Department of Entomology, Virginia Tech
George, P., Sharakhova, M. V., Sharakhov, I. V. High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization. J. Vis. Exp. (64), e4007, doi:10.3791/4007 (2012).
Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place over the next 5 years. For example, the sequencing of the genomes for 15 malaria mosquitospecies is currently being done using an Illumina platform3,4. This Anopheles species cluster includes both vectors and non-vectors of malaria. When the genome assemblies become available, researchers will have the unique opportunity to perform comparative analysis for inferring evolutionary changes relevant to vector ability. However, it has proven difficult to use next-generation sequencing reads to generate high-quality de novo genome assemblies5. Moreover, the existing genome assemblies for Anopheles gambiae, although obtained using the Sanger method, are gapped or fragmented4,6.
Success of comparative genomic analyses will be limited if researchers deal with numerous sequencing contigs, rather than with chromosome-based genome assemblies. Fragmented, unmapped sequences create problems for genomic analyses because: (i) unidentified gaps cause incorrect or incomplete annotation of genomic sequences; (ii) unmapped sequences lead to confusion between paralogous genes and genes from different haplotypes; and (iii) the lack of chromosome assignment and orientation of the sequencing contigs does not allow for reconstructing rearrangement phylogeny and studying chromosome evolution. Developing high-resolution physical maps for species with newly sequenced genomes is a timely and cost-effective investment that will facilitate genome annotation, evolutionary analysis, and re-sequencing of individual genomes from natural populations7,8.
Here, we present innovative approaches to chromosome preparation, fluorescent in situ hybridization (FISH), and imaging that facilitate rapid development of physical maps. Using An. gambiae as an example, we demonstrate that the development of physical chromosome maps can potentially improve genome assemblies and, thus, the quality of genomic analyses. First, we use a high-pressure method to prepare polytene chromosome spreads. This method, originally developed for Drosophila9, allows the user to visualize more details on chromosomes than the regular squashing technique10. Second, a fully automated, front-end system for FISH is used for high-throughput physical genome mapping. The automated slide staining system runs multiple assays simultaneously and dramatically reduces hands-on time11. Third, an automatic fluorescent imaging system, which includes a motorized slide stage, automatically scans and photographs labeled chromosomes after FISH12. This system is especially useful for identifying and visualizing multiple chromosomal plates on the same slide. In addition, the scanning process captures a more uniform FISH result. Overall, the automated high-throughput physical mapping protocol is more efficient than a standard manual protocol.
1. High-pressure Polytene Chromosome Preparation from Ovarian Nurse Cells
2. FISH Using an Automated Slide Staining System
Program is set up with the Xmatrx automated slide staining system to run the following steps, except the probe labeling. The detailed nick-translation labeling protocol is described in the documentation provided with DNA polymerase from Fermentas.
3. Slide Reading with an Automatic Fluorescent Imaging System
This section details the use of the Duet software, which comes standard with the ACCORD PLUS automated scanning system. Instructions begin after turning on in this order: Olympus U-RFL-T power supply for a halogen bulb, computer Dell precision T3500, microscope Olympus BX61 with a connected camera Olympus U-CMAD3.
4. Representative Results
Figure 1 graphically depicts the scheme of the high-pressure chromosome preparation. This step involves the process of squashing and flattening chromosomes using a Dremel tool and mechanical vise, as well as chromosome visualization using a phase-contrast microscope. Figure 2 illustrates the hybridization of a fluorescently labeled probe to target DNA on the chromosome slide preparations using an automated slide staining system, the use of an automated scanning microscope for visualizing and mapping the probes after the FISH experiment, and placing and orienting genomic scaffolds on the chromosomes.
Preparations of ovarian nurse cell polytene chromosomes from females of An. gambiae were made using the high-pressure technique. This method does not damage or change most of the chromosome structure (Figure 3). It flattens bended chromosomes and, thus, reveals hidden fine bands that are not seen on regular preparations (Figure 4).
The probes used in this protocol are genomic BAC DNA clones obtained from a ND-TAM BAC library generated from An. gambiae PEST strain DNA. The genomic DNA for this library was extracted from newly hatched first instar larvae of both sexes. Figure 5 shows the results of FISH using BAC clones hybridized to polytene chromosomes of An. gambiae. This procedure was performed using the Xmatrx automated slide staining system. Using a cytogenetic photomap for An. gambiae10, two BAC clones, 102B24 (GenBank: BH372701, BH372694) and BAC 142O19 (GenBank: BH368703, BH368698), were localized in subdivisions 16C and 16D of the 2R arm, respectively. However, the BLAST search against the An. gambiae PEST strain AgamP3 assembly 14 identified the sequences homologous to 102B24 and 142O19 in subdivisions 16B and 17A of the 2R arm, respectively (Table 1). Therefore, the correspondence between the genomic coordinates and the cytogenetic subdivisions can now be adjusted according to our mapping data. The BLAST search against the An. gambiae M form and S form genome assemblies found that the two BAC clones are located in different contigs, but in the same scaffolds within each form (Table 1). The identified contigs can now be associated with specific chromosomal locations. Moreover, the identified scaffolds can now be properly oriented within the cytological subdivisions 16CD. Interestingly, the distances between the 102B24 and 142O19 sequences in the scaffolds are 1,892,981 bp, 1,658,391 bp, and 1,688,426 bp in the PEST strain, M-form, and S-form genome assemblies, respectively. Because, the PEST strain is a hybrid between the M and S forms, this difference is likely due to incorrect assembly of the PEST genome.
Table 1. BLAST results of the BAC-end sequences against the genome assemblies of An. gambiae.
Figure 1. Schematic representation of high-pressure chromosome preparation. Mosquito ovaries are shown at the correct stage of development.
Figure 2. A scheme representing automated FISH, slide scanning, and chromosome mapping of genomic scaffolds.
Figure 3. A spread of polytene chromosome 3 of An. gambiae obtained using the high-pressure technique. 3L and 3R mark left and right arms at their telomeres. PH and IH indicate pericentric and intercalary heterochromatin, respectively.
Figure 4. Comparison of An. gambiae polytene chromosomes prepared using the traditional (top) and high-pressure (bottom) technique. The images cover subdivisions 29A-30E of arm 3R (A) and 43D-46D of arm 3L (B).
Figure 5. FISH of BAC clones to polytene chromosomes of An. gambiae. A) Hybridization of 102B24 (red signal) with the 2R arm. B) Dual-color FISH of 102B24 (red signal) and 142O19 (blue signal) to subdivisions 16C and 16D of the stretched 2R arm, respectively. Arrows indicate signals of hybridization of BAC clones labeled with Cy3 (red) and Cy5 (blue). C-the centromeric region. a/+ shows the heterozygote 2La inversion. Chromosomes were counterstained with the fluorophore YOYO-1.
The most critical step of the high-pressure procedure is the proper squashing of ovarian nurse cells isolated at Christophers' III stage of ovarian development13. Improper squashing can lead to insufficiently spread chromosomes, which can cause problems when trying to determine probe locations after FISH. If the chromosomes are over-squashed, they can be become broken or elongated to the point where banding patterns are lost. Production of multiple slides should lead to a consistency when attempting to squash the slides using the Dremel tool, which will increase overall slide production efficiency. The high-pressure technique was first developed for freshly isolated salivary glands of D. melanogaster9. However, ovaries of mosquitoes are routinely preserved in modified Carnoy's fixative solution (3 methanol: 1 glacial acetic acid by volume) before they are used for chromosomal preparations. Therefore, we modified the existing high-pressure protocol to make it suitable for the fixed ovarian nurse cell polytene chromosomes of the malaria mosquito An. gambiae. Because the high pressure is applied using a precision vise possessing a highly parallel work surface of the entire slide, it takes significantly less time to prepare the chromosome squash than using a traditional tapping technique with a pencil's eraser.
Other important steps include sufficiently soaking follicles in 50% propionic acid and heating the slide. Both of these steps are essential in helping to flatten the chromosomes. If they are neglected, chromosomes can appear shiny after dehydration, which potentially leads to an overabundance of background that can be mistaken for a signal in FISH. The high-pressure squashing technique also works well using the same solutions when making mitotic chromosome preparations. The additional spreading and flattening helps to better express the chromosomes from their nuclei. Adding equal pressure should flatten them accordingly to allow for measurements and potentially better resolution of bands visible from staining. The details about isolating mitotic chromosomes from mosquitoes are given elsewhere15. Although regular squash preparations are sufficient for many purposes including FISH 16 and immunostaining17, the high-pressure method not only lowers potential variance from one slide to the next, but also increases overall chromosome quality, leading to higher detail when mapping chromosomes. This procedure can also be used for preparing polytene chromosome membrane slides for laser-capture microdissection.
Limitations to the high-pressure method include slide breakage and overstretching of the chromosomes. Slide breakage caused by placing too much stress on the slide via the vise is possible, but is limited by using the pressures denoted in the article. For overstretching of the chromosomes, if the Dremel tool is applied to the slide for an extended period of time, there is the possibility that the chromosomes can become too stretched out and lose resolution. This can be remedied by applying the tool for a brief period of time, checking under the microscope, and applying more time with the Dremel tool if the chromosomes are insufficiently spread.
A traditional FISH protocol includes a number of washing and incubation steps, which are usually 5-20 min long and require almost full attention of a researcher for the whole duration of the experiment. Moreover, the number of slides that can be handled manually is usually limited to a few slides in a given experiment. In contrast, an automated slide staining system performs all steps (including washing, incubation, probe application, denaturation, hybridization, coverslip application and removal) automatically. This frees up to 6-8 working hours for a researcher in a FISH experiment. Moreover, an automated FISH system can significantly increase the throughput. For example, the Xmatrx system is capable of processing up to 40 assays on single preparation slides and up to 80 assays on dual preparation slides simultaneously. Among limitations of this system is that it is not efficient for a small number of FISH experiments as it requires preparing large volumes of solutions. In addition, the FISH protocol programmed in the system may require modifications and adjustments for new applications.
A slide scanning system is an automated version of a fluorescent microscope. Automated stage moving and a simple microscope control panel free a researcher's time and make operating the microscope extremely simple. For instance, the software in the ACCORD PLUS scanning system allows for easy capturing of multiple channels of fluorescence, and easy to manage image acquisition. An example of this is the inclusion of z-stack capturing, rather than taking a single image; the software captures a configurable z-stack of images to ensure that at least one image remains in focus. Although this system is made more for multiple image acquisition (a lot of cells on a single slide), it still makes finding chromosomes on the slide much easier than navigating through the slide manually.
Both the automated slide staining system and the scanning system are routinely used for FISH diagnostics of chromosomal abnormalities in human cells by cytogenetics clinics. In this study, we developed a protocol that utilizes these automated systems for a basic research application. The automated high-throughput physical mapping protocol can facilitate rapid development of physical chromosome maps for any species of interest. Although, in this report, we used polytene chromosomes for physical mapping, mitotic chromosomes can also be utilized with these systems. It would be useful for the protocol to be adjusted to mitotic chromosomes because the majority of organisms do not develop readable polytene chromosomes. However, the polytene chromosomes, when available, can provide useful information about the correspondence of functional genome domains with the chromosome structure at the highest resolution. The organizational principles of polytene chromosomes, such as patterns of bands and interbands, have recently been likened to that of regular nonpolytene (interphase) chromosomes18. Therefore, the detailed physical mapping performed on high-pressure chromosome preparations has the potential to link DNA sequences to specific chromosomal structures such as bands, interbands, puffs, centromeres, telomeres, and heterochromatin; thus, creating chromosome-based genome assemblies.
No conflicts of interest declared.
This work was supported by the grant from National Institutes of Health 1R21AI094289 to Igor V. Sharakhov. The Xmatrx Automated Slide Staining System and ACCORD PLUS Automated Scanning System were purchased with the help of the Equipment Trust Fund program, Fralin Life Science Institute, Department of Entomology and Department of Biochemistry of Virginia Tech.
|22x22 mm microscope coverslips||Fisher||12-544-10|
|18x18 mm microscope coverslips||Fisher||12-553-402|
|Dremel 200 rotary tool with a Flex-Shaft attachment||Rand||3" Multi-purpose Mini Bench Grinder (with rate limiter)|
|Specially designed 200 μl plastic tips (beaded plastic edges) for Dremel tool||Pipetman||F171300||Cut and heat fat end of pipet tips|
|Tin coated rapid vise||Avenger Gold Toolmaker||MTC-200-1||Precision ground square and parallel to within 0.00025|
|ThermoBrite Slide Denaturation/Hybridization System||Abbott Molecular||30-144110|
|25 mm barrier slides||Abbott Molecular||XT108-SL|
|25 mm coverslips||Abbott Molecular||XT122-90X|
|Xmatrx Automated Slide Staining System||Abbott Molecular||08L46-001|
|MZ6 Leica stereomicroscope||Leica||VA-OM-E194-354||A different stereomicroscope can be used|
|Olympus CX41 Phase Microscope||Olympus||CX41RF-5||A different phase microscope can be used|
|ACCORD PLUS Automated Scanning System||BioView (USA)||BV-5000-ACCP|
|10% NBF (neutral buffered formalin)||Sigma-Aldrich||HT501128|
|99% formamide||Fisher Scientific||BP227500|
|Dextran sulfate sodium salt||Sigma||D8906|
|20x SSC buffer||Invitrogen||AM9765|
|50x Denhardt’s solution||Sigma-Aldrich||D2532|
|1 mM YOYO-1 iodide (491/509) solution in DMSO||Invitrogen||Y3601|
|ProLong Gold antifade reagent||Invitrogen||P36930|
|dATP, dCTP, dGTP, dTTP||Fermentas||R0141, R0151, R0161, R0171|
|Cy3-dUTP, Cy5-dUTP||GE Healthcare||PA53022, PA55022|
|DNA polymerase I||Fermentas||EP0041|