Name: optogenetic random mutagenesis using histone-minisog in c. elegans
Uploaded: 2016-11-14T15:00:00.000+00:00
Duration: 4 min 51 s
Description: Watch this Scientific Journal Video about Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans at JoVE.com

The work is supported by HHMI. We thank our lab members for their help with testing the protocol and revising the manuscript.

1. Construction of the LED Illuminator
NOTE: The required LED equipment is summarized in the List of Materials. The entire LED setup is small and can be placed anywhere in the lab, although we recommend it be placed in a dark room to control light exposure of worms4.
<ol>
	<li>Connect the LED to a controller with cables (Figure&#160;1A,&#160;D).</li>
	<li>Connect the controller to a digital function generator/amplifier with a BNC cable (Figure 1A, C, D).</li>
	<li>Fix the LED light 10 cm above the stage using a custom holder (Figure 1A). 
	NOTE: We made the holder with metal parts.</li>
	<li>Cover the LED illumination setup partially, but avoid heat accumulation (Figure 1B), which makes worms sick. 
	NOTE: We use a custom-made hard plastic cover with open top and bottom (Figure 1A). The cover is not required for the mutagenesis itself, but is used to limit unnecessary exposure of the blue light to the surroundings. We advise to not cover the LED setup entirely (Figure 1B) to prevent overheating the worms.</li>
	<li>Wear laser-protective glasses. 
	NOTE: We wear laser-protective glasses because the light is very bright. Sunglasses may work as well.</li>
	<li>Switch the lever of the LED controller to &#39;Int.&#39;&#160;for continuous illumination (Figure 1D) and set it at 65% of the maximum power (Figure 1C).</li>
	<li>Use a photometer to measure the blue light intensity on the stage where the worms are placed using manufacturer&#39;s protocol. The setup gives 2.0 mW/mm2 under continuous illumination.</li>
</ol>
2. Blue Light Treatment
NOTE: Use the strain CZ20310 juSi164[Pmex-5-HIS-72::miniSOG-3&#39;UTR(tbb-2) + Cb-unc-119(+)] unc-119(ed3)III4. The strain is available at the Caenorhabditis Genetics Center (CGC, http://www.cgc.cbs.umn.edu/).
<ol>
	<li>Maintain the His-mSOG strain in the dark on standard nematode growth medium (NGM) plates with E. coli OP50 following a standard protocol10,11. 
	NOTE: Under ambient light, juSi164-containing strains do not display a mutation rate above a spontaneous rate4. Routine strain passage using a dissecting scope with a halogen lamp generally does not cause mutations as well. Nonetheless, care should be taken to avoid unnecessary exposure to light, e.g., keeping the strains in a standard dark-inside incubator, or covering the strains with foil. It is advised to freeze aliquots of the strain after receiving it in case unnecessary mutations accumulate over time.</li>
	<li>Pick gravid young adult (approximately 12 h&#160;post-L4) animals with a worm pick11. 
	NOTE: Younger animals show less mutagenic effect4, while older animals may have lower brood size.</li>
	<li>Cut a filter paper to fit a 60 mm plate with a 25 mm x 25 mm square hole and place onto an unseeded plate (Figure 1E). 
	NOTE: A square punch can be used but the&#160;size of the punch does not have to be precise.</li>
	<li>Drop 100 &#181;L of 100 mM CuCl2 on the filter paper evenly to restrict worms within the square hole on the plate.</li>
	<li>Pick desired number of gravid young adult hermaphrodites (see Step 3 for an example) and transfer to the center of the CuCl2 plate.</li>
	<li>Wear laser protective glasses.</li>
	<li>Switch ON the LED and the function generator.</li>
	<li>Switch the lever of the LED controller to &#39;Ext.&#39;&#160;to control it by the function generator (Figure 1D).</li>
	<li>Set the controller at 65% of the maximum power and the function generator as sine wave, 4 Hz (Figure 1C).</li>
	<li>Place the CuCl2 plate from Step 2.5 without a lid on the stage under the LED (Figure 1A).</li>
	<li>Illuminate the animals with blue light for 30 min.</li>
	<li>Transfer healthy looking worms to a seeded plate as P0. 
	NOTE: The desired number of P0 depends on the type of screen. See Step 3 for an example. The P0 worms may appear to be uncoordinated immediately after light treatment and will recover over time.</li>
	<li>Keep worms covered using foil in an incubator or in the dark; and follow the standard procedure to begin screening desired phenotypes among their progeny2,10.</li>
</ol>
3. Example of a Forward Genetic Screen
NOTE: This is an example of the clonal screen with 120 haploid genomes to check if the LED and strain are working. Figure 2 shows a schematic of the procedure.
<ol>
	<li>[Day1] After blue light treatment, pick five healthy looking worms onto a NGM plate with OP50 as P0, keep them in a 20 &#176;C incubator, and let them lay eggs for 1 d (&#39;Day1&#39; plate).</li>
	<li>[Day2] Transfer P0 onto a new seeded plate (&#39;Day2&#39; plate).</li>
	<li>[Day3] Pick and kill P0 on the &#39;Day2&#39; plate by burning them.</li>
	<li>[Day4] Pick 30 gravid young adult F1 from the &#39;Day1&#39; plate and transfer them onto individual plates (30 F1 plates for &#39;Day1&#39;).</li>
	<li>[Day5] Repeat Step 3.4 for the &#39;Day2&#39; plate (30 F1 plates for &#39;Day2&#39;).</li>
	<li>[Day7-9] Check the abnormal morphologies and/or behavior of F2 worms compared to the WT strain (Figure 3A) under a standard stereomicroscope when they become adults. 
	NOTE: A heritable mutation displays about a quarter of worms with the same phenotype among F2 when it is recessive with high penetrance. However, some mutants may grow slowly and/or show partial penetrance. Therefore, it is possible that less than a quarter of the mutants can be found on a plate.</li>
	<li>Pick worms with visible phenotypes individually and check the heritability of the phenotype in the next generation.</li>
</ol>
4. Example of an Extrachromosomal Transgene Integration
<ol>
	<li>Inject the DNA of interest into CZ20310 juSi164[Pmex-5-HIS-72::miniSOG-3&#39;UTR(tbb-2) + Cb-unc-119(+)] unc-119(ed3) and prepare transgenic worms with a low transmission rate (10-30%) following a standard injection protocol12,13. 
	NOTE: Alternatively, established extrachromosomal arrays can be crossed into the juSi164 strains. To avoid genotyping for juSi164, juSi164 heterozygotes may be used as the P0 although efficiency might be decreased. The genotyping method for juSi164 is described in Section 5.</li>
	<li>[Day1] Treat ~20 transgenic animals as P0 with blue light as described in Step 2. 
	NOTE: More P0 worms are necessary depending on the transmission rate. Transgenes can be identified by phenotypes or coinjection marker12,13.</li>
	<li>Pick 10 healthy looking P0 onto individual plates, keep them in a 20 &#176;C incubator, and let them lay eggs for a few days (P0 plates)</li>
	<li>[Day4] Single out 20 transgenic F1 worms from each P0 plate onto individual plates and let them lay eggs. (20 F1 x 10 P0 plates= 200 F1 plates in total.)</li>
	<li>[Day7] Find F1 plates with &#62;75% F2 having transgenes.</li>
	<li>Pick five transgenic animals from the &#62;75% transmission rate F1 plates (F2 plates).</li>
	<li>[Day10] Keep F2 plates with 100% transmission rate as potential integrated lines.</li>
	<li>Outcross the potential integrated lines using a standard procedure10 to check the Mendelian segregation and to remove potential background mutations.</li>
</ol>
5. Genotyping
<ol>
	<li>Lyse 20 outcrossed F2 worms in lysis buffer using a standard protocol14.</li>
	<li>Perform PCR using the primers for MosSCI insertion9, juSi164, (Table 1) with an annealing temperature of 58 &#176;C following a standard protocol14. 
	NOTE: It is unnecessary to genotype unc-119(ed3) because unc-119(ed3) can be detected by uncoordinated behavior after removing the rescuing transgene (juSi164) in the following Steps 5.4 and 5.5.</li>
	<li>Run agarose gel electrophoresis15 and examine the band sizes (Table 1) to find the WT&#160;for juSi164.</li>
	<li>Examine if the F2 progeny has paralyzed worms due to the existence of unc-119(ed3).</li>
	<li>If uncoordinated worms are found on step 5.4, single several nonuncoordinated worms to obtain all nonuncoordinated worms in the next generation.</li>
</ol>

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The authors have nothing to disclose.

Here, we describe a detailed procedure of optogenetic mutagenesis using His-mSOG expressed in the germline and provide an example of a small scale forward genetic screen. Compared to standard chemical mutagenesis, this optogenetic mutagenesis method has several advantages. Firstly, it is nontoxic, thereby keeping lab personnel away from toxic chemical mutagens such as EMS, ENU and trimethylpsoralen with ultraviolet light (TMP/UV). Secondly, the experimental procedure of mutagenesis can be used for a small number of the P0 animals and can be finished within one hour. Thirdly, the optogenetic mutagenesis produces a broad spectrum of mutations including nucleotide substitutions&#160;and chromosome deletions. Finally, the optogenetic mutagenesis can be used for integration of extrachromosomal transgenes.
The critical step in this mutagenesis protocol is to ensure that the LED setup works on His-mSOG containing strains at the expected efficiency. We recommend performing a pilot clonal screen, as described in Protocol Section 3, with about 100 haploid genomes. The mutagenesis is easily scalable from the small scale pilot screens to nonclonal screens by increasing the number of P0 for the light treatment. For large-scale nonclonal screens, we suggest synchronizing the worms to gravid young adults and transferring hundreds of them with M9 solution onto a CuCl2 plate for light treatment. The standard condition described in Steps 2 and 3 is estimated to have approximately 4.4 times lower efficiency than the widely used 50 mM EMS mutagenesis method4. The mutation rate may be improved by increasing the light exposure time and/or increasing the intensity of the light. However, these modifications can cause phototoxicity and lead to small brood size and sickness of light-treated worms. To increase the efficiency, it may be possible to use miniSOG derivatives with higher ROS yield such as miniSOG(Q103L) although it may increase the background mutagenesis effect without light treatment16,17. For the modification of the DNA construct, the Pmex-5-HIS-72::miniSOG-3&#39;UTR(tbb-2) plasmid (pCZ886) is available commercially.
Under the conditions described here, integrated transgenes are generated at approximately one&#160;line per 200 F1 on average4 when using an extrachromosomal transgenic strain with a transmission rate of 10 - 30%. This efficiency may be increased by using extrachromosomal transgenic strains with high transmission rates as previously reported for a UV radiation method18.
His-mSOG-mediated optogenetic mutagenesis induces mutations in a wide range of genes. Also, the frequency of loci recovered from the rpm-1 suppressor screen using optogenetic mutagenesis is similar to those from the same screen using EMS4. However, it is possible that optogenetic mutagenesis using a histone gene may have a bias towards certain genes, due to unknown factors such as chromatin structure. To address this issue, it is necessary to analyze many optogenetically induced mutations by whole genome sequencing.
His-mSOG-mediated optogenetic mutagenesis causes a wide spectrum of mutations including single nucleotide substitutions and deletions ranging from 1 bp to a few hundred bp4. The spectrum of single nucleotide variants suggests that miniSOG-induced reactive oxygen species are the cause of the mutations4. It is also possible that miniSOG damages histone proteins and causes chromatin instability because miniSOG is used to inactivate proteins as well19. It may be possible to manipulate the type of mutations by using light of different intensities. Since optogenetic mutagenesis causes deletions, it is important to analyze deletions as well as single nucleotide changes in whole genome sequencing data&#160;for mapping20.
It was previously reported that an epifluorescent microscope is used for miniSOG-mediated cell ablation21,22. The light intensity of a standard epifluorescent compound microscope was measured to be ~0.5 mW/mm2 without a lens, which is about 4x lower than that used in the standard optogenetic mutagenesis protocol. Thus, it is highly advisable to empirically determine the efficiency when using an epi-fluorescent microscope as a light source. One advantage of the LED is the stability over thousands of hours. Previous studies have shown that miniSOG is more potent in ROS-mediated cell ablation under pulsed light22, although the precise mechanism remains to be determined. We used a digital function generator/amplifier to make pulsed light. Another option is to connect the light source to a computer using a USB-TTL interface (Figure 1A) to regulate the LED light by a program. The LED system described here can also be used for other purposes, such as optogenetic control of excitable cells using channelrhodopsin23, cell ablation using mitochondria- or cell-membrane-targeted miniSOG17,22, and Chromophore-Assisted Light Inactivation (CALI) of proteins19. Furthermore, this optogenetic mutagenesis has the potential to be applied to other organisms, such as bacteria, yeast, fly, and zebrafish, as well as mammalian cell lines.

Forward genetic screens have been widely used to generate genetic mutants disrupting various biological processes in model organisms such as Caenorhabditis elegans (C. elegans)1. Analyses of such mutants lead to the discovery of functionally important genes and their signaling pathways1,2. Traditionally, mutagenesis in C. elegans is achieved using mutagenic chemicals, radiation, or transposons2. Chemicals such as ethyl methanesulfonate (EMS) and N-ethyl-N-nitrosourea (ENU) are toxic to humans; gamma-ray or ultraviolet (UV) radiation mutagenesis requires special equipment; and transposon-active strains, such as mutator strains3, can cause unnecessary mutations during maintenance. We have developed a simple approach to induce heritable mutations using a genetically-encoded photosensitizer4.
Reactive Oxygen Species (ROS) can damage DNA5. The mini Singlet Oxygen Generator (miniSOG) is a green fluorescent protein of 106 amino acids that was engineered from the LOV (Light, Oxygen, and Voltage) domain of Arabidopsis phototropin 2 6. Upon exposure to blue light (~450 nm), miniSOG generates ROS including singlet oxygen with &#64258;avin mononucleotide as a cofactor6-8, which is present in all cells. We constructed a His-mSOG fusion protein by tagging miniSOG to the C-terminus of histone-72, a C. elegans variant of Histone 3. We generated a single-copy transgene9 to express His-mSOG in the germline of&#160;C. elegans. Under normal culture conditions in the dark, the His-mSOG transgenic worms have normal brood size and life span4. Upon exposure to blue LED light, the His-mSOG worms produce heritable mutations among their progeny4. The spectrum of induced mutations includes nucleotide changes, such as G:C to T:A and G:C to C:G, and small chromosome deletions4. This mutagenesis procedure is simple to perform, and requires minimal lab safety setup. Here, we describe the LED illumination setup and procedures for optogenetic mutagenesis.

<table><tbody><tr><td>Ultra High Power LED light source</td><td>Prizmatix</td><td>UHP-mic-LED-460</td><td>460 &amp;plusmn; 5 nm</td></tr><tr><td>LED controller</td><td>Prizmatix</td><td>UHPLCC-01</td><td></td></tr><tr><td>Digital function generator/amplifier</td><td>PASCO</td><td>PI-9587C</td><td>PI-9587C is no longer available. The replacement is PI-8127.</td></tr><tr><td>BNC cable male/male</td><td>THORLABS</td><td>CA3136</td><td></td></tr><tr><td>USB-TTL interface</td><td>Prizmatix</td><td></td><td>Optional</td></tr><tr><td>Photometer</td><td>THORLABS</td><td>PM50 and Model D10MM</td><td></td></tr><tr><td>Filter paper</td><td>Whatman</td><td>1001-110</td><td></td></tr><tr><td>Stereomicroscope</td><td>Leica</td><td>MZ95</td><td></td></tr><tr><td>NGM plate</td><td></td><td></td><td>Dissolve 5 g NaCl, 2.5 g Peptone, 20 g Agar, 10 &amp;micro;g/mL cholesterol in 1 L H&lt;sub&gt;2&lt;/sub&gt;O. After autoclaving, add 1 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, 1 mM MgSO&lt;sub&gt;4&lt;/sub&gt;, 25 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;(pH 6.0).&amp;nbsp;</td></tr><tr><td>Lysis solution</td><td></td><td></td><td>10 mM Tris(pH 8.8), 50 mM KCl, 0.1% Triton X-100, 2.5 mM MgCl&lt;sub&gt;2&lt;/sub&gt;, 100 &amp;micro;g/mL Proteinase K</td></tr></tbody></table>

optogenetic random mutagenesis using histone-minisog in c. elegans

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most mutagenesis-based screens, researchers have relied on the use of toxic chemicals, carcinogens, or irradiation, which requires designated equipment, safety setup, and/or disposal of hazardous materials. We have developed a simple approach to induce heritable mutations in C. elegans using germline-expressed histone-miniSOG, a light-inducible potent generator of reactive oxygen species. This mutagenesis method is free of toxic chemicals and requires minimal laboratory safety and waste management. The induced DNA modifications include single-nucleotide changes and small deletions, and complement those caused by classical chemical mutagenesis. This methodology can also be used to induce integration of extrachromosomal transgenes. Here, we provide the details of the LED setup and protocols for standard mutagenesis and transgene integration.

We performed a forward genetic screen with CZ20310 juSi164 unc-119(ed3) using 30 min light exposure following the standard protocol described in Sections 2 and 3. Among 60 F1 plates corresponding to 120 mutagenized haploid genomes, we found 8 F1 plates with F2 worms displaying visible phenotypes such as body size defect (Figure 3B), uncoordinated movement (Figure 3C), and adult lethality (Figure 3D). We did not notice any difference in the number of mutants between &#39;Day1&#39;&#160;and &#39;Day2&#39; plates. All progeny of singled small F2 worms and uncoordinated F2 worms showed the corresponding phenotypes, suggesting that they are heritable mutants. The full observation is summarized in Table 2. On average, a clonal screen with 120 haploid genomes will result in about 10 F1 plates containing worms with visibly detectable phenotypes such as body shape and locomotion that easily spotted under a stereomicroscope. More quantitative analysis is necessary to determine the mutation rate precisely4. The expected number of visible mutants from this screen suggests that the strain and setup are working properly for optogenetic mutagenesis.
<img alt="Figure 1" src="/files/ftp_upload/54810/54810fig1.jpg" /> 
Figure 1: The LED Setup. (A) The whole system. The LED was mounted on a custom-made holder. Details are found in the List of Materials. (B) The cover is made of plastic. It does not cover the system completely to avoid heat accumulation. (C) Controller and function generator. (D) The back side of the system. (E) An unseeded plate with CuCl2-soaked filter paper for blue light treatment. <a href="http://cloudflare.jove.com/files/ftp_upload/54810/54810fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54810/54810fig2.jpg" /> 
Figure 2: Experimental Procedure for a Clonal Screen. After blue light treatment, P0 worms are transferred to a seeded plate and allowed to lay eggs. P0 worms are transferred to a new seeded plate 1 d after light treatment and killed 2 d&#160;after light treatment. F1 worms are singled from the P0 plates. Phenotypes of F2 worms are examined after they become adults. <a href="http://cloudflare.jove.com/files/ftp_upload/54810/54810fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54810/54810fig3.jpg" /> 
Figure 3: Example of Mutants. (A) Original strain for optogenetic mutagenesis: CZ20310 juSi164 unc-119(ed3). This strain has normal body shape, behavior, and brood size. Scale bar: 0.5 mm. (B - D) F2 progeny showing body size defect&#160;(B), uncoordinated (C), and lethal (D) phenotypes. <a href="http://cloudflare.jove.com/files/ftp_upload/54810/54810fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Table 1" src="/files/ftp_upload/54810/54810table1.jpg" /> 
Table 1: Genotyping of juSi164 and unc-119(ed3).
<img alt="Table 2" src="/files/ftp_upload/54810/54810table2.jpg" /> 
Table 2: Example of Phenotypes from a Clonal Screen.

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Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Genetically-encoded histone-miniSOG induces genome-wide heritable mutations in a blue&#160;light-dependent manner. This mutagenesis method is simple, fast, free of toxic chemicals, and well-suited for forward genetic screening and transgene integration.

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most ...

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9.0K Views.  University of California in San Diego and Howard Hughes Medical Institute. Genetically-encoded histone-miniSOG induces genome-wide heritable mutations in a blue light-dependent manner. This mutagenesis method is simple, fast, free of toxic chemicals, and well-suited for forward genetic screening and transgene integration.

Video: Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

We describe a PCR- and homologous recombination-based system for generating targeted mutations in histone genes in budding yeast cells. The resulting mutant alleles reside at their endogenous genomic sites and no exogenous DNA sequences are left in the genome following the procedure. Since in haploid yeast cells each of the four core histone proteins is encoded by two non-allelic genes with highly homologous open reading frames (ORFs), targeting mutagenesis specifically to one of two genes encoding a particular histone protein can be problematic. The strategy we describe here bypasses this problem by utilizing sequences outside, rather than within, the ORF of the target genes for the homologous recombination step. Another feature of this system is that the regions of DNA driving the homologous recombination steps can be made to be very extensive, thus increasing the likelihood of successful integration events. These features make this strategy particularly well-suited for histone gene mutagenesis, but can also be adapted for mutagenesis of other genes in the yeast genome.

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

A strategy for generating mutations in histone genes at their endogenous location in Saccharomyces cerevisiae is presented.

We describe a PCR- and homologous recombination-based system for generating targeted mutations in histone genes in budding yeast cells. The resulting ...

Gene-targeted Random Mutagenesis to Select Heterochromatin-destabilizing Proteasome Mutants in Fission Yeast

Random mutagenesis of a target gene is commonly used to identify mutations that yield the desired phenotype. Of the methods that may be used to achieve random mutagenesis, error-prone PCR is a convenient and efficient strategy for generating a diverse pool of mutants (i.e., a mutant library). Error-prone PCR is the method of choice when a researcher seeks to mutate a pre-defined region, such as the coding region of a gene while leaving other genomic regions unaffected. After the mutant library is amplified by error-prone PCR, it must be cloned into a suitable plasmid. The size of the library generated by error-prone PCR is constrained by the efficiency of the cloning step. However, in the fission yeast, Schizosaccharomyces pombe, the cloning step can be replaced by the use of a highly efficient one-step fusion PCR to generate constructs for transformation. Mutants of desired phenotypes may then be selected using appropriate reporters. Here, we describe this strategy in detail, taking as an example, a reporter inserted at centromeric heterochromatin.

This article describes a detailed methodology for random mutagenesis of a target gene in fission yeast. As an example, we target rpt4+, which encodes a subunit of the 19S proteasome, and screen for mutations that destabilize heterochromatin.

Random mutagenesis of a target gene is commonly used to identify mutations that yield the desired phenotype. Of the methods that may be used to achieve ...

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been co-opted as a powerful tool for precise eukaryotic genome engineering. Here, we present a rapid and simple method using chimeric single guide RNAs (sgRNA) and CRISPR-Cas9 Ribonucleoproteins (RNPs) for the efficient and precise generation of genomic point mutations in C. elegans. We describe a pipeline for sgRNA target selection, homology-directed repair (HDR) template design, CRISPR-Cas9-RNP complexing and delivery, and a genotyping strategy that enables the robust and rapid identification of correctly edited animals. Our approach not only permits the facile generation and identification of desired genomic point mutant animals, but also facilitates the detection of other complex indel alleles in approximately 4 - 5 days with high efficiency and a reduced screening workload.

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Here, we present a method to engineer the genome of C. elegans using CRISPR-Cas9 ribonucleoproteins and homology dependent repair templates.

The clustered regularly interspersed palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) prokaryotic adaptive immune defense system has been ...

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain and protect cellular identities. The conversion of germ cells into specific neurons in the nematode Caenorhabditis elegans (C. elegans) is a powerful tool for performing genetic screens in order to dissect regulatory pathways that safeguard established cell identities. Reprogramming of germ cells to a specific type of neurons termed ASE requires transgenic animals that allow broad over-expression of the Zn-finger transcription factor (TF) CHE-1. Endogenous CHE-1 is expressed exclusively in two head neurons and is required to specify the glutamatergic ASE neurons fate, which can easily be visualized by the gcy-5prom::gfp reporter. A trans gene containing the heat-shock promoter-driven che-1 gene expression construct allows broad mis-expression of CHE-1 in the entire animal upon heat-shock treatment. The combination of RNAi against the chromatin-regulating factor LIN-53 and heat-shock-induced che-1 over-expression leads to reprogramming of germ cell into ASE neuron-like cells. We describe here the specific RNAi procedure and appropriate conditions for heat-shock treatment of transgenic animals in order to successfully induce germ cell to neuron conversion.

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

This protocol describes how to study cellular processes during cell fate conversion in Caenorhabditis elegans in vivo. Using transgenic animals, allowing heat-shock promoter-driven overexpression of the neuron fate-inducing transcription factor CHE-1 and RNAi-mediated depletion of the chromatin-regulating factor LIN-53 germ cell to neuron reprogramming can be observed in vivo.

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain ...

Developmental Biology

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Neuroscience

An Introduction to Worm Lab: from Culturing Worms to Mutagenesis

This protocol describes procedures to maintain nematodes in the laboratory and how to mutagenize them using two alternative methods: ethyl methane sulfonate (EMS) and 4, 5', 8-trimethylpsoralen combined with ultraviolet light (TMP/UV). Nematodes are powerful biological systems for genetics studies because of their simple body plan and mating system, which is composed of self-fertilizing hermaphrodites and males that can generate hundreds of progeny per animal. Nematodes are maintained in agar plates containing a lawn of bacteria and can be easily transferred from one plate to another using a pick. EMS is an alkylating agent commonly used to induce point mutations and small deletions, while TMP/UV mainly induces deletions. Depending on the species of nematode being used, concentrations of EMS and TMP will have to be optimized. To isolate recessive mutations of the nematode Pristionchus pacificus, animals of the F2 generation were visually screened for phenotypes. To illustrate these methods, we mutagenized worms and looked for Uncoordinated (Unc), Dumpy (Dpy) and Transformer (Tra) mutants.

Screening for mutants with phenotypic defects is a straightforward method for identifying genes that function in a given biological process. In this article we describe how to culture free living worms (e.g., Pristionchus pacificus) in the laboratory and show two different mutagenesis methods, EMS and TMP/UV.

This protocol describes procedures to maintain nematodes in the laboratory and how to mutagenize them using two alternative methods: ethyl methane ...

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using fluorescent-labeled reporter proteins or differential interference contrast (DIC) microscopy, allows for the examination of the temporal progression of these dynamic events which is otherwise inferred from analysis of fixed samples1,2. Moreover, the study of the developmental regulations of cellular processes necessitates conducting time-lapse experiments on an intact organism during development. The Caenorhabiditis elegans embryo is light-transparent and has a rapid, invariant developmental program with a known cell lineage3, thus providing an ideal experiment model for studying questions in cell biology4,5and development6-9. C. elegans is amendable to genetic manipulation by forward genetics (based on random mutagenesis10,11) and reverse genetics to target specific genes (based on RNAi-mediated interference and targeted mutagenesis12-15). In addition, transgenic animals can be readily created to express fluorescently tagged proteins or reporters16,17. These traits combine to make it easy to identify the genetic pathways regulating fundamental cellular and developmental processes in vivo18-21. In this protocol we present methods for live imaging of C. elegans embryos using DIC optics or GFP fluorescence on a compound epifluorescent microscope. We demonstrate the ease with which readily available microscopes, typically used for fixed sample imaging, can also be applied for time-lapse analysis using open-source software to automate the imaging process.

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

The C. elegans embryo is a powerful system for studying cell biology and development. We present a protocol for live imaging of C. elegans embryos utilizing DIC optics or fluorescence using readily available epifluorescent microscopes and open-source software.

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using ...

A Rapid Protocol for Integrating Extrachromosomal Arrays With High Transmission Rate into the C. elegans Genome

Microinjecting DNA into the cytoplasm of the syncytial gonad of Caenorhabditis elegans is the main technique used to establish transgenic lines that exhibit partial and variable transmission rates of extrachromosomal arrays to the next generation. In addition, transgenic animals are mosaic and express the transgene in a variable number of cells. Extrachromosomal arrays can be integrated into the C. elegans genome using UV irradiation to establish nonmosaic transgenic strains with 100% transmission rate of the transgene. To that extent, F1 progenies of UV irradiated transgenic animals are screened for animals carrying a heterozygous integration of the transgene, which leads to a 75% Mendelian transmission rate to the F2 progeny. One of the challenges of this method is to distinguish between the percentage of transgene transmission in a population before (X% transgenic animals) and after integration (&#8805;75% transgenic F2 animals). Thus, this method requires choosing a nonintegrated transgenic line with a percentage of transgenic animals that is significantly lower than the Mendelian segregation of 75%. Consequently, nonintegrated transgenic lines with an extrachromosomal array transmission rate to the next generation &#8804;60% are usually preferred for integration, and transgene integration in highly transmitting strains is difficult. Here we show that the efficiency of extrachromosomal arrays integration into the genome is increased when using highly transmitting transgenic lines (&#8805;80%). The described protocol allows for easy selection of several independent lines with homozygous transgene integration into the genome after UV irradiation of transgenic worms exhibiting a high rate of extrachromosomal array transmission. Furthermore, this method is quite fast and low material consuming. The possibility of rapidly generating different lines that express a particular integrated transgene is of great interest for studies focusing on gene expression pattern and regulation, protein localization, and overexpression, as well as for the development of subcellular markers.

A Rapid Protocol for Integrating Extrachromosomal Arrays With High Transmission Rate into the C. elegans Genome

This protocol describes a rapid and low material consuming procedure for the integration of transgenic extrachromosomal arrays into the Caenorhabditis elegans genome using ultra violet (UV) irradiation. Furthermore, this protocol is particularly well suited for transgenic lines that transmit extrachromosomal arrays at a high rate.

Microinjecting DNA into the cytoplasm of the syncytial gonad of Caenorhabditis elegans is the main technique used to establish transgenic lines that ...

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These age-associated disorders are characterized by the appearance of protein aggregates with fibrillary structure in the brains of these patients. Exactly why normally soluble proteins undergo an aggregation process remains poorly understood. The discovery that protein aggregation is not limited to disease processes and instead part of the normal aging process enables the study of the molecular and cellular mechanisms that regulate protein aggregation, without using ectopically expressed human disease-associated proteins. Here we describe methodologies to examine inherent protein aggregation in Caenorhabditis elegans through complementary approaches. First, we examine how to grow large numbers of age-synchronized C. elegans to obtain aged animals and we present the biochemical procedures to isolate highly-insoluble-large aggregates. In combination with a targeted genetic knockdown, it is possible to dissect the role of a gene of interest in promoting or preventing age-dependent protein aggregation by using either a comprehensive analysis with quantitative mass spectrometry or a candidate-based analysis with antibodies. These findings are then confirmed by in vivo analysis with transgenic animals expressing fluorescent-tagged aggregation-prone proteins. These methods should help clarify why certain proteins are prone to aggregate with age and ultimately how to keep these proteins fully functional.

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

The goal of the method presented here is to explore protein aggregation during normal aging in the model organism C. elegans. The protocol represents a powerful tool to study the highly insoluble large aggregates that form with age and to determine how changes in proteostasis impact protein aggregation.

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These ...

RNAi Feeding in Liquid Culture: A High-Throughput Method to Knockdown Gene Expression in C. elegans

Source: Groh, N. et al. <a href="https://www.jove.com/video/56464/methods-to-study-changes-inherent-protein-aggregation-with-age">Methods to Study Changes in Inherent Protein Aggregation with Age in&#160;Caenorhabditis elegans</a>.&#160;J. Vis. Exp.&#160;(2017).
RNAi is powerful genetic tool available to C. elegans researchers. This video introduces a method to treat more worms with RNAi than plate feeding by utilizing liquid culture conditions.

Source: Groh, N. et al. Methods to Study Changes in Inherent Protein Aggregation with Age in&#160;Caenorhabditis elegans.&#160;J. Vis. Exp.&#160;(2017).
...

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

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Chapters in this video

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

Gene-targeted Random Mutagenesis to Select Heterochromatin-destabilizing Proteasome Mutants in Fission Yeast

A Rapid and Facile Pipeline for Generating Genomic Point Mutants in C. elegans Using CRISPR/Cas9 Ribonucleoproteins

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

In vivo Neuronal Calcium Imaging in C. elegans

An Introduction to Worm Lab: from Culturing Worms to Mutagenesis

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

A Rapid Protocol for Integrating Extrachromosomal Arrays With High Transmission Rate into the C. elegans Genome

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

RNAi Feeding in Liquid Culture: A High-Throughput Method to Knockdown Gene Expression in C. elegans