Name: a fast air-dry dropping chromosome preparation method suitable for fish in plants
Uploaded: 2015-12-16T14:00:00.000+00:00
Duration: 9 min 8 s
Description: Watch this Scientific Journal Video about A Fast Air-dry Dropping Chromosome Preparation Method Suitable for FISH in Plants at JoVE.com

We gratefully thank the DFG for financial support (HO 1779/21-1) as well as Katrin Kumke and Dr. Veit Schubert (IPK, Gatersleben) for technical advice.

1. Chromosome Preparation
<ol>
	<li>Seed germination and fixation of root tips
	<ol>
		<li>Germinate 10-20 barley seeds on two layers of moist filter paper in a Petri dish under dark conditions for 2 days at 22-24 &#176;C. Cut off vigorous roots with the length of 1-2 cm from the seed by using a razor blade.</li>
		<li>Prepare ice-cold water by placing a 500 ml glass bottle containing cold tap water into crushed ice-water. Aerate the ice-cold water and immerse root tips for 20 hr to increase the frequency of metaphase cells.</li>
		<li>Transfer roots from water to 50 ml of ethanol: acetic acid (3:1) fixative to fix them at RT&#160;for 2 days. Store roots in a freshly prepared ethanol: acetic acid (3:1) fixative at 4 &#176;C until use up to a year.</li>
	</ol>
	</li>
	<li>Washing and enzyme treatment
	<ol>
		<li>Wash the 10-20 roots with 30 ml ice-cold tap water for 5 min twice using a 50 ml glass beaker. Use binocular microscope. Transfer roots one by one into 30 ml 0.01 M citrate buffer (0.01 M citric acid + 0.01 M sodium citrate, pH 4.8) using forceps and wash by shaking the glass beaker for 5 min twice. Place roots on filter paper to remove the liquid completely and cut-off undesired non-meristematic tissue using a razor blade.</li>
		<li>Incubate up to 20 root tips in 1 ml enzyme mixture at 37 &#176;C for about 50 min to soften the plant tissue (Table 1) in a watch glass. Enzyme mixture contains 0.7% cellulase R10, 0.7% cellulase, 1% pectolyase and 1% cytohelicase diluted in 0.01 M citrate buffer. Store the enzyme mixture at -20 &#176;C and reused up to five times.</li>
		<li>Remove the enzyme by pipetting and wash the root tips on ice with 5 ml 0.01 M citrate buffer twice to replace the residual enzyme.</li>
	</ol>
	</li>
	<li>Root maceration
	<ol>
		<li>Wash root tips with 1 ml 96% ethanol twice carefully in the same watch glass. Replace ethanol with freshly prepared fixative (75% acetic acid : 25% ethanol). Use 10-15 &#181;l fixative per root tip.</li>
		<li>Transfer root tips together with the fixative into a 2 ml tube and disintegrate root meristems with a dissecting needle or forceps. Tap the tube 20 times to re-suspend cells to obtain a cell suspension. Store the cell suspension at -20 &#176;C up to two months.</li>
	</ol>
	</li>
	<li>Dropping of the cell suspension
	<ol>
		<li>Place 2-3 layers of water-soaked paper tissue on a hot plate at 50 &#176;C. Immerse microscopic slides in ice-cold tap water in the fridge for 30 min. And place slides on top of the moist paper tissue.</li>
		<li>Pipette 7-10 &#181;l of cell suspension and drop it from a distance of 20 cm onto the cooled slide placed on the hot plate. Pipette 10 &#181;l of acetic acid-ethanol mixture on the same place as cell suspension on the slide and keep the slide on the hot plate for additional 2 min. Place the slide on the hot plate without the wet tissue and let it dry for 1 min.</li>
	</ol>
	</li>
	<li>Quality control and storage of slides
	<ol>
		<li>Check slides using a phase-contrast microscope to control the quality of the chromosome spread. Use slides either the same day or store by immersing in 96% ethanol in a Coplin jar at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Pretreatment of slides before FISH; all steps are carried out at RT
	<ol>
		<li>Place slides in a Coplin jar containing 50 ml of 2x SSC (20x SSC contains 3 M NaCl and 300 mM trisodium citrate) for 5 min. Using forceps, transfer slides to a Coplin jar containing 50 ml of 45% acetic acid for 3-10 min.</li>
		<li>Transfer slides to a Coplin jar containing 50 ml of 2x SSC for 10 min. Transfer slides to a Coplin jar containing 50 ml of 4% formaldehyde (in 2x SSC) and immerse slides for 10 min to fix chromosomes.</li>
		<li>Remove formaldehyde by rinsing the slides 3 times for 4 min each, in a Coplin jar containing 50 ml 2x SSC. Dehydrate slides in a Coplin jar for 2 min in series of 70%, 90% and 100% ethanol, respectively and dry slides in a vertical position.</li>
	</ol>
	</li>
</ol>
2. Fluorescent In Situ Hybridization (FISH)
<ol>
	<li>For each slide, prepare a hybridization solution of 20 &#181;l in total using 10 &#181;l of deionized formamide, 5 &#181;l of 4x hybridization buffer (200 &#181;l buffer contains 80 &#181;l 20x SSC, 8 &#181;l 1 M Tris-HCl pH 8.0, 1.6 &#181;l 0.5 M EDTA, 11.2 &#181;l 10 &#181;g/ &#181;l salmon sperm and 99.2 &#181;l DNase-free water), 3 &#181;l of the probe and 2 &#181;l of DNase-free water.</li>
	<li>Add 20 &#181;l of hybridization solution per slide and cover with a 24 x 32 mm cover slip and arrest the cover slip with rubber cement. Denature slides with probes simultaneously at 80 &#176;C for 2 min on a hot plate.</li>
	<li>Transfer slides to a moist chamber and incubate slides at 37 &#176;C O/N avoiding light. Remove cover slips by rinsing the slides in a Coplin jar with 2x SSC. Place slides in a Coplin jar containing 55-60 &#176;C 2x SSC and incubate for 20 min.</li>
	<li>Place the slides to 2x SSC in a Coplin jar for 2 min at RT. Dehydrate slides in a Coplin jar for 2 min in series of 70%, 90% and 100% ethanol, respectively.</li>
	<li>Air-dry the slides and counterstain with 1 &#181;g/ml 4&#39;,6-diamidino-2-phenylindoline (DAPI) in antifade mounting medium, avoid intense light.</li>
</ol>
3. Microscopic Analysis and Storage
<ol>
	<li>Analyze the slides using an epifluorescence microscope. The selection of filter depends on the fluorochrome used for probe labeling. If necessary, store slides at 4 &#176;C under dark conditions up to a year.</li>
</ol>

<ol>
	<li>Geber, G., Schweizer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytochemical+heterochromatin+differentiation+in+Sinapis+alba+(Cruciferae)+using+a+simple+air-drying+technique+for+producing+chromosome+spreads.">Cytochemical heterochromatin differentiation in Sinapis alba (Cruciferae) using a simple air-drying technique for producing chromosome spreads.</a> Pl Syst Evol. 158 (2-4), 97-106 (1988).</li><li>Andras, S. C., 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=A+drop-spreading+technique+to+produce+cytoplasm-free+mitotic+preparations+from+plants+with+small+chromosomes.">A drop-spreading technique to produce cytoplasm-free mitotic preparations from plants with small chromosomes.</a> Chromosome Res. 7 (8), 641-647 (1999).</li><li>Rothfels, K. H., Siminovitch, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+air-drying+technique+for+flattening+chromosomes+in+mammalian+cells+grown+in+vitro.">An air-drying technique for flattening chromosomes in mammalian cells grown in vitro.</a> Stain Technology. 33 (2), 73-77 (1958).</li><li>Martin, R., Busch, W., Herrmann, R. G., Wanner, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+preparation+of+plant+chromosomes+for+high-resolution+scanning+electron+microscopy.">Efficient preparation of plant chromosomes for high-resolution scanning electron microscopy.</a> Chromosome Res. 2 (5), 411-415 (1994).</li><li>Kato, A., Albert, P. S., Vega, J. M., Birchler, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sensitive+fluorescence+in+situ+hybridization+signal+detection+in+maize+using+directly+labeled+probes+produced+by+high+concentration+DNA+polymerase+nick+translation.">Sensitive fluorescence in situ hybridization signal detection in maize using directly labeled probes produced by high concentration DNA polymerase nick translation.</a> Biotech. Histochem. 81 (2-3), 71-78 (2006).</li><li>Kirov, I., Divashuk, M., Van Laere, K., Soloviev, A., Khrustaleva, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+easy+'SteamDrop'+method+for+high+quality+plant+chromosome+preparation.">An easy 'SteamDrop' method for high quality plant chromosome preparation.</a> Mol. Cytogenet. 7, 21 (2014).</li><li>Spurbeck, J. L., Zinsmeister, A. R., Meyer, K. J., Jalal, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+chromosome+spreading.">Dynamics of chromosome spreading.</a> Am J Med Genet. 61 (4), 387-393 (1996).</li><li>Ma, L., 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=Synteny+between+Brachypodium+distachyon+and+Hordeum+vulgare+as+revealed+by+FISH.">Synteny between Brachypodium distachyon and Hordeum vulgare as revealed by FISH.</a> Chromosome Res. 18 (7), 841-850 (2010).</li><li>Kato, A., Lamb, J. C., Birchler, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chromosome+painting+using+repetitive+DNA+sequences+as+probes+for+somatic+chromosome+identification+in+maize.">Chromosome painting using repetitive DNA sequences as probes for somatic chromosome identification in maize.</a> Proc. Natl. Acad. Sci. U S A. 101 (37), 13554-13559 (2004).</li><li>Pan, W. H., Houben, A., Schlegel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+effective+cell+synchronization+in+plant-roots+by+hydroxyurea+and+amiprophos-methyl+or+colchicine.">Highly effective cell synchronization in plant-roots by hydroxyurea and amiprophos-methyl or colchicine.</a> Genome. 36 (2), 387-390 (1993).</li><li>Kim, J. 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=Integrated+karyotyping+of+sorghum+by+in+situ+hybridization+of+landed+BACs.">Integrated karyotyping of sorghum by in situ hybridization of landed BACs.</a> Genome. 45 (2), 402-412 (2002).</li><li>Lapitan, N. L. V., Brown, S. E., Kennard, W., Stephens, J. L., Knudson, D. 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The authors have nothing to disclose.

The chromosome preparation experiment has been carried out using young roots of cereals belonging to the grass family (Poaceae). All analyzed species have 14 relatively long mitotic metaphase chromosomes (11-15 &#181;m) in the diploid genome set and belong to large-genome species (5.1-7.9 Gbp).
Length of germinated roots was not more than 2 cm to obtain a maximum of meristematic tissue. Synchronization of dividing cells was achieved by a 20 hr long ice-water treatment that improved the quantit...

Fluorescence in situ hybridization (FISH) is an effective tool for the physical mapping of single and high copy sequences at the chromosomal level. Prerequisite is the preparation of high quality chromosome spreads. There is no general chromosome preparation protocol that would be equally suitable for animal and plant cells. Preparation of plant chromosomes is particularly challenging due to the rigid cell wall and various cytoplasm consistency within different species. One of the favorable methods for the preparation of plant chromosomes is a so-called drop technique also known as drop-spreading technique and air-drying technique 1,2. This method was first introduced in 1958 by Rothfels and Siminovitch for in vitro grown mammalian cells 3. Later Martin et al. 4 and Kato et al. 5 adapted this method for plants.
More recently, a method named &#39;SteamDrop&#39; was developed which used water steam for the preparation of non-overlapping chromosomes 6. Although, the positive influence of high humidity was observed earlier 7, &#39;SteamDrop&#39; delivers a controlled workflow of high-quality chromosome preparations 6. The steam treatment causes stretching of chromosomes probably connected to some modifications of chromosomal proteins. The quality of resulting metaphase spreads is very high, although retaining of sufficient number of complete metaphase spreads for subsequent FISH experiments demands technical expertise.
Here we present a protocol for the preparation of mitotic cereal chromosomes suitable for the FISH detection of single and high copy probes 5,8. This method is an improved variant of the air-dry dropping method described by Kato 9 performed under relative humidity of 50%-55% (Figure 1). This protocol comprises a reduced number of washing steps making its application easy, efficient and reproducible. Using this protocol we obtained high-quality chromosome spreads and FISH results for Hordeum vulgare, H. bulbosum, H. marinum, H. murinum, H. pubiflorum and Secale cereale.

<table><tbody><tr><td>Hot Plate</td><td>MEDAX GmbH</td><td>12603</td><td></td></tr><tr><td>Cellulase R10</td><td>Duchefa</td><td>C8001</td><td></td></tr><tr><td>Cellulase&amp;nbsp;</td><td>CalBioChem</td><td>219466</td><td></td></tr><tr><td>Pectolyase</td><td>Sigma</td><td>P3026</td><td></td></tr><tr><td>Cytohelicase</td><td>Sigma</td><td>C8274</td><td></td></tr><tr><td>Texas Red-12-dUTP</td><td>Invitrogen</td><td>C3176</td><td>direct fluorochrome&amp;nbsp;</td></tr><tr><td>Fluor488-5-dUTP</td><td>Invitrogen</td><td>C11397</td><td>direct fluorochrome&amp;nbsp;</td></tr><tr><td>Fluorecsence microscope</td><td>Olympus BX61</td><td>BX61</td><td></td></tr><tr><td>CCD camera</td><td>Orca ER, Hamamatsu</td><td>C10600</td><td></td></tr><tr><td>4&amp;rsquo;,6-diamidino-2-phenylindole (DAPI)&amp;nbsp;</td><td>Vector Laboratories</td><td>H-1200</td><td>fluorecsent dye</td></tr></tbody></table>

a fast air-dry dropping chromosome preparation method suitable for fish in plants

Preparation of chromosome spreads is a prerequisite for the successful performance of fluorescence in situ hybridization (FISH). Preparation of high quality plant chromosome spreads is challenging due to the rigid cell wall. One of the approved methods for the preparation of plant chromosomes is a so-called drop preparation, also known as drop-spreading or air-drying technique. Here, we present a protocol for the fast preparation of mitotic chromosome spreads suitable for the FISH detection of single and high copy DNA probes. This method is an improved variant of the air-dry drop method performed under a relative humidity of 50%-55%. This protocol comprises a reduced number of washing steps making its application easy, efficient and reproducible. Obvious benefits of this approach are well-spread, undamaged and numerous metaphase chromosomes serving as a perfect prerequisite for successful FISH analysis. Using this protocol we obtained high-quality chromosome spreads and reproducible FISH results for Hordeum vulgare, H. bulbosum, H. marinum, H. murinum, H. pubiflorum and Secale cereale.

Microscopic slides with the mitotic metaphase spreads were prepared by the fast air-dry dropping chromosome preparation method described above (Suppl. Figure 1). FISH analysis was carried out using both, repetitive and single-copy sequences. Images were obtained by a epifluorescence microscope with a set of filters enabling excitation of corresponding fluorophores and captured by a high-sensitivity CCD monochrome camera. For the image acquisition we used a computer with an image acquisition software. Res...

Watch this Scientific Journal Video about A Fast Air-dry Dropping Chromosome Preparation Method Suitable for FISH in Plants at JoVE.com

A Fast Air-dry Dropping Chromosome Preparation Method Suitable for FISH in Plants

A protocol is described for the preparation of high-quality mitotic plant chromosome spreads by a fast air-dry dropping method suitable for the FISH detection of single and high copy DNA probes.

Preparation of chromosome spreads is a prerequisite for the successful performance of fluorescence in situ hybridization (FISH). Preparation of high ...

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Research

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Biology

13.2K Views.  Leibniz Institute of Plant Genetics and Crop Plant Research (IPK). A protocol is described for the preparation of high-quality mitotic plant chromosome spreads by a fast air-dry dropping method suitable for the FISH detection of single and high copy DNA probes.

Video: A Fast Air-dry Dropping Chromosome Preparation Method Suitable for FISH in Plants

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

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

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

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

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

Genetics

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, specifically the generation of homozygous gynogenetic diploids is useful in genetic analysis because it allows the direct production of homozygotes from a single heterozygous mother. This article describes a modified protocol for ploidy duplication based on a heat pulse during the first cell cycle, Heat Shock 2 (HS2). Through inhibition of centriole duplication, this method results in a precise cell division stall during the second cell cycle. The precise one-cycle division stall, coupled to unaffected DNA duplication, results in whole genome duplication. Protocols associated with this method include egg and sperm collection, UV treatment of sperm, in vitro fertilization and heat pulse to cause a one-cell cycle division delay and ploidy duplication. A modified version of this protocol could be applied to induce ploidy changes in other animal species.

A modified protocol for ploidy manipulation uses a heat shock to induce a one-cycle stall in cytokinesis in the early embryo. This protocol is demonstrated in the zebrafish but may be applicable to other species.

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, ...

Developmental Biology

Functional Manipulation of Maternal Gene Products Using In Vitro Oocyte Maturation in Zebrafish

Cellular events that take place during the earliest stages of animal embryonic development are driven by maternally derived gene products deposited into the developing oocyte. Because these events rely on maternal products which typically act very soon after fertilization-that preexist inside the egg, standard approaches for expression and functional reduction involving the injection of reagents into the fertilized egg are typically ineffective. Instead, such manipulations must be performed during oogenesis, prior to or during the accumulation of maternal products. This article describes in detail a protocol for the in vitro maturation of immature zebrafish oocytes and their subsequent in vitro fertilization, yielding viable embryos that survive to adulthood. This method allows the functional manipulation of maternal products during oogenesis, such as the expression of products for phenotypic rescue and tagged construct visualization, as well as the reduction of gene function through reverse-genetics agents.

Functional Manipulation of Maternal Gene Products Using In Vitro Oocyte Maturation in Zebrafish

An optimized protocol for the in vitro maturation of zebrafish oocytes used for the manipulation of maternal gene products is presented here.

Cellular events that take place during the earliest stages of animal embryonic development are driven by maternally derived gene products deposited into ...

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

Immunology and Infection

Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

Meiosis is the key cellular process required to create haploid gametes for sexual reproduction. Model organisms have been instrumental in understanding the chromosome events that take place during meiotic prophase, including the pairing, synapsis, and recombination events that ensure proper chromosome segregation. While the mouse has been an important model for understanding the molecular mechanisms underlying these processes, not all meiotic events in this system are analogous to human meiosis. We recently demonstrated the exciting potential of the zebrafish as a model of human spermatogenesis. Here we describe, in detail, our methods to visualize meiotic chromosomes and associated proteins in chromosome spread preparations. These preparations have the advantage of allowing high resolution analysis of chromosome structures. First, we describe the procedure for dissecting testes from adult zebrafish, followed by cell dissociation, lysis, and spreading of the chromosomes. Next, we describe the procedure for detecting the localization of meiotic chromosome proteins, by immunofluorescence detection, and nucleic acid sequences, by fluorescence in situ hybridization (FISH). These techniques comprise a useful set of tools for the cytological analysis of meiotic chromatin architecture in the zebrafish system. Researchers in the zebrafish community should be able to quickly master these techniques and incorporate them into their standard analyses of reproductive function.

Nuclear surface spreads are an indispensable tool for studying chromosome events during meiosis. Here we demonstrate a method to prepare and visualize meiotic chromosomes during prophase I from zebrafish spermatocytes.

Meiosis is the key cellular process required to create haploid gametes for sexual reproduction. Model organisms have been instrumental in understanding ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Co-expression of Multiple Chimeric Fluorescent Fusion Proteins in an Efficient Way in Plants

Information about the spatiotemporal subcellular localization(s) of a protein is critical to understand its physiological functions in cells. Fluorescent proteins and generation of fluorescent fusion proteins have been wildly used as an effective tool to directly visualize the protein localization and dynamics in cells. It is especially useful to compare them with well-known organelle markers after co-expression with the protein of interest. Nevertheless, classical approaches for protein co-expression in plants usually involve multiple independent expression plasmids, and therefore have drawbacks that include low co-expression efficiency, expression-level variation, and high time expenditure in genetic crossing and screening. In this study, we describe a robust and novel method for co-expression of multiple chimeric fluorescent proteins in plants. It overcomes the limitations of the conventional methods by using a single expression vector that is composed of multiple semi-independent expressing cassettes. Each protein expression cassette contains its own functional protein expression elements, and therefore it can be flexibly adjusted to meet diverse expression demand. Also, it is easy and convenient to perform the assembly and manipulation of DNA fragments in the expression plasmid by using an optimized one-step reaction without additional digestion and ligation steps. Furthermore, it is fully compatible with current fluorescent protein derived bio-imaging technologies and applications, such as FRET and BiFC. As a validation of the method, we employed this new system to co-express fluorescently fused vacuolar sorting receptor and secretory carrier membrane proteins. The results show that their perspective subcellular localizations are the same as in previous studies by both transient expression and genetic transformation in plants.

We have developed a novel method for co-expressing multiple chimeric fluorescent fusion proteins in plants to overcome the difficulties of conventional methods. It takes advantage of using a single expression plasmid that contains multiple functionally independent protein expressing cassettes to achieve protein co-expression.

Information about the spatiotemporal subcellular localization(s) of a protein is critical to understand its physiological functions in cells. Fluorescent ...

Chemistry

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to the original protocol that both streamline the procedure and increase sample uniformity.  First, we normalize all sperm volumes using freezing media that does not contain the cryoprotectant. Second, cryopreserved sperm are stored in cryovials instead of capillary tubes.  The rates of sperm freezing and thawing (&delta;&deg;C/time) are probably the two most critical variables to control in this procedure.  For this reason, do not substitute different tubes for those specified.  Working in teams of 2 it is possible to freeze the sperm of 100 males per team in ~2 hrs.  Sperm cryopreserved using this protocol has an average of 25% fertility (measured as the number of viable embryos generated in an in vitro fertilization divided by the total number of eggs fertilized) and this percent fertility is stable over many years.

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a high-throughput sperm cryopreservation protocol for zebrafish. Sperm cryopreserved using this protocol has an average of 25% fertility in subsequent vitro fertilization and is stable over many years.

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to ...

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

A Fast Air-dry Dropping Chromosome Preparation Method Suitable for FISH in Plants

Summary

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Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Co-expression of Multiple Chimeric Fluorescent Fusion Proteins in an Efficient Way in Plants

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish