This work was supported by the ERC HGTCODONUSE (ERC-2015-CoG-682819) to S.B. Data used in this work were (partly) produced through the GenSeq technical facilities of the Institut des Sciences de l&#39;Evolution de Montpellier with the support of LabEx CeMEB, an ANR &#34;Investissements d&#39;avenir&#34; program (ANR-10-LABX-04-01).

Setting up this protocol requires precise knowledge of the insertion, deletion, inversion, or duplication point within the ancestral sequence. This information is usually obtained by whole-genome sequencing (WGS) of the end or intermediate point samples. In the following protocol, the general principle for the case of an insertion mutation is given for each step, alongside a representative case where the frequency of an IS10 insertion in the mutS gene in an experimental evolution population of E. coli is followed. In this population, WGS of the endpoint population identified the insertion of a 1,329 bp IS10 between positions 2,463 and 2,471, resulting in the duplication of this insertion site. This method is applicable to the three other SV types, and the specificities of each case are given in the discussion.
1. Design of triplet primers
<ol>
	<li>Use classic primer design practices (18-24 bp, 40%-60% GC content, start/end with G/C pairs where possible, Tm difference &#60; 5 &#176;C) to produce primers FW1 and RV1. Design primers to amplify a short amplicon on the WT allele around the mutant allele insertion site (Figure 1). 
	NOTE: The amplicon size can range from 100 bp to up to 3,000 bp, in line with the DNA size ladder used in the capillary electrophoresis. In this example, a 155 bp amplicon was amplified. The small fragment size chosen here prevents&#160;off-target amplification of the whole IS10 insertion sequence (see section 2).</li>
	<li>Design a second forward primer FW2 within the insertion sequence, to produce a second amplicon that is approximately 5% larger or smaller than the WT amplicon (Figure 1). This 5% size difference is the minimum size difference that the parallel capillary electrophoresis device could reliably distinguish. Therefore, design the primers such that the two amplicons have a difference in size, which is above but as close as possible to the relative threshold. 
	NOTE: Be sure to minimize the Tm difference and primer dimer formation. In this example, a second forward primer was designed to produce a 226 bp amplicon, 71 bp larger than the WT amplicon. In this representative example, the primer sequences are as follows: 
	FW1:AAAGCATTTCGCCGAACGCC 
	RV1: GCGATAAATCCACTCCAGCGCC 
	FW2: AGTTCGCTTAGGCATGGAAG</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64709/64709fig01.jpg" style="opacity: 0.9;" /> 
Figure 1: Schema of triplet primer design on mutS WT gene and mutant mutS IS10 insertion.&#160;The black triangle represents the IS10 insertion site in the&#160;mutS&#160;gene. The WT gene is in blue, and the IS10 is in orange. Primers FW1 and RV1 flag the IS10 insertion site and produce a 155 bp WT amplicon. The RV1 primer and the intra-IS10 primer FW2 produce a second 226 bp amplicon.&#160;<a href="https://www.jove.com/files/ftp_upload/64709/64709fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Optimization of PCR conditions
<ol>
	<li>Grow an overnight culture of fixed WT and mutant allele clones.</li>
	<li>Extract the DNA using any kit.</li>
	<li>Quantify the DNA.</li>
	<li>Prepare the sample of DNA by diluting the extraction of WT and mutant DNA to 5 ng/&#181;L. Mix the two DNA samples in a 50/50 ratio.</li>
	<li>Amplify 10 ng of the three DNA samples (WT, mutant, 50/50 mix) using a 2x ready-to-use PCR master mix, 0.5 &#181;M FW1 primer, 1 &#181;M RV1 primer, and 0.5 &#181;M FW2 primer in a 20 &#181;L reaction volume. Use a 2% agarose gel to migrate the PCR product by classical electrophoresis, and determine the optimal PCR conditions. 
	NOTE: Elongation times should be minimized to prevent formation of the FW1 and RV1 amplicon on the mutant allele. The annealing temperature should be adjusted to minimize biased amplification of the alleles and non-specific amplification.
	<ol>
		<li>To follow the program in this example, use the following settings: 98 &#176;C for 10 s, followed by 25 cycles of 98 &#176;C for 1 s, 58 &#176;C for 15 s, 72 &#176;C for 8 s, and a final elongation step at 72 &#176;C for 1 min. 
		&#8203;NOTE: The elongation time was reduced to 8 s to prevent amplification of the &#62;1,000 bp product from the forward primer and the RV1 primer on the mutant allele (mutS with IS10 insertion).</li>
	</ol>
	</li>
</ol>
3. Calibration curve
<ol>
	<li>Mix the two DNA samples, WT and mutant, in 10/90, 25/75, 40/60, 50/50, 60/40, 75/25, and 90/10 ratios. 
	NOTE: Biological replicates are prepared from independent overnight bacterial cultures.</li>
	<li>Amplify using optimized PCR conditions (see section 2).</li>
	<li>Quantify the amplicon product.</li>
	<li>Dilute the PCR products to 0.1 ng/&#181;L.</li>
	<li>Prepare the parallel capillary electrophoresis instrument.
	<ol>
		<li>Mix fresh gel and dye (NGS quantitative analysis kit (22, 33, or 55); HS NGS fragment 1-6,000 bp for this example). 
		NOTE: See the parallel capillary electrophoresis HS NGS fragment guide for detailed instructions (see the Table of Materials).</li>
	</ol>
	</li>
	<li>Replace the capillary storage solution and the inlet buffer, and place the rinse buffer plate in the correct drawer locations of the parallel capillary electrophoresis instrument.</li>
	<li>Add 2 &#181;L of HS diluent marker to 22 &#181;L of each diluted sample in a 96-well plate.</li>
	<li>Add a size ladder (DNA size ladder; range 1-6,000 bp) from a HS NGS quantitative analysis kit to one well of the 96-well plate.</li>
	<li>Place the 96-well plate in the correct drawer of the parallel capillary electrophoresis instrument, and select Run on the parallel capillary electrophoresis instrument software.</li>
	<li>Analyze the results using the data analysis software, which detects and identifies each peak of the size ladder, assigning each peak of the samples to their known actual size.
	<ol>
		<li>When using a quantitative kit, use the software to determine the DNA quantity of each fragment by integrating the area under the peak, as in chromatography data analysis. Again, compare samples with known amounts in standards to quantify each peak from a sample, and calculate the ratios between the different peaks detected in the samples.</li>
		<li>Construct a calibration curve (Figure 2), linking the known proportion of the mutant allele (DNA mix) to the one measured using the parallel capillary electrophoresis instrument. This calibration curve allows the reliability of the method to be evaluated and corrected for minor amplification bias.</li>
	</ol>
	</li>
</ol>
4. Sample preparation
<ol>
	<li>Grow time point samples overnight in standard conditions.</li>
	<li>Extract the DNA.</li>
	<li>Quantify the DNA.</li>
	<li>Amplify the samples using optimized PCR conditions (see section 2).</li>
	<li>Run the samples in the parallel capillary electrophoresis instrument (see steps 3.5-3.10).</li>
</ol>
5. Allele quantification
<ol>
	<li>Extract mutant allele quantities from the parallel capillary electrophoresis instrument data with the software, and calculate the actual proportions by plotting these values on the calibration curve.</li>
</ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome-wide+determination+of+barriers+to+horizontal+gene+transfer.">Genome-wide determination of barriers to horizontal gene transfer.</a> bioRxiv. , (2022).</li><li>Slomka, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+evolution+of+Bacillus+subtilis+reveals+the+evolutionary+dynamics+of+horizontal+gene+transfer+and+suggests+adaptive+and+neutral+effects.">Experimental evolution of Bacillus subtilis reveals the evolutionary dynamics of horizontal gene transfer and suggests adaptive and neutral effects.</a> Genetics. 216 (2), 543-558 (2020).</li><li>Choudhury, D., Saini, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolution+of+Escherichia+coli+in+different+carbon+environments+for+2,000+generations.">Evolution of Escherichia coli in different carbon environments for 2,000 generations.</a> Journal of Evolutionary Biology. 32 (12), 1331-1341 (2019).</li><li>Tenaillon, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tempo+and+mode+of+genome+evolution+in+a+50,000-generation+experiment.">Tempo and mode of genome evolution in a 50,000-generation experiment.</a> Nature. 536 (7615), 165-170 (2016).</li><li>Behringer, M. 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The authors have no conflicts of interest to disclose.

Here, we have proposed a cost-effective method that allows the dynamics of emerging adaptive SV alleles in experimental evolution populations to be followed. This method couples classic PCR techniques and automated parallel capillary electrophoresis, allowing for the relative quantities of two alleles to be determined. Once set up, it permits the quantification of allele proportions in many samples in parallel, and is much less expensive than WGS. This method can be seen as an equivalent to amplicon sequencing for non-SN...

Structural variants (SVs) are alterations of the genomic sequence, generally affecting 50 bp or more. The four categories of described SVs are large insertions, large deletions, inversions, and duplications. Until recently, more attention has been devoted to single-nucleotide variants (SNVs) than to structural variants, in terms of their phenotypic effects and their role as genetic determinants of disease, or their contribution to adaptation. This is probably because it is easier to both detect SNVs and predict their phenotypic effects. However, short- and long-read deep sequencing technologies have strongly improved the detection of SVs, at least in single individual or clonal genomes1. In parallel, their phenotypic effects have been better characterized, and many examples of their implication as genetic determinants of human disease2,3 or adaptation to a new environment4 have been documented.
Deletions and insertions, often due to mobile genetic element (MGE) insertions, are much more disruptive than single nucleotide polymorphisms (SNPs) and lead to frameshift mutations and protein structure modifications. Deletions and MGE insertions within genes almost always result in gene inactivation, and insertions into non-coding regions can lead to repression or constitutive expression of adjacent genes when insertion sequences (ISs) contain promoter or termination sequences5. While the knockout of essential genes leads to clear detrimental effects on bacterial fitness, the loss of non-essential genes is beneficial in some cases. Despite their inherent costs, duplications can also be advantageous, and participate in adaptation as they lead to a change in gene dosage; an increase in the activity of a specific protein can be advantageous depending on the conditions6.
Microbial experimental evolution populations are usually started with clones. This initial absence of genetic diversity, combined with the &#34;closed environment&#34; characteristic of test tubes, leads to a very limited potential of evolution by gene gain through horizontal gene transfer and recombination. In these specific conditions, the contribution to adaptation of deletions, duplications, and intragenomic MGE insertion is particularly important; bacteria often adapt through loss-of-function mutations (mainly due to deletions or MGE insertions), affecting genes that are not useful in stable, often nutrient-rich, monoculture artificial environments7. In the longest running E. coli evolution experiment, IS150 insertions are particularly frequent amongst populations evolved after 50,000 generations, with IS elements representing 35% of mutations that reach high frequency in populations that retain their ancestral point mutation rate8.
Evolve and resequence studies couple experimental evolution and next-generation sequencing (NGS) technologies to investigate how bacteria adapt, at the phenotypic and genomic levels, to different environmental conditions and stresses, such as different carbon and energy sources, antibiotics, and osmotic stress9,10,11. These studies typically obtain genomic information on the evolved populations or clones solely at the experimental end point, and in some cases, at a number of intermediate time points12,13,14. These data provide insight into genes and pathways involved in the adaptation to a given environment, but rarely allow researchers to follow the dynamics of de novo emerging and sweeping alleles over time.
One approach to follow these dynamics is to choose a limited number of segregating alleles of interest (because of the function of the genes they affect, because they sweep in parallel in independent populations, etc.) and use amplicon sequencing to quantify the allele proportion, pooling many time points in the same sequencing run15. This method has been successfully used to follow the dynamics of small size variants (SNPs or 1 bp indels) in experimental16 and natural17 populations of microbes. However, in the case of larger indels or MGE insertions, the size difference of the amplicons induces PCR efficiency differences, which distort the relationship between read and allele proportions. In certain cases, the size difference between the two alleles is superior to the classical length of the amplicon. Here, we coupled a triplet PCR technique with automated parallel capillary electrophoresis to quantify the relative frequency of an insertion allele based on size discrimination. This approach allows the exploitation of underused experimental time points to determine the dynamics of an emerging mutant allele and to follow its frequency to fixation or loss, in a cost-effective manner. We applied this method to track emerging mutS-&#160;alleles, mutated through an IS10 insertion, providing the mutated genotype with a hypermutator phenotype.
This method requires two target alleles with a &#8805;5% difference in size. First, primer triplets are designed to produce similarly sized fragments, which share a common primer. Second, PCR conditions are optimized, and a calibration curve is produced using mixes of wild-type (WT) and mutant gDNA. Lastly, samples are amplified by PCR, and the relative frequency of each allele is quantified by parallel quantitative capillary electrophoresis.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>96 Well Skirted PCR Plate</td>
			<td>4titude</td>
			<td>4Ti - 0740</td>
			<td>PCR</td>
		</tr>
		<tr>
			<td>Agarose molecular biology grade</td>
			<td>Eurogentec</td>
			<td>EP-0010-05</td>
			<td>Agarose gel electrophoresis&#160;</td>
		</tr>
		<tr>
			<td>Agilent DNF-474 HS NGS Fragment Kit Quick Guide for the Fragment Analyzer Systems</td>
			<td>Agilent</td>
			<td></td>
			<td>PDF instruction guide</td>
		</tr>
		<tr>
			<td>Buffer TBE</td>
			<td>Panreac appliChem</td>
			<td>A4228,5000Pc</td>
			<td>Agarose gel electrophoresis&#160;</td>
		</tr>
		<tr>
			<td>Calibrated Disposable Inoculating Loops and Needles</td>
			<td>LABELIANS</td>
			<td>8175CSR40H</td>
			<td>Bacterial culture</td>
		</tr>
		<tr>
			<td>Dneasy Blood and Tissue Kit</td>
			<td>Qiagen</td>
			<td>69506</td>
			<td>DNA extraction</td>
		</tr>
		<tr>
			<td>Electrophoresis power supply</td>
			<td>Amilabo</td>
			<td>ST606T</td>
			<td>Agarose gel electrophoresis&#160;</td>
		</tr>
		<tr>
			<td>Fragment Analyzer Automated CE System</td>
			<td>Agilent</td>
			<td></td>
			<td>Parallel capillary electrophoresis</td>
		</tr>
		<tr>
			<td>Fragment DNA Ladder</td>
			<td>Agilent</td>
			<td>DNF-396, range 1-6000bp</td>
			<td>Parallel capillary electrophoresis</td>
		</tr>
		<tr>
			<td>GENTAMICIN SULFATE SALT BIOREAGENT</td>
			<td>Sigma-Aldrich</td>
			<td>G1264-1G</td>
			<td>Bacterial culture</td>
		</tr>
		<tr>
			<td>High Sensitivity diluent marker</td>
			<td>Agilent</td>
			<td>DNF-373</td>
			<td>Parallel capillary electrophoresis</td>
		</tr>
		<tr>
			<td>High Sensitivity NGS quantitative analysis kit</td>
			<td>Agilent</td>
			<td>DNF-474</td>
			<td>Parallel capillary electrophoresis</td>
		</tr>
		<tr>
			<td>Ladder quick load 1 kb plus DNA ladder</td>
			<td>NEB</td>
			<td>N0469S</td>
			<td>Agarose gel electrophoresis&#160;</td>
		</tr>
		<tr>
			<td>LB Broth, VegitoneNutriSelect Plus</td>
			<td>Millipore</td>
			<td>28713</td>
			<td>Bacterial culture</td>
		</tr>
		<tr>
			<td>Master Mix PCR High Fidelity Phusion Flash</td>
			<td>Thermo Fisher Scientific</td>
			<td>F548L</td>
			<td>PCR</td>
		</tr>
		<tr>
			<td>Primers</td>
			<td>Eurogentec</td>
			<td></td>
			<td>PCR</td>
		</tr>
		<tr>
			<td>Prosize data analysis software v.4</td>
			<td>Agilent</td>
			<td>V.4</td>
			<td>Parallel capillary electrophoresis</td>
		</tr>
		<tr>
			<td>Qubit assays</td>
			<td>Invitrogen</td>
			<td>MAN0010876</td>
			<td>DNA quantification</td>
		</tr>
		<tr>
			<td>Qubit dsDNA HS Assay Kit</td>
			<td>LIFE TECHNOLOGIES SAS</td>
			<td>Q32854</td>
			<td>DNA quantification</td>
		</tr>
		<tr>
			<td>Thermocycler</td>
			<td>Eppendorf</td>
			<td>Ep gradients</td>
			<td>PCR</td>
		</tr>
		<tr>
			<td>UVbox, eBOX VX5</td>
			<td>Vilber Lourmat</td>
			<td></td>
			<td>Agarose gel electrophoresis visualisation</td>
		</tr>
		<tr>
			<td>Water for injectable preparation</td>
			<td>Aguettant</td>
			<td>PROAMP</td>
			<td>PCR</td>
		</tr>
	</tbody>
</table>

following the dynamics of structural variants in experimentally evolved populations

Structural variants (SVs) (i.e., deletions, insertions, duplications, and inversions) are now known to play an important role in phenotypic variation, and consequently in processes such as disease determination or adaptation to a new environment. However, single-nucleotide variants receive much more attention than SVs, probably because they are easier to detect, and their phenotypic effects are easier to predict. The development of short- and long-read deep sequencing technologies have strongly improved the detection of SVs, but the quantification of their frequency from pooled sequencing (poolseq) data is still technically complex and expensive.
Here, we present a rather simple and inexpensive method, which allows researchers to follow the dynamics of SV allele frequency. As an example of application, we follow the frequency of an insertion sequence (IS) insertion in experimental evolution populations of bacteria. This method is based on the design of triplets of primers around the structural variant borders, such that the amplicons produced by amplification of the wild-type (WT) and derived alleles differ in size by at least 5%, and that their amplification efficiency is similar. The quantity of each amplicon is then determined by parallel capillary electrophoresis and normalized to a calibration curve. This method can be easily extended to the quantification of the frequency of other structural variants (deletions, duplications, and inversions) and to pool-seq approaches of natural populations, including within-patient pathogen populations.

Using DNA extracted from an ancestral clone and a hypermutator clone isolated from the S2.11 population at generation 1,000, we established the calibration curve shown in Figure 2. The actual mutant proportions from laboratory-prepared DNA mixes and measured by the parallel capillary electrophoresis instrument were linked by a linear relationship of slope 1.0706, with an R2 of 0.9705. Additionally, there was a good agreement between biological replicates; the standard deviation wa...

Watch this Scientific Journal Video about Following the Dynamics of Structural Variants in Experimentally Evolved Populations at JoVE.com

Following the Dynamics of Structural Variants in Experimentally Evolved Populations

We developed a cost-effective method to follow non-single nucleotide polymorphism allele dynamics that can easily be adapted to experimental evolution frozen archives. A triplet PCR technique was coupled with automated parallel capillary electrophoresis to quantify the relative frequency of an insertion allele over the course of experimental evolution.

Structural variants (SVs) (i.e., deletions, insertions, duplications, and inversions) are now known to play an important role in phenotypic variation, and ...

following-dynamics-structural-variants-experimentally-evolved

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Biology

896 Views.  University of Montpellier, CNRS, EPHE, IRD, Montpellier, France. We developed a cost-effective method to follow non-single nucleotide polymorphism allele dynamics that can easily be adapted to experimental evolution frozen archives. A triplet PCR technique was coupled with automated parallel capillary electrophoresis to quantify the relative frequency of an insertion allele over the course of experimental evolution.

Video: Following the Dynamics of Structural Variants in Experimentally Evolved Populations

Obtaining High Quality RNA from Single Cell Populations in Human Postmortem Brain Tissue

We proposed to investigate the gray matter reduction in the superior temporal gyrus seen in schizophrenia patients, by interrogating gene expression profiles of pyramidal neurons in layer III. It is well known that the cerebral cortex is an exceptionally heterogeneous structure comprising diverse regions, layers and cell types, each of which is characterized by distinct cellular and molecular compositions and therefore differential gene expression profiles. To circumvent the confounding effects of tissue heterogeneity, we used laser-capture microdissection (LCM) in order to isolate our specific cell-type i.e pyramidal neurons.
Approximately 500 pyramidal neurons stained with the Histogene staining solution were captured using the Arcturus XT LCM system. RNA was then isolated from captured cells and underwent two rounds of T7-based linear amplification using Arcturus/Molecular Devices kits. The Experion LabChip (Bio-Rad) gel and electropherogram indicated good quality a(m)RNA, with a transcript length extending past 600nt required for microarrays. The amount of mRNA obtained averaged 51&mu;g, with acceptable mean sample purity as indicated by the A260/280 ratio, of 2.5. Gene expression was profiled using the Human X3P GeneChip probe array from Affymetrix.

We describe a process using laser-capture microdissection to isolate and extract RNA from a homogeneous cell population, pyramidal neurons, in layer III of the superior temporal gyrus in postmortem human brains. We subsequently linearly amplify (T7-based) mRNA, and hybridize the sample to the Affymetrix human X3P microarray.

We proposed to investigate the gray matter reduction in the superior temporal gyrus seen in schizophrenia patients, by interrogating gene expression ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

Photobleaching Assays (FRAP &amp; FLIP) to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with good spatial and temporal resolution. Here we describe how to perform FRAP and FLIP assays of chromatin proteins, including H1 and HP1, in mouse embryonic stem (ES) cells. In a FRAP experiment, cells are transfected, either transiently or stably, with a protein of interest fused with the green fluorescent protein (GFP) or derivatives thereof (YFP, CFP, Cherry, etc.). In the transfected, fluorescing cells, an intense focused laser beam bleaches a relatively small region of interest (ROI). The laser wavelength is selected according to the fluorescent protein used for fusion. The laser light irreversibly bleaches the fluorescent signal of molecules in the ROI and, immediately following bleaching, the recovery of the fluorescent signal in the bleached area - mediated by the replacement of the bleached molecules with the unbleached molecules - is monitored using time lapse imaging. The generated fluorescence recovery curves provide information on the protein's mobility. If the fluorescent molecules are immobile, no fluorescence recovery will be observed. In a complementary approach, Fluorescence Loss in Photobleaching (FLIP), the laser beam bleaches the same spot repeatedly and the signal intensity is measured elsewhere in the fluorescing cell. FLIP experiments therefore measure signal decay rather than fluorescence recovery and are useful to determine protein mobility as well as protein shuttling between cellular compartments. Transient binding is a common property of chromatin-associated proteins. Although the major fraction of each chromatin protein is bound to chromatin at any given moment at steady state, the binding is transient and most chromatin proteins have a high turnover on chromatin, with a residence time in the order of seconds. These properties are crucial for generating high plasticity in genome expression1. Photobleaching experiments are therefore particularly useful to determine chromatin plasticity using GFP-fusion versions of chromatin structural proteins, especially in ES cells, where the dynamic exchange of chromatin proteins (including heterochromatin protein 1 (HP1), linker histone H1 and core histones) is higher than in differentiated cells2,3.

Photobleaching Assays FRAP & FLIP to Measure Chromatin Protein Dynamics in Living Embryonic Stem Cells

We describe photobleaching methods including Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) to monitor chromatin protein dynamics in embryonic stem (ES) cells. Chromatin protein dynamics, which is considered to be one of the means to study chromatin plasticity, is enhanced in pluripotent cells.

Fluorescence Recovery After Photobleaching (FRAP) and Fluorescence Loss In Photobleaching (FLIP) enable the study of protein dynamics in living cells with ...

Selective Capture of 5-hydroxymethylcytosine from Genomic DNA

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene expression, maintenance of genome integrity, parental imprinting, X-chromosome inactivation, regulation of development, aging, and cancer1. Recently, the presence of an oxidized 5-mC, 5-hydroxymethylcytosine (5-hmC), was discovered in mammalian cells, in particular in embryonic stem (ES) cells and neuronal cells2-4. 5-hmC is generated by oxidation of 5-mC catalyzed by TET family iron (II)/&alpha;-ketoglutarate-dependent dioxygenases2, 3. 5-hmC is proposed to be involved in the maintenance of embryonic stem (mES) cell, normal hematopoiesis and malignancies, and zygote development2, 5-10. To better understand the function of 5-hmC, a reliable and straightforward sequencing system is essential. Traditional bisulfite sequencing cannot distinguish 5-hmC from 5-mC11. To unravel the biology of 5-hmC, we have developed a highly efficient and selective chemical approach to label and capture 5-hmC, taking advantage of a bacteriophage enzyme that adds a glucose moiety to 5-hmC specifically12.
Here we describe a straightforward two-step procedure for selective chemical labeling of 5-hmC. In the first labeling step, 5-hmC in genomic DNA is labeled with a 6-azide-glucose catalyzed by &beta;-GT, a glucosyltransferase from T4 bacteriophage, in a way that transfers the 6-azide-glucose to 5-hmC from the modified cofactor, UDP-6-N3-Glc (6-N3UDPG). In the second step, biotinylation, a disulfide biotin linker is attached to the azide group by click chemistry. Both steps are highly specific and efficient, leading to complete labeling regardless of the abundance of 5-hmC in genomic regions and giving extremely low background. Following biotinylation of 5-hmC, the 5-hmC-containing DNA fragments are then selectively captured using streptavidin beads in a density-independent manner. The resulting 5-hmC-enriched DNA fragments could be used for downstream analyses, including next-generation sequencing.
Our selective labeling and capture protocol confers high sensitivity, applicable to any source of genomic DNA with variable/diverse 5-hmC abundances. Although the main purpose of this protocol is its downstream application (i.e., next-generation sequencing to map out the 5-hmC distribution in genome), it is compatible with single-molecule, real-time SMRT (DNA) sequencing, which is capable of delivering single-base resolution sequencing of 5-hmC.

Described is a two-step labeling process using &beta;-glucosyltransferase (&beta;-GT) to transfer an azide-glucose to 5-hmC, followed by click chemistry to transfer a biotin linker for easy and density-independent enrichment. This efficient and specific labeling method enables enrichment of 5-hmC with extremely low background and high-throughput epigenomic mapping via next-generation sequencing.

5-methylcytosine (5-mC) constitutes ~2-8% of the total cytosines in human genomic DNA and impacts a broad range of biological functions, including gene ...

Robust 3D DNA FISH Using Directly Labeled Probes

3D DNA FISH has become a major tool for analyzing three-dimensional organization of the nucleus, and several variations of the technique have been published. In this article we describe a protocol which has been optimized for robustness, reproducibility, and ease of use. Brightly fluorescent directly labeled probes are generated by nick-translation with amino-allyldUTP followed by chemical coupling of the dye. 3D DNA FISH is performed using a freeze-thaw step for cell permeabilization and a heating step for simultaneous denaturation of probe and nuclear DNA. The protocol is applicable to a range of cell types and a variety of probes (BACs, plasmids, fosmids, or Whole Chromosome Paints) and allows for high-throughput automated imaging. With this method we routinely investigate nuclear localization of up to three chromosomal regions.

We describe a robust and versatile protocol for analyzing nuclear architecture by 3D DNA FISH using directly labeled fluorescent probes.

3D DNA FISH has become a major tool for  analyzing three-dimensional organization of the nucleus, and several variations  of the technique have been ...

The Slice Culture Method for Following Development of Tooth Germs In Explant Culture

Explant culture allows manipulation of developing organs at specific time points&#160;and is therefore an important method for the developmental biologist. For many organs it is difficult to access developing tissue to allow monitoring during ex vivo culture. The slice culture method allows access to tissue so that morphogenetic movements can be followed and specific cell populations can be targeted for manipulation or lineage tracing.
In this paper we describe a method of slice culture that has been very successful for culture of tooth germs in a range of species. The method provides excellent access to the tooth germs, which develop at a similar rate to that observed in vivo, surrounded by the other jaw tissues. This allows tissue interactions between the tooth and surrounding tissue to be monitored. Although this paper concentrates on tooth germs, the same protocol can be applied to follow development of a number of other organs, such as salivary glands, Meckel&#39;s cartilage, nasal glands, tongue, and ear.

Here we detail a method to culture tooth germs in mandible slices using a tissue chopper. This method allows unique access to the tooth during development, providing excellent opportunity for manipulation and lineage tracing, not available using more traditional culture methods.

Explant culture allows manipulation of developing organs at specific time points&#160;and is therefore an important method for the developmental ...

Aip1p Dynamics Are Altered by the R256H Mutation in Actin

Mutations in actin cause a range of human diseases due to specific molecular changes that often alter cytoskeletal function. In this study, imaging of fluorescently tagged proteins using total internal fluorescence (TIRF) microscopy is used to visualize and quantify changes in cytoskeletal dynamics. TIRF microscopy and the use of fluorescent tags also allows for quantification of the changes in cytoskeletal dynamics caused by mutations in actin. Using this technique, quantification of cytoskeletal function in live cells valuably complements in vitro studies of protein function. As an example, missense mutations affecting the actin residue R256 have been identified in three human actin isoforms suggesting this amino acid plays an important role in regulatory interactions. The effects of the actin mutation R256H on cytoskeletal movements were studied using the yeast model. The protein, Aip1, which is known to assist cofilin in actin depolymerization, was tagged with green fluorescent protein (GFP) at the N-terminus and tracked in vivo using TIRF microscopy. The rate of Aip1p movement in both wild type and mutant strains was quantified. In cells expressing R256H mutant actin, Aip1p motion is restricted and the rate of movement is nearly half the speed measured in wild type cells (0.88 &#177; 0.30 &#956;m/sec in R256H cells compared to 1.60 &#177; 0.42 &#956;m/sec in wild type cells, p &#60; 0.005).

Disease-causing mutations in actin can alter cytoskeletal function. Cytoskeletal dynamics are quantified through imaging of fluorescently tagged proteins using total internal fluorescence microscopy. As an example, the cytoskeletal protein, Aip1p, has altered localization and movement in cells expressing the mutant actin isoform, R256H.

Mutations in actin cause a range of human diseases due to specific molecular changes that often alter cytoskeletal function. In this study, imaging of ...

Isolating Potentiated Hsp104 Variants Using Yeast Proteinopathy Models

Many protein-misfolding disorders can be modeled in the budding yeast Saccharomyces cerevisiae. Proteins such as TDP-43 and FUS, implicated in amyotrophic lateral sclerosis, and &#945;-synuclein, implicated in Parkinson&#8217;s disease, are toxic and form cytoplasmic aggregates in yeast. These features recapitulate protein pathologies observed in patients with these disorders. Thus, yeast are an ideal platform for isolating toxicity suppressors from libraries of protein variants. We are interested in applying protein disaggregases to eliminate misfolded toxic protein conformers. Specifically, we are engineering Hsp104, a hexameric AAA+ protein from yeast that is uniquely capable of solubilizing both disordered aggregates and amyloid and returning the proteins to their native conformations. While Hsp104 is highly conserved in eukaryotes and eubacteria, it has no known metazoan homologue. Hsp104 has only limited ability to eliminate disordered aggregates and amyloid fibers implicated in human disease. Thus, we aim to engineer Hsp104 variants to reverse the protein misfolding implicated in neurodegenerative disorders. We have developed methods to screen large libraries of Hsp104 variants for suppression of proteotoxicity in yeast. As yeast are prone to spontaneous nonspecific suppression of toxicity, a two-step screening process has been developed to eliminate false positives. Using these methods, we have identified a series of potentiated Hsp104 variants that potently suppress the toxicity and aggregation of TDP-43, FUS, and &#945;-synuclein. Here, we describe this optimized protocol, which could be adapted to screen libraries constructed using any protein backbone for suppression of toxicity of any protein that is toxic in yeast.

Yeast proteinopathy models are valuable tools to assess the toxicity and aggregation of proteins implicated in disease. Here, we present methods for screening Hsp104 variant libraries for toxicity suppressors. This protocol could be adapted to screen any protein library for toxicity suppressors of any protein that is toxic in yeast.

Many protein-misfolding disorders can be modeled in the budding yeast Saccharomyces cerevisiae. Proteins such as TDP-43 and FUS, implicated in amyotrophic ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Studying Ribonucleotide Incorporation: Strand-specific Detection of Ribonucleotides in the Yeast Genome and Measuring Ribonucleotide-induced Mutagenesis

The presence of ribonucleotides in nuclear DNA has been shown to be a source of genomic instability. The extent of ribonucleotide incorporation can be assessed by alkaline hydrolysis and gel electrophoresis as RNA is highly susceptible to hydrolysis in alkaline conditions. This, in combination with Southern blot analysis can be used to determine the location and strand into which the ribonucleotides have been incorporated. However, this procedure is only semi-quantitative and may not be sensitive enough to detect small changes in ribonucleotide content, although strand-specific Southern blot probing improves the sensitivity. As a measure of one of the most striking biological consequences of ribonucleotides in DNA, spontaneous mutagenesis can be analyzed using a forward mutation assay. Using appropriate reporter genes, rare mutations that results in the loss of function can be selected and overall and specific mutation rates can be measured by combining data from fluctuation experiments with DNA sequencing of the reporter gene. The fluctuation assay is applicable to examine a wide variety of mutagenic processes in specific genetic background or growth conditions.

Ribonucleotides are among the most abundant non-canonical nucleotides incorporated into the genome during eukaryotic nuclear DNA replication. If not properly removed, ribonucleotides can cause DNA damage and mutagenesis. Here, we present two experimental approaches that are used to assess the abundance of ribonucleotide incorporation into DNA and its mutagenic effects.

The presence of ribonucleotides in nuclear DNA has been shown to be a source of genomic instability. The extent of ribonucleotide incorporation can be ...