Name: laser microirradiation to study in vivo cellular responses to simple and complex dna damage
Uploaded: 2018-01-31T12:30:00
Description: Watch this Scientific Journal Video about Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage at JoVE.com

We thank Dr. Akira Yasui at Tohoku University, Japan for the GFP-NTH1 expression plasmid, and Dr. Eros Lazzerini Denchi at the Scripps Research Institute, La Jolla, California for the TRF2-YFP and EGFP-53BP11220-1711 expression plasmids. This work was supported by the Air Force Office of Scientific Research (FA9550-04-1-0101) and the Beckman Laser Institute Inc. Foundation (to M.W.B), the Ford Foundation Fellowship from the National Academy of Sciences (to B.A.S), and NSF MCB-1615701 and CRCC CRR-17-426665 (to K. Y.).

1. Basic Cell Preparation
Note: This step is for standard immunofluorescence detection of the endogenous protein recruitment or modification, and for the use of a cell line stably expressing a fluorescently tagged recombinant protein. For example, Potorustridactylus (PtK2) marsupial kidney epithelial cells stably expressing EGFP-53BP11220-1711 or TRF2-YFP were used in our previous study (Figure 1)32. In the former case, th...

The advantages of using laser microirradiation for DDR research are:
1. It is feasible to induce different types and amounts of DNA damage from simple strand breaks to complex DNA damage, and to detect different repair factors at the DNA damage sites by adjusting the laser irradiation parameters. It is also possible to inflict damage multiple times in the same cell nucleus in order to evaluate trans effects (as in Figure 3).
2....

In Vivo DNA Damage Signaling is Not Well Understood 
In vivo, DNA is complexed with histones and other factors to form chromatin fibers. Regulation of chromatin structure is of paramount importance for DNA metabolism. For example, the histone variant H2AX is phosphorylated by ataxia-telangiectasia mutated (ATM) and other kinases following double-strand breaks (DSB) induction, and is important for DSB damage signal amplification as well as providing a docking site for other factors. Spreading of damage signaling and repair pathway choice appear to be critically influenced by local chromatin structure at damage sites1. A number of chromatin remodeling factors, histone chaperones, and histone modifying enzymes are indeed recruited to damage sites and are important for efficient DNA repair, highlighting the significance of chromatin regulation in the DDR and repair2,3,4. Furthermore, damage site clustering or repositioning was observed in yeast and drosophila5,6,7,8, reminiscent of relocalization of gene loci in the subnuclear compartment associated with gene regulation9,10. Recent studies in mouse and human cells also revealed mobilization of DSB sites, which influences repair fidelity and pathway choice11,12. This raises the possibility that DDR/repair may also be intimately linked to nuclear architecture, higher-order chromatin organization, and chromosome dynamics in the cell nucleus. Thus, it is critical to develop high-resolution methods to study DDR and repair processes in the context of the endogenous nuclear environment in a live cell in order to understand short- and long-term consequences of DNA damage.
Critical Role of PAR Polymerase (PARP) in Gauging the Extent and Type of Damage and Regulating Protein Assembly at the Damage Site 
PARP1 is a DNA nick sensor rapidly activated by DNA damage that plays a critical role in DNA repair13. PARP1 was originally thought to function together with X-Ray Repair Cross Complementing 1 (XRCC1) to facilitate base excision repair (BER), but recent studies reveal its role in other DNA repair pathways, including DSB repair14. Activated PARP1 uses nicotinamide adenine dinucleotide (NAD+) as a substrate to ADP-ribosylate multiple target proteins, including itself. This enzyme and other family members have attracted much attention in recent years as PARP inhibitors have emerged as promising therapeutic drugs for cancers. Although PARP inhibitors were initially found to be effective in breast cancer gene (BRCA)-mutated breast cancer cells, there is now a plethora of evidence for their effects in mono- and combination therapies together with DNA damaging agents/irradiation against a broad spectrum of cancers with mutations not limited to BRCA15,16,17,18,19,20.
At the molecular level, PARP activation was shown to play critical roles in organizing the local chromatin structure at damage sites. PAR-dependent recruitment of chromatin modification enzymes facilitates DSB repair and dictates repair pathway choices, suggesting the important scaffolding role of PAR modification at damage sites. 13,21,22,23,24,25,26,27,28,29,30,31 We recently demonstrated the exclusion of p53-binding protein 1 (53BP1) from damage sites by PAR 32, providing an alternative explanation for 53BP1-dependent hyperactivation of non-homologous endjoining (NHEJ) by PARP inhibitor and highlighting the significance of PARP in DSB repair pathway choice33,34. PARP1 also directly PARylates and affects the activity of multiple DNA repair factors14.
Using Laser Microirradiation as a Tool to Study DDR/Repair In Vivo 
Laser microirradiation to produce sub-micron alterations on individual chromosomes was first described in 196935 and reviewed in detail in 198136. Several decades later, laser microirradiation was shown to induce DNA damage at a defined submicrometer region in the cell nucleus, and was proven to be a valuable technique to study recruitment or modifications of various factors to DNA lesions in vivo13,37,38,39,40,41. This method allows detection of those factors that do not form distinct irradiation-induced foci (IRIF) at damage sites39,42. It is also possible to examine the spatiotemporal dynamics of chromatin structural changes both at damage sites and in the rest of the nucleus. We carefully compared the DDRs induced by different laser systems and input powers to evaluate the relationship between the type of DNA damage and the microirradiation conditions32,43,44,45. The aberrant recruitment patterns of 53BP1 and telomeric repeat binding factor 2 (TRF2) were observed in the previous laser damage studies, which provided the basis for the recurring concern for the &#34;non-physiological&#34; nature of laser-induced damage46,47,48,49. We found that these apparent discrepancies could now be explained by differential PARP signaling which gauges the amount and complexity of induced damage32. We confirmed that: 1) laser-microirradiated cells (even after high input-power irradiation) are arrested in interphase in a damage checkpoint control-dependent manner and remain viable (at least up to 48 h)32,50; and 2) repair factor recruitment/modifications faithfully recapitulate those observed with treatment with conventional DNA damaging agents and DSB induction by endonucleases32,39,42,44,50,51,52. These results strongly support the physiological relevance of studying laser damage-induced cellular responses.

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laser microirradiation to study in vivo cellular responses to simple and complex dna damage

DNA damage induces specific signaling and repair responses in the cell, which is critical for protection of genome integrity. Laser microirradiation became a valuable experimental tool to investigate the DNA damage response (DDR) in vivo. It allows real-time high-resolution single-cell analysis of macromolecular dynamics in response to laser-induced damage confined to a submicrometer region in the cell nucleus. However, various laser conditions have been used without appreciation of differences in the types of damage induced. As a result, the nature of the damage is often not well characterized or controlled, causing apparent inconsistencies in the recruitment or modification profiles. We demonstrated that different irradiation conditions (i.e., different wavelengths as well as different input powers (irradiances) of a femtosecond (fs) near-infrared (NIR) laser) induced distinct DDR and repair protein assemblies. This reflects the type of DNA damage produced. This protocol describes how titration of laser input power allows induction of different amounts and complexities of DNA damage, which can easily be monitored by detection of base and crosslinking damages, differential poly (ADP-ribose) (PAR) signaling, and pathway-specific repair factor assemblies at damage sites. Once the damage conditions are determined, it is possible to investigate the effects of different damage complexity and differential damage signaling as well as depletion of upstream factor(s) on any factor of interest.

Using PtK2 cells stably expressing EGFP-53BP11220-1711 or TRF2-YFP, laser input power titration was carried out to determine the optimal condition for their recruitment (Figure 1)32.
At high input-power laser damage sites, the significant clustering of CPD (crosslinking damage typically generated by ultraviolet (UV) damage) as well as GFP-NTH1 DNA glycosylase that specifically recognizes base damage were observed. In addition, hi...

Watch this Scientific Journal Video about Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage at JoVE.com

Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage

The goal of this protocol is to describe how to use laser microirradiation to induce different types of DNA damage, including relatively simple strand breaks and complex damage, to study DNA damage signaling and repair factor assembly at damage sites in vivo.

Laser Microirradiation to Study In Vivo Cellular Responses to Simple and Complex DNA Damage

DNA damage induces specific signaling and repair responses in the cell, which is critical for protection of genome integrity. Laser microirradiation ...

The overall goal of this protocol is to describe how to use laser microirradiation to induce different types of DNA damage, including simple strand breaks, and complex damage, to study DNA damage signaling and repair factor assembly in vivo. In the 1960s and 70s, we focused a laser beam through a microscope that you can see behind me, and was able to damage a small region of a chromosome. The cell divided, survived, and we couldn't understand why.Well, 40 years later, we are now using this method to selectively study DNA repair, and that's what you're going to see today. This method can help answer key questions in the DNA repair field, such as how DNA damage is recognized and processed and how damage signal propagates in the cell. Laser microirradiation allows for real time, high resolution, single cell analysis of macromolecular dynamics in response to DNA damage confounded to a several micrometer region within the cell nucleus.However, various laser conditions have been used without appreciation for differences in the types of damage induced, causing inconsistencies in cellular responses, factor recruitment and modifications. To express fluorescently-tagged proteins in Hela cells, on day one, seed six to eight times 10 to the fourth cells in 0.5 milliliters of culture medium in one well of a 24-well plate. Then place the plate in the tissue culture incubator.On day two, perform liposome-mediated DNA transfection by diluting 0.3 micrograms of a mammalian expression plasmid DNA encoding a fluorescently tagged DNA repair factor into 25 microliters of serum-free culture medium. Here, we use the damage site targeting domain of 53BP1 fused to GFP for double strand break damage, or DNA glycosylase NTH1 fused to GFP for base damage. In another 1.5 milliliter tube, dilute one microliter of liposome-based DNA transfection reagent into 25 microliters of serum-free culture medium, then mix the content of two tubes, diluted DNA and transfection reagent together and incubate the tube at room temperature for 10 minutes to form DNA-lipid complexes.Rinse the cells once with 0.5 milliliters of culture medium, then remove the medium and add 0.5 milliliters of fresh culture medium to the cells. Add the DNA-lipid complexes to the cells and gently mix by rocking the plate a few times. Then return the cells to the incubator.After four to six hours, remove the culture medium from the cells, and add 300 microliters of 0.5%Trypsin-EDTA. Remove the solution, then add 200 microliters of Trypsin-EDTA and incubate the cells at 37 degrees Celsius for three to four minutes. After confirming that the cells are detached, add 800 microliters of culture medium and resuspend the cells by gently pipeting to break up the cell clumps.Transfer the cell suspension to a 35-millimeter culture dish with a gridded coverslip, and add culture medium to a final volume of two milliliters. After 48 hours, the cells will be at 30 to 60%confluency with fluorescent protein expressed, and ready for laser damage induction. Turn on the NIR laser and allow the laser system to warm up for one hour before use.Place a dish of cells on a 37 degree Celsius heated stage in a chamber that allows carbon dioxide and humidity control. For live-cell analysis of fluorescently tagged proteins, use a high magnification oil immersion objective and an appropriate fluorescence filter combination depending on the tag. Search for cells that have comparable fluorescent protein expression.Avoid cells that have too weak or too strong expression in order to reduce cell-to-cell variability in fluorescence measurements. For confocal microscopy, use a 100x 1.3 numerical aperture oil objective lens and the software bleach function to target linear tracks inside the cell nucleus. Adjust the laser transmission percentage to 5%without changing any other settings.Then place a cell in the center of the field. Click the region of interest, and using the mouse, draw a line or box in the nucleus approximately 50 by four pixels. Avoid touching the nuclear membrane or nucleoli.Then click the bleach button to induce DNA damage. Here, the difference between the peak recruitment of GFP-53BP1 and GFP-NTH1 are evident. To carry out quantitative fluorescence analysis of live cells with the confocal microscope, use 1x magnification, and an argon laser for GFP FITC.Set a resolution of 512 x 512 pixels, a scan speed of five, and number one for quick scan or two plus for averaging over multiple scans to improve signal-to-noise ratio. Then capture images. Draw the bleach ROY for damage induction, then select the acquired Time Series sub menu, set number at 20, and the cycle delay at 30 seconds.Choose bleach once after first scan, and then click start B.In this case, the first image is acquired immediately before damage induction, followed by the acquisition of 19 additional images at 30 second intervals. Here, you can see that the GFP-NTH1 is rapidly and transiently recruited to damage sites, whereas GFP-53BP1 accumulates slowly, and persists at damage sites. Some repair factors may be recruited to damage sites in a cell cycle specific manner.To identify G1 phase cells for damage induction, under an inverted microscope, using a 10x objective, and 10x ocular lenses, identify metaphase cells which are slightly lifted and rounded in shape. Acquire images to record the position of metaphase cells on the gridded coverslip. After three to four hours, confirm cell division has resulted in two daughter cells that are ready for damage induction at the recorded position.To prepare SG2 phase cells, use a double thymidine block protocol. Confirm synchronization efficiency by DNA profiling and or detection of Rad51 at damage sites. At high-input power laser damage, clustering of CPD UV damage and GFP-NTH1 DNA glycosylase that specifically recognizes base damage were more clearly observed compared to low input power.In addition, higher levels of XRCC1 and CtIP were observed, reflecting the increased number of single and double strand breaks, and thus indicating that complex DNA damage was induced. As demonstrated here, variation of laser input power results in changes in damage signaling. PARP1 mediates poly ADP-ribosylation or PAR of target proteins at damage sites which is significantly induced by high input power laser but only weakly by low laser power.53BP1 is recruited to low, but not high input power damage sites. Interestingly, the recruitment to low input power damage was suppressed by the presence of the second high input power damage in the same cell nucleus. Inhibition was alleviated by the PARP Inhibitor.Conversely, the inhibitor of PARP that breaks down PAR, enhanced the PAR signal, and effectively suppressed 53BP1 recruitment even at the low input power damage sites. This indicates that PARP signaling, and not the nature of the damage, per say, dictates 53BP1 recruitment. This is an example in which laser titration revealed differential activation of PARP1 gauging the amount and complexity of DNA damage, and controlling the damage site accessibility.Once mastered, this technique can be done in about five days, including cell preparation, laser system setup, DNA damage induction, imaging of live or immunosilent cells, and the data analysis. It may take a longer with cells synchronization or time cost analysis. While attempting this procedure, it's important to remember that titration of laser input power may deviate over time, even in the same laser system, especially after system realignment, thus it is important to periodically check the repair factor recruitment pattern.Following this procedure, results from other DNA damaging methods, such as analyzing irradiation, chemical agents, or expression of endonucleases, can be performed and compared in order to confirm that the results are not due to laser specific artifacts. After it's development, this technique paved the way for researcher's in the field of DNA repair to explore the DNA damage response with regard to nuclear chromatin, and to identify new factors and pathways of DNA damage, recognition, processing, and signaling.

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Research

JoVE Journal

Genetics

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the replacement of the entire gene with a selectable marker via homologous recombination. During homologous recombination, exchange of DNA takes place between sequences with high similarity. Therefore, linear genomic sequences flanking a target gene can be used to specifically direct a selectable marker to the desired integration site. Blunt ends of the deletion construct activate the cell&#39;s DNA repair systems and thereby promote integration of the construct either via homologous recombination or by non-homologous-end-joining. In organisms with efficient homologous recombination, the rate of successful gene deletion can reach more than 50% making this strategy a valuable gene disruption system. The smut fungus Ustilago maydis is a eukaryotic model microorganism showing such efficient homologous recombination. Out of its about 6,900 genes, many have been functionally characterized with the help of deletion mutants, and repeated failure of gene replacement attempts points at essential function of the gene. Subsequent characterization of the gene function by tagging with fluorescent markers or mutations of predicted domains also relies on DNA exchange via homologous recombination. Here, we present the U. maydis strain generation strategy in detail using the simplest example, the gene deletion.

Genetic Manipulation of the Plant Pathogen Ustilago maydis to Study Fungal Biology and Plant Microbe Interactions

We describe a robust gene replacement strategy to genetically manipulate the smut fungus Ustilago maydis. This protocol explains how to generate deletion mutants to investigate infection phenotypes. It can be extended to modify genes in any desired way, e.g., by adding a sequence encoding a fluorescent protein tag.

Gene deletion plays an important role in the analysis of gene function. One of the most efficient methods to disrupt genes in a targeted manner is the ...

Tools to Study the Role of Architectural Protein HMGB1 in the Processing of Helix Distorting, Site-specific DNA Interstrand Crosslinks

High mobility group box 1 (HMGB1) protein is a non-histone architectural protein that is involved in regulating many important functions in the genome, such as transcription, DNA replication, and DNA repair. HMGB1 binds to structurally distorted DNA with higher affinity than to canonical B-DNA. For example, we found that HMGB1 binds to DNA interstrand crosslinks (ICLs), which covalently link the two strands of the DNA, cause distortion of the helix, and if left unrepaired can cause cell death. Due to their cytotoxic potential, several ICL-inducing agents are currently used as chemotherapeutic agents in the clinic. While ICL-forming agents show preferences for certain base sequences (e.g., 5&#39;-TA-3&#39; is the preferred crosslinking site for psoralen), they largely induce DNA damage in an indiscriminate fashion. However, by covalently coupling the ICL-inducing agent to a triplex-forming oligonucleotide (TFO), which binds to DNA in a sequence-specific manner, targeted DNA damage can be achieved. Here, we use a TFO covalently conjugated on the 5&#39; end to a 4&#39;-hydroxymethyl-4,5&#39;,8-trimethylpsoralen (HMT) psoralen to generate a site-specific ICL on a mutation-reporter plasmid to use as a tool to study the architectural modification, processing, and repair of complex DNA lesions by HMGB1 in human cells. We describe experimental techniques to prepare TFO-directed ICLs on reporter plasmids, and to interrogate the association of HMGB1 with the TFO-directed ICLs in a cellular context using chromatin immunoprecipitation assays. In addition, we describe DNA supercoiling assays to assess specific architectural modification of the damaged DNA by measuring the amount of superhelical turns introduced on the psoralen-crosslinked plasmid by HMGB1. These techniques can be used to study the roles of other proteins involved in the processing and repair of TFO-directed ICLs or other targeted DNA damage in any cell line of interest.

Targeted DNA damage can be achieved by tethering a DNA damaging agent to a triplex-forming oligonucleotide (TFO). Using modified TFOs, DNA damage-specific protein association, and DNA topology modification can be studied in human cells by the utilization of modified chromatin immunoprecipitation assays and DNA supercoiling assays described herein.

High mobility group box 1 (HMGB1) protein is a non-histone architectural protein that is involved in regulating many important functions in the genome, ...

Peptide-derived Method to Transport Genes and Proteins Across Cellular and Organellar Barriers in Plants

The capacity to introduce exogenous proteins and express (or down-regulate) specific genes in plants provides a powerful tool for fundamental research as well as new applications in the field of plant biotechnology. Viable methods that currently exist for protein or gene transfer into plant cells, namely Agrobacterium and microprojectile bombardment, have disadvantages of low transformation frequency, limited host range, or a high cost of equipment and microcarriers. The following protocol outlines a simple and versatile method, which employs rationally-designed peptides as delivery agents for a variety of nucleic acid- and protein-based cargoes into plants. Peptides are selected as tools for development of the system due to their biodegradability, reduced size, diverse and tunable properties as well as the ability to gain intracellular/organellar access. The preparation, characterization and application of optimized formulations for each type of the wide range of delivered cargoes (plasmid DNA, double-stranded DNA or RNA, and protein) are described. Critical steps within the protocol, possible modifications and existing limitations of the method are also discussed.

Existing methods for the modification of plants have limited applicability. The novel peptide-derived technology proposed here promises both simplicity and efficiency in the introduction of exogenous protein or genes into the desired intracellular compartments of intact plants.

The capacity to introduce exogenous proteins and express (or down-regulate) specific genes in plants provides a powerful tool for fundamental research as ...

Analysis of Cap-binding Proteins in Human Cells Exposed to Physiological Oxygen Conditions

Translational control is a focal point of gene regulation, especially during periods of cellular stress. Cap-dependent translation via the eIF4F complex is by far the most common pathway to initiate protein synthesis in eukaryotic cells, but stress-specific variations of this complex are now emerging. Purifying cap-binding proteins with an affinity resin composed of Agarose-linked m7GTP (a 5' mRNA cap analog) is a useful tool to identify factors involved in the regulation of translation initiation. Hypoxia (low oxygen) is a cellular stress encountered during fetal development and tumor progression, and is highly dependent on translation regulation. Furthermore, it was recently reported that human adult organs have a lower oxygen content (physioxia 1-9% oxygen) that is closer to hypoxia than the ambient air where cells are routinely cultured. With the ongoing characterization of a hypoxic eIF4F complex (eIF4FH), there is increasing interest in understanding oxygen-dependent translation initiation through the 5' mRNA cap. We have recently developed a human cell culture method to analyze cap-binding proteins that are regulated by oxygen availability. This protocol emphasizes that cell culture and lysis be performed in a hypoxia workstation to eliminate exposure to oxygen. Cells must be incubated for at least 24 hr for the liquid media to equilibrate with the atmosphere within the workstation. To avoid this limitation, pre-conditioned media (de-oxygenated) can be added to cells if shorter time points are required. Certain cap-binding proteins require interactions with a second base or can hydrolyze the m7GTP, therefore some cap interactors may be missed in the purification process. Agarose-linked to enzymatically resistant cap analogs may be substituted in this protocol. This method allows the user to identify novel oxygen-regulated translation factors involved in cap-dependent translation.

Here, we present human cell culture protocols to analyze translation initiation factors that bind the 5' cap of mRNA during physiological oxygen conditions. This method utilizes an Agarose-linked m7GTP cap analog and is suitable to investigate cap-binding factors and their interacting partners.

Translational control is a focal point of gene regulation, especially during periods of cellular stress. Cap-dependent translation via the eIF4F complex ...

Chromatin Immunoprecipitation (ChIP) Protocol for Low-abundance Embryonic Samples

Chromatin immunoprecipitation (ChIP) is a widely-used technique for mapping the localization of post-translationally modified histones, histone variants, transcription factors, or chromatin-modifying enzymes at a given locus or on a genome-wide scale. The combination of ChIP assays with next-generation sequencing (i.e., ChIP-Seq) is a powerful approach to globally uncover gene regulatory networks and to improve the functional annotation of genomes, especially of non-coding regulatory sequences. ChIP protocols normally require large amounts of cellular material, thus precluding the applicability of this method to investigating rare cell types or small tissue biopsies. In order to make the ChIP assay compatible with the amount of biological material that can typically be obtained in vivo during early vertebrate embryogenesis, we describe here a simplified ChIP protocol in which the number of steps required to complete the assay were reduced to minimize sample loss. This ChIP protocol has been successfully used to investigate different histone modifications in various embryonic chicken and adult mouse tissues using low to medium cell numbers (5 x 104 - 5 x 105 cells). Importantly, this protocol is compatible with ChIP-seq technology using standard library preparation methods, thus providing global epigenomic maps in highly relevant embryonic tissues.

Chromatin Immunoprecipitation ChIP Protocol for Low-abundance Embryonic Samples

Here, we describe a chromatin immunoprecipitation (ChIP) and ChIP-seq library preparation protocol to generate global epigenomic profiles from low-abundance chicken embryonic samples.

Chromatin immunoprecipitation (ChIP) is a widely-used technique for mapping the localization of post-translationally modified histones, histone variants, ...

The Nematode Caenorhabditis Elegans - A Versatile In Vivo Model to Study Host-microbe Interactions

We demonstrate a method using Caenorhabditis elegans as a model host to study microbial interaction. Microbes are introduced via the diet making the intestine the primary location for disease. The nematode intestine structurally and functionally mimics mammalian intestines and is transparent making it amenable to microscopic study of colonization. Here we show that pathogens can cause disease and death. We are able to identify microbial mutants that show altered virulence. Its conserved innate response to biotic stresses makes C. elegans an excellent system to probe facets of host innate immune interactions. We show that hosts with mutations in the dual oxidase gene cannot produce reactive oxygen species and are unable to resist microbial insult. We further demonstrate the versatility of the presented survival assay by showing that it can be used to study the effects of inhibitors of microbial growth. This assay may also be used to discover fungal virulence factors as targets for the development of novel antifungal agents, as well as provide an opportunity to further uncover host-microbe interactions. The design of this assay lends itself well to high throughput whole-genome screens, while the ability to cryo-preserve worms for future use makes it a cost-effective and attractive whole animal model to study.

The Nematode Caenorhabditis Elegans - A Versatile In Vivo Model to Study Host-microbe Interactions

Here, we present the nematode Caenorhabditis elegans as a versatile host model to study microbial interaction.

We demonstrate a method using Caenorhabditis elegans as a model host to study microbial interaction. Microbes are introduced via the diet making the ...

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Simultaneous Video-EEG-ECG Monitoring to Identify Neurocardiac Dysfunction in Mouse Models of Epilepsy

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can potentially increase risk of sudden unexpected death in epilepsy (SUDEP). Electroencephalography (EEG) and electrocardiography (ECG) are widely used clinical diagnostic tools to monitor for abnormal brain and cardiac rhythms in patients. Here, a technique to simultaneously record video, EEG, and ECG in mice to measure behavior, brain, and cardiac activities, respectively, is described. The technique described herein utilizes a tethered (i.e., wired) recording configuration in which the implanted electrode on the head of the mouse is hard-wired to the recording equipment. Compared to wireless telemetry recording systems, the tethered arrangement possesses several technical advantages such as a greater possible number of channels for recording EEG or other biopotentials; lower electrode costs; and greater frequency bandwidth (i.e., sampling rate) of recordings. The basics of this technique can also be easily modified to accommodate recording other biosignals, such as electromyography (EMG) or plethysmography for assessment of muscle and respiratory activity, respectively. In addition to describing how to perform the EEG-ECG recordings, we also detail methods to quantify the resulting data for seizures, EEG spectral power, cardiac function, and heart rate variability, which we demonstrate in an example experiment using a mouse with epilepsy due to Kcna1 gene deletion. Video-EEG-ECG monitoring in mouse models of epilepsy or other neurological disease provides a powerful tool to identify dysfunction at the level of the brain, heart, or brain-heart interactions.

Here, we present a protocol to record brain and heart bio signals in mice using simultaneous video, electroencephalography (EEG), and electrocardiography (ECG). We also describe methods to analyze the resulting EEG-ECG recordings for seizures, EEG spectral power, cardiac function, and heart rate variability.

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can ...

Investigation of Protein Recruitment to DNA Lesions Using 405 Nm Laser Micro-irradiation

The DNA Damage Response (DDR) uses a plethora of proteins to detect, signal, and repair DNA lesions. Delineating this response is critical to understand genome maintenance mechanisms. Since recruitment and exchange of proteins at lesions are highly dynamic, their study requires the ability to generate DNA damage in a rapid and spatially-delimited manner. Here, we describe procedures to locally induce DNA damage in human cells using a commonly available laser-scanning confocal microscope equipped with a 405 nm laser line. Accumulation of genome maintenance factors at laser stripes can be assessed by immunofluorescence (IF) or in real-time using proteins tagged with fluorescent reporters. Using phosphorylated histone H2A.X (&#947;-H2A.X) and Replication Protein A (RPA) as markers, the method provides sufficient resolution to discriminate locally-recruited factors from those that spread on adjacent chromatin. We further provide ImageJ-based scripts to efficiently monitor the kinetics of protein relocalization at DNA damage sites. These refinements greatly simplify the study of the DDR dynamics.

Studying DNA damage repair kinetics requires a system to induce lesions at defined sub-nuclear regions. We describe a method to create localized double-stranded breaks using a laser-scanning confocal microscope equipped with a 405 nm laser and provide automated procedures to quantify the dynamics of repair factors at these lesions.

The DNA Damage Response (DDR) uses a plethora of proteins to detect, signal, and repair DNA lesions. Delineating this response is critical to understand ...

A Simple, Rapid, and Quantitative Assay to Measure Repair of DNA-protein Crosslinks on Plasmids Transfected into Mammalian Cells

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in mammalian cell lines. Rather than globally assaying removal of xenobiotic-induced or spontaneous chromosomal DPC removal, this assay examines the repair of a homogeneous, chemically defined lesion specifically introduced at one site within a plasmid DNA substrate. Importantly, this approach avoids the use of radioactive materials and is not dependent on expensive or highly-specialized technology. Instead, it relies on standard recombinant DNA procedures and widely available real-time, quantitative polymerase chain reaction (qPCR) instrumentation. Given the inherent flexibility of the strategy utilized, the size of the crosslinked protein, as well as the nature of the chemical linkage and the precise DNA sequence context of the attachment site can be varied to address the respective contributions of these parameters to the overall efficiency of DPC repair. Using this method, plasmids containing a site-specific DPC were transfected into cells and low molecular weight DNA recovered at various times post-transfection. Recovered DNA is then subjected to strand-specific primer extension (SSPE) using a primer complementary to the damaged strand of the plasmid. Since the DPC lesion blocks Taq DNA polymerase, the ratio of repaired to un-repaired DNA can be quantitatively assessed using qPCR. Cycle threshold (CT) values are used to calculate percent repair at various time points in the respective cell lines. This SSPE-qPCR method can also be used to quantitatively assess the repair kinetics of any DNA adduct that blocks Taq polymerase.

The goal of this protocol is to quantify the repair of defined DNA-protein crosslinks on plasmid DNA. Lesioned plasmids are transfected into recipient mammalian cell lines and low-molecular weight harvested at multiple time points post-transfection. DNA repair kinetics are quantified using strand-specific primer extension followed by qPCR.

The purpose of this method is to provide a flexible, rapid, and quantitative technique to examine the kinetics of DNA-protein crosslink (DPC) repair in ...