This work was supported by the USDA National Institute of Food and Agriculture, Hatch project 1016879 and Maryland Agricultural Experiment Station via MAES Grant No. 2956952.

1. Preparation of dosage curve for effective mutagenesis
<ol>
	<li>Soak 100 seeds with the genotype of interest in six 250 mL glass flasks (100 in each flask) containing 50 mL of distilled water. Shake at 100 rpm for 8 h at room temperature (RT) for imbibition by the seeds.</li>
	<li>In a fume hood, prepare 50 mL of 0.4%, 0.6%, 0.8%, 1.0%, and 1.2% (w/v) ethyl methanesulfonate (EMS) solution by dissolving 0.167, 0.249, 0.331, 0.415, and 0.498 mL of EMS in distilled water, respectively. 
	NOTE: EMS is liquid at RT with a density of 1.206 g/mL. 
	CAUTION: Use appropriate personal protective equipment (PPE) while handling EMS.</li>
	<li>Decant the water out of five flasks and add 50 mL of EMS solution in each flask containing imbibed seed so that there are six different treatments with 0.0%, 0.4%, 0.6%, 0.8%, 1.0%, and 1.2% EMS solution. Shake flasks for 16 h at 75 rpm and RT.</li>
	<li>Decant the EMS solution and collect the treated seeds separately for each treatment using cheese cloth. Inactivate the used EMS solution by adding one volume of EMS-inactivating solution (0.1 M NaOH, 20% w/v Na2S2O3) for 24 h. Also treat the contaminated flasks and pipette tips with the EMS-inactivating solution for 24 h.</li>
	<li>Wash the EMS-treated seeds under running tap water for 2 h. Transplant each seed individually into root trainers containing potting soil.</li>
	<li>Grow plants at 20&#8211;25 &#176;C under a 16 h light period.</li>
	<li>Record data on plant survival after 15 days of transplantation. Use the following equation to calculate the survival rate for each treatment: 
	<img alt="Equation 1" src="/files/ftp_upload/59743/59743eq1.jpg" /> 
	NOTE: If the germination rate is lower than 100% in controls, accurate survival rates in all the treatments should be calculated after subtracting the number of seeds that failed to germinate in the controls. A survival rate of 40%&#8211;60% is desirable for effective mutagenesis. It may be required to perform a second round of dosage optimization with a modified concentration according to the survival of the treated seeds until achieving the desirable lethality rate of 40%&#8211;60%.</li>
</ol>
2. Mutagenesis and maintenance of mutant population
<ol>
	<li>Soak the final batch of at least 3,000 seeds dividing equally (600 seeds each) in five 1,000 mL flasks containing 300 mL distilled water. Shake for 8 h at 100 rpm under RT for imbibition. 
	NOTE: The final size of the desirable population will depend on the mutation frequency and ploidy level of the genotype, but it is advisable to use at least 3,000 seeds for hexaploids, 4,000 seeds for tetraploids, and more than 7,000 seeds for diploids.</li>
	<li>In a fume hood, prepare 1,500 mL of the optimized EMS concentration solution in distilled water.</li>
	<li>Decant the water out of the flasks and add 300 ml of the optimal concentration of EMS solution in each flask containing 600 imbibed seeds. Shake the flasks for 16 h at 75 rpm and RT.</li>
	<li>Decant the EMS and collect the treated seeds in cheese cloth. Inactivate the EMS solution and treatment containers with EMS-inactivating solution as done in step 1.4.</li>
	<li>Wash the EMS-treated seeds under running tap water for 2 h. Transplant each EMS-treated M1 seed individually into root trainers.</li>
	<li>Grow M1 plants (derived from M0 seeds) at 20&#8211;25 &#176;C under a 16 h light period. 
	NOTE: It may be required to vernalize the seedlings at the two leaf stage for 6 weeks at 4 &#176;C, if the genotype of interest has a winter-type growth habit.</li>
	<li>Allow the M1 plants to self-pollinate, and harvest the M2 seeds separately for each fertile M1 plant. 
	NOTE: To avoid chances of potential outcrossing, cover the spikes of M1 plants with pollination bags before anthesis.</li>
	<li>Plant a single M2 seed from each M1 plant to avoid genetic redundancy.</li>
	<li>Collect tissue from M2 plants at the two-leaf stage in 1.1 mL of racked 96 well microtubes. Collect around 80 mm of leaf tissue from each plant and record the ID of each sample in a tissue collection plan.</li>
	<li>Freeze-dry the leaf tissue using a lyophilizer and store at -80 &#176;C.</li>
	<li>Maintain the M2 plants at 20&#8211;25 &#176;C under a 16 h light period.</li>
	<li>Record data on mutant phenotypes of the M2 plants at regular intervals. The expected phenotypes are albino, chlorina, grassy shoot, variegated, partially fertile, sterile, etc.</li>
	<li>Allow the M2 plants to self-fertilize and mature. Separately harvest and save the M3 seeds of the M2 plants (Figure 1). 
	NOTE: M seeds are used for validating the phenotype in reverse genetics studies. Therefore, M should be carefully catalogued and stored under cool and dry conditions. Alternatively, if the seed must be increased in field, head rows can be planted for each M plant, and M seeds from each row can be harvested and saved separately as the TILLING population resource.</li>
</ol>
3. Cel-1 assay for genetic characterization of mutants
<ol>
	<li>Extract DNA from the leaf tissue of M2 using a plant DNA extraction kit with a DNA purification system (see Table of Materials) following the manufacturer&#8217;s recommendations.</li>
	<li>Quantify DNA using a spectrophotometer and normalize the DNA concentrations to 25 ng/&#181;L with nuclease-free water in 96 well blocks. 
	NOTE: It is important to check the quality of the DNA by running it on a gel, as low-quality DNA (smeared DNA) may result in false negatives in pooled samples.</li>
	<li>Create 4x DNA pools by combining DNA from four 96 well blocks into one plate, while maintaining the row and column identity of each sample. Add 50 &#181;L of DNA from each individual sample into the pool plate so that each pool plate well contains a total 200 &#181;L of DNA from four different 96 well blocks.</li>
	<li>Catalogue the identity of pooled DNA in the format of Pool Plate-Row-Column (e.g., Pool 1 A1 = Box1A1 + Box2A1 + Box3A1 + Box4A1).</li>
	<li>Design genome-specific primers [A1] for the gene of interest using the Genome Specific Primers (GSP) page &#60;https://probes.pw.usda.gov/GSP&#62; with default settings for polyploid species. For diploids, use Primer 3 &#60;http://primer3.ut.ee/&#62; for designing primers using default settings. Design multiple primers, if needed, to cover the entire coding region of the gene of interest. 
	NOTE: The latest IWGSC assembly can be used to obtain sequences for the gene of interest in wheat using the URGI BLAST tool &#60;https://wheat-urgi.versailles.inra.fr/Seq-Repository/BLAST&#62;. The optimal amplicon length is in the range of 800&#8211;1,500 bp. Table 1 shows examples of primers for waxy genes in hexaploid wheat.</li>
	<li>Run a PCR for gene-specific primers on pooled DNA as follows:
	<ol>
		<li>Add 5 &#181;L of PCR buffer, 2 &#181;L (each) of 4 &#181;M forward and reverse primers, 0.1 &#181;L of DNA polymerase (see Table of Materials), 5 &#181;L of pooled DNA template, then increase the volume to 25 &#181;L using nuclease-free water.</li>
		<li>Use a touch down profile to run the PCR reaction on a thermal cycler as follows: 95 &#176;C for 1 min, seven cycles of 95 &#176;C for 1 min, 67 &#176;C to 60 &#176;C for 1 min with temperature decreases of 1 &#176;C per cycle and 72 &#176;C for 2 min, followed by 30 cycles of 95 &#176;C for 1 min, 60 &#176;C for 1 min, 72 &#176;C for 2 min, and final extension at 72 &#176;C for 7 min. 
		NOTE: Above PCR profile works on wheat DNA template for most of the primers designed using primer 3 default settings. In case of non-specific amplification, PCR profile should be made stringent before moving to the next steps.</li>
	</ol>
	</li>
	<li>Generate heteroduplexes between mismatched DNA by incubating PCR products in thermal cycler using profile as follows: 95 &#176;C for 2 min, five cycles of 95 &#176;C for 1 s, 95 &#176;C to 85 &#176;C for 1 min with temperature decreases of 2 &#176;C per cycle, and 60 cycles of 85 &#176;C to 25 &#176;C with decreases of 1 &#176;C per cycle.</li>
	<li>Add 2.5 &#181;L of homemade Cel-1 endonuclease to the heteroduplexed PCR products and incubate for 45 min at 45 &#176;C. Terminate the Cel-1 reaction by adding 2.5 &#181;L of 0.5 M EDTA (pH 8.0). 
	NOTE: Cel-1 endonuclease can be extracted from fresh celery stalks using the protocol performed by Till et al.19 It is very important to test the activity and optimum amount of Cel-1 endonuclease, which can be tested using previously characterized mutants or a commercially available mutation detection kit.</li>
	<li>Run the Cel-1 treated products on a 3.0% agarose gel at 100 V for 2.5 h. The wells containing smaller and unique cleaved band(s), in addition to full-length uncleaved bands, will contain the mutant DNA sample.</li>
	<li>Denconvolute mutant pools.
	<ol>
		<li>Follow steps 3.6, 3.7, 3.8, and 3.9 for individual M2 DNA samples constituting the mutant pools identified in step 3.9.</li>
		<li>To determine the zygosity of mutants, run two PCR (as described in step 3.6) for individual M2 DNA, in which the first reaction contains 2.5 &#181;L of M2 DNA and 2.5 &#181;L of wild-type DNA and the second reaction contains only 5 &#181;L of M2 DNA. If the mutation is heterozygous, an additional cleaved band will be present in both reactions. On the other hand, additional cleaved bands will be found only in the first reaction if the mutant is homozygous.</li>
	</ol>
	</li>
	<li>To identify the nature of the mutation, sequence the PCR products of the confirmed mutants using a Sanger sequencing platform following the manufacturer&#8217;s instructions.</li>
</ol>
4. Calculation of mutation frequency
NOTE: Mutation frequency of a TILLING population refers to the average physical distance in which one mutation occurs in the individuals of that population. For example, a mutation frequency of 1/35 kb in a TILLING population means that an average individual of that population possesses 1 mutation per every 35 kb in the genome.
<ol>
	<li>To determine the mutation frequency of a TILLING population, calculate the total number of bases screened.</li>
	<li>To calculate the total number of bases screened, multiply the PCR product size by the total number of individuals screened.</li>
	<li>Divide the total number of bases screened by the number of unique mutations observed using the following equation, which will yield the physical region possessing 1 mutation in the given TILLING population: 
	<img alt="Equation 2" src="/files/ftp_upload/59743/59743eq2.jpg" /> 
	NOTE: To account for the limitation in resolution of 50 bp at both the ends based on an agarose gel-based platform, subtract 100 bp from the product size in the calculation.</li>
</ol>

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Authors declare no competing financial interests.

TILLING is a highly valuable reverse genetics tool for gene validation, especially for small grains where transformation-based approaches have serious bottlenecks11. Developing a mutagenized population with a high mutation frequency is one of the critical steps in conducting functional genomics studies. The most important step in developing a robust TILLING population is to determine the optimal concentration of EMS. The 40%-60% survival rate in the M1 has been found to be a good indica...

Point mutations in genomes can serve many useful purposes for researchers. Depending on their nature and location, these mutations can be used to assign functions to genes or even distinct domains of proteins of interest. On the other hand, as a source of novel genetic variation, useful mutations can be selected for desired traits using phenotyping screens and further used in crop improvement. TILLING is a powerful reverse genetics tool that includes chemical mutagenesis and detection of sequence variation in the target gene. First developed in Arabidopsis1 and Drosophilia melanogaster2, TILLING populations have been developed and utilized in many small grain crops such as hexaploid bread wheat (Triticum aestivum)3, barley (Hordeum vulgare)4, tetraploid durum wheat (T. dicoccoides durum)5, diploid wheat (T. monococcum)6 and the &#34;D&#34; genome progenitor of wheat Aegilops tauschii7. These resources have been used to validate the roles of genes in regulating abiotic and biotic stress tolerance8, regulating flowering time9, and developing nutritionally superior crop varieties5.
TILLING, along with the use of alkylating mutagenic agents such as ethyl methanesulfonate (EMS), sodium azide, N-methyl-N-nitrosourea (MNU), and methyl methanesulfonate (MMS), has advantages over other reverse genetics tools for several reasons. First, mutagenesis can be conducted on practically any species or variety of plant10 and is independent of the transformation bottleneck, which is particularly challenging in the case of small grains11. Second, in addition to generating knockout mutations that can be obtained by other gene validation approaches, a range of missense and splicing mutations can be induced, which can discern functions of individual domains of the proteins of interest12. Moreover, TILLING generates an immortal collection of mutations throughout the genome; thus, a single population can be used for functional validation of multiple genes. In contrast, other reverse genetics tools generate resources specific to only the gene under study13. Useful mutations identified through TILLING can be deployed for breeding purposes and are not subject to regulation, unlike gene editing, whose non-transgenic classification is still uncertain in many countries. This becomes especially relevant to small grains that are internationally traded14.
TILLING is a simple and efficient gene validation strategy and requires mutagenized populations to be developed for investigating genes of interest. Developing an effective mutagenized population is key to determining the efficiency of a TILLING-based gene validation study. A TILLING population with a low overall mutation frequency indicates that an impractically large population must be screened for desired mutations, whereas a high mutagen concentration leads to high mortality in the population and an insufficient number of mutagenized individuals. Once a good population is developed, there are multiple ways to detect mutations in the genes of interest, and the choice of platform depends on the experimental scale and availability of resources. Whole genome sequencing and exome sequencing has been used to characterize all mutations in TILLING populations in plants with small genomes15,16. Exome sequencing of two TILLING populations has been performed in bread and durum wheat and is available to the public for identifying desirable mutations and ordering mutant lines of interest17. It is a great public resource in terms of availability of desirable mutations; however, in gene validation studies, the wild-type line should possess the candidate gene of interest. Unfortunately, it is still cost-prohibitive to sequence the exome of the entire TILLING population for reverse genetics-based validation of a few candidate genes in another background. Amplicon sequencing and Cel-1-based assays have been used in detecting mutations in targeted populations in wheat, and Cel-1 assays are simpler, requiring no computational knowledge, and are especially suitable for validation of a small number of genes with basic lab equipment6,18.
In the present article, described are methods for the development of a good TILLING population, including preparation of the dosage curve, mutagenesis and maintenance of the mutant population, and screening of the mutant population using the PCR-based Cel-1 assay. This protocol has already been implemented successfully in developing and utilizing mutagenized populations of Triticum aestivum, Triticum monoccocum6, barley, Aegilops tauchii7, and several others. Included are explicit details of these methods along with useful tips that will help researchers develop TILLING populations, using EMS as a mutagen in any small grain plant of choice.

<table><tbody><tr><td>96 well 1.1 ml microtubes in microracks</td><td>National Scientific</td><td>TN0946-08R</td><td>For collecting leaf tissues</td></tr><tr><td>Agarose I biotechnology grade</td><td>VWR</td><td>0710-500G</td><td></td></tr><tr><td>Biosprint 96 DNA Plant Kit</td><td>Qiagen</td><td>941558</td><td>Kit for DNA extraction</td></tr><tr><td>Cel-1 endonuclease</td><td>Extracted as described by Till et al 2006</td><td></td><td>Single strand specific endonuclease</td></tr><tr><td>Centrifuge 5430 R</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>Ethyl methanesulfonate</td><td>Sigma Aldrich</td><td>M-0880-25G</td><td>EMS, Chemical mutagen</td></tr><tr><td>Freeze Dry/Shell freeze system</td><td>Labconco</td><td></td><td>For lyophilization of leaf tissue</td></tr><tr><td>Kingfisher Flex purification system</td><td>Thermo fisher scientific</td><td>5400610</td><td>High throughput DNA extraction robot</td></tr><tr><td>My Taq DNA Polymerase</td><td>Bioline</td><td>BIO-21107</td><td></td></tr><tr><td>Nuclease free water</td><td>Sigma aldrich</td><td>W4502-1L</td><td></td></tr><tr><td>NuGenius gel imaging system</td><td>Syngene</td><td></td><td></td></tr><tr><td>Orbit Environ-shaker</td><td>Lab-line</td><td></td><td></td></tr><tr><td>SPECTROstar Nano</td><td>BMG LABTECH</td><td></td><td>Nano drop for DNA quantification</td></tr><tr><td>T100 Thermal cycler</td><td>BIO-RAD</td><td>1861096</td><td></td></tr></tbody></table>

development of targeting induced local lesions in genomes (tilling) populations in small grain crops by ethyl methanesulfonate mutagenesis

Targeting Induced Local Lesions IN Genomes (TILLING) is a powerful reverse genetics tool that includes chemical mutagenesis and detection of sequence variation in target genes. TILLING is a highly valuable functional genomics tool for gene validation, especially in small grains in which transformation-based approaches hold serious limitations. Developing a robust mutagenized population is key to determining the efficiency of a TILLING-based gene validation study. A TILLING population with a low overall mutation frequency indicates that an impractically large population must be screened to find desired mutations, whereas a high mutagen concentration leads to high mortality in the population, leading to an insufficient number of mutagenized individuals. Once an effective population is developed, there are multiple ways to detect mutations in a gene of interest, and the choice of platform depends upon the experimental scale and availability of resources. The Cel-1 assay and agarose gel-based approach for mutant identification is convenient, reproducible, and a less resource-intensive platform. It is advantageous in that it is simple, requiring no computational knowledge, and it is especially suitable for validation of a small number of genes with basic lab equipment. In the present article, described are the methods for development of a good TILLING population, including preparation of the dosage curve, mutagenesis and maintenance of the mutant population, and screening of the mutant population using the PCR-based Cel-1 assay.

Figure 2 shows the dosage curve of hexaploid bread wheat cultivar Jagger, diploid wheat Triticum monococcum6, and a genome donor of wheat Aegilops tauschii7. The EMS doses for desired 50% survival rates were about 0.25%, 0.6% and 0.7% for T. monococcum, Ae. tauschii, and T. aestivum, respectively. The higher EMS tolerance of hexaploid wheat is due to its genome buffer...

Watch this Scientific Journal Video about Development of Targeting Induced Local Lesions IN Genomes TILLING Populations in Small Grain Crops by Ethyl Methanesulfonate Mutagenesis at JoVE.com

Development of Targeting Induced Local Lesions IN Genomes TILLING Populations in Small Grain Crops by Ethyl Methanesulfonate Mutagenesis

Described is a protocol for developing a Targeting Induced Local Lesions IN Genomes (TILLING) population in small grain crops with use of ethyl methanesulfonate (EMS) as a mutagen. Also provided is a protocol for mutation detection using the Cel-1 assay.

Development of Targeting Induced Local Lesions IN Genomes (TILLING) Populations in Small Grain Crops by Ethyl Methanesulfonate Mutagenesis

Targeting Induced Local Lesions IN Genomes (TILLING) is a powerful reverse genetics tool that includes chemical mutagenesis and detection of sequence ...

development-targeting-induced-local-lesions-genomes-tilling

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

11.5K Views.  University of Maryland, College Park. Described is a protocol for developing a Targeting Induced Local Lesions IN Genomes (TILLING) population in small grain crops with use of ethyl methanesulfonate (EMS) as a mutagen. Also provided is a protocol for mutation detection using the Cel-1 assay.

Video: Development of Targeting Induced Local Lesions IN Genomes TILLING Populations in Small Grain Crops by Ethyl Methanesulfonate Mutagenesis

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.

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.

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

Genetics

An Array-based Comparative Genomic Hybridization Platform for Efficient Detection of Copy Number Variations in Fast Neutron-induced Medicago truncatula Mutants

Mutants are invaluable genetic resources for gene function studies. To generate mutant collections, three types of mutagens can be utilized, including biological such as T-DNA or transposon, chemical such as ethyl methanesulfonate (EMS), or physical such as ionization radiation. The type of mutation observed varies depending on the mutagen used. For ionization radiation induced mutants, mutations include deletion, duplication, or rearrangement. While T-DNA or transposon-based mutagenesis is limited to species that are susceptible to transformation, chemical or physical mutagenesis can be applied to a broad range of species. However, the characterization of mutations derived from chemical or physical mutagenesis traditionally relies on a map-based cloning approach, which is labor intensive and time consuming. Here, we show that a high-density genome array-based comparative genomic hybridization (aCGH) platform can be applied to efficiently detect and characterize copy number variations (CNVs) in mutants derived from fast neutron bombardment (FNB) mutagenesis in Medicago truncatula, a legume species. Whole genome sequence analysis shows that there are more than 50,000 genes or gene models in M. truncatula. At present, FNB-induced mutants in M. truncatula are derived from more than 150,000 M1 lines, representing invaluable genetic resources for functional studies of genes in the genome. The aCGH platform described here is an efficient tool for characterizing FNB-induced mutants in M. truncatula.

An Array-based Comparative Genomic Hybridization Platform for Efficient Detection of Copy Number Variations in Fast Neutron-induced Medicago truncatula Mutants

This protocol provides experimental steps and information about reagents, equipment, and analysis tools for researchers who are interested in carrying out whole genome array-based comparative genomic hybridization (CGH) analysis of copy number variations in plants.

Mutants are invaluable genetic resources for gene function studies. To generate mutant collections, three types of mutagens can be utilized, including ...

Identifying Mutations by High Resolution Melting in a TILLING Population of Rice

Target Induced Local Lesions In Genomes (TILLING) is a strategy of reverse genetics for the high-throughput screening of induced mutations. However, the TILLING system has less applicability for insertion/deletion (Indel) detection and traditional TILLING needs more complex steps, like CEL I nuclease digestion and gel electrophoresis. To improve the throughput and selection efficiency, and to make the screening of both Indels and single base substitions (SBSs) possible, a new high-resolution melting (HRM)-based TILLING system is developed. Here, we present a detailed HRM-TILLING protocol and show its application in mutation screening. This method can analyze the mutations of PCR amplicons by measuring the denaturation of double-stranded DNA at high temperatures. HRM analysis is directly performed post-PCR without additional processing. Moreover, a simple, safe and fast (SSF) DNA extraction method is integrated with HRM-TILLING to identify both Indels and SBSs. Its simplicity, robustness and high throughput make it potentially useful for mutation scanning in rice and other crops.

In this article, we present the protocol that is described as high-resolution melting analysis (HRM)-based Target Induced Local Lesions In Genomes (TILLING). This method utilizes fluorescence changes during the melting of the DNA duplex and is suitable for high-throughput screening of both insertion/deletion (Indel) and single base substition (SBS).

Target Induced Local Lesions In Genomes (TILLING) is a strategy of reverse genetics for the high-throughput screening of induced mutations. However, the ...

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 ...

Biology

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

It is becoming common for plant scientists to develop projects that require the genotyping of large numbers of plants. The first step in any genotyping project is to collect a tissue sample from each individual plant. The traditional approach to this task is to sample plants one-at-a-time. If one wishes to genotype hundreds or thousands of individuals, however, using this strategy results in a significant bottleneck in the genotyping pipeline. The Ice-Cap method that we describe here provides a high-throughput solution to this challenge by allowing one scientist to collect tissue from several thousand seedlings in a single day 1,2. This level of throughput is made possible by the fact that tissue is harvested from plants 96-at-a-time, rather than one-at-a-time.
The Ice-Cap method provides an integrated platform for performing seedling growth, tissue harvest, and DNA extraction. The basis for Ice-Cap is the growth of seedlings in a stacked pair of 96-well plates. The wells of the upper plate contain plugs of agar growth media on which individual seedlings germinate. The roots grow down through the agar media, exit the upper plate through a hole, and pass into a lower plate containing water. To harvest tissue for DNA extraction, the water in the lower plate containing root tissue is rapidly frozen while the seedlings in the upper plate remain at room temperature. The upper plate is then peeled away from the lower plate, yielding one plate with 96 root tissue samples frozen in ice and one plate with 96 viable seedlings. The technique is named "Ice-Cap" because it uses ice to capture the root tissue. The 96-well plate containing the seedlings can then wrapped in foil and transferred to low temperature. This process suspends further growth of the seedlings, but does not affect their viability. Once genotype analysis has been completed, seedlings with the desired genotype can be transferred from the 96-well plate to soil for further propagation. We have demonstrated the utility of the Ice-Cap method using Arabidopsis thaliana, tomato, and rice seedlings. We expect that the method should also be applicable to other species of plants with seeds small enough to fit into the wells of 96-well plates.

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

The Ice-Cap method allows one to grow plants in 96-well plates and non-destructively harvest root tissue from each seedling. DNA extracted from this root tissue can be used for genotyping reactions. We have found that Ice-Cap works well for Arabidopsis thaliana, tomato, and rice seedlings.

It is becoming common for plant scientists to develop projects that require the genotyping of large numbers of plants. The first step in any genotyping ...

Fluorescence-microscopy Screening and Next-generation Sequencing: Useful Tools for the Identification of Genes Involved in Organelle Integrity

This protocol describes a fluorescence microscope-based screening of Arabidopsis seedlings and describes how to map recessive mutations that alter the subcellular distribution of a specific tagged fluorescent marker in the secretory pathway. Arabidopsis is a powerful biological model for genetic studies because of its genome size, generation time, and conservation of molecular mechanisms among kingdoms. The array genotyping as an approach to map the mutation in alternative to the traditional method based on molecular markers is advantageous because it is relatively faster and may allow the mapping of several mutants in a really short time frame. This method allows the identification of proteins that can influence the integrity of any organelle in plants. Here, as an example, we propose a screen to map genes important for the integrity of the endoplasmic reticulum (ER). Our approach, however, can be easily extended to other plant cell organelles (for example see1,2), and thus represents an important step toward understanding the molecular basis governing other subcellular structures.

A fundamental quest in cell biology is to define the mechanisms that underlie the identity of the organelles that make eukaryotic cells. Here we propose a method to identify the genes responsible for the morphological and functional integrity of plant organelles using fluorescence microscopy and next-generation sequencing tools.

This protocol describes a fluorescence microscope-based screening of Arabidopsis seedlings and describes how to map recessive mutations that alter the ...

CRISPR-Based Efficient Genome Editing for Resistant Starch in Rice Breeding

Breeding by Design for Functional Rice with Genome Editing Technologies

The conventional approaches to crop breeding, which rely predominantly on time-consuming and labor-intensive methods such as traditional hybridization and mutation breeding, face challenges in efficiently introducing targeted traits and generating diverse plant populations. Conversely, the emergence of genome editing technologies has ushered in a paradigm shift, enabling the precise and expedited manipulation of plant genomes to intentionally introduce desired characteristics. One of the most widespread editing tools is the CRISPR/Cas system, which has been used by researchers to study important biology-related problems. However, the precise and effective workflow of genome editing has not been well-defined in crop breeding. In this study, we demonstrated the entire process of breeding rice varieties enriched with high levels of resistant starch (RS), a functional trait that plays a crucial role in preventing diseases such as diabetes and obesity. The workflow encompassed several key steps, such as the selection of functional SBEIIb gene, designing the single-guide RNA (sgRNA), selecting an appropriate genome editing vector, determining the vector delivery method, conducting plant tissue culture, genotyping mutation and phenotypic analysis. Additionally, the time frame necessary for each stage of the process has been clearly demonstrated. This protocol not only streamlines the breeding process but also enhances the accuracy and efficiency of trait introduction, thereby accelerating the development of functional rice varieties.

Author Spotlight: Streamlining Rice Breeding with CRISPR/Cas for Obtaining Optimal Phenotypic and Agronomic Traits 

The protocol describes breeding resistant starch rice varieties by design using genome editing technologies in a precise, efficient, and technically simple way.

The conventional approaches to crop breeding, which rely predominantly on time-consuming and labor-intensive methods such as traditional hybridization and ...

Bioengineering

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases (TALENs)

Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs. Towards this end, much recent work has focused on identifying candidate genes for the evolution of traits in a variety of species. However, until recently it has been challenging to functionally validate interesting candidate genes. Recently developed tools for genetic engineering make it possible to manipulate specific genes in a wide range of organisms. Application of this technology in evolutionarily relevant organisms will allow for unprecedented insight into the role of candidate genes in evolution. Astyanax mexicanus (A. mexicanus) is a species of fish with both surface-dwelling and cave-dwelling forms. Multiple independent lines of cave-dwelling forms have evolved from ancestral surface fish, which are interfertile with one another and with surface fish, allowing elucidation of the genetic basis of cave traits. A. mexicanus has been used for a number of evolutionary studies, including linkage analysis to identify candidate genes responsible for a number of traits. Thus, A. mexicanus is an ideal system for the application of genome editing to test the role of candidate genes. Here we report a method for using transcription activator-like effector nucleases (TALENs) to mutate genes in surface A. mexicanus. Genome editing using TALENs in A. mexicanus has been utilized to generate mutations in pigmentation genes. This technique can also be utilized to evaluate the role of candidate genes for a number of other traits that have evolved in cave forms of A. mexicanus.

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases TALENs

Gene-targeting mutagenesis is now possible in a wide range of organisms using genome editing techniques. Here, we demonstrate a protocol for targeted gene mutagenesis using transcription activator like effector nucleases (TALENs) in Astyanax mexicanus, a species of fish that includes surface fish and cavefish.

Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs.  Towards this end, much recent ...

Agrobacterium-Mediated Immature Embryo Transformation of Recalcitrant Maize Inbred Lines Using Morphogenic Genes

Demonstrated here is a detailed protocol for Agrobacterium-mediated genetic transformation of maize inbred lines using morphogenic genes Baby boom (Bbm) and Wuschel2 (Wus2). Bbm is regulated by the maize phospholipid transferase gene (Pltp) promoter, and Wus2 is under the control of a maize auxin-inducible (Axig1) promoter. An Agrobacterium strain carrying these morphogenic genes on transfer DNA (T-DNA) and extra copies of Agrobacterium virulence (vir) genes are used to infect maize immature embryo explants. Somatic embryos form on the scutella of infected embryos and can be selected by herbicide resistance and germinated into plants. A heat-activated cre/loxP recombination system built into the DNA construct allows for removal of morphogenic genes from the maize genome during an early stage of the transformation process. Transformation frequencies of approximately 14%, 4%, and 4% (numbers of independent transgenic events per 100 infected embryos) can be achieved for W22, B73, and Mo17, respectively, using this protocol.

Agrobacterium-Mediated Immature Embryo Transformation of Recalcitrant Maize Inbred Lines Using Morphogenic Genes

Plant morphogenic genes can be used to improve genetic transformation of recalcitrant genotypes. Described here is an Agrobacterium-mediated genetic transformation (QuickCorn) protocol for three important public maize inbred lines.

Demonstrated here is a detailed protocol for Agrobacterium-mediated genetic transformation of maize inbred lines using morphogenic genes Baby boom (Bbm) ...

Generating Transposon Insertion Libraries in Gram-Negative Bacteria for High-Throughput Sequencing

Transposon sequencing (Tn-seq) is a powerful method that combines transposon mutagenesis and massive parallel sequencing to identify genes and pathways that contribute to bacterial fitness under a wide range of environmental conditions. Tn-seq applications are extensive and have not only enabled examination of genotype-phenotype relationships at an organism level but also at the population, community and systems levels. Gram-negative bacteria are highly associated with antimicrobial resistance phenotypes, which has increased incidents of antibiotic treatment failure. Antimicrobial resistance is defined as bacterial growth in the presence of otherwise lethal antibiotics. The &#8220;last-line&#8221; antimicrobial colistin is used to treat Gram-negative bacterial infections. However, several Gram-negative pathogens, including Acinetobacter baumannii can develop colistin resistance through a range of molecular mechanisms, some of which were characterized using Tn-seq. Furthermore, signal transduction pathways that regulate colistin resistance vary within Gram-negative bacteria. Here we propose an efficient method of transposon mutagenesis in A. baumannii that streamlines generation of a saturating transposon insertion library and amplicon library construction by eliminating the need for restriction enzymes, adapter ligation, and gel purification. The methods described herein will enable in-depth analysis of molecular determinants that contribute to A. baumannii fitness when challenged with colistin. The protocol is also applicable to other Gram-negative ESKAPE pathogens, which are primarily associated with drug resistant hospital-acquired infections.

We describe a method to generate saturating transposon mutant libraries in Gram-negative bacteria and subsequent preparation of DNA amplicon libraries for high-throughput sequencing. As an example, we focus on the ESKAPE pathogen, Acinetobacter baumannii, but this protocol is amenable to a wide range of Gram-negative organisms.

Transposon sequencing (Tn-seq) is a powerful method that combines transposon mutagenesis and massive parallel sequencing to identify genes and pathways ...

Development of Targeting Induced Local Lesions IN Genomes (TILLING) Populations in Small Grain Crops by Ethyl Methanesulfonate Mutagenesis

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Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

An Array-based Comparative Genomic Hybridization Platform for Efficient Detection of Copy Number Variations in Fast Neutron-induced Medicago truncatula Mutants

Identifying Mutations by High Resolution Melting in a TILLING Population of Rice

An Introduction to Worm Lab: from Culturing Worms to Mutagenesis

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

Fluorescence-microscopy Screening and Next-generation Sequencing: Useful Tools for the Identification of Genes Involved in Organelle Integrity

Breeding by Design for Functional Rice with Genome Editing Technologies

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases (TALENs)

Agrobacterium-Mediated Immature Embryo Transformation of Recalcitrant Maize Inbred Lines Using Morphogenic Genes

Generating Transposon Insertion Libraries in Gram-Negative Bacteria for High-Throughput Sequencing