline function following chemical exposure is monitored using a fluorescent reporter specific to aneuploid embryos.

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the ...

Developmental Biology

Detection of Inter-chromosomal Stable Aberrations by Multiple Fluorescence In Situ Hybridization (mFISH) and Spectral Karyotyping (SKY) in Irradiated Mice

Ionizing radiation (IR) induces numerous stable and unstable chromosomal aberrations. Unstable aberrations, where chromosome morphology is substantially compromised, can easily be identified by conventional chromosome staining techniques. However, detection of stable aberrations, which involve exchange or translocation of genetic materials without considerable modification in the chromosome morphology, requires sophisticated chromosome painting techniques that rely on in situ hybridization of fluorescently labeled DNA probes, a chromosome painting technique popularly known as fluorescence in situ hybridization (FISH). FISH probes can be specific for whole chromosome/s or precise sub-region on chromosome/s. The method not only allows visualization of stable aberrations, but it can also allow detection of the chromosome/s or specific DNA sequence/s involved in a particular aberration formation. A variety of chromosome painting techniques are available in cytogenetics; here two highly sensitive methods, multiple fluorescence in situ hybridization (mFISH) and spectral karyotyping (SKY), are discussed to identify inter-chromosomal stable aberrations that form in the bone marrow cells of mice after exposure to total body irradiation. Although both techniques rely on fluorescent labeled DNA probes, the method of detection and the process of image acquisition of the fluorescent signals are different. These two techniques have been used in various research areas, such as radiation biology, cancer cytogenetics, retrospective radiation biodosimetry, clinical cytogenetics, evolutionary cytogenetics, and comparative cytogenetics.

Detection of Inter-chromosomal Stable Aberrations by Multiple Fluorescence In Situ Hybridization mFISH and Spectral Karyotyping SKY in Irradiated Mice

The present protocol describes the usefulness of multiple fluorescence in situ hybridization (mFISH) and spectral karyotyping (SKY) in identifying inter-chromosomal stable aberrations in the bone marrow cells of mice after exposure to total body irradiation.

Ionizing radiation (IR) induces numerous stable and unstable chromosomal aberrations. Unstable aberrations, where chromosome morphology is substantially ...

Rapid Analysis of Chromosome Aberrations in Mouse B Lymphocytes by PNA-FISH

Defective DNA repair leads to increased genomic instability, which is the root cause of mutations that lead to tumorigenesis. Analysis of the frequency and type of chromosome aberrations in different cell types allows defects in DNA repair pathways to be elucidated. Understanding mammalian DNA repair biology has been greatly helped by the production of mice with knockouts in specific genes. The goal of this protocol is to quantify genomic instability in mouse B lymphocytes. Labeling of the telomeres using PNA-FISH probes (peptide nucleic acid - fluorescent in situ hybridization) facilitates the rapid analysis of genomic instability in metaphase chromosome spreads. B cells have specific advantages relative to fibroblasts, because they have normal ploidy and a higher mitotic index. Short-term culture of B cells therefore enables precise measurement of genomic instability in a primary cell population which is likely to have fewer secondary genetic mutations than what is typically found in transformed fibroblasts or patient cell lines.

DNA repair pathways are essential for maintenance of genomic integrity and preventing mutation and cancer. The goal of this protocol is to quantify genomic instability by direct observation of chromosome aberrations in metaphase spreads from mouse B cells using fluorescent in situ hybridization (FISH) for telomeric DNA repeats.

Defective DNA repair leads to increased genomic instability, which is the root cause of mutations that lead to tumorigenesis. Analysis of the frequency ...

Immunology and Infection

An Automated Method to Perform The In Vitro Micronucleus Assay using Multispectral Imaging Flow Cytometry

The in vitro micronucleus (MN) assay is often used to evaluate cytotoxicity and genotoxicity but scoring the assay via manual microscopy is laborious and introduces uncertainty in results due to variability between scorers. To remedy this, automated slide-scanning microscopy as well as conventional flow cytometry methods have been introduced in an attempt to remove scorer bias and improve throughput. However, these methods have their own inherent limitations such as inability to visualize the cytoplasm of the cell and the lack of visual MN verification or image data storage with flow cytometry. Multispectral Imaging Flow Cytometry (MIFC) has the potential to overcome these limitations. MIFC combines the high resolution fluorescent imagery of microscopy with the statistical robustness and speed of conventional flow cytometry. In addition, all collected imagery can be stored in dose-specific files. This paper describes the protocol developed to perform a fully automated version of the MN assay on MIFC. Human lymphoblastoid TK6 cells were enlarged using a hypotonic solution (75 mM KCl), fixed with 4% formalin and the nuclear content was stained with Hoechst 33342. All samples were run in suspension on the MIFC, permitting acquisition of high resolution images of all key events required for the assay (e.g. binucleated cells with and without MN as well as mononucleated and polynucleated cells). Images were automatically identified, categorized and enumerated in the MIFC data analysis software, allowing for automated scoring of both cytotoxicity and genotoxicity. Results demonstrate that using MIFC to perform the in vitro MN assay allows statistically significant increases in MN frequency to be detected at several different levels of cytotoxicity when compared to solvent controls following exposure of TK6 cells to Mitomycin C and Colchicine, and that no significant increases in MN frequency are observed following exposure to Mannitol.

The in vitro micronucleus assay is a well-established method for evaluating genotoxicity and cytotoxicity but scoring the assay using manual microscopy is laborious and suffers from subjectivity and inter-scorer variability. This paper describes the protocol developed to perform a fully automated version of the assay using multispectral imaging flow cytometry.

The in vitro micronucleus (MN) assay is often used to evaluate cytotoxicity and genotoxicity but scoring the assay via manual microscopy is laborious and ...

Bioengineering

The Lambda Select cII Mutation Detection System

A number of transgenic animal models and mutation detection systems have been developed for mutagenicity testing of carcinogens in mammalian cells. Of these, transgenic mice and the Lambda (&#955;) Select cII Mutation Detection System have been employed for mutagenicity experiments by many research groups worldwide. Here, we describe a detailed protocol for the Lambda Select cII mutation assay, which can be applied to cultured cells of transgenic mice/rats or the corresponding animals treated with a chemical/physical agent of interest. The protocol consists of the following steps: (1) isolation of genomic DNA from the cells or organs/tissues of transgenic animals treated in vitro or in vivo, respectively, with a test compound; (2) recovery of the lambda shuttle vector carrying a mutational reporter gene (i.e., cII transgene) from the genomic DNA; (3) packaging of the rescued vectors into infectious bacteriophages; (4) infecting a host bacteria and culturing under selective conditions to allow propagation of the induced cII mutations; and (5) scoring the cII-mutants and DNA sequence analysis to determine the cII mutant frequency and mutation spectrum, respectively.

The Lambda Select cII Mutation Detection System

We describe a detailed protocol for the Lambda Select cII mutation assay in cultured cells of transgenic rodents or the corresponding animals treated with a chemical/physical agent of interest. This approach has been widely used for mutagenicity testing of carcinogens in mammalian cells.

A number of transgenic animal models and mutation detection systems have been developed for mutagenicity testing of carcinogens in mammalian cells. Of ...

Chromosomics: Detection of Numerical and Structural Alterations in All 24 Human Chromosomes Simultaneously Using a Novel OctoChrome FISH Assay

Fluorescence in situ hybridization (FISH) is a technique that allows specific DNA sequences to be detected on metaphase or interphase chromosomes in cell nuclei1. The technique uses DNA probes with unique sequences that hybridize to whole chromosomes or specific chromosomal regions, and serves as a powerful adjunct to classic cytogenetics. For instance, many earlier studies reported the frequent detection of increased chromosome aberrations in leukemia patients related with benzene exposure, benzene-poisoning patients, and healthy workers exposed to benzene, using classic cytogenetic analysis2. Using FISH, leukemia-specific chromosomal alterations have been observed to be elevated in apparently healthy workers exposed to benzene3-6, indicating the critical roles of cytogentic changes in benzene-induced leukemogenesis. 
Generally, a single FISH assay examines only one or a few whole chromosomes or specific loci per slide, so multiple hybridizations need to be conducted on multiple slides to cover all of the human chromosomes. Spectral karyotyping (SKY) allows visualization of the whole genome simultaneously, but the requirement for special software and equipment limits its application7. Here, we describe a novel FISH assay, OctoChrome-FISH, which can be applied for Chromosomics, which we define here as the simultaneous analysis of all 24 human chromosomes on one slide in human studies, such as chromosome-wide aneuploidy study (CWAS)8. The basis of the method, marketed by Cytocell as the Chromoprobe Multiprobe System, is an OctoChrome device that is divided into 8 squares, each of which carries three different whole chromosome painting probes (Figure 1). Each of the three probes is directly labeled with a different colored fluorophore, green (FITC), red (Texas Red), and blue (Coumarin). The arrangement of chromosome combinations on the OctoChrome device has been designed to facilitate the identification of the non-random structural chromosome alterations (translocations) found in the most common leukemias and lymphomas, for instance t(9;22), t(15;17), t(8;21), t(14;18)9. Moreover, numerical changes (aneuploidy) in chromosomes can be detected concurrently. The corresponding template slide is also divided into 8 squares onto which metaphase spreads are bound (Figure 2), and is positioned over the OctoChrome device. The probes and target DNA are denatured at high-temperature and hybridized in a humid chamber, and then all 24 human chromosomes can be visualized simultaneously. 
OctoChrome FISH is a promising technique for the clinical diagnosis of leukemia and lymphoma and for detection of aneuploidies in all chromosomes. We have applied this new Chromosomic approach in a CWAS study of benzene-exposed Chinese workers8,10.

A novel fluorescence in situ hybridization (FISH) method that simultaneously examines both numerical and structural chromosome alterations, particularly the specific chromosomal translocations associated with leukemia and lymphoma, of all 24 human chromosomes on a single device in one hybridization, is described.

Fluorescence in situ hybridization (FISH) is a technique that allows specific DNA sequences to be detected on metaphase or interphase chromosomes in cell ...

Biology

Quantification of three DNA Lesions by Mass Spectrometry and Assessment of Their Levels in Tissues of Mice Exposed to Ambient Fine Particulate Matter

DNA adducts and oxidized DNA bases are examples of DNA lesions that are useful biomarkers for the toxicity assessment of substances that are electrophilic, generate reactive electrophiles upon biotransformation, or induce oxidative stress. Among the oxidized nucleobases, the most studied one is 8-oxo-7,8-dihydroguanine (8-oxoGua) or 8-oxo-7,8-dihydro-2&#39;-deoxyguanosine (8-oxodGuo), a biomarker of oxidatively induced base damage in DNA. Aldehydes and epoxyaldehydes resulting from the lipid peroxidation process are electrophilic molecules able to form mutagenic exocyclic DNA adducts, such as the etheno adducts 1,N2-etheno-2&#39;-deoxyguanosine (1,N2-&#949;dGuo) and 1,N6-etheno-2&#39;-deoxyadenosine (1,N6-&#949;dAdo), which have been suggested as potential biomarkers in the pathophysiology of inflammation. Selective and sensitive methods for their quantification in DNA are necessary for the development of preventive strategies to slow down cell mutation rates and chronic disease development (e.g., cancer, neurodegenerative diseases). Among the sensitive methods available for their detection (high performance liquid chromatography coupled to electrochemical or tandem mass spectrometry detectors, comet assay, immunoassays, 32P-postlabeling), the most selective are those based on high performance liquid chromatography coupled to tandem mass spectrometry (HPLC-ESI-MS/MS). Selectivity is an essential advantage when analyzing complex biological samples and HPLC-ESI-MS/MS evolved as the gold standard for quantification of modified nucleosides in biological matrices, such as DNA, urine, plasma and saliva. The use of isotopically labeled internal standards adds the advantage of corrections for molecule losses during the DNA hydrolysis and analyte enrichment steps, as well as for differences of the analyte ionization between samples. It also aids in the identification of the correct chromatographic peak when more than one peak is present.
We present here validated sensitive, accurate and precise HPLC-ESI-MS/MS methods that were successfully applied for the quantification of 8-oxodGuo, 1,N6-dAdo and 1,N2-dGuo in lung, liver and kidney DNA of A/J mice for the assessment of the effects of ambient PM2.5 exposure.

We describe here methods for sensitive and accurate quantification of the lesions 8-oxo-7,8-dihydro-2&#39;-deoxyguanosine (8-oxodGuo), 1,N6-etheno-2&#39;-deoxyadenosine (1,N6-dAdo) and 1,N2-etheno-2&#39;-deoxyguanosine (1,N2-dGuo) in DNA. The methods were applied to the assessment of the effects of ambient fine particulate matter (PM2.5) in tissues (lung, liver and kidney) of exposed A/J mice.

DNA adducts and oxidized DNA bases are examples of DNA lesions that are useful biomarkers for the toxicity assessment of substances that are ...

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage checkpoint signaling as a surveillance mechanism to sense DNA damage and direct cellular responses to DNA damage. There are several groups of proteins called sensors, transducers and effectors involved in DNA damage checkpoint signaling (Figure 1). In this complex signaling pathway, ATR (ATM and Rad3-related) is one of the major kinases that can respond to DNA damage and replication stress. Activated ATR can phosphorylate its downstream substrates such as Chk1 (Checkpoint kinase 1). Consequently, phosphorylated and activated Chk1 leads to many downstream effects in the DNA damage checkpoint including cell cycle arrest, transcription activation, DNA damage repair, and apoptosis or senescence (Figure 1). When DNA is damaged, failing to activate the DNA damage checkpoint results in unrepaired damage and, subsequently, genomic instability. The study of the DNA damage checkpoint will elucidate how cells maintain genomic integrity and provide a better understanding of how human diseases, such as cancer, develop.
Xenopus laevis egg extracts are emerging as a powerful cell-free extract model system in DNA damage checkpoint research. Low-speed extract (LSE) was initially described by the Masui group1. The addition of demembranated sperm chromatin to LSE results in nuclei formation where DNA is replicated in a semiconservative fashion once per cell cycle.
The ATR/Chk1-mediated checkpoint signaling pathway is triggered by DNA damage or replication stress 2. Two methods are currently used to induce the DNA damage checkpoint: DNA damaging approaches and DNA damage-mimicking structures 3. DNA damage can be induced by ultraviolet (UV) irradiation, &gamma;-irradiation, methyl methanesulfonate (MMS), mitomycin C (MMC), 4-nitroquinoline-1-oxide (4-NQO), or aphidicolin3, 4. MMS is an alkylating agent that inhibits DNA replication and activates the ATR/Chk1-mediated DNA damage checkpoint 4-7. UV irradiation also triggers the ATR/Chk1-dependent DNA damage checkpoint 8. The DNA damage-mimicking structure AT70 is an annealed complex of two oligonucleotides poly-(dA)70 and poly-(dT)70. The AT70 system was developed in Bill Dunphy's laboratory and is widely used to induce ATR/Chk1 checkpoint signaling 9-12.
Here, we describe protocols (1) to prepare cell-free egg extracts (LSE), (2) to treat Xenopus sperm chromatin with two different DNA damaging approaches (MMS and UV), (3) to prepare the DNA damage-mimicking structure AT70, and (4) to trigger the ATR/Chk1-mediated DNA damage checkpoint in LSE with damaged sperm chromatin or a DNA damage-mimicking structure.

Study of the DNA Damage Checkpoint using Xenopus Egg Extracts

Xenopus egg extract is a useful model system to investigate the DNA damage checkpoint. This protocol is for the preparation of Xenopus egg extracts and DNA damage checkpoint inducing reagents. These techniques are adaptable to a variety of DNA damaging approaches in the study of the DNA damage checkpoint signaling.

On a daily basis, cells are subjected to a variety of endogenous and environmental insults. To combat these insults, cells have evolved DNA damage ...

Chromosome Preparation From Cultured Cells

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as fluorescence in situ hybridization (FISH), this assay has the powerful ability to analyze individual cells for aberrations that involve gains or losses of portions of the genome&#160;and rearrangements involving one or more chromosomes. In humans, chromosome abnormalities occur in approximately 1 per 160 live births1,2, 60-80% of all miscarriages3,4, 10% of stillbirths2,5, 13% of individuals with congenital heart disease6, 3-6% of infertility cases2, and in many patients with developmental delay&#160;and birth defects7. Cytogenetic analysis of malignancy is routinely used by researchers and clinicians, as observations of clonal chromosomal abnormalities have been shown to have both diagnostic and prognostic significance8,9.&#160; Chromosome isolation is invaluable for gene therapy and stem cell research of organisms including nonhuman primates and rodents10-13.
Chromosomes can be isolated from cells of live tissues, including blood lymphocytes, skin fibroblasts, amniocytes, placenta, bone marrow, and tumor specimens. Chromosomes are analyzed at the metaphase stage of mitosis, when they are most condensed and therefore more clearly visible. The first step of the chromosome isolation technique involves the disruption of the spindle fibers by incubation with Colcemid, to prevent the cells from proceeding to the subsequent anaphase stage. The cells are then treated with a hypotonic solution and preserved in their swollen state with Carnoy&#39;s fixative. The cells are then dropped on to slides and can then be utilized for a variety of procedures. G-banding involves trypsin treatment followed by staining with Giemsa to create characteristic light and dark bands. The same procedure to isolate chromosomes can be used for the preparation of cells for procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY)14,15.

Chromosomes can be isolated from live cells such as lymphocytes or skin fibroblasts, and from organisms including humans or mice. These chromosome preparations can be further utilized for routine G-banding and molecular cytogenetic procedures such as fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH), and spectral karyotyping (SKY).

Chromosome (cytogenetic) analysis is widely used for the detection of chromosome instability. When followed by G-banding and molecular techniques such as ...

Measuring DNA Damage and Repair in Mouse Splenocytes After Chronic In Vivo Exposure to Very Low Doses of Beta- and Gamma-Radiation

Low dose radiation exposure may produce a variety of biological effects that are different in quantity and quality from the effects produced by high radiation doses. Addressing questions related to environmental, occupational and public health safety in a proper and scientifically justified manner heavily relies on the ability to accurately measure the biological effects of low dose pollutants, such as ionizing radiation and chemical substances. DNA damage and repair are the most important early indicators of health risks due to their potential long term consequences, such as cancer. Here we describe a protocol to study the effect of chronic in vivo exposure to low doses of &#947;- and &#946;-radiation on DNA damage and repair in mouse spleen cells. Using a commonly accepted marker of DNA double-strand breaks, phosphorylated histone H2AX called &#947;H2AX, we demonstrate how it can be used to evaluate not only the levels of DNA damage, but also changes in the DNA repair capacity potentially produced by low dose in vivo exposures. Flow cytometry allows fast, accurate and reliable measurement of immunofluorescently labeled &#947;H2AX in a large number of samples. DNA double-strand break repair can be evaluated by exposing extracted splenocytes to a challenging dose of 2 Gy to produce a sufficient number of DNA breaks to trigger repair and by measuring the induced (1 hr post-irradiation) and residual DNA damage (24 hrs post-irradiation). Residual DNA damage would be indicative of incomplete repair and the risk of long-term genomic instability and cancer. Combined with other assays and end-points that can easily be measured in such in vivo studies (e.g., chromosomal aberrations, micronuclei frequencies in bone marrow reticulocytes, gene expression, etc.), this approach allows an accurate and contextual evaluation of the biological effects of low level stressors.

Measuring DNA Damage and Repair in Mouse Splenocytes After Chronic In Vivo Exposure to Very Low Doses of Beta- and Gamma-Radiation

A protocol to evaluate changes in DNA damage levels and DNA repair capacity that may be induced by chronic in vivo low dose irradiation in mouse spleen lymphocytes, by measuring phosphorylated histone H2AX, a marker of DNA double-strand breaks, using flow cytometry is presented.

Low dose radiation exposure may produce a variety of biological effects that are different in quantity and quality from the effects produced by high ...

Analysis of the Ambient Particulate Matter-induced Chromosomal Aberrations Using an In Vitro System

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